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Masters Theses 1911 - February 2014
1998
The environmental fate of fungicides used to manage Magnaporthe poae, the causal agent of summer patch /
Jeffery J. Doherty University of Massachusetts Amherst
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THE ENVIRONMENTAL FATE OF FUNGICIDES USED TO MANAGE
MAGNAPORTHE POAE, THE CAUSAL AGENT OF SUMMER PATCH
A Thesis Presented
by
JEFFERY J. DOHERTY
Submitted to the Graduate School of the University of Massachusetts Amherst in partial fulfillment of the requirements for the degree of
MASTER OF SCIENCE
May 1998
Department of Plant Pathology THE ENVIRONMENTAL FATE OF FUNGICIDES USED TO MANAGE
MAGNAPORTHE POAE, THE CAUSAL AGENT OF SUMMER PATCH
A Thesis Presented
by
JEFFERY J. DOHERTY
Approved as to style and content by:
Gail L. Schumann, Chair
Mark S. Mount, Member oJ* John M. Clark, Member
Bruce B. Clarke, Member
Mark S. Mount, Department Head Plant Pathology ACKNOWLEDGMENTS
A Masters thesis is actually the end product of the work of many people. I would like to thank Michael Johnson, Dwight Jesseman, Bess Dicklow, Lori Bickford, Tania
Fernandez and Sharon Sousa for assisting me in the laborious task of preparing samples.
The personnel at the Massachusetts Pesticide Analysis Laboratory - Dan, Ray, Gerry,
Laura and Andy - provided invaluable technical assistance (and put up with my inveterate whining). I am indebted to the members of my committee - Dr. Schumann, Dr.
Clark, Dr. Mount and Dr. Clarke - for their advice and feedback. The Umass Turfgrass
Research Facility and the Massachusetts Pesticide Analysis Lab provided facilities without which this project could not have been accomplished.
Research funding and support of my graduate education is gratefully acknowledged to the following organizations: Golf Course Superintendents Association of New England
Inc., Massachusetts Turf and Lawngrass Council Inc., Ciba-Geigy Corporation (now
Novartis), DowElanco (now Dow Agrosciences), E.I. duPont de Nemours and Co., and
Mobay Corporation (now Bayer Corporation).
I owe a special debt of gratitude to my adviser. Dr. Gail Schumann, who saw in me some potential and gave me a chance. For this and her support throughout this process I am very grateful.
Special thanks go out to Dr. J.M. Clark, without whose advice and encouragement I never would have survived this arduous process.
Ill ABSTRACT
THE ENVIRONMENTAL FATE OF FUNGICIDES USED TO MANAGE
MAGNAPORTHE POAE, THE CAUSAL AGENT OF SUMMER PATCH
MAY 1998
JEFFERY J. DOHERTY, B.S., UNIVERSITY OF MASSACHUSETTS AMHERST
M.S. UNIVERSITY OF MASSACHUSETTS AMHERST
Directed by: Professor Gail Schumann
Magnaporthe poae , an ectotrophic, root-infecting fungus, is the causal agent of summer patch, a very damaging disease of Poa and Festuca species. Chemical control of summer patch is both erratic and expensive, suggesting a difficulty in delivering the fungicides to the root zone.
The most important barrier to efficacious delivery of the fungicides to the roots is the thatch layer, a tightly intermingled layer of living and dead stems, leaves, and roots of grass that develops between the verdure and the soil surface. Three different application technologies, a boom sprayer, a high pressure injector, and a modified slicer/seeder, were compared to determine if any were superior in delivering the fungicides fenarimol, propiconazole, and triadimefon to the root zone without increasing the possibility of groundwater contamination. Four different matrices of the turfgrass system, leaf, thatch, soil, and roots, were analyzed for the presence of the fungicides. None of the application methods allowed any of the three fungicides to reach the roots in any appreciable amount, principally due to their high affinity for the thatch. Regardless of application type, the majority of the fungicides (40-80%) were found associated with the thatch layer.
IV Additionally, none of the fungicides were ever found below the top 5.1 cm of the soil, so the threat of groundwater contamination appears minimal over the time course of this experiment (1 month).
Triadimefon is rapidly hydrolyzed to triadimenol in turfgrass plants, thatch, soil, and roots. Triadimenol poses a greater threat to groundwater than triadimefon because it is more water soluble, has a greater half-life, a lower organic carbon/water partition coefficient and greater mammalian toxicity than triadimefon. Triadimefon was rapidly converted to triadimenol in all four matrices, and was not detectable at the end of the experiment (28 days). Triadimenol was the only compound detected in the roots in any appreciable amount
( 0.04 - 0.19% of the applied triadimefon). However, triadimenol was never detected below the top 5.1 cm of the soil profile, so its threat to groundwater over the course of the experiment appears minimal.
Benomyl was applied to determine if post-application irrigation had any effect on its delivery to the root zone. Post-application irrigation appeared to allow benomyl to move into the thatch and roots more quickly. Immediately after application, benomyl residues in the root tissue of the irrigated plot were 2.2 times greater than those found in the non- irrigated plot. By the end of the experiment (14 days), however, there was no difference between the irrigated and the non-irrigated plots.
Because a large proportion of the applied fenarimol, propiconazole, and the triadimefon metabolite triadimenol remained associated with the thatch at the end of the study, a longer experimental time frame should be examined that includes spring and fall recharge events in future research. TABLE OF CONTENTS
Page
ACKNOWLEDGMENTS.iii
ABSTRACT.iv
LIST OF TABLES.x
LIST OF FIGURES.xi
Chapter
1. INTRODUCTION AND LITERATURE REVIEW.I
1.1 Ecology and Epidemiology of Magnaporthe poae.2 1.2 Environmental Effects on Summer Patch Development.6 1.3 Management of Summer Patch.8 1.4 Sterol-Biosynthesis Inhibiting Fungicides.12
1.4.1 Mode of Action of Sterol-Biosynthesis Inhibiting Fungicides. 12 1.4.2 Classes of Sterol-Biosynthesis Inhibiting Fungicides.13
1.4.2.1 Azoles.13 1.4.2.2 Pyrimidines.15
1.5 Benzimidazole Fungicides.15
1.5.1 Benomyl Mode of Action. 16
1.6 Environmental Fate of Fungicides Applied to Manage Patch Diseases. 18
1.6.1 Drift. 19 1.6.2 Photochemical Degradation. 19 1.6.3 Volatility. 22 1.6.4 Runoff.. 24 1.6.5 Plant Uptake. 25 1.6.6 Thatch Layer. 29 1.6.7 Degradation. 33 1.6.8 Mobility. 36
VI 2. ENVIRONMENTAL FATE OF FENARIMOL, PROPICONAZOLE, AND TRIADIMEFON APPLIED TO KENTUCKY BLUEGRASS UTILIZING THREE APPLICATION SCHEMES.41
2.1 Introduction. 41 2.2 Materials and Methods.47
2.2.1 Experimental Site. 47 2.2.2 Fungicides. 47 2.2.3 Fungicide Application. 47 2.2.4 Sampling. 50 2.2.5 Storage Samples. 52 2.2.6 Analytical Procedures. 52 2.2.7 Instrumental Analysis. 54 2.2.8 Quality Assurance/Quality Control. 55
2.2.8.1 Instrument Calibration.55 2.2.8.2 Sample Controls.55
2.3 Results. 57
2.3.1 Fungicide Concentration. 57
2.3.1.1 Leaf Tissue. 57 2.3.1.2 Thatch. 61 2.3.1.3 Soil. 65 2.3.1.4 Root Tissue. 69
2.3.2 Total Accumulated Residue. 72
2.3.2.1 Leaf Tissue. 72 2.3.2.2 Thatch. 72 2.3.2.3 Soil. 72 2.3.2.4 Root Tissue. 74
2.3.3 Percent of Applied Fungicide Recovered. 74
2.3.3.1 Leaf Tissue...,. 74 2.3.3.2 Thatch. 82 2.3.3.3 Soil. 84 2.3.3.4 Root Tissue. 85
2.4 Discussion. 86
Vll 3. FATE OF TRIADIMEFON AND TRIADIMENOL IN A TURFGRASS SYSTEM.93
3.1 Introduction. 93 3.2 Materials and Methods.94
3.2.1 Experimental Site. 94 3.2.2 Fungicides. 96 3.2.3 Fungicide Application. 96 3.2.4 Sampling. 97 3.2.5 Storage Samples. 98 3.2.6 Analytical Procedures. 98 3.2.7 Instrumental Analysis. 100 3.2.8 Quality Assurance/Quality Control. 101
3.2.8.1 Instrument Calibration. 101 3.2.8.2 Sample Controls. 101
3.3 Results. 102
3.3.1 Fungicide Concentration. 102
3.3.1.1 Leaf Tissue. 102 3.3.1.2 Thatch. 105 3.3.1.3 Soil. 109 3.3.1.4 Root Tissue. 112
3.3.2 Percent of Applied Fungicide Recovered. 115
3.3.2.1 Leaf Tissue. 115 3.3.2.2 Thatch. 120 3.3.2.3 Soil. 121 3.3.2.4 Root Tissue. 122
3.4 Discussion. 123
4. FATE OF BENOMYL IN A TURFGRASS SYSTEM WITH AND WITHOUT POST-APPLICATION IRRIGATION.128
4.1 Introduction. 128 4.2 Materials and Methods. 128
4.2.1 Experimental Site. 128 4.2.2 Fungicide Application. 130 4.2.3 Sampling. 130
viii 4.2.4 Storage Samples. 131 4.2.5 Analytical Procedures. 132 4.2.6 Quality Assurance/Quality Control. 134
4.2.6.1 Instrument Calibration. 134 4.2.6.2 Sample Controls. 135
4.3 Results. 136
4.3.1 Fungicide Concentration. 136
4.3.1.1 Leaf Tissue. 136 4.3.1.2 Thatch. 136 4.3.1.3 Soil. 141 4.3.1.4 Root Tissue. 141
4.3.2 Percent of Applied Fungicide Recovered. 146
4.3.2.1 Leaf Tissue. 146 4.3.2.2 Thatch. 149 4.3.2.3 Soil. 149 4.3.2.4 Root Tissue. 150
4.4 Discussion. 150
5. SUMMARY. 154
5.1 Conclusions. 159
BIBLIOGRAPHY. 160
IX LIST OF TABLES
Table Page
1. Physical properties, environmental characteristics, application rates, and toxicities of three SBI fungicides...43
2. Average recoveries (with standard deviations) from the four turfgrass matrices amended with fungicide on the day of extraction (concurrent) and at the time of sampling (storage).56
3. Total accumulated residues (TAR) associated with leaf, thatch, soil, and root following application by boom sprayer, high pressure injector, and modified slicer/seeder.73
4. Physical properties, environmental characteristics, application rates, and toxicities of triadimefon and triadimenol.95
5. Physical properties, environmental characteristics, application rates, and toxicities of benomyl and carbendazim.129
6. Average recoveries (with standard deviations) of benomyl from the four turfgrass matrices amended with fungicide on the day of extraction (concurrent) or at the time of sampling (storage).133 LIST OF FIGURES
Figure Page
1. Irrigation (I) and rainfall (R) events (in mm of H2O) to the study plots in South Deerfield, MA from 24 August to 21 September 1992.49
2. Concentration (in ppm) of fenarimol, propiconazole, and triadimefon in leaf tissue pre-application (pre),immediately post-application ( Day 0), and 1,3,7,10,14,21, and 28 days following application with a boom sprayer (A), high pressure injector (B), and a modified slicer/seeder (C).59
3. Concentration (in ppm) of fenarimol, propiconazole and triadimefon in thatch pre-application (pre), immediately post-application ( Day 0), and 1,3,7,10,14,21, and 28 days following application with a boom sprayer (A), high pressure injector (B), and a modified slicer/seeder (C).63
4. Concentration (in ppb) of fenarimol, propiconazole and triadimefon in soil pre-application (pre), immediately post-application ( Day 0), and 1,3,7,10,14,21, and 28 days following application with a boom sprayer (A), high pressure injector (B), and a modified slicer/seeder (C).67
5. Concentration (in ppm) of fenarimol, propiconazole and triadimefon in roots pre-application (pre), immediately post-application ( Day 0), and 1,3,7,10,14,21, and 28 days following application with a boom sprayer (A), high pressure injector (B), and a modified slicer/seeder (C).71
6. Percent of total applied fenarimol recovered from all matrices (total) and from leaf, thatch, roots, and soil for each day from Day 0 to day 28 following application with the boom sprayer (A) and high pressure injection (B).76
7. Percent of total applied propiconazole recovered from all matrices (total) and from leaf, thatch, roots, and soil for each day from Day 0 to day 28 following application with the boom sprayer (A), high pressure injection (B), and modified slicer/seeder (C).78
XI 8. Percent of total applied triadimefon recovered from all matrices (total) and from leaf, thatch, roots, and soil for each day from Day 0 to day 28 following application with the boom sprayer (A) and high pressure injection (B).80
9. Chemical structure of triadimefon and triadimenol.94
10. Concentration of triadimefon and triadimenol in leaf tissue (in ppm) pre-application (pre), immediately post-application ( Day 0), and 1,3,7,10,14,21, and 28 days following application with a boom sprayer (A), high pressure injector (B).104
11. Concentration of triadimefon and triadimenol in thatch (in ppm) pre-application (pre), immediately post-application (Day 0), and 1,3,7,10,14,21, and 28 days following application with a boom sprayer (A), high pressure injector (B).107
12. Concentration of triadimefon and triadimenol in soil (in ppb) pre-application (pre), immediately post-application ( Day 0), and 1,3,7,10,14,21, and 28 days following application with a boom sprayer (A), high pressure injector (B).111
13. Concentration of triadimefon and triadimenol in root tissue (in ppm) pre-application (pre), immediately post-application (Day 0), and 1,3,7,10,14,21, and 28 days following application with a boom sprayer (A), high pressure injector (B).114
14. Percent of total applied triadimefon (A) and triadimenol (B) recovered from all matrices (total) and from leaf, thatch, roots, and soil for each day from Day 0 to day 28 following application with the boom sprayer. 117
15. Percent of total applied triadimefon (A) and triadimenol (B) recovered from all matrices (total) and from leaf, thatch, roots, and soil for each day from Day 0 to day 28 following application with the high pressure injector. 119
16. Concentration (in ppm) of benomyl/carbendazim in leaf tissue immediately post-application (Day 0), and 1,2,5,9, and 14 days following application by boom sprayer with and without post-application irrigation. 138
XU 17. Concentration (in ppm) of benomyl/carbendazim in thatch immediately post-application (Day 0), and 1,2,5,9, and 14 days following application by boom sprayer with and without post-application irrigation. 140
18. Concentration (in ppb) of benomyl/carbendazim in soil immediately post-application (Day 0), and 1,2,5,9, and 14 days following application by boom sprayer with and without post-application irrigation. 143
19. Concentration (in ppm) of benomyl/carbendazim in root tissue immediately post-application (Day 0), and 1,2,5,9, and 14 days following application by boom sprayer with and without post-application irrigation. 145
20. Percent of total applied benomyl/carbendazim recovered from all matrices (total) and from leaf, thatch, roots, and soil for each day from Day 0 to day 14 following application with the boom sprayer without (A) and with (B) post-application irrigation. 148 CHAPTER 1
INTRODUCTION AND LITERATURE REVIEW
Magnaporthe poae is the causal agent of summer patch (Landschoot and Jackson, 1989), a very damaging disease of Kentucky bluegrass (Poa pratensisk annual bluegrass (Poa annual and fine fescue (jFestuca spp). Summer patch was known as Fusarium blight
(Couch and Bedford, 1966) until 1984, when it was differentiated from Fusarium blight and reported as Phialophora graminicola (Smiley, 1984; Smiley and Craven Fowler; 1984).
These isolates were later classified as a Phialophora state of a Magnaporthe sp.
(Landschoot and Jackson 1987) which was eventually named hi poae (Landschoot and
Jackson, 1989). Smiley and Craven Fowler (1984) also reported a similar patch disease on
P, pratensis. shown to be caused by Leptosphaeria korrae. which was later named necrotic ring spot (Worf et al., 1986). Therefore, care must be taken when consulting the literature prior to 1984 because reports on Fusarium blight may be dealing with either one of these causal agents.
Chemical control of summer patch is both erratic and expensive (Demoeden, 1993).
Fungicides are generally more effective against summer patch when applied in large (800-
1000 L H2O ha'^) amounts of water (Demoeden, 1989a; Demoeden and Minner; 1981,
Demoeden and Nash; 1982, Hartman et al.; 1989, Landschoot and Clarke, 1989; Plumley et al. 1992; Smiley, 1980). This may indicate a problem in delivering the fungicide to the target area, the top 8 cm of the soil, where the fungus survives (B. Clarke, personal communication). By analyzing the various compartments in the turf system (turf foliage.
1 thatch, soil, and roots) it is hoped that the main barriers to effective fungicide delivery can be elucidated.
1.1 Ecology and Epidemiology of Magnaporthc poae
Summer patch is an increasingly important turfgrass disease of intensively managed turf.
These turfs are frequently of similar genotype with the plants being grown in close proximity to one another. They are also subjected to numerous stresses such as mowing, compaction, temperature and moisture extremes, and the non-target affects of pesticides
(Endo, 1972). A turfgrass sward also contains a thatch layer, which is made up of decomposing leaves, stems, and roots, between the roots and the soil. This thatch layer provides both a habitat and food source for microorganisms, as well as acting as a barrier to the delivery of lipophilic compounds, such as sterol biosynthesis inhibiting (SBI) fungicides, to the soil and the roots of the plants.
Magnaporthe poae is a member of a group of ectotrophic, root-infecting (ERI) fimgi.
Members of this group have been identified as turfgrass pathogens including
Gaeumannomyces graminis var. avenae (take-all patch), korrae (necrotic ring spot) and others. These fimgi appear well adapted to the turfgrass ecosystem and can persist either on or in roots for a long period of time (Landschoot et al., 1993). The pathogen is able to invade the vascular tissue of the roots under favorable circumstances, leading to death of the plant (Landschoot et al., 1993).
There is little known about the factors affecting the ecology and epidemiology of ERI fungi because most of these diseases and causal agents have only recently been identified as
2 turfgrass pathogens. A literature base does exist for ERJs in wheat, and this may be useful
towards understanding these diseases.
A common characteristic of all ERI fungi is their ectotrophic growth habit, described by
Garrett (1956) as the continuous external growth of fungal hyphae over root surfaces that
can occur prior to the penetration of the root cortical cells. The growth of graminis var.
tritici (take-all of wheat) and G* graminis var. avenae (take-all of bentgrass and other small
grains) have been studied extensively compared to other ERI fungi. Garrett (1970) has discussed at length the ectotrophic growth habit of G^ graminis (reported as Ophiobolus graminis). Though there are probably some differences with other ERIs, the mechanisms described by Garrett can provide a starting point.
The ectotrophic hyphae found on wheat begin as single hyphae and fuse laterally after 24 to 30 hours to form dark brown runner hyphae (Weste, 1972). These runner hyphae then grow along the roots. Under conditions conducive to infection this ectotrophic mycelium can then infest turfgrass roots, crowns, stolons, rhizomes, stem bases, and leaf sheaths
(Endo et al., 1985; Nilsson and Smith, 1981; Smiley Craven Fowler, 1984; Smith, 1965;
Worf et al., 1986). The mycelia may form hypophodia which develop infection hyphae capable of penetrating cortical tissue (Skou, 1981).
While a decline in plant vigor has been shown to increase ectotrophic growth (Garrett,
1970) and patch disease development (Davis and Demoeden, 1991; McCarty et al., 1991;
Smiley et al., 1985a; Smiley et al., 1985b), there is little evidence of a correlation between ectotrophic growth and an increase in patch disease symptoms (Landschoot et al., 1993).
Garrett (1970) discusses the idea that vigorously growing plants restrict colonization to the
3 ectotrophic habit. Weakening host defenses then lead to infection. Alternatively, Garrett
(1970) suggested that the ectotrophic hyphae may initiate several infection loci in an attempt to overcome plant defenses. In the event of a complete collapse of the plant’s defense mechanisms ERI fungi could be outcompeted by less specialized pathogens.
Therefore a weakened, but not debilitated, plant provides the best host for ERI flmgi
(Landschoot et al., 1993).
Root-infecting fungi can survive in the soil in the absence of living host tissue as dormant resting structures or as saprophytes (Garrett, 1970). G, graminis. and most likely other ERI fungi, rarely produce dormant resting structures to ensure survival in the soil (Garrett, 1970;
Shipton, 1981; Walker, 1981). Though runner hyphae and pseudoparenchymatous tissue may function in this role, they are not thought to be important due to their limited ability to infect seedlings (Hornby, 1981). Evidence points to G^ graminis surviving saprophytically in plant tissue already colonized through parasitism. It is thought that many specialized root pathogens compete better with more saprophytically competent organisms in tissue they colonized before the death of the plant (Garrett, 1970). While direct isolation from organic debris has been rare (Gams and Domsch, 1967; Warcup, 1957), G^ graminis has been detected using trap hosts such as wheat (Hornby, 1969). graminis has been found to have fairly poor saprophytic abilities (Butler, 1953; Rao, 1959). Therefore, as stated by Garrett
(1970), saprophytic colonization without prior parasitism appears to be of minor importance in the survival of Gr graminis. The situation of ERI fungi in turf may also be the same, but the perennial nature of turfgrass presents a continuous survival mechanism.
4 Although gr^niinig can survive adverse soil conditions, its ability to do so is limited.
In situations conducive to microbial activity available organic substrates are rapidly exhausted (Garrett, 1938, 1970), reducing the survival of G. graminis due to its poor competitiveness. Under conditions inimical to microbial activity, however, saprophytic survival increases.
Spread of graminis can occur when healthy tissue comes into direct contact with infected tissue or colonized debris (Skou, 1981). The fiingus also has the ability to grow from a colonized substrate a short distance to host tissue (Brown and Hornby, 1971; Pope and Jackson, 1973; Skou, 1981). Studies on the role of ascospores in the dissemination of
Gi graminis have shown that they are able to infect only when there is little competition
(Brooks, 1964, 1965; Garrett, 1939). Fruiting structures of M. poae have not been found in nature, so ascospores are not considered important as a means of dissemination of this fungus at present.
The movement of infested tissue during cultivation may account for a large portion of the disseminated inoculum. It has been demonstrated that spring dead spot (another ERI patch disease) can be spread by transferring pathogen-infested turf cores to areas previously free of the disease (Pair et al., 1986). It is also possible that inoculum can be transported by core aerators, dethatching machines, verticutters, slicer/seeders, or even the spikes of golf shoes. Although infested seed has not been shown to be a means by which ERI fungi are disseminated, it is possible the pathogens could be spread from one location to another through the transport of infected sod (Smith et al., 1989).
5 The entire life cycle of M. poae appears to take place in the soil. Fungicides must therefore be delivered to the soil in order to effectively manage this disease. The poor competitive ability of this fungus is interesting but not really useful from a practical standpoint, as crop rotation is not an option in a turfgrass area.
1.2 Environmental Effects on Summer Patch Development
Summer patch symptoms usually occur in periods of hot, rainy weather (Kackley et al.,
1990b; Smiley et al., 1992) between June and September on closely mowed turf (Smith et al., 1989). Initially drought stress was believed to be an important factor predisposing turf to
Fusarium blight syndrome (Bean, 1966,1969; Couch and Bedford, 1966; Endo and
Coolbaugh, 1974), from which summer patch was separated in 1984. It was noted by
Smiley (1980), conversely, that Fusarium blight was more closely associated with excessive moisture or alternating periods of rainfall and drought. These conflicting reports were made before poae was identified as the causal agent of summer patch. Therefore, any data relating to the environmental effects on Fusarium blight syndrome must be questioned.
M. poae grows best at lower osmotic potentials (Kackley et al., 1990a; Smiley et al.,
1985b). Summer patch is more severe on turf maintained at higher osmotic potentials
(Kackley et al., 1990c) with symptoms most severe on non-moisture stressed R pratensis and on mildly stressed R annua. A field study by Kackley et al. (1990b) found that summer patch was not increased by drought stress but was, in fact, more severe on plots that were not drought stressed. Kackley et al. therefore concluded that temperature and water potential, but not drought stress, play a key role in summer patch development.
6 Optimal growth of M. poae was found to occur around 30°C (Kackley et al., 1990a;
Landschoot and Jackson, 1990; Smiley et al., 1985b), while the optimal temperatures for root growth of P. pratensis are between 10 and 18.5®C (Beard, 1973) with a considerable reduction in growth at 26°C (Youngner, 1969).
It was speculated by Kackley et al. (1990a) that in the warm, moist conditions under which Ml poae thrives, host root development is impaired, and the host is, therefore, more susceptible to the disease. With cooler (less than 25°C) moist soil conditions, the host is favored and disease development minimal, with patch development occurring when the turf is stressed by drought. Summer patch development then is a function of pathogen growth, host root development, and stress (Smiley et al., 1985b).
G. graminis can tolerate temperatures as high as 60-70°C for one hour on each of several days (Fellows, 1941) and can survive 45 weeks under conditions that are dry or moist and cool (15°C). Dry and hot (35°C) or wet and cool conditions, on the other hand, reduce viability. In addition, four weeks in a wet, hot environment completely eliminated the viability of Gl graminis (MacNish, 1973).
Nitrogen availability also strongly influences the survival of graminis in the soil
(Shipton, 1981). Garrett (1938,1940) reported that in soils amended with nitrogen the pathogen was viable for longer periods of time than in unamended soils. It has been suggested that in order to assimilate the carbohydrate reserves in organic substrates the fungus requires sufficient nitrogen (Garrett, 1970). Summer patch is most often seen on poorly drained, compacted soils (Smiley et al., 1992). Beyond this, little is known of the way edaphic factors (pH, fertility, soil type, etc.) affect the development of summer patch.
7 1.3 Management of Summer Patch
According to Demoeden (1993) there are four reasons why ERJ fimgi cause such destructive diseases: (1) limited availability of disease resistant germplasm; (2) incomplete understanding of pathogen biology and disease etiology; (3) few field studies investigating the effects of cultural practices on disease severity; and (4) inconsistent level of disease control with fungicides.
Turgeon (1991) stated that a reduced mowing height often reduces the ability of a turf to resist diseases. In a field study on Fusarium blight, Turgeon and Meyer (1974) found that disease severity was greater at the lower cut (1.9 cm) for R pratensis cv. Pennstar, but conversely was greater at the higher cut (3.8 cm) for cultivars Merion and Nugget. Mowing height was found to have no effect on disease severity for cultivars Fylking or Kenblue.
Studies on summer patch have shown that disease severity is greater at lower mowing heights. P, pratensis was found to suffer greater damage from summer patch when maintained at 7.6 cm than 1.9 cm (Davis and Demoeden, 1991). Unmowed R pratensis was found to sustain less damage from poae than turf maintained at 2.0 cm in a greenhouse study (Smiley et al., 1985b). Plumley et al. (1992) found that the height of cut had no correlation with the movement of M. poae through the soil profile of a R pratensis cv.
Baron cut at 4.0 and 8.0 cm.
It is generally believed that Fusarium blight and summer patch are more severe under high nitrogen fertility levels. Turgeon and Meyer (1974) found that heavy fertilization in
» spring increased the severity of Fusarium blight for all but one cultivar of P. pratensis. Nash
8 (1988) found summer patch to be more severe on plots treated with urea than those treated
with slow release forms of nitrogen.
Davis and Demoeden (1991) tested the effect of several sources of nitrogen, selected for their ability to affect soil pH, on the severity of summer patch. The authors found summer patch to be least severe in plots treated with sulfur coated urea (SCU, a slow release, acid reacting nitrogen source), and most severe in plots treated with sodium nitrate (a quick release, neutral reacting nitrogen source). The authors hypothesized that the slow release and acidifying attributes of SCU help reduce disease, while the sodium nitrate increased disease through a reduction in root growth and an inability to modify soil pH.
Thompson et al. (1992) found that ammonium nitrate slightly reduced summer patch severity in the first year of treatment and significantly reduced it in the second when compared to a turf treated with calcium nitrate or unfertilized. The authors speculated that a reduction in rhizosphere pH played a role. A related study by Thompson et al. (1995) found that ammonium chloride, ammonium sulfate, and SCU reduced disease severity compared to unfertilized turf, while calcium nitrate and potassium nitrate increased disease symptoms.
Resistant grasses are available. Perennial ryegrass, tall fescue, creeping bentgrass and all warm season grasses are resistant or possibly immune to summer patch in the field
(Demoeden, 1989c). poae primarily affects Kentucky bluegrass, annual bluegrass, and fine fescues (Demoeden, 1989c; Smiley, 1987). Strong creeping red fescue cultivars
Pennlawn and Flyer are considered susceptible to summer patch in Maryland, but the hard fescue cultivar Aurora is considered resistant. Kemp et al. (Kemp, 1991, Kemp et al.,
1990a, 1990b) state that strong creeping red fescues are less susceptible to summer patch
9 than hard fescues or slender creeping fescues. Chewings fescue was found by the authors to be moderately susceptible.
There are some Kentucky bluegrass cultivars that possess good summer patch resistance.
Among these are Adelphi, Aspen, Enmundi, Rugby, Sydsport, and Touchdown, while cultivars such as Dormie, Fylking, S-21, Merion, and Windsor are considered highly susceptible (Smiley and Craven Fowler, 1986). Susceptibility is also dependent on many complex soil, cultural and climatic factors which affect root infection and disease development (Demoeden, 1993). M:. poae may coexist with other ERI fungi such as K korrae and ^ incrustans. which can further complicate varietal susceptibility (Demoeden,
1993).
Demoeden (1993) described chemical control as 'often erratic and expensive'. The timing, rate, and the amount of water in which the fungicide is applied all influence efficacy
(Demoeden, 1989c). There is an extensive review of the fungicides used to manage
Fusarium blight by Smiley (Smiley, 1980). Some fungicides commonly used to manage summer patch are the benzimidazoles, benomyl and thiophanate-methyl, and the sterol biosynthesis inhibitor (SBI) fungicides, triadimefon, propiconazole and fenarimol
(Demoeden, 1989c). When applied in 800-1000 L H2O ha'^ prior to symptom expression, the SBI fungicides generally provided better control than the benzimidazoles (Demoeden,
1989c; Demoeden and Minner, 1981; Demoeden and Nash, 1982; Hartman et al., 1989;
Landschoot and Clarke, 1989; Plumley et al., 1991; Smiley, 1980). The benzimidazoles, however, may provide some post-infection activity (Demoeden and Minner, 1981;
Demoeden and Nash, 1982). The efficacy of the benzimidazoles may increase when applied
10 in 2000-4000 L H2O ha*’ compared to 800 L (Landschoot and Clarke, 1989). Other SBI fungicides such as cyproconazole, myclobutanil, and terbuconazole have been shown to provide control ofM. poae in plate assays (Thompson and Clarke, 1991). Importantly, chlorothalonil and iprodione have been shown to increase summer patch in certain
Kentucky bluegrass cultivars (Demoeden, 1993).
Demoeden (1993) offers some general recommendations on managing summer patch.
Cultural practices such as raising mowing height, watering deeply and infrequently, use of slow release nitrogen, aerification to reduce compaction, and overseeding with resistant varieties are all useful tools in the management of summer patch. For preventative management Demoeden (1993) recommends one of the SBI fungicides be applied 21 to 28 days prior to symptom development. For curative action, one of the benzimidazoles applied in a large volume of water may provide an acceptable level of control. In annual bluegrass putting greens, where summer patch is a severe problem, multiple applications of these fungicides at 21 to 28 day intervals are recommended. Demoeden (1993) notes that although many fungicides can reduce the severity of summer patch, the level of control varies from year to year and, under very high disease pressure, the level of control may be unacceptable.
The increases in efficacy when fungicides are applied in large amounts of water indicates a difficulty in delivering these fungicides to the target zone, the top 8 cm of the soil. There are many barriers to the application of fungicides to the soil under turf including the dense plant canopy and the thatch layer. It is possible that available equipment such as a high
11 pressure injector or a modified slicer/seeder could be used to break through these barriers to better deliver these fungicides to the soil.
1.4 Sterol Biosynthesis Inhibiting Fungicides
1.4.1 Mode of Action of Sterol Biosynthesis Inhibiting Fungicides
Sterol biosynthesis in all living organisms begins with acetic acid, with mevalonic acid and squalene being key intermediates (Bloch, 1965; Gaylor, 1974). The first cyclization product is lanosterol in mammals and fungi, whereas in plants it is cycloartenol (Nes, 1977).
In yeasts and fimgi there are several alternative pathways by which lanosterol is converted to ergosterol, the end product (Barton et al, 1973; Fryberg et al., 1973). One important step in this conversion is the demethylation of the three methyl groups at C-4 and C-14 (Kato,
1986). The methyl group is converted to hydroxymethyl and then formyl intermediates and is finally removed as fomiic acid (Alexander et al., 1972). Cytochrome P-450 is involved in this oxidative elimination (Alexander et al., 1974; Aoyama and Yoshida, 1978a; Aoyama and Yoshida, 1978b; Aoyama et al., 1984; Ohba et al., 1978; Yoshida and Aoyama, 1984).
The primary mode of action of the sterol biosynthesis inhibiting (SBI) fungicides is inhibition of demethylation at C-14, with demethylation at C-4 also occurring at a reduced rate. This leads to the accumulation of lanosterol and many abnormal sterols such as 24- methylenedihydrolanosterol, obtusifol, and other compounds (Kato, 1986; Ragsdale, 1975;
Schmitt and Benveniste, 1979).The formation of ergosterol is concurrently reduced
(Buchenauer, 1976; Leroux and Gredt, 1976; Ragsdale and Sisler, 1972), and all of these accumulated sterol precursors possess extra methyl group(s) at C-4 and C-14 (Kato, 1986).
12 The three SBI fungicides used in this study are fenarimol, propiconazole, and triadimefon. These fungicides inhibit demethylation by binding to the cytochrome P-450 component of the C-14 demethylase. Studies on another SBI fungicide, buthiobate, in a reconstituted enzyme system of cytochrome P-450 and NADPH-cytochrome P-450 showed that the fungicide induced a Type II spectral change of the cytochrome (Aoyama et al.,
1984). This suggests that the SBI binds to the cytochrome and occupies the active site, thereby inhibiting catalytic activity. As mentioned above, C-14 demethylation occurs through the formation of hydroxymethyl and formyl intermediates (Alexander et al., 1972).
The fact that no such intermediates were detected in the inhibited cultures indicates that buthiobate may block the initial hydroxylation by the cytochrome P-450 involved enzyme system (Kato, 1986).
1.4.2 Classes of Sterol Biosynthesis Inhibiting Fungicides
1.4.2.1 Azoles
Azole fungicides are a group of 1-substituted and 1,2,4-triazoles. This group is one of the largest classes of SBI fungicides today (Kramer, 1986). There are eleven imidazole and triazole products on the market today, including triadimefon (Bayleton'^^) triadimenol
(Baytan'T'^) and propiconazole (Banner^^). In 1983, the estimated market turnover for this
group of fungicides was 220 million dollars (Kramer, 1986). Western Europe accounted for
85% of these sales, primarily for use on cereals. Since 1983, use of these compounds in the
U.S. has most likely increased, especially in the turfgrass market. Several companies are
developing new compounds in this class as fungicides and also as plant growth regulators.
13 Structure activity relationship studies on azoles have shown that to achieve good activity the compounds must not only possess suitable lipophilicity but also the proper stereochemical parameters (Kramer, 1986). This is due to the fact that the lipophilic part of the azole molecule must bind to the lipophilic binding site of the cytochrome P-450, which is normally occupied by ergosterol precursors during sterol biosynthesis (Gadher et ah,
1983 ).
Azoles may contain different functional groups such as keto, ketal, hydroxy, or nitrilo groups, or they may have no functional group attached at all. One of the SBI fungicides in this study, triadimefon, contains a keto group. This class of azoles consists of compounds used for many purposes, from systemic, broad spectrum fungicides to growth regulators to antidandruff shampoos (Kramer, 1986). The melting point of triadimefon is 82.3°C, and the
vapor pressure is 1.50 x 10 ^ mbar at 20°C. The acute oral LD50 for rats is 1000 mg kg body weight. Triadimefon is soluble in a number of organic solvents including methylene chloride and toluene, but is relatively insoluble in hexane. Its water solubility is 64 mg L‘‘ at 20°C (Pesticide Manual, 1994).
Triadimenol is a metabolite of triadimefon, but it is also marketed as a fungicide in its own right in Europe. It belongs to the class of azoles containing hydroxy groups. The majority of the compounds in this class are used as systemic fungicides. Triadimenol is
slightly soluble in water (120 mg L"') and is soluble in many organic solvents such as
methylene chloride and toluene but is relatively insoluble in hexane. The oral LD50 for rats is 700 mg kg \ The vapor pressure of triadimenol is less than 7.5 x 10*^ mbar at 20°C and its melting point is 112°C (Pesticide Manual, 1994).
14 Propiconazole is a representative of the class of azoles with ketal groups. This group consists mainly of highly active fungicides and antimycotics. Propiconazole has a water solubility of 100 mg L' at 20 C, a vapor pressure of 4.2 x 10" at 20°C, and a boiling point
of 180°C at 0.13 mbar. The acute oral LD50 of propiconazole for rats is 1517 mg kg‘^
Propiconazole is soluble in many organic solvents including methylene chloride and toluene, but is relatively insoluble in hexane (Pesticide Manual, 1994).
1.4.2.2 Pyrimidines
This class of fungicides is the most active in inhibiting sterol biosynthesis (Sisler et al.,
1984). This class also contains ancymidol, the first plant growth regulator whose mode of action is based on inhibition of gibberellic acid biosynthesis (Coolbaugh et al., 1982; Sisler et al., 1984). Fenarimol, the third SBI to be studied, is a member of this class. The melting point of fenarimol is 117-119®C, and its water solubility is 13.7 ppm at 25°C at pH 7.0. The
vapor pressure of fenarimol is less than 4.88 x 10 ^ mm Hg at 25°C. The acute oral LD50 for rats is 2500 mg kg'' (Pesticide Manual, 1994). Fenarimol degrades in the presence of UV light (Kramer, 1986).
1.5 Benzimidazole Fungicides
Benomyl is a member of the benzimidazole group of fungicides which are derivatives of a substituted 2 carbon atom of the heterocyclic benzimidazole molecule. There are two types of fungicidal derivatives of benzimidazoles. The first type includes thiabendazole and suberidazole, where the heterocyclic rings are joined by a C-C bond. The second type
belong to a group of compounds where methylbenzimidazole 2-yl carbamic acid
(carbendazim) is the fungitoxic moiety, including benomyl and thiophanate-methyl. With
15 benomyl, the butylcarbamoyl substituent on one of the ring nitrogen atoms is readily hydrolyzed to leave carbendazim and other products. Carbendazim is considered partly or even wholly responsible for the fungitoxicity of benomyl. Butyl isocyanate, a transient toxicant, is also produced during the conversion of benomyl to carbendazim, but is considered of limited practical significance (Singh et al., 1989). Systemic fungicidal
activities of various 2-aminobenzimidazoles were first reported by Klopping in 1960
(Pesticide Manual, 1979). This led to the development of benomyl by E. I. Du Pont de
Nemours and Company and its introduction in the United States in 1968.
Benomyl is very sparingly soluble in water (4 mg L ^) but is soluble in chloroform. The vapor pressure of benomyl is less than 3.7 x 10 * mbar at 20°C and it decomposes without
melting. The acute oral LD50 of benomyl for rats is greater than 10,000 mg kg‘^ The vapor pressure of carbendazim is less than 100 nPa at 20°C, and it has a melting point of 310® C.
Carbendazim is slightly soluble in water (29 mg kg'^) and has an acid dissociation constant
(pka) of 4.48. It is soluble in acetone and ethanol. The acute oral LD50 for rats is greater than 15,000 mg kg'* (Pesticide Manual, 1994).
1.5.1 Benomyl Mode of Action
Benomyl is rapidly converted to carbendazim in aqueous solution (Clemons and Sisler,
1969). Carbendazim is also found in plants treated with benomyl (Peterson and Edgington,
1969, Sims et al., 1969), and is considered responsible for the fungitoxicity of the compound (Clemons and Sisler, 1969; Maxwell and Brody, 1971; Selling et al., 1970;
Vonk and Kaars Sijpestein, 1971). Carbendazim is also sold as a fungicide in its own right.
16 Clemons and Sisler (1971) reported important early evidence on the mode of action of carbendazim when they discovered that a solution of carbendazim caused an 85% inhibition of DNA synthesis in 8 hours, while RNA and protein synthesis were not affected until later.
Later studies found that carbendazim inhibits mitosis and causes a disorganization of the fine structure of the fungal hyphae (Borck, 1973; Davidse, 1973; Hammerschlag and Sisler,
1973; Howard and Aist, 1976; Richmond and Philips, 1975). These effects are similar to those caused by colchicine, which binds to tubulin preventing the formation of the microtubule subunits into spindle fibers (Wilson and Bryan, 1974). Later studies found that carbendazim prevents the assembly of tubulin into microtubules (Davidse, 1973, 1975,
1979; Davidse and Flach, 1977,1978) by binding to tubulin (Davidse, 1975; Davidse and
Flach, 1977), possibly to the beta subunit (Sheir-Neiss et al., 1978). Fungal tubulin has a high affinity for carbendazim, and carbendazim binding is completely inhibited by colchicine (Davidse and Flach, 1977). The tubulin of more sensitive strains of fungi has a higher affinity for carbendazim, and the tubulin of resistant strains exhibits no carbendazim binding (Davidse and Flach, 1977). Selectivity of carbendazim may be based on a different affinity of the fimgal tubulin for the compound.
Early reports on the mode of action of benomyl suggested that inhibition of respiration was an important target site. Hammerschlag and Sisler (1972) attribute this to the formation of a transient toxicant, butyl isocyanate, formed during the conversion of benomyl to carbendazim. Butyl isocyanate causes a rapid inhibition of respiration and a concurrent
reduction in DNA and RNA synthesis as well as growth. However, butyl isocyanate is very
unstable in water, rapidly breaking down to butylamine and CO2 (Hammerschlag and
17 Sisler, 1972; Watkins, 1976). Therefore, carbendazim ultimately causes greater cell death by disrupting microtubule formation (Langcake et ah, 1983). Another postulated mechanism is through the benzimidazole structural resemblance to the DNA purine bases adenine and guanine, which may inhibit DNA synthesis (Cremlyn, 1977). This hypothesis is based on the fact that addition of low concentrations of purines reduces the fungitoxicity of benomyl and thiabendazole to Fusarium oxysporum (Cremlyn, 1977).
1.6 Environmental Fate of Fungicides Applied to Manage Patch Diseases
The efficacy of any fungicide is directly dependent on its environmental fate. A key factor in either preventative or curative fungicide applications is to place a sufficient amount of the compound in the right place at the right time. In the case of summer patch the fungicide must be delivered to the top 8 cm of the soil (where the fungus is found) in a timely marmer and in an amount sufficient to manage the disease. This section will deal with impediments to placing the fungicide in this 'active zone' including drift, photochemical degradation, volatility, runoff, plant uptake, binding, degradation, and
leaching. Some specific pesticide-soil-water-crop interactions influence the fate of these
compounds including:
1) Chemical properties: solubility, saturated vapor density, chemical diffusion coefficient,
lipophilicity, and adsorption coefficient.
2) Soil properties: bulk density, hydraulic properties, percentage organic matter, proportion
of clay minerals, and soil structure.
3) Boundary conditions at the soil surface: irrigation and rainfall frequency, rates, and
duration, pesticide timing, rate formulation, and depth of incorporation.
18 4) Plant characteristics: root and cover growth, transpiration, nutrient and chemical uptake patterns (Wagenet and Hutson, 1990).
1.6.1 Drift
While the concept of drift is ill defined it can be considered the aerial, windbome
movement of the spray mixture (pesticides, adjuvants, carrier, and solvent) away from the
target site. According to Lin and Graney (1992) drift is most important for aerial
applications but can occur during ground applications. Drift is usually associated with
hydraulic sprayers that are not used in applying turfgrass pesticides. Turf pesticides are
applied close to the ground over a dense canopy of plants. Aerial drift during turf pesticide
application can therefore be considered minimal.
1.6.2 Photochemical Degradation
Photochemical reactions are a contributing factor in the abiotic degradation of pesticides
in the environment. Testing of photochemical degradation is most advanced for the gas
phase of the atmosphere, less developed for surface waters, cloud droplets, and aerosols,
and least developed for the soil compartment (Klopffer, 1992). The difficulty in working
with non-homogeneous surfaces is a major factor in the incomplete development of testing
photochemical degradation in the soil compartment.
While biodegradation may dominate the environmental fate of pesticides in the soil,
abiotic degradation controls the fate of pesticides in the air and water. All abiotic
degradation, except for hydrolysis and abiotic reduction, is initiated by solar radiation and
may be classified as a photochemical processes (Klopffer et al., 1982; Wayne, 1988).
Whether in the atmosphere, water, or soil there are two main types of photochemical
19 degradation: photochemical oxidative degradation (indirect photochemical reactions) and direct photochemical reactions (formerly referred to as photolysis) (Klopffer et al., 1982). In water, direct photochemical reactions usually only occur near the surface of the water or in cloud or fog droplets.
In the soil, only surfaces exposed to solar radiation can contribute to photodegradation
(Kotzias and Parlar, 1985). In the management of summer patch, the fungicides are applied in a large quantity of water and then subsequently irrigated in an attempt to deliver them deeper into the soil. This should reduce their residence time on the surface. However, in hot, dry conditions, soil water may move back toward the surface, transporting the fungicide to the soil surface where it may undergo photochemical degradation (Klein et al., 1991).
There are many factors which make the study of photodegradation in soils problematic.
These include competitive sunlight absorption by soil chromophores, variable sorption of pesticides on organic and inorganic colloids, competing biotic and abiotic transformation processes, and thermal heating of the soil surface which enhances the convective and evapotranspirative transport of hydrophobic organics to the surface, all of which complicate the study of photochemical kinetics in soil (Hebert and Miller, 1990). There are very few data on photodegradation on soil surfaces. In laboratory studies, Hebert and Miller (1990) found that direct photodegradation was restricted to the photic depth of the soils (0.2-0.4 mm), with indirect photodegradation extending slightly deeper. Irradiation with natural sunlight resulted in photodegradation at greater depths (up to 2 mm).
Indirect photochemical processes can also affect the fate of surface applied pesticides.
Soil surfaces exposed to sunlight can generate a variety of oxidants which could transform
20 xenobiotics (Pohlman and Mill, 1983). A study by Gohre and Miller (1986) found that irradiated soils can generate a significant amount of singlet oxygen, thought to be able to diffuse to depths approaching 1mm.
Buchenauer et al. (1973) commented that thiophanate-methyl was converted to carbendazim by UV light but that carbendazim itself was hardly affected by irradiation.
However, it was later found that under higher light intensities greater than 90% of the carbendazim was inactivated in two hours, and was converted to another fungitoxic compound with a bimodal pattern of inhibition (Watkins, 1974). Carbendazim apparently breaks down to dimethyl oxalate, dicarbomethoxyguanidine, monocarbomethoxyguanidine, and possibly an acid salt of guanidine (Watkins, 1974).
Abdou et al. (1986) exposed carbendazim in methanol to UV light in the presence of singlet oxygen for 600 hours and found 60% of the carbendazim transformed to the first three compounds mentioned above as well as 2-guanidinobenzamidazole, 2- aminobenzimidazole and benzimidazole. Irradiation of carbendazim in aqueous HCl (15%) for 200 hours converted 75% of the compound to 2-guanidinobenzimidazole, benzimidazole, 2,4'-bibenzimidazole and 2,5'-bibenzimidazole. Benomyl irradiated in a 1% methanol solution yielded dicarbomethoxyguanidine, carbendazim, monocarbomethoxyguanidine, monocarbomethoxyurea, 2-guanidinobenzimidazole, 2- aminobenzimidazole and benzimidazole (Abdou et al., 1986) after 300 hours, at which time all benomyl had disappeared. When benomyl was irradiated in aqueous HCl, it yielded the same products as those produced by carbendazim in HCl. Sixty percent of the carbendazim in methanol was transformed in 600 hours, while 75% was transformed in 200 hours when
21 the compound was irradiated as a hydrochloride salt (Abdou et al., 1986). The nature of the reaction medium therefore plays a decisive role in the singlet oxygen photodegradation of carbendazim and benomyl.
1.6.3 Volatility
Volatilization plays a major role in pesticide dispersion in the environment, and there is much literature on the many different pesticides that have been found at various times in the atmosphere (Glotfelty et al., 1984; Lee, 1976; Scharf et al., 1992). For extremely persistent pesticides volatility may be the major pathway by which pesticide residues in plants and soil are decreased (Dorffler et al., 1991). Transport following volatilization can lead to considerable wet and dry deposition of pesticides (Glotfelty et al., 1987; Scharf et al.,
1992). Some pesticides are apparently transported long distances before deposition (Atlas and Giam, 1981; Eisenreich et al., 1981; Harder et al., 1980; Zell and Ballschmitter, 1980).
Even pesticides with a low vapor pressure can reach the atmosphere by volatilization. For example, in Germany during one rain event the rainwater was found to contain 0.295ug L ^ of propiconazole (Scharf et al., 1992).
Soil water content is the most important determinant of volatilization (Glotfelty et al.,
1984; Spencer et al., 1973). Many studies have found a correlation between soil moisture and volatility (Glotfelty et al., 1984; Majewski et al, 1989,1990; Spencer and Cliath, 1974).
Majewski et al. (1989) found that air concentration dropped off dramatically during the first few hours after sunrise and usually by noon pesticide residues in the air were below the detection limits. The reduction in volatility may be due to the drying out of the top few millimeters of soil which effectively stopped pesticide evaporation (Glotfelty et al., 1984;
22 Spencer et al., 1973,). High flux values frequently coincide with the presence of dew
(Majewski et al, 1989, 1990) or rain (Majewski et al., 1989).
Increased vaporization of pesticides from moist soil is due to the increased vapor pressure caused by the displacement of the chemical from the soil surface by water (Spencer and Cliath, 1969, 1975). Pesticides and water are in competition for sorption sites in the soil. The more strongly a pesticide is adsorbed, the greater the moisture content of the soil must be in order to achieve the maximum pesticide vapor density in the soil. Organic matter also affects volatility by increasing the capacity of the soil to adsorb pesticides (Dorffler et al., 1991; Spencer and Cliath, 1974). Glotfelty et al., (1984) found that weakly polar pesticides become more strongly adsorbed as soil organic matter content increases. This stronger adsorption was found to restrict vapor loss. An increase in soil temperature is usually associated with increased volatility, but soil temperature has a much less dramatic effect than water (Majewski et al, 1989), and may, in fact, reduce volatility by drying out the surface of the soil (Spencer and Cliath, 1969).
Soil incorporation has been shown to restrict losses by volatilization (Taylor, 1978;
White et al, 1977). If the fungicides can be placed deeper in the soil one would expect volatility to be decreased. The function of two of the pieces of application equipment to be used in this study (the high pressure injector and the modified slicer/seeder) is to deliver the fungicide just below the soil surface. One would expect little volatilization from these plots.
The traditional boom sprayer, on the other hand, applies the compounds on top of the turf canopy, which intercepts much of the fungicide. In this case one would expect volatility to be more important.
23 1.6.4 Runoff
Loss through runoff is not generally considered a major pathway for the loss of turfgrass pesticides. Soil runoff from a turf stand is minimal, due to the density of the plants which protect the soil surface and act as a filter to restrict soil transport (Lin and Graney, 1992). It is therefore likely that the most important transport pathway is via pesticides in the runoff water.
The pesticide characteristics important in determining propensity for runoff include volatilization rate, plant uptake through the leaves, partitioning coefficient, foliar decay rate and wash off, and soil decay rate (Lin and Graney, 1992). Important environmental parameters include the kinetic energy, intensity, and duration of rainfall and the slope, texture, strength and permeability of the soil (Ahuja et al., 1981). These factors all influence pesticide contamination of runoff water.
The turfgrass system consists of a dense canopy of plants and a thatch layer above the soil. The thatch layer has a high organic matter content (Hurto et al., 1980) which can bind and adsorb many hydrophobic pesticides (Niemczyk and Krueger, 1982). While the thatch
layer may initially slow down water infiltration, even completely dried thatch slowed water
infiltration for only 10 minutes, after which time the infiltration rate equaled that for soil
without a thatch layer (Taylor and Blake, 1982). Thatch also has a high water-holding
capacity (Beard, 1973), and this can affect the hydrology of a runoff event. The depth of the
thatch layer may range from 0.5 to 5 cm, depending on age, grass species, and cultural
practices (Hamilton, 1990; Hurto et al., 1980; Taylor and Blake, 1982).
24 Ahuja et al. (1981) found that the rainfall-runoff-soil interactions are limited to the top
0.2 to 0.3 cm on bare, sieved soil. One might expect this interaction zone to be even less in an established turfgrass stand because the dense plant canopy would reduce the kinetic energy of the raindrops while the thatch/soil has good permeability and strength.
Both the SBI fungicides and benomyl are strongly adsorbed to both the soil and organic matter and are poorly soluble in water. As mentioned above, the turfgrass system practically eliminates the loss of soil and organic matter particles in runoff, and therefore any adsorbed chemicals. Runoff and mobility studies by Rhodes and Long (1974) found no benomyl or carbendazim in the runoff from greenhouse turf plots that received daily applications of water to runoff. Ninety-nine percent of the compounds were found within 10 cm of the
application site and although runoff occurred on these turf plots there was no soil erosion
(Rhodes and Long, 1974). In addition, runoff from turf plots occurs only in extreme rainfall
events (Watschke, personal communication). Kenimer et al. (1992) found that chemical
concentrations in runoff water were significantly lower than those in the soil solution,
indicating that equilibrium between pore water and surface water is not instantaneous.
These factors combine to lessen the significance of the transport of pesticides in runoff
water on turf By far the most important fate pathways for systemic compounds in turf is
uptake by the foliage, binding in the thatch, and degradation.
1.6.5 Plant Uptake
Penetration of the cuticle and uptake by the plant is an important pathway in the fate of t-
SBI fungicides and benomyl. Foliar uptake is a very complex process about which little is
known. It has been found that the relative uptake of compounds differs between plant
25 species (Stevens et al., 1988) making it impossible to extrapolate findings from one species to another.
Cuticular penetration is affected by a pesticide's physicochemical properties, but there is no simple relationship between physicochemical properties of compounds and their uptake
(Baker et al., 1992). Lipophilicity is an important characteristic, as foliar penetration of structurally analogous compounds frequently increases with increasing lipophilicity (Norris and Bukovac, 1966; Sargent et al., 1969). This is most likely due to the hydrophobic nature of the epicuticular wax (EW) and cuticular membrane (Baker, 1982; Holloway, 1982). The
EW acts as both a barrier to polar chemicals and a sink for hydrophobic compounds
(Stevens et al., 1988). In general, lipophilic compounds with low melting points are taken up the fastest. These compounds, which include triadimefon and triadimenol, were found to form amorphous deposits that partially merged with the EW (Baker et al., 1992).
Molecular size and solubility in both aqueous and non-polar media may also affect uptake. Molecular size may affect the rate of diffusion through the cuticle, and its effects have been seen in both isolated cuticles (Schonherr, 1979) and whole plants (Price and
Anderson, 1985). Uptake occurs primarily with materials in solution, so uptake may be related to both water solubility and the octanol/water partition coefficient (Kq^)? although
by itself is not a good predictor (Stevens et al., 1988). The cuticle/water partition coefficient is more accurate, and it is directly related to molecular size (Sabljic et al., 1990).
Compounds of intermediate lipophilicity (log K^w^l) exhibit the most rapid uptake
(Stevens and Baker, 1987; Stevens et al., 1988).
26 Plant structures also affect uptake. Composition of the EW is at least partially
responsible for different rates of uptake by plants. Uptake of fenarimol was shown to be
faster in rape (whose EW consists of hydrocarbons and long chain alkyl ketones) and
strawberry (EW consisting of pentacyclic triterpenoid acids) than in maize and sugar beet
(EW predominantly primary alcohols) (Stevens et al., 1988). Baker et al. (1992) found that penetration of triadimefon in maize and sugar beet was much greater than that of triadimenol, but uptake by rape and strawberry was not statistically different. The maize leaf cuticle apparently represents a significant barrier to foliar entry, as uptake was lower for all non-polar chemicals tested, including triadimefon and triadimenol. Uptake was also found to be more rapid in younger, expanding apple leaves than older ones (O’Leary and
Jones, 1987).
Uptake through abaxial leaf surface is greater than that for the adaxial surface (O’Leary and Jones, 1987), possibly due to the numerous ectodesmata in the cells at the base of trichomes, which have thinner cuticles (Crafts and Foy, 1962; Franke, 1967). These cells have been shown to preferentially absorb compounds placed on the leaf (Hull, 1970). It is also possible that the thickness and orientation of the waxes in the cuticle of the adaxial surface present a more formidable barrier (Norris and Bukovac, 1969). Though some workers believed that a temperature coefficient of greater than 2 indicated active uptake
(Sargent et al., 1969), it is now believed that uptake increased at higher temperatures is due to changes in the properties of the EW (Norris and Bukovac, 1969) and possibly an increase in the kinetic energy of the diffusing molecule, allowing it to overcome potential energy barriers between the liquid phase of the droplet and the wax layers of the cuticle.
27 Uptake is also dependent on concentration (O’Leary and Jones, 1987), indicating a dependence on the establishment of a concentration gradient across the cuticle. In contrast to earlier studies (Franke, 1967; Kamimura and Goodman, 1964). O'Leary and Jones (1987) found that uptake of the compounds studied was not influenced by light.
Although benomyl is rapidly converted to carbendazim in water (Peterson and
Edgington, 1969) and plants (Baude et al., 1973; Fuchs et al., 1974; Peterson and
Edgington, 1970; Upham and Delp, 1973), it appears to be relatively stable on the leaf surface (Baude et al., 1973; Upham and Delp, 1973). Baude et al. (1973) found that 49 hours after application, 48 to 77% of the combined residues on the leaves were found to be benomyl, indicating that benomyl remains on the foliage as a rather stable residue and is only gradually converted to carbendazim.
Benomyl is taken up through the cuticle more rapidly than carbendazim (Solel and
Edgington, 1973; Upham and Delp, 1973). Once inside the plant, benomyl is rapidly converted to carbendazim and, in fact, intact benomyl is rarely reported to be recovered from plants. Most studies agree that any systemic translocation and fimgitoxic effects removed from the site of application should be attributed to carbendazim (Baude et al.,
1973; Clemons and Sisler, 1969; Fuchs et al., 1972; Hammerschlag and Sisler, 1972, 1973;
Peterson and Edgington, 1970; Siegel and Zabia, 1972; Sims et al., 1969; Upham and Delp,
1973).
The dense plant canopy found in the turfgrass system provides a significant barrier to
applying fungicides to the root zone. In intensively managed turfs, where summer patch is a
serious problem, the plant density is greatest. The rapid uptake, of SBI fungicides and
28 benomyl is considered a benefit when treating foliar diseases, but it is problematic when attempting to deliver fungicides to the root zone. The inconsistent chemical control of this disease could be due to differences in plant density and/or the thickness of the thatch layer.
1.6.6 Thatch Layer
Once the fungicides penetrate the canopy they encounter another barrier, the thatch layer.
The thatch layer is a tightly intermingled layer of living and dead stems, leaves, and roots of grass that develops between the verdure and the soil surface (Beard, 1973). The thatch layer is therefore rich in humic materials. Humic substances have many effects on the movement of pesticides including binding of the compounds, increasing their solubility, and possibly increasing the rate of microbial degradation through an increase in microbial activity.
Humus is derived primarily from higher plants during microbial decomposition of the original plant constituents and from new substances synthesized by soil microorganisms
(Haider, 1980). Humic compounds have widely varying chemical structures due to the many different sources from which they are derived and the influence of climatic factors and the mineral structure of the soil (Haider, 1980).
Humic materials are divided into fractions based on their behavior in dilute acids and bases. Humic acid (HA) is soluble in dilute alkali and is precipitated by acidification, while fulvic acid (FA) remains in solution when the alkaline extract is acidified. The third fraction is humin, which cannot be extracted with dilute acid and base (Khan, 1980a). Some characteristics of humic substances arer resistance to microbial degradation, formation of water insoluble complexes with metal ions and hydrous oxide, and the ability to interact with clay minerals and hydrophobic organic compounds such as pesticides (Khan, 1980a)
29 through partitioning, adsorption, catalysis, hydrolysis, dealkylation, photosensitation
(Senesi and Chen, 1989) and incorporation (Bollag et al., 1992). The polydispersed, polyelectrolyte character, surface activity properties, various reactive functional groups, free radical moieties, and hydrophobic and hydrophilic sites in the molecular structure are important factors in the interaction between pesticides and organic matter (Langvik et al.,
1994).
Humic substances are open flexible structures perforated by voids of varying dimensions that can trap or fix organic or inorganic compounds that fit into these voids, acting as a molecular sieve (Khan, 1980a), in addition to reacting via the oxygen containing functional groups.
It has long been known that some pesticides remain associated with the soil for extended periods of time (Bollag et al., 1992), and organic matter content is the most important soil factor relating to the binding of pesticides (Bollag et al., 1992; Senesi, 1992; Singh et al.,
1989). Binding is also affected by the chemical and structural characteristics of the pesticide
(Bollag et al., 1992; Parris, 1980). Binding can occur with either the parent compound or a metabolite. Compounds may become bound through either abiotic or biotic agents, with the reactions and processes similar to those responsible for the humification process (Bollag and Bollag, 1989; Bollag et al, 1992). Pesticide binding can occur through sorption (Van der Waals forces, hydrogen bonding, hydrophobic bonding), electrostatic interactions
(charge transfer, ion exchange, ligand exchange), covalent bonding, or a combination of these factors (Bollag et al., 1992; Khan, 1978; Pignatello, 1989; Senesi, 1992; Senesi and
Chen, 1989). SBI fungicides are not ionic at normal soil pH, so ionic mechanisms are
30 probably not important, but all other mechamsms may apply. Carbendazim, with an acid
disassociation constant (pka) of 4.48, may be ionized in acidic soils.
A portion of the compounds bound by sorption remains available to interact with the soil
environment, indicating reversibility (Bollag et al., 1992; Parris, 1980). Over time, bound
residues become more stable and resistant to extraction or degradation. This may be the
result of a redistribution of the pesticide from a weaker to a stronger site and/or the
formation of covalent bonds (Calderbank, 1989). Covalent bonding leads to more persistent
complexes that are very stable, resisting acid or base hydrolysis, thermal treatment, and
microbial degradation (Bollag et al., 1978; Helling and Krivonak, 1978; Hsu and Bartha,
1974; Saxena and Bartha, 1983).
Oxidative coupling reactions are believed to be responsible for the incorporation of pesticides into humic substances (Bollag and Bollag, 1989; Bollag et al., 1992). These reactions are mediated by plant and microbial enzymes, inorganic chemicals, clay, and soil extracts (Bollag et al., 1983; Shindo and Huang, 1983; Wang et al., 1986). These reactions eventually lead to polymerization. Enzymes that are able to catalyze oxidative coupling reactions include peroxidases and monophenol monooxygenases [such as a laccase from
Rhizoctonia praticola] (Bollag and Bollag, 1989), both of which require the presence of oxygen but no coenzyme for activity. The presence of a reactive electron donor, such as ferulic or syringic acid, can cause even relatively inert aromatic structures to be oxidized
(Bollag et al., 1992; Shuttleworth and Bollag, 1986).
The binding of compounds reduces their toxic effects (Bollag and Bollag, 1989;
Richards and Shieh, 1986) by decreasing the amount of fungicide available to the biota.
31 reducing the amount available and, therefore, its relative toxicity (McCarthy and Jiminez,
1985; Ogram et al., 1985). Binding is generally thought to reduce the leaching and transportability of compounds (Bollag et al., 1992), and it can also result in the formation of insoluble precipitates which can be removed from aquatic environs by filtration or sedimentation (Klibanov, 1980; Maloney et al., 1986).
The release of bound xenobiotics is not well understood, but it is believed to be due primarily to microbial activity. It is known that certain soil fimgi such as Penicillium frequantans can degrade humic substances and, by doing so, may free up bound residues
(Mathur and Paul, 1967). Microbial release depends on the type of binding, with covalently bound materials much more resistant. Mineralization of humus has been estimated at 3 to
5% a year in temperate climates. This corresponds to the rates of mineralization found in several studies of bound pesticides (Bollag et al., 1992; Hsu and Bartha, 1974, 1976;
MacRae, 1986).
Benomyl is rapidly converted to carbendazim in the soil (Baude et al., 1974; Khan,
1980b; Woodcock, 1978), and carbendazim is known to adsorb to organic matter (Helweg,
1977). A soil thin layer chromatography study (Rhodes and Long, 1974) found that carbendazim had no movement at all when the plate was run using a muck soil (85% organic matter) even though this soil had the highest pH, a condition under which carbendazim is believed to be more mobile (Aharonson and Kafkafi, 1975; Schreiber et al.,
1971). Other studies have shown that uptake of carbendazim increases with decreasing organic matter content of the soil (Schreiber et al., 1971), indicating greater availability of carbendazim in soils with a low organic matter content.
32 The thatch layer seems expressly designed to prohibit the movement of pesticides to the
soil. The presence of heterocyclic compounds in humus indicates these compounds, or at
least their aromatic moieties, can become polymerized with HA and FA, as well as interacting with the humus through any number of binding mechanisms. The aliphatic side chains found on SBI fungicides help to impart a lipophilic character on these compounds, aiding in their partitioning into the humus fraction of the soil.
1.6.7 Degradation
In general, there is little information on the behavior of SBI fungicides in the soil, and this holds true for their degradation. This may be due to the fact that they are not usually applied directly to the soil but as foliar sprays (Patil et al., 1988). Degradation can occur through either biotic or abiotic means. The rate of degradation is influenced by the properties of the soil, the microbial population and the physicochemical properties of the pesticide.
Important soil factors include the amount and type of soil organic matter, the soil type, soil moisture, pH, and temperature. Soil pH affects degradation primarily by changing the rate of hydrolysis and also the makeup of the microbial population (Schoen, 1987). A higher content of organic matter can increase the rate of degradation for several reasons, including lowering the pH which increases the rate of reactions that are normally faster under acidic conditions, increasing the rate of surface catalyzed reactions, and providing a substrate for microorganisms which may increase cometabolism (Schoen, 1987). There is a small correlation between soil type and degradation, but, in general, increased clay content increases degradation, possibly through an increase in surface area allowing for greater
33 catalysis (Kowalska et al., 1994; Schoen, 1987). Schoen et al. (1987) state that under dry conditions surface catalysis would be expected to predominate, while moist conditions would favor reactions in solution or microbial degradation. A study on the degradation rates of a series of substituted benzyltriazoles as well as triadimefon found that the rates increased as the water content of the soil increased (Patil et al., 1988). The same study also found that degradation rates increased from 5 to 10°C but ftirther temperature increases had little effect.
The composition of the microbial population also affects the rate of degradation.
Protozoa (Murphy et al., 1982), fungi (Helweg, 1972; Nakanishi and Oku, 1969), actinomycetes (Chacko et al., 1966), and bacteria (Helweg, 1972) have all been shown to degrade fungicides. A soil with a high organic matter content, such as that found under turf, generally has a higher microbial population. This could increase the rate of microbial degradation. Enhanced degradation, defined as an increase in the degrading activity of the soil microflora towards a pesticide after repeated applications, has been shown to occur with the fungicides iprodione, vinclozolin (Walker et al., 1986) and benomyl (Yarden et al.,
1985) as well as the insecticide isofenphos in turfgrass thatch (Niemczyk and Chapman,
1987). Higher degradation rates were observed in soils previously treated with benomyl regardless of the soil type, crop, or method of application (Yarden et al., 1987). Accelerated degradation was also found in soils where only foliar applications occurred, indicating that even a low dose is enough to condition the soil.
The single most important pesticide factor related to degradation is concentration.
Degradation rates have been found to be much slower at high concentrations (Schoen,
34 1987) . Other important factors include the pesticide’s nutritive value, microbial toxicity, availability to soil microorganisms, and the properties of its metabolites (Somasundaram et al., 1991). Degradation in mixtures is usually slower (Schoen, 1987), and this may be due to microbial inhibition or an interference with soil-catalyzed reactions.
Several compounds containing triazole groups are very persistent, including the fungicide
PP450, which has a half-life of 578 days (Patil et al., 1988). Triazole compounds with electron donating groups are generally degraded more slowly than those with electron withdrawing groups, which are fairly strongly adsorbed (Patil et al., 1988). The half-life of triadimefon is 15 days but its metabolite triadimenol degrades very slowly (Patil et al.,
1988) . In fact, triadimefon has been shown to control fungi the year after application
(Demoeden, 1989b; Rawlinson et al., 1982). This prolonged period of selection pressure increases the chance of the appearance of resistance. Such persistence is obviously undesirable.
The first step in the degradation of benomyl is its conversion to carbendazim. As mentioned previously, this occurs readily in water, plants, and soil, and its rate depends on pH and temperature (Fuchs et al., 1972). Benomyl is not usually found in the soil after four weeks (Baude et al., 1974), but carbendazim is much more persistent. The half-life of carbendazim in the soil has been estimated at anywhere from three months to two years
(Austin and Briggs, 1976; Baude et al., 1974; Edwards and Thompson, 1973; Hine et al.,
1969; Janutolo and Stipes, 1978). The half-life found when applied to turf was estimated at
3 to 6 months (Baude et al., 1974). The rate of degradation is accelerated at higher pH
35 (Austin and Briggs, 1976), at higher temperatures (Li and Nelson, 1985), and at higher soil moisture contents (Fuchs et ah, 1970; Li and Nelson, 1985).
Benomyl is known to be degraded by soil microorganisms (Fuchs and DeVrie, 1978;
Gupta and Chatrath, 1979; Helweg, 1972, 1973, 1977; Yarden et ah, 1985). Fumigation, solarization, and autoclaving soil have all been shown to significantly extend the half-life of carbendazim (Yarden et ah, 1985), indicating the importance of microbial breakdown to pesticide persistence.
Although degradation can be viewed as a barrier, it is important that fungicides are not too persistent. The half-lives of propiconazole and triadimefon are acceptable at about 20 days. Triadimenol, however, has a half-life of 110 to 270 days, and fenarimol has a half-life of 365 days (Pesticide Manual, 1994). Therefore, one would expect triadimenol and fenarimol to reach the root zone in greater amounts than any of the other compounds.
1.6.8 Mobility
Fungicide behavior in the soil has generally been less studied than that of the widely applied herbicides, however, the principles of pesticide movement in soils is widely applicable to organic compounds. Many of these have already been covered in this proposal. This section will focus on movement and its relationship to chemical and soil properties.
Many soil properties affect the mobility of pesticides. Organic matter content is negatively correlated with the movement of non-ionic organic chemicals through soil
(Goss, 1992; Kowalska et al., 1994; Somasundaram et al., 1991). Clay content, cation exchange capacity (CEC), and water-holding capacity are also negatively correlated with
36 mobility. Adsorption has been found to be greater in soils with a high CEC (Somasundaram et al., 1991). The effect of all these is considerably less than the effect of organic matter, as seen above.
The pH of the soil also has an effect on mobility, with movement greater at higher pH
Somasundaram et al., 1991), possibly due to stronger adsorption at a lower pH (Renner et al., 1988). Particle size, and sulfur and oxygen content have also been shown to affect mobility for certain compounds (Garibarini and Lion, 1986; Rippen et al., 1982).
Chemical properties of the pesticide also affect its mobility. Some of these properties include water solubility (Kanazawa, 1986; Somasundaram et al., 1991), octanol/water partition coefficient (Ko^) (Jury et al., 1987; Kanazawa, 1986; Kawamoto and Urbano,
1989; Somasundaram et al., 1991), (Gustafson, 1989; Hodson and Williams, 1988; Jury et al., 1987; Kanazawa, 1986; Kawamoto and Urbano, 1989), molecular weight (Kanazawa,
1986) and half-life (Gustafson, 1989; Jury et al., 1987). The of a compound is the ratio of adsorbed chemical per unit weight of organic carbon to the concentration of the chemical in aqueous solution (Hodson and Williams, 1988).
More soluble chemicals usually have a lower soil sorption coefficient (Somasundaram et al., 1989). A significant relationship has been found between water solubility and mobility
(Kanazawa, 1986; Somasundaram et al., 1989). Solubility by itself is not a good predictor of mobility, but is adequate when combined with adsorption (Somasundaram et al., 1989).
Water solubility is influenced by temperature, pH and the composition of the soil medium
(Somasundaram et al., 1989).
37 Pesticides may be carried out on migrating colloidal particles on which they are assumed to be sorbed (Chaplain and Mills, 1992; Magee et al., 1991). Conversely, these interactions between migrating particles and the porous soil matrix may cause a reduction in mobility due to the physicochemical interactions between colloidal particles and the porous matrix.
This could lead to trapping of the pesticide in the soil (Chaplain and Mills, 1992).
The presence of macropores could possibly lead to the movement of a compound past the root zone, where degradation rates are the greatest (Isensee et al., 1990). It is well documented that macropores can rapidly transport water through the soil (Isensee et al.,
1990; Shipitalo et al., 1990). There is generally a large number of macropores under turf due to earthworm activity. Macropores have been implicated in increased mobility of pesticides in soil under no-till agriculture (Czapar et al., 1992; Isensee et al., 1990; Shipitalo et al., 1990). Rainfall timing is critical for macropore transport. An initial, light rain can move the solutes into the soil matrix thereby reducing the potential for macropore transport
(Shipitalo et al., 1990). A heavy initial rainfall immediately after application, however, may transport even strongly adsorbed chemicals past the root zone (Shipitalo et al., 1990).
Pesticide movement through soil can be considered as a chromatographic process with the compound partitioning between the moving water and the water saturated soil
(Gustafson, 1989; Jamet and Eudeline, 1992; Mestres et al., 1987). This explains the usefulness of and the organic carbon/water partition coefficient ^ study of the movement of triazole fungicides, mobility was found to be inversely related to (Jamet and Eudeline, 1992). The of compounds has been found by other authors to be a good predictor of mobility (Kanazawa, 1986; Kawamoto and Urbano, 1989; Vowles and
38 Mantoura, 1987). Estimates of can be made by using retention times on reverse phase
HPLC (Hodson and Williams, 1988; McCall et al., 1980) or on an alkylcyano column, which may better mimic sorption onto soil (Vowles and Mantoura, 1987).
Benomyl and carbendazim are considered to have low mobility in the soil (Aharonson and Kafkafl, 1975; Baude et al., 1974; Helling, 1971; Hine et al., 1969; Rhodes and Long,
1974). Austin and Briggs (1976) found neither benomyl nor carbendazim below 25 cm of the soil surface during a 10 month study of benomyl applied to bare soil. Another study using turf found most of the residues (0.5-1.1 ppm) in the top 10.16 cm of the soil, while concentrations in the 10 to 20 cm section were 0 to 0.05 ppm (Baude et al., 1974). This was following the application of 9.07 kg ai acre'^ in five foliar applications 10 days apart, followed by daily watering to percolation. Rhodes and Long (1974), in a study of the fate of benomyl in turf, found greater than 95% of the compound in the top 4 cm of the soil even under the rigorous conditions of their experiment and never detected any benomyl or carbendazim in the percolation from the turf plots. The authors also ran the compounds on soil thin layer chromatography plates and placed carbendazim in mobility class I
(immobile) and benomyl in mobility class II (slightly mobile) according to the mobility classification suggested by Helling and Turner (1968).
At present, the universally accepted way to compare mobility is by (Gustafson,
1989). Pesticides with a of greater than 300 ml g’’ are considered to be strongly adsorbed to the organic matter in the soil (Goss, 1992). There is also a relationship between
Kqc retention time on reverse phase HPLC (Kanazawa, 1986).
39 Half-life is critical in determining whether a compound will leach below the root zone
(Bottom and Funari, 1992; Goss, 1992; Jury et aL, 1987; van der Zee and Boesten, 1991). A pesticide with a long half-life has the potential for leaching even if it is strongly adsorbed in the soil. It is considered a positive attribute if the surface residence time of a compound exceeds its half-life significantly (Ghodrati and Jury; 1992, Jury et al., 1987; Rao et al.,
1985).
In general one would expect to find little of these fungicides, with the possible exception of triadimenol, in the soil. The relatively short half-life of triadimefon and propiconazole would seem to preclude their being around long enough to reach the soil in any appreciable amount. The lipophilic character of these compounds, as well as their relatively high and Kqc of the fungicides used would seem to preclude their movement into the soil. The high rates of water in which these fungicides are recommended to be applied are indicative of this problem. It is possible that the high pressure injector or the modified slicer/seeder could overcome these barriers to the delivery of fungicides to the target zone.
40 CHAPTER 2
ENVIRONMENTAL FATE OF FENARIMOL, PROPICONAZOLE, AND
TRIADIMEFON APPLIED TO KENTUCKY BLUEGRASS UTILIZING THREE
APPLICATION SCHEMES
2.1 Introduction
Public concern over the application of pesticides on golf courses has intensified in recent years due to their increased use and the ability of analysts to detect ever lower concentrations of these chemicals (Lehr, 1991), Among these concerns is the impact of turfgrass fungicides on groundwater quality. Triadimefon, a widely-used sterol biosynthesis inhibiting (SBI)-fungicide, has been demonstrated to leach through golf course greens constructed of sand, potentially reaching groundwater (Petrovic et ah,
1994). Less is known about the behavior of other SBI fungicides in soil and in turfgrass systems.
Summer patch, caused by the fungus Magnaporthe poae , is a very damaging disease of Poa and certain Festuca species (Jackson, 1993). Magnaporthe poae is a member of a group of ectotrophic, root-infecting fungi. Under favorable climatic conditions (i.e. hot and rainy weather), this pathogen is able to invade the vascular tissue of turfgrass roots, causing plant death (Landschoot et al., 1993). For turfgrass protection from this type of disease, it is imperative to deliver fungicides effectively to the roots in sufficient amounts prior to the onset of infection.
41 Chemical control of summer patch has been both erratic and expensive (Demoeden,
1993). This is due, in part, to inefficient delivery processes for fungicides. The lack of efficacious control in the absence of large amounts of water may indicate a problem in delivering fungicides to the top 8 cm of the soil where the fungus survives ( B. Clarke,
Rutgers University, personal communication). Fungicides have been shown to be more effective when delivered in a large amount of water (e.g. 1600-2000 L H2O ha'*)
(Demoeden, 1989a; Demoeden and Minner, 1981; Demoeden and Nash, 1982; Hartman et al., 1989; Landschoot and Clarke, 1989; Plumley et al., 1991; Smiley, 1980), or 800-
1000 L ha'* and followed by irrigation. Excess water, however, may provide the physical means to transport SBI fungicides from the turfgrass surface to groundwater. Such contamination would be unacceptable by today’s regulatory standards and would greatly increase the environmental impact of these fungicides.
Three SBI fungicides, fenarimol (Rubigan"^^), propiconazole (Banner"^^), and triadimefon (Bayleton"^^), were chosen for comparison in this study. Although similar in their modes of action, these fungicides vary in their water solubility, vapor pressure, environmental persistence, and toxicity (Table 1).
Fenarimol is the least water soluble of the three fungicides (13.7 mg L'*, Pesticide
Manual, 1994). Propiconazole is roughly seven times more soluble, and triadimefon is five times more soluble than fenarimol. Triadimefon has the shortest half-life in soil of the three fungicides at 6 to 18 days (Pesticide Manual, 1994). The soil half-life of propiconazole is approximately 4.5 times longer, and the half-life of fenarimol is over 30 times longer than triadimefon. Triadimefon exhibits the greatest mammalian toxicity with
42 Table 1. Physical properties, environmental characteristics, application rates, and toxicities of three SBI fungicides. w 3 3 -o > < H c/3 . O ■s .o < <1/ r^:^ 0 cC 0 Vi (U a, C GO O u O 'rt ^ ”7 ^3 U c3 U
ite in soil and water, respectively. an oral LD50 of 1000 mg/kg for rats (Pesticide Manual, 1994). Propiconazole is slightly
less toxic, with an oral LD50 of 1517 mg/kg and fenarimol is the least toxic to mammals
with an oral LD50 of 2500 mg/kg (Pesticide Manual, 1994).
There are several physical barriers that impinge on the effective delivery of fungicides to the roots in a turfgrass system. With traditional boom sprayer application, the first barrier is the leaf canopy. Fungicides, including the SBIs, are absorbed by the epicuticular wax and cuticle of the plant (Baker et al., 1992). Since these compounds are translocated primarily acropetally (i.e. root to shoot), they do not enter root tissue in sufficient concentrations following foliar applications to elicit fungicidal action in the roots (Kuck et al., 1995).
The next physical barrier encountered when attempting to deliver fungicides to the roots of turfgrass plants is the thatch layer. The organic matter content is the single most important factor affecting the binding of pesticides in environmental matrices, thereby retarding the movement of pesticides to groundwater (Bollag et al., 1992; Senesi, 1992;
Singh et al., 1989 ). Because the thatch layer associated with turfgrass has a very high organic matter content (Hurto et al., 1980), it has been shown to adsorb many hydrophobic pesticides (Lickfeldt and Branham, 1995; Niemczyk and Krueger, 1982), including the fungicide triadimefon (Dell et al., 1994). Thus, the thickness and organic matter content of thatch should have a pronounced effect on leaching, runoff, and volatilization of turfgrass pesticides.
44 Once SBI fungicides pass through the thatch layer, they enter into the soil and partition into the roots. Similar to the thatch layer, the most important factor influencing the mobility of fungicides in soil is its organic matter content (Goss, 1992; Kowalska et al., 1994; Somasundaram et al., 1991). Native soils underlying turfgrass generally have a high organic matter content, principally due to the breakdown of the thatch layer, which would retard the mobility of lipophilic pesticides.
The most important physicochemical property of a fungicide relative to its mobility in soil is its soil sorption constant (i.e. Kqc), which is based on organic carbon (Gustafson,
1989; Kawamoto and Urbano, 1989). According to the United States Environmental
Protection Agency (USEPA), any pesticide with a Kqc of less than 300 to 500 has the potential to leach through soil (USEPA, 1986). Overall, the Koc values for the SBI fungicides selected for the present study fall in the low to middle range of K^c values for turfgrass fungicides (Pesticide Manual, 1994). Fungicides with high values include benomyl (1900), chlorothalonil (1600-14000), and mancozeb (>2000). Those at the lower end of the spectrum include carbendazim (200-250), iprodione (373-1551) and vinclozalin (100-735). The relatively high Kq^ values for propiconazole and fenarimol
(i.e. propiconazole Koc= 1000; fenarimol Kq^ = 600) indicate that their rate of movement in soil of high organic matter content would be low. The relatively low value for triadimefon (i.e. Kq^ = 300) indicates that this fungicide should have greater soil mobility than either propiconazole or fenarimol (Pesticide Manual, 1994). Also, microbial metabolism and abiotic transformation of the parent fungicides can result in transformation products that have greatly enhanced environmental mobility.
45 The problems in the efficient delivery of the fungicides to the target zone of the roots of turfgrass, as discussed above, have stimulated interest in the use of alternative delivery technologies to boom sprayer application. Such novel technologies would have to allow better penetration of the thatch layer for direct delivery of the fungicides to the roots of turfgrass plants. Two novel application technologies under consideration in the current study include high pressure injection and slicer/seeder modified for pesticide application.
The present research examines these two novel application technologies to determine whether the use of such alternative application methods results in better delivery of the fungicides to the turfgrass root system in comparison with the commonly-used boom sprayer application method. The three SBI fungicides were chosen to determine if there was any difference in their mobility in the turfgrass system for fungicides of differing values when applied by various means. A number of important turf diseases are associated with root and crown infections, so these results are likely to have be applicable to other diseases in addition to summer patch. Additionally, there is concern that such alternative methods may bypass the turfgrass canopy and thatch layer and result in an increased potential for groundwater contamination via soil leaching. The turfgrass canopy and the thatch layer are known to bind many pesticides and limit their movement into the soil. Any process that bypasses these barriers could allow pesticides to reach the underlying soil in greater quantities, increasing the chance of groundwater contamination and human exposure.
46 2.2 Materials And Methods
2.2.1 Experimental Site
Three 12.2 m X 12.2 m (0.0149 ha) plots of Kentucky bluegrass of a mixture of cultivars were selected at the University of Massachusetts Turfgrass Research Center in
South Deerfield, MA. The turf was established in 1985 and overlay a Hadley silt loam
soil (coarse, silty, mixed, nonacid, mesic, Typic Udifluvent, pH 6.8). The turf was maintained at a height of 5.1 cm, and irrigated as needed to prevent drought stress (Fig.
1).
2.2.2 Fungicides
Three SBI fungicides were applied to the plots on 22 August 1992. The three fungicides
were fenarimol {(±)-2,4'-dichloro-a-pyrimidin-5-yl)benzhydryl alcohol}, propiconazole
{(±)-1 - [2-(2,4-dichlorophenyl)-4-propy 1-1,3 -dioxolan-2-ylmethy 1]-1 //-1,2,4-triazole, and
triadimefon {1 -(4-chlorophenoxy)-3,3-dimethyl-1 -(1 //-1,2,4-triazol-1 -yl)butan-2-one}.
2.2.3 Fungicide Application
The fungicides were applied using three types of applicators: 1) a standard boom
sprayer (F.E. Myers and Bro. Co., Ashland, OH); 2) a high pressure injector (Cross
Equipment, Albany, GA); and 3) a modified slicer/seeder (Cushman/Ryan, Lincoln, NE).
Each fungicide was applied at the recommended rate for summer patch disease control.
Fenarimol (Rubigan IAS, DowElanco, Indianapolis, INf) was applied at a rate of 0.76 kg
active ingredient (ai) ha \ propiconazole (Banner 1.1 EC, Ciba-Geigy, Greensboro, NC)
at 1.67 kg ai ha*^ and triadimefon (Bayleton 25 DF, Mobay, Kansas City, MO) at 3.05 kg
ai ha\
47 (N o^ o^
X) S .2 a
■£ o
0) Ol
3) ^ ii: <
48 ului ni (h) liejniBH pne (i) uopcSuJi
49 The boom sprayer was used to apply all three fungicides as a mixture to the first plot. The boom sprayer consisted of nine T-jet “9010” flat fan nozzles applying 621 L ha ^ at an operating pressure of 1925 kPa and generated a swath width of 4,6 m. The boom sprayer application was put on last and immediately followed by 1.3 cm of irrigation water applied to the entire experimental area.
The high pressure injector consisted of four 60 L tanks connected to a skid-mounted boom with 18 nozzles at a 7.2 cm spacing. Three separate passes were made across the second plot to apply each of the three individual fungicides directly into the thatch as an aqueous stream at a pressure of 11,100 kPa in 880 L ha V
The slicer/seeder was modified for pesticide application by placing a small plastic tank on top of the machine with 10 coulters spaced 5.0 cm apart leading to a point directly behind each sheer. The sheer created a furrow approximately 1.3 cm deep into which the tank mixture flowed via the gravity-fed coulters at a rate of 408 L ha'. Only propiconazole was applied with the modified sheer/seeder to the third plot.
2,2.4 Sampling
Each of the three plots was divided into sixteen 3.1 m X 3.1 m subplots that were
further subdivided into nine 1 m^ sections, one of which was randomly selected for each
sampling period. It is estimated that each square meter of the plot contained
approximately 455 g of leaf tissue (±15.7%), 11,989 g of thatch (±8.4%), 56,700 g of soil
(±3.9%) (1 m X 1 m X 5.1 cm), and 85 g of root tissue (±11.3%) (1 m X 1 m X 5.1 cm)
(all in wet weights). These calculated quantities were used to determine the percentages
50 of applied fungicides recovered. The amount of fungicide applied to each turfgrass matrix was calculated by assuming 100% of the applied fungicide was deposited in each matrix.
This number was then divided by the actual amount recovered to determine the percent of applied fungicide recovered. Leaf, thatch, roots and soil were sampled immediately before and after the applications and at 1,3,7,10,14,21, and 28 days post-application. At each collection period, 75 g of leaf tissue were collected from each of the sixteen subplots by cutting the plants at the soil line. The tissue was placed in 0.95 liter plastic Zip-Loc™ freezer bags for storage. Following this process, sixteen soil cores, one from each subplot, were taken with a 30.5 cm X 2.5 cm JMC Zero Contamination Tube Corer (Clements
Assoc., Inc., Newton, lA). The thatch layer of each core was delineated as the deepest point at which rhizomes were visible and was marked. Samples were then placed in ice chests and transported to the Massachusetts Pesticide Analysis Laboratory (MPAL) where they were stored at -20‘’C until analysis.
In the laboratory, the thatch layer was removed from the top of each soil core and the remainder of each core was divided into five 5.1 cm X 2.5 cm sections and bulked (i.e. the top 5.1 cm of the cores from each subplot for each application method during a sampling period were placed together in a 2.78 L Zip-Loc'^’^ freezer bag and mixed by rotating the bag for 3 min). The bulked samples were screened through a #10 sieve and the roots or thatch were separated out individually by hand. The soil, roots, and thatch were placed into individual plastic freezer bags for storage in a -20° C freezer for analysis.
51 2.2.5 Storage Samples
On each sampling date, equal amounts of nontreated (i.e. no applied fungicide) leaf tissue and soil cores (i.e., containing soil, roots, and thatch) were collected from an adjacent untreated area to be used as fortified storage samples (i.e., amended with the fungicides of interest to assess any loss during storage prior to analysis), concurrent fortified samples (i.e., to provide daily checks of the analytical method efficiency), and concurrent nontreated samples (i.e., to provide daily checks to assess background matrix
interferences). Following sample collection, leaf tissue was amended with 0.4 pg g \ thatch with 1.0 pg g\ roots with 2.0 pg g'^ and soil with 0.08 pg g'* of the fungicides of
interest and stored with the concurrently collected field samples to assess any loss of the
compounds during storage (i.e., fortified storage samples).
2.2.6 Analytical Procedures
Leaf: The determination of fungicide residues was based on a method developed by
Newsome and Collins (1990). Five g of leaf tissue were cut up using scissors and placed
in a 250 ml red actinic Erlenmeyer flask. Fifty ml of acetone were added to the flask and
it was shaken for 30 min on a rotary table top shaker (Eberbach) at low speed (180 rpm).
The liquid was then decanted through 13 g anhydrous sodium sulfate into a 500 ml flat
bottom boiling flask. The leaf tissue was re-extracted with an additional 50 ml acetone
that was likewise quantitatively transferred to the boiling flask. The combined extract was
reduced to approximately 3 ml under vacuum, quantitatively transferred in acetone to a
50 ml centrifuge tube, and the sample volume adjusted to 10 ml under a stream of
nitrogen. The sample volume was then readjusted to 50 ml with distilled water. Prior to
52 loading the sample extracts, C,g solid phase extraction cartridges (500 mg, J&W
Scientific, Folsom, CA) were sequentially washed with 10 ml aliquots of methanol, methylene chloride, and methanol, to remove any contaminants within the cartridge. The cartridge was activated with 25 ml distilled water. The sample extract was passed through the activated cartridge, followed by a wash step that consisted of 10 ml of a 40% solution of methanol in distilled water. The retained fungicide was eluted from the cartridge with
5 ml of methanol. Following elution, a 4 ml aliquot of the eluent was transferred to a 15 ml centrifuge tube and the volume adjusted to 10 ml with distilled water. A liquid/liquid extraction was performed in the centrifuge tube by adding 5 ml toluene and gently shaking the tube. The fungicide partitioned into the toluene (i.e. top layer), which was then removed for analysis. Each sample analysis was replicated three times using three subsamples.
Thatch: Fifty ml of acetone were added to 1 g of thatch in a 120 ml amber glass jar and the sample was homogenized twice for 20 seconds at 20,000 rpm with a Brinkmann
Polytron PT3000 homogenizer (Littau, Switzerland). The homogenized sample was shaken for 30 min on a rotary table top shaker, and the acetone decanted into a 500 ml flat-bottomed boiling flask. The extract was reduced to ca 3 ml under vacuum, quantitatively transferred in acetone to a 50 ml centrifuge tube and the sample volume
/ adjusted to 10 ml under a stream of nitrogen. The sample volume was then readjusted to
50 ml with distilled water. The Cjg cartridges were washed and activated as outlined for leaf tissue. The sample was passed through the cartridge followed by a 10 ml wash solution of 40% methanol in distilled water. The sample was eluted from the cartridge
53 with 10 ml acetone into a 13 ml centrifuge tube. The sample eluent volume was adjusted to 2 ml under a stream of nitrogen for analysis. Each sample analysis was replicated three times using three subsamples.
Soil: Fifty ml of acetone were added to 25 g of soil in a 250 ml red actinic erlenmeyer flask and shaken for 30 min on a rotary table top shaker at low speed (180 rpm). The liquid was decanted through 13 g anhydrous sodium sulfate into a 500 ml boiling flask.
The sample was re-extracted with 50 ml acetone and the extract quantitatively transferred to the boiling flask containing the first extract. The sample was reduced to ca 3 ml under vacuum. The remainder of the procedure followed that outlined for thatch above.
Roots: Twenty ml of acetone were added to 0.5 g of root tissue in a 60 ml amber glass jar. The sample was homogenized and shaken as previously described for thatch. The
acetone was decanted into a boiling flask. The sample was re-extracted with acetone and
the extract quantitatively transferred to the boiling flask containing the first extract and
reduced to ca 3 ml. The remainder of the procedure followed that outlined above for
thatch
2.2.7 Instrumental Analysis
Fungicide residues were analyzed using a Varian model 3400 gas chromatograph
equipped with a nitrogen-phosphorus detector (NPD) and a DB-5 glass megabore column
(53 mm i.d. X 15 m, J & W Scientific, Folsom, CA). Operating conditions were: injection
volume, 1.0 pi; injector temperature, 250°C ; detector temperature, 300°C; column oven
temperature, 160“C for 2 min, ramped at 15“C min' to 235“C and held for 3 min. Gas
54 flows were: helium, 10 ml min’'; hydrogen, 3.5 ml min’'; nitrogen, 20 ml min’'; and air
115 ml min’'.
2.2.8 Quality Assurance/Quality Control
2.2.8.1 Instrument Calibration
Calibration procedures for the chromatographic system used for analysis involved developing standard curves consisting of at least three concentrations for each fungicide of interest using high purity analytical standards (>99%). Triadimefon standards were provided by Mobay Corp. (Kansas City, MO). Fenarimol and propiconazole standards were provided by Riedel de Haen (Seelze, Germany). Standard curves consisting of at least three concentrations of each analyte were included at the beginning of each day of analysis. Area counts for each standard concentration (n=3) were averaged to generate a
2 calibration curve (r > 0.99).
2.2.8.2 Sample Controls
For each day of analysis, corresponding fortified storage samples and nontreated leaf, thatch, roots, and soil amended with the fungicides of interest and without (i.e. concurrent fortified and nontreated samples, respectively) were extracted and analyzed to determine
recovery efficiency and daily method performance, respectively. Reagent and matrix
blanks were also included in each analysis. Results are given in Table 2.
55 ^ ’N —^ ^ ^ (30 W5 C o cs M 1—( 00 ^ ^ B £ O -H. dd cd ed o sO eCA ^ ^ CT' t4-. ^ tN 00 r- a> O CA r-' tT vd o cd U o o^ o 00 & e p 32 43 (N vq ^ o o u, ro o 3 -*3 Tf .—I iri 00 Ov Os ^cB 'Ti -H -H -H dddld!^ 32 c sO sO N© 43 cd o vP vO ^ ^ ^ c/3 m <^ <—' ^ 2 e (3 O m’ o V o o o ci3 fc ^ o 2 o (3 c o o • O O cs o cq m rq o o' wS VO 3 iri x: 03 o o di -H -H -H vO vO 43 ^ vP •H 1 »r) H f<3 vq f4 ■§ S ^ 00 C « «d On ed t4-i 1^0 0 2 Ov ov to o
•B ^ VO o\ tn 00 r-‘ r-‘ (Nj r—I r—I C4-< •£ £ ed § 3-5 ddSdd 03 ® 03 ^ nO S© nO sO > 03 ^ CT' O *0 O 00 O o o o O (N r^' cs tsi ‘- (30 Ov o o t" OO 00 03 G (30 .p ed « B G C ^ ‘I 03 "o § B • *0 4> -o o 56 2.3 Results 2.3.1 Fungicide Concentration 2.3.1.1 Leaf Tissue Boom Sprayer Application Fenarimol: Residues of fenarimol associated with leaf tissue following the boom sprayer application were 42 ppm (± 3.1% coefficient of variance) immediately after application (Fig. 2A, Day 0) and attained a maximum concentration of 63 ppm (±3.0 %) one day after application (Fig. 2A, Day 1). Fenarimol residues declined over time to 2 ppm (±5.5 %) by 28 days post-application (Fig. 2A, Day 28). Propiconazole: Residues of propiconazole associated with leaf tissue following the boom sprayer application were 56 ppm (± 1.8 %) immediately after application (Fig. 2A, Day 0) and attained a maximum concentration of 89 ppm (± 3.3 %) one day after application (Fig.2A, Day 1). Propiconazole residues declined over time to 2 ppm (± 9.7 %) by Day 28 post-application (Fig.2A, Day 28). Triadimefon: Maximum triadimefon residues associated with leaf tissue following the boom sprayer application occurred during the first sampling period immediately after spraying (Fig. 2A, Day 0) at a concentration of 158 ppm (± 2.4 %). Triadimefon residues declined over time to 1 ppm (± 5.8 %) by Day 21 post-application (Fig. 2A, Day 21) and were not detected on Day 28 post-application. 57 Figure 2. Concentration (in ppm) of fenarimol, propiconazole, and triadimefon in leaf tissue pre-application (pre), immediately post application (Day 0), and 1,352,10,14,21, and 28 days following application with a boom sprayer (A), high pressure injector (B), and a modified slicer/seeder (C). 58 Concentration in Leaf Tissue (PPM) 200 150 100 200 150 100 50 Days Following Application 50 0 0 pre 59 21 28 21 28 Slicer/Seeder fenarimol propiconazole fenarimol propiconazole Modified triadimefon propiconazole triadimefon Pressure Injector Sprayer High Boom B A High Pressure Injection Application Fenarimol: High pressure injection application resulted in maximum fenarimol residues associated with leaf tissue (13 ppm, ± 1.3 %) immediately after application (Fig. 2B, Day 0). Fenarimol residues declined to 1 ppm (± 7.7 %) by Day 28 post-application (Fig. 2B, Day 28). Propiconazole: Maximum propiconazole residues associated with leaf tissue were 20 ppm (± 3.6 %) immediately after application with the high pressure injector (Fig. 2B, Day 0). Propiconazole residues declined to 1 ppm (± 3.5 %) by Day 28 post-application (Fig. 2B, Day 28). Triadimefon: Maximum triadimefon residues associated with leaf tissue following application with the high pressure injector occurred during the first sampling period immediately following application (Fig. 2B, Day 0) resulting in a concentration of 32 ppm (±1.3 %). Triadimefon residues declined over time to 1 ppm (±3.8 %) by Day 14 post-application ( Fig 2B, Day 14). No triadimefon residues were detected after Day 14 following application with the high pressure injector. Modified Slicer/Seeder Application Propiconazole: Residues of propiconazole associated with leaf tissue following application with the modified slicer/seeder attained a maximum concentration of 8 ppm (± 5.1 %) immediately following application (Fig. 2C, Day 0). Propiconazole residues declined to 1 ppm (± 5.4 %) by Day 21 post-application (Fig. 2C, Day 21), and were not 60 detected on Day 28 post-application. Leaf tissue samples taken in the pre-application period (pre, Fig. 2) had no detectable residues at the detection limits established. 2.3.1.2 Thatch Boom Sprayer Application Fenarimol: Residues of fenarimol associated with thatch following the boom sprayer application were 3.7 ppm (± 2.3%) immediately following application (Fig. 3A, Day 0) and attained a maximum concentration of 6.5 ppm (± 34.2%) three days after application (Fig. 3A, Day 3). Fenarimol residues declined over time to 2.62 ppm (± 5.8%) at 14 days post-application ( Fig. 3 A, Day 14) and then increased to 4.8 ppm (± 1.9%) at 28 days post-application (Fig. 3A, Day 28). Propiconazole: Residues of propiconazole associated with thatch following boom sprayer application were 5.8 ppm (+ 5.0%) immediately following application (Fig. 3A, Day 0) and attained a maximum concentration of 8.4 (± 18%) ppm three days after application (Fig. 3A, Day 3). Propiconazole residues declined over time to 3.2 ppm (± 6.6%) at 14 days post-application (Fig. 3A, Day 14) and then increased to 4.8 ppm (± 2.8%) by Day 28 post-application (Fig. 3A, Day 28). Triadimefon: Maximum triadimefon residues associated with thatch following the boom sprayer application occurred on the first sampling period immediately following application (Fig. 3 A, Day 0) resulting in a concentration of 10.7 ppm (± 2.5%). Triadimefon residues declined over time to 1.5 ppm (± 24.6%) at 3 days post- application(Fig. 3 A, Day 3). No triadimefon residues were detected following Day 3. 61 Figure 3. Concentration (in ppm) of fenarimol, propiconazole and triadimefon in thatch pre-application (pre), immediately post application ( Day 0), and 1,3,7,10,14,21, and 28 days following application with a boom sprayer (A), high pressure injector (B), and a modified slicer/seeder (C). 62 Concentration in Thatch (PPM) Days Following Application 63 fenarimol fenarimol propiconazole propiconazole triadimefon triadimefon Pressure Injector Sprayer High Boom B High Pressure Injection Application Fenarimol: Residues of fenarimol associated with thatch following the high pressure injection application were 2.1 ppm (± 11.0%) immediately following application (Fig. 3B, Day 0) and attained a maximum concentration of 4.0 ppm (±11.6%) 10 days after application (Fig. 3B, Day 10). Fenarimol residues associated with thatch declined over time to 2.6 ppm (± 19.2%) by 28 days post-application (Fig. 3B, Day 28). Propiconazole: Residues of propiconazole associated with thatch following application with the high pressure injector were 2.7 ppm (± 11.0%) immediately after application (Fig. 3B, Day 0) and attained a maximum concentration of 6.1 ppm (± 3.5%) 10 days after application (Fig. 3B, Day 10). Propiconazole residues in the thatch declined over time to 1.7 ppm (± 9.7%) by 28 days post-application (Fig. 3B, Day 28). Triadimefon: Maximum triadimefon residues associated with the thatch following high pressure injection application occurred on the first sampling period immediately after application (Fig. 3B, Day 0), resulting in a concentration of 7.4 ppm (± 26.0%). Triadimefon levels in thatch declined over time to 1.1 ppm (± 5.5%) 14 days after application (Fig. 3B, Day 14). No triadimefon residues were detected after Day 14. Modified Slicer/Seeder Application Propiconazole: Residues of propiconazole associated with thatch following application with the modified slicer/seeder were 2.1 ppm (± 3.6%) immediately after application (Fig. 3C, Day 0) and attained a maximum concentration of 7.3 ppm (± 5.3%) 64 at 7 days after application (Fig. 3C, Day 7). Propiconazole residues associated with the thatch declined over time to 1.1 ppm (± 3.4%) by 28 days post-application (Fig. 3C, Day 28). Thatch samples taken in the pre-application period (pre, Fig. 3) had no detectable residues at the detection limits established. 2.3.1.3 Soil Boom Sprayer Application Fenarimol: Residues of fenarimol associated with the soil following boom sprayer application were 28.8 ppb (± 3.9%) immediately following application (Fig. 4A, Day 0). Fenarimol levels in the soil generally increased over time and attained a maximum concentration of 76.0 ppb (± 8.2%) by 28 days post-application (Fig. 4A, Day 28). Propiconazole: Residues of propiconazole associated with soil following application with the boom sprayer were 113.6 ppb (± 5.2%) immediately following application (Fig. 4A, Day 0). Propiconazole residues associated with soil increased to 149.6 ppb (± 3.4%) by 10 days post-application (Fig. 4A, Day 10) before declining to 103.2 ppb (± 10.7%) by 28 days post-application (Fig. 4A, Day 28). Triadimefon: Residues of triadimefon associated with soil following boom sprayer application were 224.8 ppb (± 2.6%) immediately following application (Fig. 4A, Day 0). Triadimefon soil residues declined over time to 25.6 ppb (± 4.9%) at three days post¬ application (Fig. 4A, Day 3). No triadimefon residues were detected after this sampling period. 65 Figure 4. Concentration (in ppb) of fenarimol, propiconazole and triadimefon in soil pre-application (pre), immediately post application ( Day 0), and 1,3,7,10,14,21, and 28 days following application with a boom sprayer (A), high pressure injector (B), and a modified slicer/seeder (C). 66 Concentration in Soil (PPB) 250 200 250 150 200 100 50 0 Days Following Application 67 21 28 fenarimol fenarimol Slicer/Seeder propiconazole propiconazole propiconazole Modified Pressure Injector triadimefon triadimefon Sprayer Boom High C B A High Pressure Injection Application Fenarimol: Following application with the high pressure injector, fenarimol residues in the soil were not detected until 21 days after application at a level of 35.2 ppb (± 1.3%) (Fig. 4B, Day 21). Fenarimol levels in the soil then declined to 32 ppb (± 2.3%) 28 days after application (Fig. 4B, Day 28). Propiconazole: Soil residues of propiconazole following high pressure injection were not detected until 21 days post-application at a level of 42.4 ppb (± 2.6%) (Fig. 4B, Day 21). Propiconazole levels in soil then declined over time to 36.8 ppb (± 1.3%) by 28 days after application (Fig 4B, Day 28). Triadimefon: Triadimefon was not detected in soil following application with the high pressure injector. Modified Slicer/Seeder Application Propiconazole: Following application with the modified slicer/seeder, propiconazole residues were first detected 7 days post-application at 136.8 ppb (± 8.0 %) (Fig. 4C, Day 7). Propiconazole residues declined to 64.8 ppb (± 3.1%) at 14 days post-application (Fig. 4C, Day 14), and then to 55.2 ppb (± 1.7%) by 28 days post-application (Fig. 4C, Day 28). Soil samples taken in the pre-application period (pre. Fig 4) had no detectable residues of the fungicides at the detection limits established. 68 2.3.1.4 Root Tissue Boom Sprayer Application Fenarimol: No fenarimol residues were detected in root tissue at any time following application with the boom sprayer (Fig. 5A, Days 0~28). Propiconazole: No propiconazole residues were detected in root tissue at any time following application with the boom sprayer (Fig. 5A, Days 0-28). Triadimefon: Residues associated with root tissue following application with the boom sprayer were 1.7 ppm (± 6.8%) immediately following application (Fig. 5A, Day 0). No residues were detected after this sampling period (Fig. 5 A, Days 1-28). High Pressure Injection Application No residues of any of the three fungicides were detected in the roots following application with the high pressure injector (Fig. 5B, Days 0-28). Modified Slicer/Seeder Application Propiconazole: Following application with the modified slicer/seeder, propiconazole residues in root tissue were not detected until 7 days after application (Fig. 5C, Day 7) at which time a concentration of 2.2 ppm (± 7.3%) was determined. No propiconazole residues were detected in the roots at any other sampling period. 69 Figure 5. Concentration (in ppm) of fenarimol, propiconazole and triadimefon in roots pre-application (pre), immediately post application (Day 0), and 1,3,7,10,14,21, and 28 days following application with a boom sprayer (A), high pressure injector (B), and a modified slicer/seeder (C). 70 Concentration in Root Tissue (PPM) Days Following Application pre 0,3^ P« 01 71 fenarimol fenarimol / triadimefon propiconazole propiconazole triadimefon Sprayer Pressure Injector Boom High B Root tissue samples taken in the pre-application period (pre, Fig. 5) had no detectable residues of any of the fungicides at the detection limits established. 2.3.2 Total Accumulated Residue 2.3.2.1 Leaf Tissue Boom sprayer application of fenarimol and triadimefon resulted in total accumulated residues (TAR, sum of all residue concentrations over days 0-28) associated with leaf tissue approximately 4 times that found following application with the high pressure injector (Table 3). Propiconazole TAR associated with leaf tissue following boom sprayer application were 3.1 times greater than those found following application with the high pressure injector and 7.7 times that found following application with the modified slicer/seeder (Table 3). 2.3.2.2 Thatch TAR associated with thatch following application of fenarimol with the boom sprayer were 1.5 times greater than those found following application with the high pressure injector (Table 3). Boom sprayer application resulted in triadimefon TAR 1.4 times those found following application with the high pressure injector (Table 3). Following application with the boom sprayer, propiconazole TAR associated with thatch were 1.4 times those found following high pressure injection application and 1.6 times those found following application with the modified slicer/seeder (Table 3). 2.3.2.3 Soil Fenarimol TAR associated with the soil were 4.8 times greater following application with the boom sprayer versus the high pressure injection application (Table 3). 72 u-> o COOD i-i 3 O u 3 a> 3 > O X •c ed s CO c/5 UhCd ,00 3 H-> x: o CDed aUrn 5 Vc o 'i o c/) rv jaa c3 CN ON VO I ir> *a a> • o o VO (N VO VO % ’S s ^ 2 ^ *0 c« Ui *3 OD O *3 +-> 3 ' u o 3 o Ui 05 l-l u (U Ui 05 M ' .X C/D o (U 3 ' c *0 o ■3 s. O cd ed •—< ca ed *—' 3 o <1^ t (U a> 4> 3 , S Oh |_c o Oh Wh o g Oh Uh O •c C/D 3 o C/D 3 C/D 3 5 o cd c/5 o S ed O C/5 *3 CO CD _ a> P cd O o S o, B 0) *0 *3 ed b o o o •c o 2: H-» H o o Oh O Oh O ' to H o H-» cn CQ PQ ffl ^ c 00 s « a a H , H P, eo the expenment ( 0-28 days). 73 Propiconazole TAR associated with the soil following boom sprayer application were 9.9 times those found following application with the high pressure injection and 2.4 times those found following application with the modified slicer/seeder (Table 3). Following application with the high pressure injector, no triadimefon was found associated with the soil, while the TAR found following application with the boom sprayer was 314 ppm (Table 3). 2.3.2.4 Root Tissue No fenarimol was detected in the root tissue following application with either the boom sprayer or the high pressure injector (Table 3). Propiconazole TAR following application with the modified slicer/seeder were 2.2 ppm, while no propiconazole was detected in root tissue following boom sprayer and high pressure injection application (Table 3). TAR of triadimefon following application with the boom sprayer were 1.7 ppm, while no residues were detected following application with the high pressure injector (Table 3). 2.3.3 Percent Of Applied Fungicide Recovered 2.3.3.1 Leaf Tissue Boom Sprayer Application Fenarimol: Immediately after application, 25.0% of the fenarimol applied by the boom sprayer was recovered from leaf tissue (Fig. 6A, Day 0). Fenarimol leaf residues increased to 37.9% one day following application and then decreased over time to 1.2% 28 days following application (Fig. 6A, Days 1-28). 74 Figure 6. Percent of total applied fenarimol recovered from all matrices (total) and from leaf, thatch, roots, and soil for each day from Day 0 to day 28 following application with the boom sprayer (A) and high pressure injection (B). 75 Percent of Applied Fenarimol Days Following Application A BoomSprayer 76 Figure 7. Percent of total applied propiconazole recovered from all matrices (total) and from leaf, thatch, roots, and soil for each day from Day 0 to day 28 following application with the boom sprayer (A), high pressure injection (B), and modified slicer/seeder (C). 77 Percent of Applied Propiconazole A B High PressureInjector Modified Slicer/Seeder Boom Sprayer 78 Figure 8. Percent of total applied triadimefon recovered from all matrices (total) and from leaf, thatch, roots, and soil for each day from Day 0 to day 28 following application with the boom sprayer (A) and high pressure injection (B). 79 Percent of Applied Triadimefon Days Following Application B HighPressureInjector 80 PropiconazQlg; Immediately after application, 16.5% of the propiconazole applied by the boom sprayer was recovered from leaf tissue (Fig. 7A, Day 0). Propiconazole leaf residues increased to 26.3% one day following application and then decreased over time to 0.7% 28 days following application (Fig. 7A, Days 1-28). Triadimefon: Immediately after application, 23.6% of the triadimefon applied by the boom sprayer was recovered from leaf tissue (Fig 8 A, Day 0). Triadimefon leaf residues then decreased over time to 0.2% 21 days following application, and was not detectable by 28 days following application (Fig 8A, Days 1-28). High Pressure Injection Application Fenarimol: Immediately after application, 7.8% of the fenarimol applied with the high pressure injector was recovered from leaf tissue (Fig. 6B, Day 0). Fenarimol leaf residues then decreased over time to 0.5% 28 days following application (Fig 6B, Days 1-28). Propiconazole: Immediately after application, 6.0% of the propiconazole applied with the high pressure injector was recovered from leaf tissue (Fig. 7B, Day 0). Propiconazole leaf residues then decreased over time to 0.4% 28 days following application (Fig. 7B, Days 1-28). Triadimefon: Immediately after application, 4.7% of the triadimefon applied with the high pressure injector was recovered from leaf tissue (Fig. 8B, Day 0). Triadimefon leaf residues decreased over time to 0.2% 14 days following application and was not detected after this sampling period (Fig 8B, Days 1-28). Modified Slicer/Seeder Application Propiconazole: Immediately after application, 2.4% of the propiconazole applied with the modified slicer/seeder was recovered from leaf tissue (Fig. 7C, Day 0). Propiconazole leaf residues then decreased over time to 0.4% 21 days following application, and was not detected 28 days following application (Fig 7C, Days 1-28). 2.3.3.2 Thatch Boom Sprayer Application Fenarimol: Immediately after application, 58.7% of the fenarimol applied with the boom sprayer was recovered from thatch (Fig. 6A, Day 0). Fenarimol residues in thatch increased to 82.0% three days following application (Fig. 6A, Day 3). Fenarimol residues then decreased to 51.7% on Day 7, and to 46.7% on Day 21 (Fig. 6A, Days 7-21). Fenarimol thatch residues then rose to 76.5% 28 days following application (Fig. 6A, Day 28). Propiconazole: Immediately after application, 45.5% of the propiconazole applied with the boom sprayer was recovered from thatch (Fig. 7A, Day 0). Propiconazole residues in thatch increased to 82.3% three days following application (Fig. 7A, Day 3). Propiconazole residues decreased to 29.8% on Day 7, and to 26.1% on day 21 before increasing to 37.7% on Day 28 (Fig. 7A, Days 7-28). Triadimefon: Immediately after application, 42.3% of the triadimefon applied with the boom sprayer was recovered from thatch (Fig 8A, Day 0). Triadimefon residues in thatch then decreased over time to 5.7% three days following application and was not detected in thatch after this sampling period (Fig. 8A, Days 3-28). 82 High Pressure Injection Application Fenarimpl, Immediately after application, 33.0% of the fenarimol applied with the high pressure injector was recovered from thatch (Fig 6B, Day 0). Fenarimol residues in thatch increased to 41.9% one day following application and then fell to 16.5% three days following application (Fig. 6B, Days 1-3). Fenarimol residues then rose to 64.1% on Day 10 before falling to 41.6% on Day 28 (Fig. 6B, Days 7-28). Propiconazole: Immediately after application, 21.4% of the propiconazole applied with the high pressure injector was recovered from thatch (Fig. 7B, Day 0). Propiconazole residues in thatch then increased to 47.3% on Day 10 before falling to 13.6% 28 days following application (Fig. 7B, Days 1-28). Triadimefon: Immediately after application, 29.0% of the triadimefon applied with the high pressure injector was recovered from thatch (Fig. 8B, Day 0). Triadimefon thatch residues then decreased over time to 4.5% on Day 14 and were not detectable after this sampling period (Fig. 8B, Days 1-28). Modified Slicer/Seeder Application Propiconazole: Immediately after application, 16.1% of the propiconazole applied with the modified slicer/seeder was recovered from thatch (Fig. 7C, Day 0). Propiconazole residues in thatch increased to 57.2% on Day 7 and then decreased to 8.4% on Day 28 (Fig. 7C, Days 1-28). 83 2.3.3.3 Soil Boom Sprayer Application Fenarimpl; Immediately after application, 2.2% of the fenarimol applied with the boom sprayer was recovered from soil (Fig, 6A, Day 0). Fenarimol was not detected in soil on Day 1 but residues rose to 3.0% on Day 3 and to 5.2% on Day 10 (Fig. 6A, Days 1-10). Except for a decrease on Day 14, fenarimol residues remained steady and were 5.8% on Day 28 (Fig. 6A, Days 14-20). Propiconazole: Immediately after application, 4.2% of the propiconazole applied with the boom sprayer was recovered from the soil (Fig. 7A, Day 0). Propiconazole residues in soil decreased over time to 2.9% on Day 7 and then increased to 3.8% on Day 28 (Fig. 7A, Days 1-28). Triadimefon: Immediately after application, 4.2% of the triadimefon applied with the boom sprayer was recovered from the soil (Fig. 8A, Day 0). Triadimefon residues in soil then decreased over time to 0.5% on Day 3 and were not detected after this sampling period (Fig. 8A, Days 1-28). High Pressure Injection Application Fenarimol: No fenarimol was recovered from the soil until 21 days following application, when 2.7% of the fenarimol applied with the high pressure injector was recovered (Fig. 6B, Days 0-21). Fenarimol residues in soil then decreased to 2.5% on Day 28 (Fig. 6B, Day 28). Propiconazole: No propiconazole was recovered from the soil until 21 days following application, when 1.5% of the propiconazole applied with the high pressure injector was 84 recovered (Fig. 7B, Days 0-21). Propiconazole residues in soil then decreased to 1.4% on Day 28 (Fig. 7B, Day 28). Triadimelpn; No triadimefon was recovered from the soil following application with the high pressure injector (Fig. 8B, Days 0-28). Modified Slicer/Seeder Propiconazole: No propiconazole was recovered from the soil until 7 days following application, when 5.1% of the propiconazole applied with the modified slicer/seeder was recovered (Fig. 7C, Days 0-7). Propiconazole was not detected in soil on Day 10 but 2.4% of the applied propiconazole was present on Day 14 (Fig. 7C, Days 10-14). Propiconazole residues in soil then decreased over time to until 28 days following application, when 2.0% of the applied propiconazole was recovered from the soil (Fig. 7C, Days 14-28). 2.3.3.4 Root Tissue Boom Sprayer Application Fenarimol: No fenarimol was recovered from the root tissue following application with the boom sprayer (Fig. 6A, Days 0-28). Propiconazole: No propiconazole was recovered from the root tissue following application with the boom sprayer (Fig. 7A, Days 0-28). Triadimefon: Immediately following application 0.05% of the triadimefon applied with the boom sprayer was recovered from the root tissue (Fig. 8 A, Day 0). No triadimefon was recovered from the root tissue following this sampling period (Fig. 8A, Days 1-28). 85 High Pressure Injection Application Fenarimol: No fenarimol was recovered from the root tissue following application with the high pressure injector (Fig, 6B). Propiconazole: No propiconazole was recovered from the root tissue following application with the high pressure injector (Fig. 7B). Triadimefon: No triadimefon was recovered from the root tissue following application with the high pressure injector (Fig 8B). Modified Slicer/Seeder Application Propiconazole: No propiconazole was recovered from the root tissue following application with the modified slicer/seeder until seven days following application, when 0.12% of the applied propiconazole was recovered (Fig. 1C, Days 0-7). No propiconazole was recovered following this sampling period (Fig 1C, Days 10-28). 2.4 Discussion The concern that fenarimol, propiconazole and triadimefon could be potential groundwater contaminants through the use of alternative turfgrass application technologies such as high pressure injection or modified slicer/seeder appears to be unfounded. None of the application methods resulted in movement of the fungicides through the turfgrass system to the roots in any appreciable amount. Even though all three fungicides are relatively water soluble and were watered into the thatch with 1.3 cm of irrigation, there was no evidence that they bypassed the top 5.1 cm of the soil since they were never found in the soil or roots in the second 5.1 cm section of the soil core. Thus, it 86 is not likely that fungicides applied to turfgrass by these means will bypass the entire root zone and contaminate groundwater over the time frame examined in the present study. The majority of the fungicides were associated with the thatch layer. Approximately 40-60% of the total applied fungicides was recovered from the thatch immediately following application with the boom sprayer, another 18-25% was recovered from the leaf tissue, and 1% to 6% was recovered from the soil. Immediately following application with the high pressure injector, 20-30% of the total applied fungicides (but 78-86% of the amount recovered) was associated with the thatch. Only 5-8% of the applied fungicides were recovered from the leaf tissue, and none were recovered from the roots or soil. Immediately following application using the modified slicer/seeder, 16% of the total applied propiconazole (but 87% of the propiconazole recovered) was associated with the thatch, 2% was found associated with the leaf, and none was recovered from the roots or soil. Obviously, thatch is a major reservoir for these SBI fungicides and it functions as a barrier to their efficacious delivery to the turfgrass root system. As a barrier, thatch also should restrict the movement of these fungicides into the groundwater. There was relatively little degradation of fenarimol (ti/2 (half-life) = 365 days, Pesticide Manual, 1994) when applied to turfgrass over the 28 days of the experiment. Nearly 84% of the fenarimol applied by boom sprayer could be accounted for in leaf, thatch, soil, and root samples taken 28 days after application. Following application with the high pressure injector, there was also no substantial decrease in the recovery of fenarimol. Immediately after application, 41% of the total applied fenarimol was recovered from the high pressure 87 injector plot and 45% of the applied fenarimol was recovered after 28 days. Apparently fenarimol is exhibiting very little breakdown over the time course of this experiment. Propiconazole (ti/2= 40-70 days, Pesticide Manual, 1994) exhibited somewhat greater breakdown over time. Following application with the boom sprayer, total recoveries for propiconazole went from 66% immediately after application to 42% 28 days post¬ application. Following application with the high pressure injector, 28% of the total applied propiconazole was recovered immediately after application and dropped to 15% by Day 28. With application by the modified slicer/seeder, 19% of the total applied propiconazole was recovered immediately following application and dropped to 10% 28 days post-application. Propiconazole is apparently degrading in the turfgrass system to a greater extent than fenarimol over the time course of this experiment. The behavior of triadimefon (ti/2 = 6-18 days. Pesticide Manual, 1994) in the turfgrass system differed greatly when compared to fenarimol and propiconazole. Immediately after application with the boom sprayer, 70% of the applied triadimefon was recovered from the leaf, thatch, soil, and root samples but by 10 days post-application less than 1% of the triadimefon was recovered. The triadimefon applied with the high pressure injector followed a similar pattern of rapid disappearance. Immediately following application, 34% of the total applied triadimefon was recovered but by Day 10 only 5% was recovered. It has been previously reported that triadimefon is rapidly hydrolyzed to triadimenol in both plants (Garcia et al., 1991) and soil (Patil et al., 1988). This rapid conversion to triadimenol likely accounts for the rapid decrease in triadimefon in turfgrass over the time course of this study and for the relatively short half-life of 88 triadimefon. Triadimenol, however, has a much longer half-life (I1/2 = 110-270 days, Pesticide Manual, 1994) than triadimefon, and therefore poses a greater potential for groundwater contamination. Also, triadimenol (acute oral LD50 for rats 700 mg/kg) possesses greater mammalian toxicity than triadimefon (acute oral LD50 for rats 1000 mg/kg) (Pesticide Manual, 1994). As expected, boom sprayer application resulted in average leaf residues that were 3 and 8 times higher than residue levels detected following high pressure injection or modified slicer/seeder application, respectively. Over time, fungicide levels decreased in the leaf tissue. Following application with the boom sprayer, 16-25% of the total fungicides applied were recovered from the leaf tissue. This amount decreased to approximately 3% of the applied fungicides 14 days following application. Following application with the high pressure injector, approximately 5-8% of the total applied fungicides were recovered from the leaf tissue immediately after application. By Day 14, approximately 1% of the total applied fungicides was recovered from the leaf tissue. As expected, the amounts of fungicides associated with the turfgrass canopy when applied with the high pressure injector (i.e. 5-8%) were less than the amounts found following boom sprayer application (i.e. 16-25%). Following modified slicer/seeder application, only approximately 2% of the applied propiconazole was recovered from the leaf tissue, and this dropped to 1% by Day 7 following application. Thus, the two alternative application methods (i.e. high pressure injection and modified slicer/seeder) are designed » to bypass the turfgrass canopy and they apparently do this effectively. 89 In contrast to the leaf tissue, fungicide residues in the soil (0-5.1 cm) generally increased over time. Following application with the boom sprayer, fenarimol residues in soil showed the greatest increase. Immediately following application, 2% of the total applied fenarimol was recovered from the soil. By 28 days post-application, 6% of the applied fenarimol was recovered from soil, a 3 fold increase. Following boom sprayer application, propiconazole residues in soil accounted for 4% of the applied propiconazole, and this level stayed relatively constant throughout the course of the experiment. The increase of fenarimol in soil over time compared to the steady levels found for propiconazole may be attributable to their different soil half lives and values. Fenarimol is approximately five times more persistent in the soil than propiconazole, and propiconazole has a Ko^that is approximately twice that of fenarimol. The shorter half-life and relatively constant concentration of propiconazole in the soil may indicate that it is degrading as it moves into the soil, while the higher K^c of propiconazole indicates that its movement from the thatch to the soil is slower than that of fenarimol. Due to the conversion of triadimefon to triadimenol, triadimefon disappeared from soil cores rapidly (not detected by Day 7) after application from a level of 4% immediately following application. With the high pressure injector, no ftmgicides were detected in the soil until 21 days after application. With the modified slicer/seeder, propiconazole did not appear in the soil until 14 days following application. Apparently, the thatch layer is acting as an effective reservoir for these SBI fungicides and is slowly releasing them into the soil layer below. The lack of recovery from the soil for fungicides applied with the high pressure injector 90 and modified slicer/seeder plots, however, could be due to sampling error as discussed below. A point of concern in the above discussion is the relatively complete mass balance obtained when the fungicides were applied by boom sprayer compared to the two alternative application methods. Both the high pressure injector and the modified slicer/seeder deposited the fungicides in discreet bands primarily in the thatch. The high pressure injector nozzles deposited streams of the tank mixture 7.2 cm apart, and the modified slicer/seeder deposited the tank mixture in slits that were 5 cm apart. The internal diameter of the soil corer used to collect turfgrass samples was 2.5 cm. Due to this disparity in size, and the apparent limited lateral movement of the fungicides, it is possible that the some turfgrass cores were removed from areas between the fungicide treated bands. Given this sampling regime, an approximation of mass balance could only be expected for the residues recovered following boom sprayer application. Total fungicide recovery for boom sprayer treated plots ranged between 66% and 98% over the first three days following application, between 7% and 47% from the high pressure injector plots, and 18% to 44% from the modified slicer/seeder plots. The low and varied recovery of fungicides treated with the high pressure injection and the slicer/seeder may indicate that the sampling methodology was not adequate to achieve mass balance. These findings also indicate a lack of lateral movement for these SBI fungicides in the turfgrass system. When using the high pressure injector for the control of white grubs, the treatment was effective only when the larvae were active (P. Vittum, personal communication). This 91 suggests that the insecticides applied by high pressure injection remained in discreet bands and the larvae would have to move through these bands in order to contact the insecticide. Additionally, it has been suggested that mole crickets are able to alter their behavior to avoid contact with the bands of insecticide deposited following application (Brandenburg, 1996). This provides further evidence that application of pesticides utilizing high pressure injection results in discreet bands of the pesticide in the soil, with little or no lateral movement of the compounds. In summary, none of the application methods appeared to effectively deliver any of the three SBI fungicides to the roots in significant amounts. The modified slicer/seeder was, in fact, so destructive to the turf that it is unlikely to have any practical usefulness. The high pressure injector is believed to deliver chemicals to the soil in discreet bands. Due to the inability to achieve mass balance following high pressure injection and the anecdotal evidence relating to grub control and mole cricket behavior, it appears that there is little lateral movement of these fungicides in turfgrass. Finally, the application of these SBI fungicides by the three methods detailed in this study resulted in little vertical movement in the turfgrass profile. As a result, they would appear to pose little danger to groundwater when applied via boom, high pressure injector, or modified slicer/seeder means over the experimental time frame examined by this study. 92 CHAPTER 3 FATE OF TRIADIMEFON AND TRIADIMENOL IN A TURFGRASS SYSTEM 3.1 Introduction Triadimefon and its metabolite triadimenol have been demonstrated to leach through the turfgrass system, possibly contaminating the groundwater (Petrovic et al, 1994). Triadimefon is a widely used sterol biosynthesis inhibiting (SBI) fungicide used against a variety of leaf, crown, and root diseases of turfgrass, including summer patch, a root disease caused by Magnaporthe poae. Delivering fungicides to the roots of turfgrass plants can be a problem. In order to reach the roots, it is necessary to deliver the fungicides through the thatch, a layer of organic matter consisting of turfgrass shoots, roots, and crowns in various states of decay. The thatch layer is known to adsorb many hydrophobic pesticides (Lickfeldt and Branham, 1995; Niemczyk and Krueger, 1982) including triadimefon (Dell et al., 1994). It is possible that by disrupting or bypassing the thatch layer it would be easier to deliver the fungicides to the root system. Delivering fungicides below the thatch, however, increases the chance of groundwater contamination. Triadimefon is known to metabolize to triadimenol (Fig. 9) in soil (Patil et al., 1988) and plants (Garcia et al., 1991). Pesticide metabolites may be as toxic or even more toxic than their parent compounds. Metabolites are also frequently more mobile in the environment than their parent molecules, leading to concerns about groundwater contamination (Coats, 1993). Hydrolysis and oxidation are common pesticide 93 Cl 0-CH-C0C(CH3)3 Cl -< Triadimefon Triadimenol Figure 9. Chemical structure of triadimefon and triadimenol transformation reactions that can result in more polar, and therefore more water soluble metabolites. More water soluble, and therefore more mobile, compounds pose a greater threat to groundwater. Pesticide metabolites such as triadimenol, which have greater mammalian toxicity, are more water soluble, more mobile, and have a greater half-life than their parent compound may pose more of a danger to groundwater and wildlife. Triadimenol is almost twice as water soluble as triadimefon, has 1.4 times greater mammalian toxicity, a slightly lower value, and a half- life roughly 16 times greater (Table 4). All these factors combine to make triadimenol more of a threat to groundwater than its parent compound triadimefon. 3.2 Materials And Methods 3.2.1 Experimental Site Three 12.2 m X 12.2 m (0.0149 ha) plots of Kentucky bluegrass of a mixture of cultivars were selected at the University of Massachusetts Turfgrass Research Center in South Deerfield, MA. The turf was established in 1985 and overlay a Hadley silt loam 94 Table 4. Physical properties, environmental characteristics, application rates, and toxicities of triadimefon and triadimenol. 'o 3 Uh (U (30 c: ca CO CO CO TJ O 3o (N oo A m O ^ O • « CO o m ^ V OO 00 rn in (N Os tin c ^3 TJ H X ^ o 2 S D (U cd kJ x> CO 00 (N 95 • ^ -C Pm C/3 "cS -*-* O cd CO (U CO c d T w CO -a 00 c CO c3 (U Cu CJ o 00 H l-l o o o (u c+_00Z 'o +-» = Not Applied soil (coarse, silty, mixed, nonacid, mesic, Typic Udifluvent, pH 6.8). The turf was maintained at a height of 5.1 cm and irrigated as needed to prevent drought stress (Fig. 1). 3.2.2 Fungicides The SBI fungicide triadimefon {l-(4-chlorophenoxy)-3,3-dimethyl-l-(l//-l,2,4-triazol- l-yl)butan-2-one}(Fig. 9) was applied to the plots on 22 August 1992. Triadimefon is known to metabolize to triadimenol {(\RS,2RS;lRS,2SR)-\-(4-ch\orophQnoxy)-3,3- dimethyl-l-(lH-l,2,4-triazol-l-yl)butan-2-ol}(Fig. 2) in soil (Patil et al., 1988) and plants (Garcia et al., 1991). 3.2.3 Fungicide Application The fungicide was applied using two types of applicators: 1) a standard boom sprayer (F.E. Myers and Bro. Co., Ashland, OH); 2) a high pressure injector (Cross Equipment, Albany, GA). Triadimefon (Bayleton 25 DF, Mobay, Kansas City, MO) was applied at the recommended rate for summer patch control, 3.05 kg ai ha’\ The boom sprayer was used to apply triadimefon to the first plot. The boom sprayer consisted of nine T-jet “9010” flat fan nozzles applying 621 L ha’^ at an operating pressure of 1925 kPa and generated a swath width of 4.6 m. The boom sprayer application was put on last and was immediately followed by 1.3 cm of irrigation water applied to the entire experimental area. The high pressure injector consisted of four 60 L tanks connected to a skid-mounted boom with 18 nozzles at a 7.2 cm spacing. Triadimefon was applied in one pass directly into the thatch as an aqueous stream at a pressure of 11,100 kPa in 880 L ha'^ . 96 3.2.4 Sampling Each of the two plots was divided into sixteen 3.1 m X 3.1 m (0.000961 ha) subplots that were further subdivided into nine 1 m (0.0001 ha) sections, one of which was randomly selected for each sampling period. It was estimated that each square meter of the plot contained approximately 455 g of leaf tissue, 11,989 g of thatch, 56,700 g of soil (1 m X 1 m X 5.1 cm), and 85 g of root tissue (1 m x 1 m x 5.1 cm) (all in wet weights). Leaf, thatch, soil and root were sampled 15 minutes before and 30 minutes after application and at 1,3,7,10,14,21, and 28 days post-application. At each collection period, 75 g of leaf tissue were collected from each of the sixteen subplots by cutting the plants at the soil line. The tissue was placed in 1.0 L plastic Zip-Loc™ freezer bags for storage. Following this process, sixteen soil cores, one from each subplot, were taken with a 30.5 cm X 2.5 cm JMC Zero Contamination Tube (Clements Assoc., Inc., Newton, lA). The thatch layer of each core was delineated as the deepest point at which rhizomes were visible and was marked. Samples were then placed in ice chests and transported to the Massachusetts Pesticide Analysis Laboratory (MPAL) where they were stored at -20°C for analysis. In the laboratory, the thatch layer was removed from the top of each soil core and the remainder of each core was divided into five 5.1 cm X 2.5 cm sections and bulked (i.e. the top 5.1 cm of the cores from each subplot for each application method during a sampling period were placed together in a 2.8 L Zip-Loc™ freezer bag and mixed by rotating the bag for 3 min). The bulked samples were screened through a #10 sieve and 97 the roots and thatch were separated out individually by hand. The soil, roots, and thatch were placed into individual plastic freezer bags for storage in a -20® C freezer. 3.2.5 Storage Samples On each sampling date, equal amounts of nontreated (i.e., no applied fungicide) leaf tissue and soil cores (i.e., containing soil, roots, and thatch) were collected from an adjacent untreated area to be used as fortified storage samples (i.e., to assess any loss during storage prior to analysis), concurrent fortified samples (i.e., to provide daily checks of the analytical method efficiency), and concurrent nontreated samples (i.e., to provide daily checks to assess background interferences). Following sample collection, leaf tissue was amended with 0.4 pg g \ thatch with 1.0 pg g'\ roots with 2.0 pg g’^ and soil with 0.08 pg g'^ of triadimefon and stored with the concurrently collected field samples to assess any degradation of the compounds during storage (i.e., fortified storage samples). 3.2.6 Analytical Procedures Leaf: The determination of triadimefon and triadimenol residues was based on a method developed by Newsome and Collins (1990). Five g of leaf tissue were cut up using scissors and placed in a 250 ml red actinic Erlenmeyer flask. Fifty ml of acetone were added to the flask and it was shaken for 30 min on a rotary table top shaker (Eberbach) at low speed (180 rpm). The liquid was decanted through 13 g anhydrous sodium sulfate into a 500 ml fiat bottom boiling flask. The leaf tissue was re-extracted with an additional 50 ml acetone that was likewise quantitatively transferred to the boiling flask. The combined extract was reduced to approximately 3 ml under vacuum. 98 quantitatively transferred in acetone to a 50 ml centrifuge tube, and the sample volume adjusted to 10 ml under a stream of nitrogen. The sample volume was then readjusted to 50 ml with distilled water. Prior to loading the sample extracts, Cjg solid phase extraction cartridges (500 mg, J&W Scientific, Folsom, CA) were sequentially washed with 10 ml aliquots of methanol, methylene chloride, and methanol to remove any contaminants. The cartridge was activated with 25 ml distilled water. The sample extract was then passed through the activated cartridge, followed by a wash step that consisted of 10 ml of a 40% solution of methanol in distilled water. The retained triadimefon and triadimenol was eluted from the cartridge with 5 ml of methanol. Following elution, a 4 ml aliquot of the eluent was transferred to a 15 ml centrifuge tube and the volume adjusted to 10 ml with distilled water. A liquid/liquid extraction was performed in the centrifuge tube by adding 5 ml toluene and gently shaking the tube. The triadimefon and triadimenol partitioned into the toluene (i.e. top layer), which was then removed for instrumental analysis. Each sample analysis was replicated three times using three subsamples. Thatch: Fifty ml of acetone were added to 1 g of thatch in a 120 ml amber jar and the sample was homogenized twice for 20 seconds at 20,000 rpm with a Brinkmann Polytron PT3000 homogenizer (Littau, Switzerland). The homogenized sample was shaken for 30 min on a rotary table top shaker, and the acetone decanted into a 500 ml flat-bottomed boiling flask. The extract was reduced to ca 3 ml under vacuum, quantitatively transferred in acetone to a 50 ml centrifuge tube and the sample volume was adjusted to 10 ml under a stream of nitrogen. The sample volume was readjusted to 50 ml with distilled water . The Cl8 cartridges were washed and activated as outlined above for leaf tissue. The 99 sample was then passed through the cartridge. This was then followed by a wash step consisting of 10 ml of a 40% solution of methanol in distilled water. The cartridge was eluted with 10 ml acetone into a 13 ml centrifuge tube and the sample eluent volume was adjusted to 2 ml under a stream of nitrogen for analysis. Each sample analysis was replicated three times using three subsamples. Soil: Fifty ml of acetone were added to 25 g of soil in a 250 ml red actinic erlenmeyer flask and shaken for 30 min. on a rotary table top shaker at low speed (180 rpm). The liquid was decanted through 13 g anhydrous sodium sulfate into a 500 ml boiling flask. The sample was extracted again with 50 ml acetone and the extract quantitatively transferred to the boiling flask containing the first extract. The sample was reduced to ca 3 ml under vacuum. The remainder of the procedure followed that outlined for thatch as described above. Roots: Twenty ml of acetone were added to 0.5 g of root tissue in a 60 ml amber jar. The sample was homogenized and shaken as previously described for thatch. The acetone was decanted into a boiling flask as described previously. The sample was re-extracted with acetone and the extract quantitatively transferred to the boiling flask containing the first extract and reduced to ca 3 ml. The remainder of the procedure followed that outlined above for thatch 3.2.7 Instrumental Analysis Triadimefon and triadimenol residues were analyzed using a Varian model 3400 gas chromatograph equipped with a nitrogen-phosphorus detector (NPD) and a DB-5 glass megabore column (53 mm i.d. X 15 m, J & W Scientific, Folsom, CA). The injector and 100 detector temperatures were 250° and 300°C respectively. Injections (Ipl) were made onto the column with an initial temperature of 160°C, which was held for 2 min, the temperature was increased to 235°C (15°C min'*) and maintained for 3 min. Gas flow was as follows: Helium, 10 ml min'*; hydrogen, 3.5 ml min'*; nitrogen, 20 ml min'*; and air, 115 ml min'*. 3.2.8 Quality Assurance/Quality Control 3.2.8.1 Instrument Calibration Calibration procedures for the chromatographic system used for analysis involved developing standard curves consisting of at least three concentrations for triadimefon and triadimenol using high purity analytical standards (>99% purity). Triadimefon and triadimenol standards were provided by Mobay Corp. (Kansas City, MO). Standard curves consisting of at least three concentrations of each analyte were included at the beginning of each day of analysis. Area counts for each standard concentration (n=3) 2 were averaged to generate a calibration curve (r > 0.99). 3.2.8.2 Sample Controls For each day of analysis, corresponding fortified storage samples and nontreated leaf, thatch, roots, and soil amended with and without the fungicides of interest (i.e. concurrent fortified and nontreated samples, respectively) were extracted and analyzed to determine recovery efficiency and daily method performance, respectively. Reagent and matrix blanks were also included in each analysis. 101 Average triadimefon recoveries and standard deviations from daily amended leaf, thatch, root, and soil samples were 102% (+17.9), 108.7% (±10.0), 101.8% (±12.1) and 101.9% (±5.6), respectively. Average triadimenol recoveries and standard deviations from daily amended leaf, thatch, root, and soil were 100.5% (± 14%), 104.9% (± 10.5%), 101.7% (± 5.3%) and 102.9% (±4.2%) and 102.3% (± 5.7%), respectively. Average triadimefon recoveries and standard deviations for fortified storage samples of leaf, thatch, root, and soil samples were 82.0% (±11.8), 99.2% (±16.3), 80.7% (±14.5) and 101.1% (±9.0), respectively. Average triadimenol recoveries and standard deviations for fortified storage samples of leaf, thatch, roots, and soil were 83.0% (± 15.7%), 102.6% (± 17.3%), 76.6% (± 8.3%) and 103.5% (± 5.3%), respectively. 3.3 Results 3.3.1 Fungicide Concentration 3.3.1.1 Leaf Tissue Boom Sprayer Application Triadimefon: Maximum triadimefon residues associated with leaf tissue following the boom sprayer application occurred during the first sampling period immediately after spraying (Fig. lOA, Day 0) at a concentration of 158 ppm (± 2.4 % coefficient of variation). Triadimefon residues declined over time to 1 ppm (± 5.8 %) by Day 21 post¬ application (Fig. lOA, Day 21) and were not detected on Day 28 post-application. 102 Figure 10. Concentration of triadimefon and triadimenol in leaf tissue (in ppm) preapplication (pre), immediately post-application ( Day 0), and 1,3,7,10,14,21, and 28 days following application with a boom sprayer (A), high pressure injector (B). 103 Concentration in Leaf Tissue (PPM) Days Following Application B A High PressureInjector Boom Sprayer 104 Triadimenol: Triadimenol first appeared in the leaf samples one day after application (Fig. lOA, Day 1) at a concentration of 54.6 ppm (± 3.2%). Triadimenol concentrations then declined over time to 18.6 ppm (± 12.4%) on Day 10 (Fig. lOA, Day 10). After Day 10 triadimenol residues in leaf increased throughout the course of the experiment reaching 44.3 ppm (± 1.2%) on Day 28 (Fig. lOA, Day 28). High Pressure Injection Application Triadimefon: Maximum triadimefon residues associated with leaf tissue following application with the high pressure injector occurred during the first sampling period immediately following application (Fig. lOB, Day 0) resulting in a concentration of 32 ppm (± 1.3 %). Triadimefon residues declined over time to 1 ppm (± 3.8 %) by Day 14 post-application ( Fig lOB, Day 14). No triadimefon residues were detected after Day 14 following application with the high pressure injector. Triadimenol: Triadimenol residues associated with leaf tissue were 11.9 ppm (± 4.6%) immediately following application with the high pressure injector ( Fig. lOB, Day 0). Triadimenol levels then increased to 15.2 ppm (±7.0%) three days following application (Fig. lOB, Day 3). Triadimenol residues associated with leaf tissue then decreased to 10.5 ppm (± 3.5%) 14 days following application (Fig. lOB, Day 14), before increasing to 18.2 (±5.5%) ppm 28 days after application (Fig. lOB, Day 28). 3.3.1.2 Thatch Boom Sprayer Application Triadimefon: Maximum triadimefon residues associated with thatch following the boom sprayer application occurred on the first sampling period immediately following 105 Figure 11. Concentration of triadimefon and triadimenol in thatch (in ppm) preapplication (pre), immediately post-application ( Day 0), and 1,3,7,10,14,21, and 28 days following application with a boom sprayer (A), high pressure injector (B). 106 Concentration in Thatch (PPM) 20 16 Days Following Application pre 013710 14 2128 Q HIGHPRESSUREINJECTOR □ triadimenol ■ triadimefon Boom Sprayer 107 application (Fig. 11 A, Day 0,) resulting in a concentration of 10.7 ppm (± 2.5%). Triadimefon residues declined over time to 1.5 ppm (± 24.6%) at 3 days post-application (Fig. 11 A, Day 3). No triadimefon residues were detected following Day 3. Triadimenol: Triadimenol residues in thatch following boom sprayer application were 3.02 ppm immediately following application (Fig. 11 A, Day 0). Residues then increased to 18.0 ppm three days following application (Fig. 1 lA, Day 3), before decreasing to 9.9 ppm (± 8.7%) 21 days following application (Fig. 11 A, Day 21). Triadimenol residues associated with thatch then increased to 17.0 ppm (± 4.0%) 28 days following application (Fig. 11 A, Day 28). High Pressure Injection Application Triadimefon: Maximum triadimefon residues associated with the thatch following high pressure injection application occurred on the first sampling period immediately after application (Fig. 1 IB, Day 0), resulting in a concentration of 7.4 ppm (± 26.0%). Triadimefon levels in thatch declined over time to 1.1 ppm (± 5.5%) 14 days after application (Fig. 1 IB, Day 14,). No triadimefon residues were detected after Day 14. Triadimenol: Triadimenol residues in thatch following application with the high pressure injector first appeared one day after application at a concentration of 3.6 ppm (± 11.6%) (Fig. 1 IB, Day 1). Triadimenol residues then increased to 10.9 ppm (± 15.2%) seven days following application (Fig. 1 IB, Day 7) before decreasing to 5.7 ppm (± 4.4%) 14 days after application (Fig. 1 IB, Day 14). Triadimenol residues then increased 108 to 11.7 ppm (± 2.6%) 21 days following application (Fig. 1 IB, Day 21) before decreasing to 7.7 ppm (± 4.7%)28 days following application (Fig. 1 IB, Day 28). 3.3.1.3 Soil Boom Sprayer Application Triadimefon: Residues of triadimefon associated with soil following boom sprayer application were 224.8 ppb (± 2.6%) immediately following application (Fig. 12A, Day 0). Triadimefon soil residues declined over time to 25.6 ppb (± 4.9%) at three days post¬ application (Fig. 12A, Day 3). No triadimefon residues were detected after this sampling period. Triadimenol: Residues of triadimenol associated with the soil immediately following boom sprayer application were 44.8 ppb (± 2.4%)(Fig. 12A, Day 0). Triadimenol levels in the soil increased over time and attained a maximum concentration of 808.8 ppb (± 1.6%) 10 days post-application (Fig. 12A, Day 10). Triadimenol residues then decreased to 352.8 (± 5.2%) ppb 14 days following application (Fig. 12A, Day 14) before increasing to 803.2 ppb (± 7.1%) 28 days following application (Fig. 12A, Day 28). High Pressure Injection Application Triadimefon: Triadimefon was not detected in soil following application with the high pressure injector. Triadimenol: Following application with the high pressure injector, triadimenol residues in the soil were not detected until one day after application at a level of 19.2 ppb (± 1.7%) (Fig. 12B, Day 1). Triadimenol levels in the soil increased over time to 324 ppb 109 Figure 12. Concentration of triadimefon and triadimenol in soil (in ppb) preapplication (pre), immediately post-application ( Day 0), and 1,3,7,10,14,21, and 28 days following application with a boom sprayer (A), high pressure injector (B). 110 Concentration in Soil (PPB) Days Following Application A High PressureInjection Boom Sprayer 111 (± 1.7%) 28 days after application (Fig. 12B, Day 28). 3.3.1.4 Root Tissue Boom Sprayer Application Triadimefon: Triadimefon residues associated with root tissue following application with the boom sprayer were 1.7 ppm (± 6.8%) immediately following application (Fig. 13 A, Day 0). No residues were detected after this sampling period (Fig. 13 A, Days 1-28). Triadimenol: Triadimenol residues were first detected in root tissue three days following application with the boom sprayer at a level of 1.6 ppm (± 3.0%) (Fig. 13A, Day 3). Triadimenol residues in root tissue then increased to 4.8 ppm (± 7.1%) on Day 10 (Fig. 13A, Day 10), before decreasing to 3.4 ppm (± 5.6%)14 days following application (Fig. 13 A, Day 14). Triadimenol residues in root tissue then increased over time to 6.7 ppm (± 5.0%) on Day 28 (Fig. 13A, Day 28). High Pressure Injection Application Triadimefon: No residues of triadimefon were detected in the roots following application with the high pressure injector (Fig. 13B, Days 0-28). Triadimenol: Triadimenol residues in root tissue following application with the high pressure injector were not detected until 14 days following application at a level of 1.0 112 Figure 13. Concentration of triadimefon and triadimenol in root tissue (in ppm) preapplication (pre), immediately post-application (Day 0), and 1,3,7,10,14,21, and 28 days following application with a boom sprayer (A), high pressure injector (B). 113 Concentration in Root Tissue (PPM) Days Following Application High PressureInjection □ triadimenol ■ triadimefon 114 ppm (± 7.1%) (Fig. 13B, Day 14). Triadimenol residues in root tissue then increased to 2.0 ppm (± 7.8%) 28 days following application (Fig. 13B, Day 28). 3.3.2 Percent Of Applied Fungicide Recovered 3.3.2.1 Leaf Tissue Boom Sprayer Application Triadimefon. Immediately after application, 23.6% of the triadimefon applied by the boom sprayer was recovered from leaf tissue (Fig 14A, Day 0). Triadimefon leaf residues then decreased over time to 0.2% of the total applied 21 days following application, and was not detectable by 28 days following application (Fig 14A, Days 1-28). Triadimenol. The percentage of triadimenol recovered was calculated by converting the amount of triadimefon applied to the equivalent amount of triadimenol (i.e completely converting 1 g of triadimefon to triadimenol would give you 1.007 g triadimenol). Following application with the boom sprayer, triadimenol was not detected in the leaf tissue until one day after application, when 8.1% of the applied triadimefon was recovered as triadimenol (Fig. 14B, Day 1). Triadimenol leaf residues then declined over time to 2.7/0 10 days after application (Fig. 14B, Day 10). Triadimenol leaf residues then increased to 6.6% of the applied triadimefon 28 days following application (Fig. 14B, Day 28). High Pressure Injection Application Triadimefon: Immediately after application 4.7% of the triadimefon applied by the boom sprayer was recovered from leaf tissue (Fig 15 A, Day 0). Triadimefon leaf residues 115 Figure 14. Percent of total applied triadimefon (A) and triadimenol (B) recovered from all matrices (total) and from le^, thatch, roots, and soil for each day from Day 0 to day 28 following application with the boom sprayer. 116 Percent of Applied Fungicide B Triadimenol Boom Sprayer Triadimefon 117 Figure 15. Percent of total applied triadimefon (A) and triadimenol (B) recovered from all matrices (total) and from leaf, thatch, roots, and soil for each day from Day 0 to day 28 following application with the high pressure injector. 118 Percent of Applied Fungicide Days Following Application B A High PressureInjector Triadimenol Triadimefon 119 then decreased over time to 0.2% 14 days following application, and were not detectable by 28 days following application (Fig 15A, Days 1-28). Triadimcnpl: Immediately after application 1.7% of the applied triadimefon was recovered as triadimenol from the leaf tissue (Fig. 15B, Day 0). Triadimenol residues in leaf tissue then dropped to 0.9% of the applied triadimefon 3 days following application (Fig. 15B, Day 3). Seven days after application 2.3% of the applied triadimefon was recovered as triadimenol from the leaf tissue and the levels reached a low of 1.5% 14 days after application and then increased over time to 2.7% 28 days following application ( Fig. 15B, Day 28). 3.3.2.2 Thatch Boom Sprayer Application Triadimefon: Immediately after application, 42.3% of the triadimefon applied with the boom sprayer was recovered from thatch (Fig 14A, Day 0). Triadimefon residues in thatch then decreased over time to 5.7% 3 days following application and was not detected in thatch after this sampling period (Fig. 14A, Days 3-28). Triadimenol: Immediately following the application of triadimefon, 11.6% of the applied triadimefon was recovered from the thatch as triadimenol (Fig. 14B, Day 0). Triadimenol levels increased over time until three days after application when 69.2% of the applied triadimefon was recovered from the thatch as triadimenol (Fig. 14B, Days 1- 3). The percentage of triadimenol recovered generally decreased until 21 days after application when 38.2% of the applied triadimefon was recovered as triadimenol (Fig. 120 14B, Days 7-21). Triadimenol levels in thatch then increased to 52.4% of the applied triadimefon 28 days after application (Fig. 14B, Day 28). High Pressure Injection Application Triadimefon: Immediately after application, 29.0% of the triadimefon applied with the high pressure injector was recovered from thatch (Fig. 15A, Day 0). Triadimefon thatch residues then decreased over time to 4.5% on Day 14 and were not detectable after this sampling period (Fig. 15A, Days 1-28). Triadimenol: No triadimenol was recovered from the thatch immediately following application of triadimefon with the high pressure injector (Fig. 15B, Day 0). One day after application, 13.7% of the applied triadimefon was recovered from the thatch as triadimenol (Fig. 15B, Day 1). Triadimenol residues increased over time to 42.1% of the applied triadimefon 7 days after application (Fig. 15B, Days 3-7), decreased to 21.8% 14 days following application (Fig. 15B, Days 10-14) and increased to 45.0% of the applied triadimefon 21 days after application (Fig. 15B, Day 21). Triadimenol residues in thatch then decreased to 29.7% of the applied triadimefon 28 days after application (Fig. 15B, Day 28). 3.3.2.3 Soil Boom Sprayer Application Triadimefon: Immediately after application, 4.2% of the triadimefon applied with the boom sprayer was recovered from the soil (Fig. 14A, Day 0). Triadimefon residues in soil then decreased over time to 0.5% on Day 3 and were not detected after this sampling period (Fig. 14A, Days 1-28). 121 TriadimenQl: Immediately following application of triadimefon with the boom sprayer 0.8% of the triadimefon was recovered from the soil as triadimenol (Fig. 14B, Day 0). Triadimenol residues then increased over time to 14.9% of the applied triadimefon 10 days after application (Fig. 14B, Days 1-10). Triadimenol then decreased to 6.5% of the applied triadimefon 14 days following application (Fig. 14B, Day 14), before increasing to 14.8% 28 days after application (Fig. 14B, Days 21-28). High Pressure Injection Application Triadimefon: No triadimefon was recovered from the soil following application with the high pressure injector (Fig. 15A, Days 0-28). Triadimenol: No triadimenol was recovered from the soil immediately following application of triadimefon with the high pressure injector (Fig. 15B, Day 0). One day after application, 0.4% of the applied triadimefon was recovered from the soil as triadimenol (Fig 15B, Day 1). Triadimenol soil residues generally increased over time to 6.0% of the applied triadimefon 28 days after application (Fig 15B, Days 3-28). 3.3.2.4 Root Tissue Boom Sprayer Application Triadimefon: Immediately following application, 0.05% of the triadimefon applied with the boom sprayer was recovered from the root tissue (Fig. 14A, Day 0). No triadimefon was recovered from the root tissue following this sampling period (Fig. 14A, Days 1-28). Triadimenol: Triadimenol was not detected in the roots until three days following application of triadimefon with the boom sprayer, when 0.04% of the applied triadimefon 122 was recovered as triadimenol(Fig 14B, Days 0-3). Triadimenol levels generally increased over time, reaching 0.19% of the applied triadimefon 28 days after application (Fig. 14B, Days 7-28). High Pressure Injection Application Triadimefon; No triadimefon was recovered from the root tissue following application with the high pressure injector (Fig 15A, Days 1-28). Triadimenol: No triadimenol was recovered from the roots until 14 days following application of triadimefon with the high pressure injector when 0.03% of the applied triadimefon was recovered as triadimenol (Fig. 15B, Days 0-14). Triadimenol levels then increased over time to 0.06% of the applied triadimefon 28 days after application (Fig 15B, Days 21-28). 3.4 Discussion It appears likely that triadimefon, through its primary metabolite triadimenol, may pose a threat to groundwater when applied with either the boom sprayer or the high pressure injector. Triadimenol exhibits 1.4 times greater mammalian toxicity, is twice as water soluble, has a slightly lower a soil half-life roughly 15 times that of triadimefon. Although it appears that triadimenol did not bypass the root zone during the time-course of this experiment it was still present in large amounts at the last sampling period. It is possible that during spring and fall recharge the triadimenol still present in the turfgrass system could migrate to the groundwater. The majority of the triadimefon applied (42.3%) was found associated with the thatch layer immediately after application with the boom sprayer. Another 26.3% was found 123 associated with the leaf tissue, 4,2% with the soil, and 0.05% associated with the roots. Following application with the high pressure injector 29.0% of the applied triadimefon (but 86.1% of the amount recovered) was found associated with the thatch. Another 4.7% (13.9% of total)was found associated with the leaf, and no triadimefon was found in the soil or roots. The thatch is apparently acting as both a barrier to the movement of and a reservoir for triadimefon. Metabolism of triadimefon to triadimenol occurred fairly rapidly in all four matrices. Immediately following application with the boom sprayer 70% of the applied triadimefon was recovered from all four matrices. Ten days following application, less than 1% of the triadimefon was recovered. The triadimefon applied with the high pressure injector followed a similar pattern of rapid disappearance. Immediately following application, 34% of the total applied triadimefon was recovered, but by day 10 only 5% of the fungicide was recovered. It has been previously reported that triadimefon is rapidly hydrolyzed to triadimenol in both plants (Garcia et al, 1991) and soil (Patil et al, 1988). This rapid conversion to triadimenol likely accounts for the rapid decrease in triadimefon in turfgrass over the time course of this study and for the relatively short half-life of triadimefon. Triadimenol, however, has a much longer half-life (ti/2 = 110-270 days. Pesticide Manual, 1994) than triadimefon, and therefore poses a much greater potential for groundwater contamination. Triadimenol (acute oral LD50 for rats 700 mg/kg) also possesses greater mammalian toxicity than triadimefon (acute oral LD50 for rats 1000 mg/kg) (Pesticide Manual, 1994). 0 124 One day after application only 34% of the triadimefon found associated with leaf tissue immediately after application with the boom sprayer was still present in the leaf tissue, and triadimenol was first detected in the leaf tissue on this sampling date. On Day 1 triadimefon concentrations in leaf tissue were at their highest level and then decreased throughout the course of the experiment. Triadimenol levels in leaf tissue increased from Day 0 to Day 1, after which they declined slightly and thereafter remained relatively steady. A similar pattern was found following application with the high pressure injector. This could be explained by the loss of both compounds by removal of leaf tissue via mowing, followed by uptake of triadimenol after it passed through the thatch and into the soil, where it became available for uptake by the plant roots. Triadimefon residues in the soil decreased rapidly following both applications, while triadimenol residues in the soil generally increased throughout the course of the experiment. Following boom sprayer application triadimenol residues reached a maximum 10 days following application and, except for a decrease on Day 14, remained fairly constant. Following application with the high pressure injector triadimenol levels generally increased throughout the course of the experiment. This is consistent with the conversion of triadimefon to triadimenol. Triadimenol residues following application with the high pressure injector were not detected until 14 days after application, and then increased until Day 28. Triadimenol residues were found in the root tissue immediately following application with the boom sprayer but then rapidly disappeared and was not found at any time after this. Triadimenol was first detected three days after application and generally increased 125 throughout the course of the experiment. The lack of detection of either of the compounds one day following application could be due to the triadimefon passing through the thatch on application being taken up by the plant and then moving upward to the leaf, thereby disappearing from the root tissue. It may then have taken some time for the triadimenol in the thatch to move downward and be taken up by the root tissue. This would explain the increase throughout the course of the experiment, as triadimenol slowly leached from the thatch to be taken up by the roots. A point of concern in the above discussion is the relatively good mass balance obtained when the fungicides were applied by boom sprayer compared to the two alternative application methods. The high pressure injector deposited the fungicides in discreet bands primarily in the thatch. The high pressure injector nozzles deposited streams of the tank mixture 7.2 cm apart. The internal diameter of the soil corer used to collect turfgrass samples was 2.5 cm. Due to this disparity in size, it is possible that the some turfgrass cores were removed from areas between the fungicide treated bands. Given this sampling regime, an approximation of mass balance was achieved only for the residues recovered following boom sprayer application. Recoveries for fungicides applied by boom sprayer ranged between 81% and 109% over the first three days following application, and between 7% and 47% for the high pressure injector. The low and varied recoveries may indicate that the sampling methodology was inadequate for the high pressure injection application. These findings also indicate a lack of lateral movement of these SBI fungicides, which is supported by anecdotal evidence concerning other lipophilic pesticides. When using the high pressure injector fpr control of white grubs, it 126 was found that the treatment was not effective if the larvae were not active (P. Vittum, personal comm.). This suggests that the insecticides applied by high pressure injection remained in discreet bands and the larvae would have to move through these bands in order to contact the insecticide and achieve control. There was little vertical movement of either compound in the turfgrass profile following either application method. Neither compound was ever detected below the top 5.1 cm of the soil profile. It is therefore unlikely that they would pose a threat to the groundwater. However, the persistence of triadimenol in the soil may lead to a problem after repeated applications. 127 CHAPTER 4 FATE OF BENOMYL IN A TURFGRASS SYSTEM WITH AND WITHOUT POST-APPLICATION IRRIGATION 4.1 Introduction Benomyl (Table 5) is the least water soluble of the fungicides discussed in this thesis (4 mg L’^), which is less than 33% of the solubility of fenarimol. Benomyl is rapidly converted to carbendazim (Table 5) (Clemons and Sisler, 1969) in soil, plants (Baude et ah, 1974; Fuchs et al. 1972, 1974) and water (Singh and Chiba, 1985, Singh et al. 1990). (ti/2 =19 hours in soil). Carbendazim, with a pKa of 4.2, is twice as water soluble as benomyl at pH 7, and more than seven times as water soluble at pH 4. The ti/2 of carbendazim in soil is 6-12 months, close to that of triadimenol (110-270 days) and fenarimol (>365 days). The mammalian toxicity (oral LD50, rats) of benomyl (10,000 mg kg’^) and carbendazim (15,000 mg kg"') are much less than the SBI fungicides discussed earlier (1000 - 2000 mg kg’^). 4.2 Materials And Methods 4.2.1 Experimental Site Two 12.2 m X 12.2 m (0.0149 ha) plots of Kentucky bluegrass were selected at the University of Massachusetts Turfgrass Research Center in South Deerfield, MA. The turf was established in 1985 and overlay a Hadley silt loam soil (coarse, silty, mixed, nonacid, mesic, Typic Udifluvent, pH 6.8). The turf was maintained at a height of 5.1 cm, and irrigated as needed to prevent drought stress (Fig. 1). 128 Table 5. Physical properties, environmental characteristics, application rates, and toxicities of benomyl and carbendazim. ‘ob X) Ui > 2^ _o < 4-> H 5 H ^ < a 52 cd (zi O 5 D ^ c o (D O 00 •*-< c O U rsu ::: % X) O A B 129 X ro (N ON (N o m m (N (U c3 X o, ^ A B I DO ,0 3 x> X X ^ X •u .S C 'C c: !§ Oh X o x ^ U td 2^ o H o ^ o CQ .g ^ ■§ U ‘o CO o z CO »-i (u Dh a I 2 .2 a, o O D O -o (U o B X) O 3 3 (U 3 3 fc »- _ • X X X •I o X X DO <3 ^ NA = Not Applied. . t 4.2.2 Fungicide Application Benomyl (methyl l-(butylcarbamoyl) benzimidazol-2-ylcarbamate) was applied to two plots on 17 July 1991, using a standard boom sprayer (F.E. Meyers and Bro. Co., Ashland, OH). The boom sprayer consisted of nine T-jet “9010” flat fan nozzles applying 621 L ha"' at an operating pressure of 1925 kPa and generated a swath width of 4.6 m. Benomyl was applied at the recommended rate for summer patch control (12.2 kg active ingredient ha'*). The first plot received 1.3 cm of irrigation 15 min after application. The other plot received no post-application irrigation. 4.2.3 Sampling Each of the two plots was divided into sixteen 3.1 m X 3.1 m subplots that were further subdivided into nine 1 m^ sections, one of which was randomly selected for each sampling period. It is estimated that each square meter of the plot contained approximately 455 g of leaf tissue (±15.7%), 11,989 g of thatch (±8.4%), 56,700 g of soil (±3.9%) (1 m X 1 m X 5.1 cm), and 85 g of root tissue (±11.3%) (1 m X 1 m X 5.1 cm) (all in wet weights). These calculated quantities were used to determine the percentages of applied fungicides recovered. The amount of benomyl applied to each turfgrass matrix was calculated by assuming 100% of the applied benomyl was deposited in each matrix. This number was then divided by the actual amount recovered to determine the percent of applied benomyl recovered. Leaf, thatch, roots and soil were sampled immediately before and after the applications and at 1,2,5,9, and 14 days post-application. At each collection period, 75 g of leaf tissue were collected from each of the sixteen subplots by cutting the plants at the soil line. The tissue was placed in 0.95 liter plastic Zip-LocTM freezer bags 130 for storage. Following this process, sixteen soil cores, one from each subplot, were taken with a 30.5 cm X 2.5 cm JMC Zero Contamination Tube Corer (Clements Assoc., Inc., Newton, lA). The thatch layer of each core was delineated as the deepest point at which rhizomes were visible and was marked. Samples were then placed in ice chests and transported to the Massachusetts Pesticide Analysis Laboratory (MPAL) where they were stored at -20°C until analysis. In the laboratory, the thatch layer was removed from the top of each soil core and the remainder of each core was divided into three 10.2 cm X 2.5 cm sections and bulked (i.e. the top 5.1 cm of the cores from each subplot for each application method during a sampling period were placed together in a 2.78 L Zip-Loc™ freezer bag and mixed by rotating the bag for 3 min). The bulked samples were screened through a #10 sieve and the roots or thatch were separated out individually by hand. The soil, roots, and thatch were placed into individual plastic freezer bags for storage in a -20° C freezer for analysis. 4.2.4 Storage Samples On each sampling date, equal amounts of nontreated (i.e. no applied benomyl) leaf tissue and soil cores (i.e., containing soil, roots, and thatch) were collected from an adjacent untreated area to be used as fortified storage samples (i.e., amended with benomyl to assess any loss during storage prior to analysis), concurrent fortified samples (i.e., to provide daily checks of the analytical method efficiency), and concurrent nontreated samples (i.e., to provide daily checks to assess background matrix interferences). Following sample collection, leaf tissue was amended with 0.4 pg g''. 131 thatch with 1.0 |a.g g'*, roots with 2.0 jj,g g’^ and soil with 0.08 ^g g’* of benomyl and stored with the concurrently collected field samples to assess any loss of the compounds during storage (i.e., fortified storage samples). Results are given in Table 6. 4.2.5 Analytical Procedures Leaf: Benomyl residues in leaf tissue were determined using an immunoassay kit from Quantix (Quantix Systems, Cinnaminson, NJ). The immunoassay detected both benomyl and its primary metabolite, carbendazim. Fifty ml of acetonitrile were added to 0.5 g of leaf tissue and shaken for 15 min on a table top shaker (Eberbach) at low speed . The tissue was allowed to settle for 5 min and 100 pi of supernatant was removed and diluted 1000 fold in distilled, deionized water. The immunoassay was performed using the manufacturers instructions. All solutions and buffers were provided with the immunoassay kit. Two hundred pi of diluted supernatant and QA/QC samples were then pipetted into the wells of a 96 well microtiter plate (leaving the first two wells empty for blanking). Fifty pi of an enzyme conjugate were then pipetted into the wells and shaken for 10 min on an microplate shaker (IKA Schuttler). The wells were washed 5 times with wash buffer solution. Two hundred pi of substrate solution were then added to each well and the plate was again shaken for 10 min. Fifty pi of stop solution were then added to each well and mixed for 10 sec. Optical density (OD) was determined for each well utilizing a UV Max plate reader (Molecular Devices) at 650 nm. Thatch: Benomyl residues in thatch were determined by shaking 0.25 g of thatch in 10 ml acetonitrile for 15 min on a table top shaker (Eberbach) at low speed. The thatch was allowed to settle for 5 min, and 100 pi of supernatant were removed and diluted 1000 fold 132 oo uo 00 Os o O -H o' so o 00 oo oo o C/D ON o' o OS Os CN m o' Os (N 00 t(-l«3 U 5^ hJ O' 00 o' CN OO (Z) C (U (Ul fc cd0/1 ±3 CIh g Vi o c 1—> o 1/5 (/2 o 133 in distilled, deionized water. The rest of the procedure followed that outlined for leaf tissue above. Soil: Benomyl residues in soil were analyzed by extracting 8.4 g of soil in 16.8 ml methanol in a 120 ml amber glass bottle shaken for 15 min. The soil was allowed to settle for 5 min, 100 pi of supernatant were removed and diluted 1000 fold in distilled, deionized water. The rest of the procedure followed that outlined for leaf tissue above. Root: Analysis of root tissue for benomyl followed those described for thatch above. 4.2.6 Quality Assurance/Quality Control 4.2.6.1 Instrument Calibration Calibration procedures for the immunoassay system used for benomyl analysis involved developing standard curves consisting of three concentrations of benomyl using a high purity standard (> 99%). A record was kept of: 1) the source of the standards, 2) lot numbers, 3) dates of arrival, 4) storage conditions. A standard curve (1, 5, and 10 pg mf^) was included for each day of analysis (r^ = 0.9978). Calibrations and standard curves were recorded to observe trends or changes over time. Control limits were based upon a particular accuracy associated with routine determination of the analyte. A batch of samples was rejected if certain accuracy limitations were not met. Control limits were defined as the ability of any sample determination to fall within ± 20% of the true value of a quality control set. QC samples were introduced with the normal sample load. QA/QC samples represented about 20% of the sample load. If the QC samples were determined to be 20% above or below their true 134 value the entire batch of samples was rerun. The control samples were in the same matrix as the samples. 4.2.6.2 Sample Controls On each sampling date, equal amounts of nontreated (i.e. no applied benomyl) leaf tissue and soil cores (i.e. containing soil, roots, and thatch) were collected from an adjacent untreated area to be used as fortified storage samples (i.e. amended with benomyl to assess any loss during storage prior to analysis), concurrent fortified samples (i.e. to provide daily checks of the analytical method efficiency), and concurrent nontreated samples (i.e. to provide daily checks to assess background matrix interferences). Following sample collection, leaf tissue was amended with 4.0 pg g'', thatch with 8.0 pg g , roots with 8.0 pg g * and soil with 0.24 pg g ^ of benomyl and stored with the concurrently collected field samples to assess any loss of benomyl during storage (i.e., fortified storage samples). For each day of analysis, corresponding fortified storage samples and nontreated leaf, thatch, roots, and soil amended with benomyl and without (i.e. concurrent fortified and nontreated samples, respectively) were extracted and analyzed to determine recovery efficiency and daily method performance, respectively. Reagent and matrix blanks were also included in each analysis. 135 4.3 Results 4.3.1 Fungicide Concentration 4.3.1.1 Leaf Tissue No Post-Application Irrigation Residues of benomyl associated with leaf tissue following boom sprayer application with no post-application irrigation were 352 ppm (±155 ppm) immediately following application (Fig. 16, Day 0) and attained a maximum concentration of 434 ppm (±91 ppm) on Day 2 (Fig. 16, Day 2). Benomyl residues then declined to 65 ppm (±1.7) on Day 14 (Fig.l6, Day 14). Post-Application Irrigation Residues of benomyl associated with leaf tissue following boom sprayer application and post-application irrigation were 206 ppm (± 24.2 ppm) immediately after application (Fig. 16, Day 0) and attained a maximum concentration of 234 ppm (± 16.1 ppm) on Day 2 (Fig. 16, Day 2). Benomyl residues then declined to 50 ppm (±21 ppm) on Day 14 (Fig. 16, Day 14). 4.3.1.2 Thatch No Post-Application Irrigation No benomyl was found associated with the thatch immediately following application with the boom sprayer with no post-application irrigation. One day following application benomyl residues were 4.1 ppm (±5.8 ppm) (Fig. 17, Day 1), and increased to 8.0 ppm (±1.7 ppm) on Day 2 (Fig. 17, Day 2). Benomyl residues then decreased to 1.3 ppm (±2.9 ppm) on Day 5 (Fig. 17, Day 5) before increasing to 8.9 ppm on Day 9 (Fig. 17, Day 9). 136 -o c: a r\ ONc\ ri (N no c3C Q c o "S o "Oh ajDh c/5 Sh C ^ O (U ccj 15 .SP •-50) ^ -I o b c/5 'E, C/5 -4-> CC 'tJ c/5 CD O CX ZT 'Z T^3 *1!> § ’§ b O -(-> Ul O >%CD b cc3 Vi X)CD a, C^-H c/5 o B S o D- X) >. C d §1 II Sc Oi^ 8 ^ d cj) o d ^ o cu td c/5 •1-N(SO ^CO pH "O 137 Days Post-Application PMJd - n! 138 v\ 03 C o3 r\ 0\r> CN T3 C o3 o3 Q c .2 "S o 'Dh -t->m O Ph • ^ O (U .2trt tix) T30) -CP £I« o •P ■’—' 43 .2 2 Ph ’B ^ .2 4I. c o Ph ■§ IIp •33 43 Os -13^ ;Pi p >> o3 S O .'p p ^ X) *-' £ ^ O ^ »-l SPC/3 h P.^ £d d O w Xi g >. .2 ^ B p -b .2 p -d 8 .1 o 'B. '-! « (DO 1—< p (U >-l o 00p _P3 !4h pi 139 TD CN h Days Post-Application PMJd - qajKRl ni i^nionsg 140 Benomyl then decreased to 7.3 ppm (±2.3 ppm) on Day 14 (Fig. 17, Day 14). Post-Application Irrigation Residues of benomyl associated with thatch following boom sprayer application and post-application irrigation were 14.0 ppm (±3.3 ppm) immediately following application (Fig. 17, Day 0) and declined to 9.0 ppm (±2.7 ppm) by Day 1 (Fig. 17, Day 1). Benomyl residues then increased to 15.6 ppm (±4.2 ppm)on Day 2 (Fig. 17, Day 2) before declining to 5.2 ppm (±3.1 ppm) on Day 14 (Fig. 17, Day 14). 4.3.1.3 Soil No Post-Application Irrigation Following application by the boom sprayer with no post-application irrigation benomyl was not detected in the soil at any time. Post-Application Irrigation Benomyl residues following application with the boom sprayer and post-application irrigation were only detected on Day 2 at a concentration of 244 ppb (±106 ppb) (Fig. 18, Day 2). Benomyl residues were detected at no other time in the top 10.2 cm or any 10.2 cm section below this. 4.3.1.4 Root Tissue No Post-Application Irrigation Residues of benomyl associated with root tissue immediately following application by the boom sprayer with no post-application irrigation were 1.3 ppm (1.5 ppm) (Fig. 19, Day 0), and increased to 4.5 ppm (±0.5 ppm) by Day 2 (Fig. 19, Day 2). Benomyl residues 141 c/3 CCS "O -ra a a o^rv Q c o 'S o * ’E, CTi r2 B..2 E .SP .-H •’“' nd c (U o 15 o •'—' Cu c«o ac3 C -M .i-H t/3 • s a N -S •rt O I c>s ■§> a > 142 TD O o o add - nos «*! I^raonag 143 O1/2 Dh c ^•2 ■-3 2 g S *2 2 'S ^ (/2 OX) ^ IJh T3 144 , * ; ^1,»<,” - . -'”^1 X J ■ ,r_,, ^ . ■, -V .• • ■.}- ■' A.-X o^ •^TP—;'~'7*r^ ‘r^--», 00 I\[dd - »00)I UI iXuiouag 145 then dropped to 2.0 ppm (± 2.3 ppm) on Day 5 (Fig. 19, Day 5) before increasing to 4.9 ppm (± 1.2 ppm) on Day 9 (Fig. 19, Day 9). Benomyl then decreased to 3.9 ppm (± 2.3 ppm) on Day 14 (Fig. 19, Day 14). Post-Application Irrigation Residues of benomyl associated with root tissue immediately following application by the boom sprayer with post-application irrigation were 2.9 ppm (±0.2 ppm) (Fig. 19, Day 0) Benomyl residues then increased to 6.0 ppm (±0.8 ppm) on Day 2 (Fig. 19, Day 2) and then decreased to 5.0 ppm (±0.1 ppm) on Day 14. 4.3.2 Percent Of Applied Benomyl Recovered 4.3.2.1 Leaf Tissue Post-Application Irrigation Immediately after application, 7.7% of the benomyl applied by the boom sprayer with post-application irrigation was recovered from leaf tissue (Fig. 20A, Day 0). Benomyl increased to 8.7% on Day 2 (Fig. 20A, Day 2), before declining to 1.9% on Day 14 (Fig. 20A, Day 14). No Post-Application Irrigation Immediately after application, 13.2% of the benomyl applied by the boom sprayer with no post-application irrigation was recovered from leaf tissue (Fig. 20B, Day 0). Benomyl increased to 16.2% on Day 2 (Fig. 20B, Day 2), before declining to 2.4% on Day 14 (Fig. 20B, Day 14). 146 Figure 20. Percent of total applied benomyl/carbendazim recovered from all matrices (total) and from leaf, thatch, roots, and soil for each day from Day 0 to day 14 following application with the boom sprayer without (A) and with (B) post-application irrigation. 147 Percent of Applied Percent of Applied Percent ofAppliedBenomylRecovered- Percent ofAppliedBenomylRecovered- 148 4.3.2.2 Thatch Post-Application Irrigation Immediately after application, 13.8% of the benomyl applied by the boom sprayer with irrigation was recovered from thatch (Fig. 20B, Day 0). Benomyl increased to 15.4% on Day 2 (Fig. 20A, Day 2), and then declined to 5.1% on Day 14 (Fig. 20A, Day 14). No Post-Application Irrigation No benomyl was recovered from the thatch immediately after application by the boom sprayer with no post-application irrigation (Fig. 20B, Day 0). Benomyl was first detected on Day 1, when 4.0 % of the applied benomyl was recovered (Fig. 20B, Day 1). Benomyl then increased to 7.9% on Day 2 before decreasing to 1.3% on Day 5 (Fig. 20B, Days 2- 3). Benomyl then increased to 8.8% on Day 9 before decreasing to 7.2% on Day 14 (Fig. 20B, Days 9-14). 4.3.2.3 Soil Post-Application Irrigation No benomyl was recovered from the soil immediately after application by the boom sprayer with irrigation (Fig. 20B, Day 0). Benomyl was detected on Day 2, when 4.5% of the applied benomyl was recovered (Fig. 20A, Day 2). No benomyl was recovered after this sampling period. No Post-Application Irrigation No benomyl was recovered from the soil at any time following application with the boom sprayer without irrigation. 149 4.3.2.4 Root Tissue Post-Application Irrigation Iminediately after application, 0.040% of the benomyl applied by the boom sprayer with irrigation was recovered from roots (Fig. 20A, Day 0). Benomyl decreased to 0.036% on Day 1 before increasing to 0.084% on Day 2 (Fig. 20A, Days 1-2). Benomyl decreased to 0.054% on Day 9 and was 0.068% on Day 14 (Fig. 20A, Days 9-14). No Post-Application Irrigation Immediately after application, 0.018% of the benomyl applied by the boom sprayer without irrigation was recovered from root (Fig. 20B, Day 0). Benomyl increased to 0.062% on Day 2 before decreasing to 0.028% on Day 5 (Fig. 20B, Days 2-5). Benomyl then increased to 0.068% on Day 9 and then decreased to 0.054% on Day 14 (Fig. 20B, Days 9-14). 4.4 Discussion The concern that benomyl, or its breakdown product carbendazim, could be a potential groundwater contaminant following application in a large amount of water and with post¬ application irrigation appears to be unfounded. Benomyl was recovered from the top 10.2 cm of the soil on only one sampling period in the plot that received post-application irrigation. Benomyl was found in the top 10.2 cm of roots on every sampling date but was never recovered from the soil or roots below this layer. Thus it is not likely that benomyl applied to turfgrass by these means will bypass the entire root zone and contaminate groundwater over the experimental time frame examined. 150 Following application of the benomyl without irrigation, the majority of the benomyl was found associated with leaf tissue. Leaf tissue contained 13.2% of the applied benomyl, roots contained 0.018%, and none was recovered from the thatch or the soil. Immediately following application of benomyl with irrigation, the majority of the benomyl was recovered from thatch, where 13.8% of the applied benomyl was recovered. At this sampling period, 7.7% was recovered from leaves and 0.04% was recovered from root. As expected, post-application irrigation was responsible for the movement of some of the benomyl from the leaf surface its deposition it in thatch and roots. The purpose of the post-application irrigation was to move the benomyl to the target zone, the roots, and this was accomplished, though the levels of benomyl are probably inadequate for control of fungal root pathogens. Though the half-life of benomyl is short (19 hours), that of carbendazim is significantly longer (3 to 6 months). Since the immunoassay detected both benomyl and carbendazim residues together, there is no way to estimate the conversion of benomyl to carbendazim. However, the short half-life of benomyl would lead one to expect that over the course of several days the majority of the benomyl would be converted to carbendazim. Despite the long half-life of carbendazim and the relatively short duration of this experiment (14 days), a decrease in the total amount recovered was detected. Following benomyl application without irrigation, a maximum level of 24.1% of the applied benomyl was recovered on Day 2, which then decreased to 9.6% on Day 14. 151 As expected, post-application irrigation reduced leaf residues immediately following application by 41%, as compared with no irrigation. By Day 5, however, there was no statistically significant difference between the irrigated and non-irrigated plots. Post- application irrigation correspondingly increased benomyl residues in thatch from samples taken in the early collection periods. Immediately following application, benomyl residues were 14 ppm in the irrigated plot while no benomyl was recovered from the plot that received no post-application irrigation. On Day 1, the thatch in the irrigated plots contained benomyl residues that were 2.2 times that found in the plot that received no irrigation, and this increased to a 7.3 fold difference by Day 5. After this period, there was no statistically significant difference between the two plots. Likewise, post-application irrigation also increased the amount of benomyl reaching the roots, though the difference was only significantly different on Days 2 and 5. Immediately following application, benomyl levels in root of the irrigated plots were 1.7 times those from the non-irrigated plots. Five days following application, the root tissue from the irrigated plots contained 2.9 times more benomyl than plots that received no post-application irrigation. On Day 9, however, the non-irrigated plots contained 1.3 times more benomyl than roots from plots that received irrigation. By Day 14, however, the root tissue from the irrigated plots contained benomyl residues 1.3 times those found in the non-irrigated plots. The post-application irrigation was apparently effective in delivering the fungicides to the roots. Benomyl was detected in the top 10.2 cm of soil only on Day 2, when a concentration of 244 ppb was recovered from the irrigated plot. Benomyl w^ also never recovered 152 from the soil of the plots that received no irrigation. Benomyl was never recovered from either plot below the top 10.2 cm section of the soil core. A point of concern in the above discussion was the poor mass balance achieved in the present study. Only 24.1% of the total applied benomyl was recovered from the plot that was not irrigated (Day 2), and 24.7% was recovered from the plot that received irrigation (Day 2). Benomyl was applied with three other fungicides, fenarimol, propiconazole, and triadimefon. Benomyl application rates were 16 times that of fenarimol, 8 times that of propiconazole, and 4 times that of triadimefon. None of these other fungicides were recovered from the thatch, roots, or soil following either application. Despite a pre¬ application test of compatibility as a tank mix, there was a yellow residue in the spray tank at the end of the application. This indicates a formulation incompatibility among the three SBI fungicides and benomyl. In this experiment we were not allowed to perform our own quality control during the boom sprayer application (e.g. calibration of the sprayer, application). Following this problem we were allowed to perform our own validation (e.g. calibration of the sprayer) in the second year. In the second year, benomyl was dropped from the study, and the other three SBI fungicides were applied using the boom sprayer. A relatively good mass balance was achieved for the fungicides following boom sprayer application. 153 CHAPTER 5 SUMMARY Attempts to manage diseases affecting turfgrass root systems, such as summer patch (causal agent, Magnaporthe poae), require that fungicides be delivered effectively to the root zone. The thatch layer, a hydrophobic mat of partially decomposed plant material lying at the top of the soil profile, provides a barrier to the movement of lipophilic compounds to the turfgrass roots. This thatch layer must be penetrated or bypassed in order to efficiently deliver fungicides to the roots. Without the thatch layer to retard the movement of fungicides, however, there is an increased chance of groundwater contamination. In this study, a high pressure injector and a modified slicer/seeder were compared to a traditional boom sprayer application in order to determine if any method is superior in delivering fungicides to the root zone. The majority of the fungicides were found associated with the thatch layer. Approximately 40-60% of the total applied fungicides were recovered from the thatch, 18-25% from the leaf, and 10-20% was recovered from the soil immediately following application with the boom sprayer. Immediately following application with the high pressure injector, 20-30% of the total applied fungicides (but 78-86% of the total amount recovered) were recovered from the thatch, 5-8% was recovered from the leaf tissue and none was recovered from the soil or roots. Immediately following application with the modified slicer/seeder, 16% of the total applied 154 propiconazole (but 87% of the propiconazole recovered) was associated with the thatch, 2% was found associated with the leaf and none was recovered from the soil or roots. Propiconazole and fenarimol exhibited little breakdown over the 28 days of the experiment, whereas triadimefon exhibited fairly rapid breakdown. Boom sprayer application resulted in leaf residues 3 and 8 times greater than residue levels detected following application with the high pressure injector and modified slicer/seeder, respectively, with fungicide levels generally decreasing over time. In contrast to the leaf tissue, fungicide levels in the soil generally increased over time. Following application with the boom sprayer, maximum fenarimol concentrations (determined as a percentage of the total applied fenarimol) were as follows: leaf tissue- 26.3% one day post¬ application (Day 1); thatch- 82.0% Day 3; soil- 5.8% Day 28; fenarimol was never detected in the roots following boom sprayer application. Maximum propiconazole recoveries following boom sprayer application as follows; leaf- 26.3% Day 1; thatch- 82.3% Day 3; soil- 4.2% Day 0; no propiconazole was recovered from the roots following application with the boom sprayer. Maximum triadimefon concentrations following application with the boom sprayer were as follows; leaf- 23.6% Day 0; thatch- 42.3% Day 0; soil- 4.2% Day 0; root- 0.05% Day 0. Nevertheless, none of the application methods allowed the fungicides to move through the turfgrass system and reach the roots in any appreciable amount. None of the fungicides, in fact, were ever found below the top 5.1 cm of the soil, so the threat of groundwater contamination appears to be minimal in the short time interval (1 month) following application. 155 Triadimefon is rapidly metabolized to triadimenol in turfgrass plants, thatch, soil, and roots. Triadimenol poses a greater threat to groundwater than triadimefon because it is more water soluble, has a greater half-life, a lower value, and greater mammalian toxicity. Immediately following boom sprayer application, leaf tissue residues were 24% of the applied triadimefon. These levels declined until triadimefon was no longer detectable in leaf tissue after 28 days. Triadimenol was first recovered one day after application, when 8% of the residues were recovered as triadimenol. Triadimenol levels first declined in leaf tissue until 10 days following application (3% of applied), after which they increased to 7% of the applied triadimefon after 28 days. Immediately after application with the high pressure injector, only 5% of the applied triadimefon was recovered from leaf tissue. These residues declined until triadimefon was not detectable after 21 days. Immediately following application, 2% of the applied triadimefon was recovered from leaf tissue as triadimenol, and these levels increased to 3% at. Thatch residues immediately following application of triadimefon with the boom sprayer were 42% of the applied triadimefon, and 12% was recovered as triadimenol. Triadimefon levels declined and were not detected in the thatch after 7 days. Triadimenol levels increased to 70% of the applied triadimefon after 5 days. These levels dropped slowly until the end of the experiment, when 52% of the applied triadimefon was recovered as triadimenol. Immediately following application with the high pressure injector, 29% of the applied triadimefon was found in the thatch. Triadimenol was first detected one day after application, when 14% of the applied triadimefon was recovered as triadimenol. Triadimefon levels then decreased and were not detectable in the thatch after 156 21 days. Triadimenol levels increased until seven days after application, when 42% of the applied triadimefon was recovered as triadimenol. Triadimenol levels then slowly decreased to 30% of the applied triadimefon after 28 days. Soil residues immediately following application with the boom sprayer were 4.2% of the applied triadimefon. Triadimefon residues declined to 0.5% of the amount applied 3 days following application, and were not detectable in the soil after 7 days. Immediately following application, 0.8% of the applied triadimefon was recovered as triadimenol. Triadimenol residues in soil generally increased to 14.8% of the applied triadimefon after 28 days. Triadimefon was never detected in the soil following application. Triadimenol was first detected in the soil one day following application, when 0.4% of the applied triadimefon was recovered as triadimenol. Triadimenol residues increased to 6.0% of the applied triadimefon after 28 days. Root residues following application with the boom sprayer were 0.05% of the applied triadimefon immediately after application, but not after that time. Triadimenol was first detected in the roots after 3 days, when 0.04% of the applied triadimefon was recovered as triadimenol. Triadimenol levels increased to 0.19% of the applied triadimefon at 28 days.Triadimefon was not detected in the root tissue following application with the high pressure injector. Triadimenol was first detected in the roots at 14 days, when 0.03% of the applied triadimefon was recovered as triadimenol. These levels increased to 0.06% of the applied triadimefon at 28 days. 157 Tnadimefon was converted to the more mobile triadimenol fairly rapidly in all four matrices. Though this would appear to increase its threat to the groundwater, neither compound was ever detected below the first 5 cm of the soil profile. Benomyl levels in the leaf tissue of the plot that received no irrigation were 1.7 times those found in the plots that received post-application irrigation immediately following application. Benomyl levels in the non-irrigated plot remained above those for the irrigated plot throughout the experiment. Immediately after application and irrigation, 13.8% of the applied benomyl was recovered from the thatch. No benomyl was recovered from the thatch in the plot that received no post-application irrigation until 2 days following application. By the end of the experiment (14 days) the benomyl residues in thatch were similar (5.1% with irrigation versus 7.2% without irrigation). Benomyl was recovered from the soil on only one sampling period. Two days following application, 4.5% of the applied benomyl was recovered from the irrigated plot, while benomyl was never detected in the plot that received no post-application irrigation. Post-application irrigation resulted in benomyl root tissue residues that were 2.2 times those found in the non-irrigated plot. Residues in the root tissue in the irrigated plot remained approximately twice that of the non-irrigated plot until 14 days following application, when root tissue in the irrigated plot contained benomyl residues 1.3 times that found in the non-irrigated plot. Thus, post-application irrigation does appear to allow benomyl to move into the thatch and roots more quickly. The fact that benomyl was 158 never recovered from the soil or roots below the 10.2 cm layer indicates that benomyl does not pose a threat to groundwater over the 14 days of the experiment. 5.1 Conclusions • The thatch layer provides a barrier to the movement of lipophilic flmgicides to the turfgrass roots. • The majority of the fungicides were found associated with the thatch layer. • Propiconazole, fenarimol and benomyl/carbendazim exhibited little breakdown over the course of the experiment. • Triadimefon is rapidly hydrolyzed to triadimenol in leaf, thatch, soil, and roots. • The triadimefon metabolite, triadimenol, was the only compound found in the roots in any appreciable amount. • Post-application irrigation increased benomyl/carbendazim residues in thatch and turfgrass roots. • Fourteen days after application, irrigation had no effect on benomyl/carbendazim levels. • None of the application methods allowed the fungicides to reach the roots in any appreciable amount. • None of the fungicides were found below the top 5.1 cm of the soil, so the threat of groundwater contamination appears minimal over the time course of the experiments. • Because a large proportion of the applied fenarimol, propiconazole, and the triadimefon metabolite, triadimenol, remained associated with the thatch at the end of the study, a longer experimental time frame should be examined that includes spring and fall recharge events in future research. 159 BIBLIOGRAPHY Abdou, W.M., Mahran, M.R., Sidky, M.M., and Wamhoff, H. 1986. 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