PURPLE NUTSEDGE (CYPERUS ROTUNDUS L.) AND YELLOW NUTSEDGE (CYPERUS ESCULENTUS L.) MANAGEMENT WITH TILLAGE AND THE HERBICIDES IMAZAPIC AND IMAZETHAPYR

Transcription

1 PURPLE NUTSEDGE (CYPERUS ROTUNDUS L.) AND YELLOW NUTSEDGE (CYPERUS ESCULENTUS L.) MANAGEMENT WITH TILLAGE AND THE HERBICIDES IMAZAPIC AND IMAZETHAPYR By DEREK DUANE HORRALL A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY UNIVERSITY OF FLORIDA

2 2010 Derek Duane Horrall 2

3 TABLE OF CONTENTS LIST OF TABLES... 5 LIST OF FIGURES... 9 ABSTRACT CHAPTER 1 INTRODUCTION page Morphology Tuber Biology Tuber Formation Photosynthesis Cultural Control Mechanical Control Herbicides History of the Imidazolinone Family of Herbicides EFFECT OF TILLAGE AND GLYPHOSATE APPLICATION ON CONTROL OF PURPLE NUTSEDGE (CYPERUS ROTUNDUS L.) AND YELLOW NUTSEDGE (C. ESCULENTUS L.) Introduction Materials and Methods Tillage Study Glyphosate and Tillage Study Results and Discussion Tillage Study Glyphosate and Tillage Study IMAZAPIC AND IMAZETHAPYR FOR PURPLE NUTSEDGE (CYPERUS ROTUNDUS L.) CONTROL Introduction Materials and Methods Results and Discussion Tuber Number Tuber Weight Tuber Germination EFFECTS OF GROWTH STAGE, SITE OF APPLICATION, AND RATE OF IMAZAPIC AND IMAZETHAPYR ON PURPLE NUTSEDGE (CYPERUS ROTUNDUS L.) AND YELLOW NUTSEDGE (C. ESCULENTUS L.) CONTROL

4 Introduction Materials and Methods Site of Application Site of Application and Nutsedge Age Root Uptake Nutsedge Growth and Tuber Production Results and Discussion Site of Application Site of Application and Nutsedge Age Root Uptake Nutsedge Growth and Tuber Production INFLUENCE OF IMAZAPIC AND IMAZETHAPYR ON PURPLE NUTSEDGE (CYPERUS ROTUNDUS L.) AND YELLOW NUTSEDGE (C. ESCULENTUS L.) GROWTH AND REPRODUCTION Introduction Materials and Methods Results and Discussion EFFECTS OF RESIDUAL CONCENTRATIONS OF SOIL-APPLIED IMAZAPIC AND IMAZETHAPYR ON THE GROWTH AND REPRODUCTION OF PURPLE NUTSEDGE (CYPERUS ROTUNDUS L.) AS A FUNCTION OF TIME Introduction Materials and Methods Results and Discussion CONCLUSIONS REFERENCES BIOGRAPHICAL SKETCH

5 LIST OF TABLES Table page 1-1 Imazapic chemistry and toxicity information Imazethapyr chemistry and toxicity information Effect of tillage on mid-season purple nutsedge tuber density, three years combined Effect of tillage on late-season purple nutsedge tuber density, three years combined Effect of tillage on mid-season purple nutsedge tuber weight, three years combined Effect of tillage on late-season purple nutsedge tuber weight, three years combined Effect of tillage on purple nutsedge tuber germination, three years combined Effect of glyphosate and tillage on purple nutsedge tubers, year one Effect of glyphosate and tillage on purple nutsedge tubers, year two Effect of glyphosate and tillage on yellow nutsedge tubers, year one Effect of glyphosate and tillage on yellow nutsedge tubers, year two Effect of herbicide treatments on mid-season purple nutsedge tuber density, year one Effect of herbicide treatments on late-season purple nutsedge tuber density, year one Effect of herbicide treatments on late-season purple nutsedge tuber density, year two Effect of herbicide treatments on mid-season purple nutsedge tuber density, year three Effect of herbicide treatments on late-season purple nutsedge tuber density, year three Effect of herbicide treatments on mid-season purple nutsedge tuber weight, year one Effect of herbicide treatments on late-season purple nutsedge tuber weight, year one Effect of herbicide treatments on late-season purple nutsedge tuber weight, year two Effect of herbicide treatments on late-season purple nutsedge tuber weight, year three Purple nutsedge tuber germination, years one, two, and three

6 4-1 Effect of foliar and soil + foliar applied herbicides on purple nutsedge growth, year one Effect of foliar and soil + foliar applied herbicides on yellow nutsedge growth, year one Effect of herbicide rate, herbicide site of application, and stage of growth at application on purple nutsedge control, 4 weeks after treatment, years one and two Effect of herbicide, herbicide rate, herbicide site of application, and stage of growth on purple nutsedge shoot height 4 weeks after treatment, years one and two Effect of herbicide, herbicide rate, herbicide site of application, and stage of growth on purple nutsedge shoot number 4 weeks after treatment, years one and two Effect of herbicide, herbicide rate, herbicide site of application, and stage of growth on purple nutsedge shoot dry weight 4 weeks after treatment, years one and two Effect of herbicide, herbicide rate, herbicide site of application, and stage of growth on purple nutsedge regrowth 4 weeks after initial harvest (8 weeks after treatment), years one and two Effect of herbicide, herbicide rate, herbicide site of application, and stage of growth at application on purple nutsedge regrowth shoot height 4 weeks after initial harvest, (8 weeks after treatment), years one and two Effect of herbicide, herbicide rate, herbicide site of application, and stage of growth at application on purple nutsedge shoot regrowth dry weight 4 weeks after initial harvest, (8 weeks after treatment), years one and two Effect of herbicide, herbicide rate, herbicide site of application, and stage of growth at application on yellow nutsedge control 4 weeks after treatment, years one and two Effect of herbicide, herbicide rate, herbicide site of application, and stage of growth on yellow nutsedge shoot height 4 weeks after treatment, years one and two Effect of herbicide, herbicide rate, herbicide site of application, and stage of growth on yellow nutsedge shoot number 4 weeks after treatment, years one and two Effect of herbicide, herbicide rate, herbicide site of application, and stage of growth on yellow nutsedge shoot dry weight 4 weeks after treatment, years one and two Effect of herbicide, herbicide rate, herbicide site of application, and stage of growth on yellow nutsedge regrowth 4 weeks after initial harvest (8 weeks after treatment), years one and two

7 4-15 Effect of herbicide, herbicide rate, herbicide site of application, and stage of growth at application on yellow nutsedge regrowth shoot height 4 weeks after initial harvest, (8 weeks after treatment), years one and two Effect of herbicide, herbicide rate, herbicide site of application, and stage of growth at application on yellow nutsedge shoot regrowth dry weight 4 weeks after initial harvest, 8 weeks after treatment, years one and two Effect of surface applied herbicide, herbicide rate, and stage of growth at application on purple nutsedge control, three years combined Effect of surface applied herbicide, herbicide rate, and stage of growth at application on purple nutsedge regrowth, three years combined Effect of surface applied herbicide, herbicide rate, and stage of growth at application on yellow nutsedge regrowth, three years combined Effect of surface applied herbicide, herbicide rate, and stage of growth at application on yellow nutsedge regrowth, three years combined Effect of herbicide on purple nutsedge control, two years combined Effect of herbicide rate of application on purple nutsedge control, two years combined Effect of plant growth stage at time of herbicide application on purple nutsedge control, two years combined Effect of herbicide and stage of growth at time of application on purple nutsedge growth, two years combined Effect of herbicide rate and stage of growth at time of application on purple nutsedge growth, two years combined Effect of herbicide on yellow nutsedge control, two years combined Effect of herbicide rate of application on yellow nutsedge control, two years combined Effect of plant growth stage at time of herbicide application on yellow nutsedge control, two years combined Effect of herbicide and stage of growth at time of application on yellow nutsedge growth, two years combined Effect of herbicide rate and stage of growth at time of application on yellow nutsedge growth, two years combined

8 5-1 Effect of preemergence herbicide treatments on purple nutsedge reproduction, years one and two Effect of early postemergence herbicide treatments on purple nutsedge reproduction, years one and two Effect of preemergence herbicide treatments on yellow nutsedge reproduction, years one and two Effect of early postemergence herbicide treatments on yellow nutsedge reproduction, years one and two Effects of residual concentrations of soil-applied imazapic and imazethapyr on purple nutsedge control and tuber number and dry weight as a function of time Effects of residual concentrations of soil-applied imazapic and imazethapyr on purple nutsedge shoot number and height, and shoot and root dry weight, as a function of time Corn root bioassay

9 LIST OF FIGURES Figure page 6-1 Imazapic dose response for corn root bioassay 1 MAT Imazethapyr dose response for corn root bioassay 1 MAT

10 Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy PURPLE NUTSEDGE (CYPERUS ROTUNDUS L.) AND YELLOW NUTSEDGE (CYPERUS ESCULENTUS L.) MANAGEMENT WITH TILLAGE AND THE HERBICIDES IMAZAPIC AND IMAZETHAPYR Chair: Barry J. Brecke Major: Agronomy By Derek Duane Horrall May 2010 Purple and yellow nutsedge (Cyperus rotundus L. and C. esculentus L.) together, rank fifth in importance among all weeds in the United States. Both of these nutsedge species reduce peanut (Arachis hypogaea L.) yields by competing for water, light, and nutrients. They also reduce crop quality due to the contamination of harvested peanuts by nutsedge tubers. Due to the prolonged use of herbicides that differentially controlled yellow nutsedge, purple nutsedge has become an even greater problem in southeastern peanut fields. Imazapic (Cadre) and imazethapyr (Pursuit) were the first herbicides to provide effective postemergence purple nutsedge control in peanuts. Field and greenhouse studies were conducted to determine the effectiveness of imazapic and imazethapyr in controlling purple and yellow nutsedge, and to evaluate the impact these herbicides have on nutsedge tuber production. Tillage was also evaluated as a means of reducing purple nutsedge growth and reproduction. Tilling at two week intervals more than twice in May and June did not result in significantly lower purple nutsedge tuber numbers in July. Tilling at two week intervals more than three times throughout the season did not significantly reduce tuber numbers by September. 10

11 Alternating tillage operations with glyphosate applications resulted in the greatest reduction in tuber number, weight, and viability at mid- and late-season sampling dates. In an herbicide screening study conducted for both purple and yellow nutsedge in the field, imazapic applied at the rate of 71g ha -1 early postemergence (EPOST) resulted in the greatest reduction of tuber numbers and tuber dry weights by July and September. Greenhouse studies indicated that EPOST applications of imazapic and imazethapyr 2 weeks after emergence (WAE) were more effective than those applied to purple and yellow nutsedge 4 and 6 WAE. Foliar-only treatments of purple and yellow nutsedge 2, 4, and 6 WAE provided better shoot control than soil-applied treatments. The greatest control of nutsedge, however, regardless of plant age, was obtained by treating both the foliage and soil. It was determined at the conclusion of a year-long greenhouse study that soil-applied imazapic provided better residual control of purple nutsedge than imazethapyr. 11

12 CHAPTER 1 INTRODUCTION Purple nutsedge (Cyperus rotundus L.) and yellow nutsedge (Cyperus esculentus L.) are flowering, monocotyledonous, perennial plants classified in the kingdom Plantae, class Angiospermae, subclass Monocotyledoneae, superorder Commelinidae, order Cyperales, and family Cyperaceae (Salisbury and Ross 1992). Cyperaceae is known as the sedge family. This family of plants consists of 146 genera and 5,315 species (Zomlefer 1994). Two hundred and twenty of these species are considered to be weeds, of which roughly 42% are among the species in the genus Cyperus (Bendixen and Nandihalli 1987; Zomlefer 1994). Purple nutsedge is also known as purple nutgrass and cocograss. Additional common names for yellow nutsedge include yellow nutgrass, northern nutgrass (Bell et al. 1962), chufa (Bendixen and Nandihalli 1987), tigernut (Addy and Eteshola 1984), earth almond, ground almond, and rush nut (Salisbury and Ross 1992). Purple nutsedge is native to India (Holm et al. 1991a), while the region of origin for yellow nutsedge is unknown (Doll 1983b). Purple and yellow nutsedge are the worst and sixteenth worst weeds in the world, respectively (Holm et al. 1991b). Purple nutsedge grows in more countries, regions, and localities than any other weed in the world (Holm et al. 1997). It occurs in Africa, Asia, Australia, Europe, North America, and South America. It is considered the world's worst weed, based on the number of countries where it is reported as a serious, principal, or common weed (Bendixen and Nandihalli 1987; Holm et al. 1991a). Purple nutsedge is an important weed in 52 crops in 92 tropical and subtropical countries. Low temperatures seem to limit its range to latitudes within 30 0 to 35 0 north and south of the equator (Holm et al. 1991a; Williams 1982). Competition with crops for light, water, and nutrients is the primary factor in determining degree of weediness. 12

13 Purple nutsedge is a serious or principal weed in rice (Oryza sativa L.), sugarcane (Saccharum officinarum L.), corn (Zea mays L.) cotton (Gossypium hirsutum L.), and vegetable crops. Purple nutsedge is listed as one of the three most noxious weeds of rice in Ghana, Indonesia, Iran, Peru, South Africa, and Taiwan; of sugarcane in Argentina, India, Indonesia, Peru, and Taiwan; of corn in Ghana and the Philippines; of cotton in Sudan, Swaziland, Turkey, and Uganda; and of vegetable crops in Brazil, Malaysia, Taiwan, and Venezuela. In some of these and other areas around the world, purple nutsedge is a significant pest in peanut (Arachis hypogaea L.), soybean (Glycine max L.), sorghum (Sorghum vulgare Pers.), and many plantation crops such as coffee (Coffea arabica L.) and tea (Thea sinensis and Camellia sinensis) (Holm et al. 1991a). The widespread distribution of purple and yellow nutsedge is largely a function of the dissemination of tubers. Although both species produce viable seeds, they are insignificant for propagational purposes in most cultivated areas, mainly due to inadequate seedling vigor (Stoller and Sweet 1987). Though it is impossible to verify the exact means by which these weeds were so widely distributed, many plausible explanations exist (Bendixen and Nandihalli 1987). Nutsedge tubers and seeds may contaminate commercial seeds and feeds, and subsequently be distributed widely. Tubers are known to develop in Irish potato (Solanum tuberosum L.) tubers and in other commercial "root" crops such as sweet potato [Ipomea batatas (L.) Poir], sugar beet (Beta vulgaris L.), and onion (Allium cepa L.). It is also known that tubers are a contaminant in the harvest and bagging of these and other crops. Consequently, nutsedge is distributed with these foods and seed stocks. Nutsedge tubers can also contaminate peanuts during harvest and shipment (Bendixen and Nandihalli 1987). 13

14 Tubers are brought to farms on tillage and harvesting implements and are moved by rainwater into drainage ditches, where they may be carried further. Tubers are also brought into new areas by floodwaters and by surface irrigation water. Dissemination of floating tubers is of major concern in paddy rice. In addition, wind can sweep tubers along the soil surface for great distances. Seeds might be able to survive being passed through the digestive tracts of birds and other animals, and thus be spread over their feeding ranges. Nutsedge often contaminates nursery stock; thus tubers might be distributed with transplanted, potted, or balled plants (Holm et al. 1991a). The many possible reasons for the widespread occurrence of purple and yellow nutsedge suggests that their distribution is limited more by environmental conditions (e.g., cold temperatures, soil moisture content, and degree of sunlight) than by a lack of means of dispersal (Bendixen and Nandihalli 1987). One theory as to how purple and yellow nutsedge spread to the United States is that tubers, rhizomes, and in some instances entire plants, were present in the soil used as ballast in the holds of ships originating from India and other regions where nutsedge was already established (Bendixen and Nandihalli 1987). Upon arrival in the U.S., the nutsedgecontaminated ballast was discarded in order for the holds of the ships to be loaded with goods for the return voyages. Purple and yellow nutsedge combined rank fifth in importance among all weeds in the United States with quackgrass [Elyfrigia repens (L.) Desv.] the only perennial weed ranking higher. Although purple nutsedge is considered to be the worst weed in the world, yellow nutsedge is more widespread and troublesome in the United States due to its ability to tolerate colder temperatures than purple nutsedge. Yellow nutsedge occurs in all 50 states, whereas purple nutsedge is seldom seen north of Arkansas, Tennessee, and Virginia (Bendixen and 14

15 Nandihalli 1987; Holm et al. 1991b). Purple nutsedge also occurs in Hawaii and central and southern California, however, where it is especially troublesome in the production of sugarcane and vegetable crops (Holm et al. 1991a). In addition to being affected by low temperatures, purple nutsedge does not tolerate extremely low or high soil moisture, soils with a high salt content, or shading. When the overstory of such crops as plantation trees and sugarcane begins to shade the soil, the leaves of purple nutsedge yellow and die. The dormant tubers remain viable, however, and as soon as an opening in the canopy appears, the sedge begins to reestablish itself. Purple nutsedge is more sensitive to drought than yellow nutsedge (Bendixen 1973). Other than the aforementioned limitations, however, purple nutsedge grows prolifically in nearly every soil type, elevation, ph, and humidity, and can withstand the highest temperatures known in agriculture (Holm et al. 1991a; Holm et al. 1991b). Under optimum growing conditions, purple nutsedge is more competitive than yellow nutsedge (Bendixen 1973; Wills 1987). In the southeastern United States, purple and yellow nutsedge are most detrimental in peanut, cotton, corn, and vegetable crops. Heavy infestations of either nutsedge species are capable of lowering peanut yields by up to 25%. While good control of yellow nutsedge can be obtained in both cotton and peanut via the use of herbicides, purple nutsedge is easier to control in cotton than in peanut (York 1994). Morphology Purple and yellow nutsedge often grow in mixed stands, and thus may be difficult to distinguish one from the other before they have flowered (Holm et al. 1991b; Wills 1987). Their common names are derived from the color of their inflorescences. Purple nutsedge has a purplish-brown inflorescence, while that of yellow nutsedge is golden brown (Doll 1983a; Wills 1987). Other distinguishing morphological characteristics include leaf color, leaf tip shape, plant 15

16 height, as well as tuber shape, size, color, and taste (Doll 1983a; Doll 1983b; Holm et al. 1991a; Holm et al. 1991b). The leaves of purple nutsedge are dark green, with blunt tips that are similar in shape to the keel of a boat. Yellow nutsedge leaves, on the other hand, are paler green and more acuminate. The leaves of both species are in three ranks with closed sheaths and without ligules. The basal leaves of purple nutsedge are shorter than the inflorescence, while those of yellow nutsedge are longer than the inflorescence. Both species have triangular stems, which are common to all true sedges. Mature plant height for the species ranges from cm for purple nutsedge to cm for yellow nutsedge (Doll 1983a; Doll 1983b; Wills 1987; Wills and Briscoe 1970). Tubers of purple nutsedge are irregular in shape, hairy, nearly black, typically 1-3 cm in size, and occur in chains on wiry rhizomes. Contrastingly, yellow nutsedge tubers tend to be spherical, smooth (i.e., devoid of hairs), generally range from cm in size, and are not linked together by rhizomes. Its tubers may be red, tan, brown, or black. Newly formed tubers of both species are often white, and become darker with age. Those that have over-wintered in the soil tend to be the darkest (Doll 1983a; Doll 1983b). Purple nutsedge tubers taste bitter, while those of yellow nutsedge are esculent; hence its species name, esculentus. Tuber Biology The most impressive characteristic of purple nutsedge is its prolific production of subterranean tubers, which are capable of remaining dormant and thus of sustaining the species through extreme environmental conditions such as drought, flooding, heat, or lack of soil aeration. The plant may grow to a depth of 100 cm in moist, fertile soils. Its rhizomes can puncture and pass completely through the roots and underground storage organs of vegetable root crops including Irish and sweet potatoes, sugar beet, onions, cassava (Manihot esculenta 16

17 Crantz), and carrots (Daucus carota L.), subsequently reducing their market value (Holm et al. 1991a). Purple nutsedge is capable of propagating at an exceptionally fast rate. In one study, a single purple nutsedge tuber placed in a field produced 1,900 plants and almost 7,000 tubers, and covered an area approximately two meters in diameter after just one year (Holm et al. 1991b). In Brazil, purple nutsedge is known as tiririca ate amamnha, which translates as until tomorrow. This refers to the plant s ability to regrow quickly following weeding (Williams 1976). Similarly, a single yellow nutsedge plant was shown to produce 7,000 tubers, with tuber populations of 1,000 per square meter in a single season (Hauser 1962a; Hauser 1962b; Horowitz 1972; Smith and Frick 1937; Stoller and Sweet 1987; Tumbleson and Kommedahl 1961). Tuber production in yellow and purple nutsedge begins four to six weeks after seedling emergence. Ninety-five percent of the tubers for both species are formed within 45cm of the soil surface (Bell et al. 1962; Stoller and Sweet 1987; Tumbleson and Kommedahl 1961). Generally, more than 80% of tubers are located in the upper 15cm of the soil profile (Bayer 1987; Stoller and Sweet 1987). Rhizomes do not penetrate as deeply in finer-textured soils such as those high in clay content. Tubers occur deeper in coarser-textured soils, or in soils frequently disturbed by cultivation (Bayer 1987; Stoller and Sweet 1987). Tuber longevity in both species is dependent upon location in the soil profile. In general, the deeper a tuber is in the soil, the longer it will survive (Bayer 1987). Yellow nutsedge propagates predominately by vegetative parts, including rhizomes, tubers, and a basal bulb (Garg et al. 1967). The basal bulb develops in young seedlings as a swelling at the junction of the mesocotyl and coleoptile (Garg et al. 1967). These bulbs consist of a stem fragment (rhizome) with compressed internodes, which have meristems for roots, 17

18 secondary rhizomes, leaves, and the flower stalk (Stoller and Sweet 1987). The basal bulb gives rise to rhizomes, which differentiate into either tubers or shoots (Garg et al. 1967). Tuber Formation The principal factor stimulating tuber production in yellow nutsedge is day length (Jansen 1971; Stoller and Sweet 1987). Photoperiods longer than 12 hours promote rhizome development and shoot production, while shorter photoperiods promote rhizome tuberization (Bell et al. 1962; Garg et al. 1967; Jansen 1971; Stoller and Sweet 1987). Thus, photoperiods shorter than 12 hours tend to cause rhizomes to differentiate into tubers, while with photoperiods longer than 12 hours, rhizomes have a propensity to differentiate into basal bulbs and shoots (Garg et al. 1967; Stoller and Sweet 1987). Bell et al. (1962) showed that tuberization of yellow nutsedge was promoted at photoperiods of 8 to 12 hours, and shoot formation at 16 hours (Jansen 1971). Tubers are produced on rhizomatous tissue containing numerous buds, a characteristic common to many stem tissues (Stoller and Sweet 1987). In yellow nutsedge grown with greater than 12 hours of daylight, buds on rhizomes sprout and initiate rhizomatous growth which develops into shoots typical of most monocots (Stoller and Sweet 1987). While photoperiod has been identified as the major stimulant for tuber production and flowering in yellow nutsedge (Jansen 1971; Stoller and Sweet 1987), controversy exists regarding the role of photoperiodism in rhizome differentiation and subsequent tuber formation in purple nutsedge. Horowitz (1972) found that natural photoperiods of hours had no apparent effect on tuberization in purple nutsedge, whereas Hammerton (1975) found that day length was the major factor influencing purple nutsedge growth and development. Similarly, Williams (1982) noted that tuber production in purple nutsedge increased as day length decreased. In addition, Berger (1966) reported that tuber formation was induced by a short photoperiod of 10 hours, and inhibited by a photoperiod of 18 hours. Tuber formation can 18

19 apparently occur throughout the year in tropical climates (Hammerton 1975; Horowitz 1972). Before tubers are formed, the plant complex usually includes many shoots interconnected by rhizomes capable of diverting resources into tubers. Thus, tuber formation in purple nutsedge may respond to excess carbohydrate levels, as well as to plant growth regulators, photoperiodism, and temperature (Stoller and Sweet 1987). In yellow nutsedge, 12 to 14 hour photoperiods promote flowering, and consequently seed production, whereas longer or shorter photoperiods are inhibitory (Jansen 1971). Purple nutsedge, on the other hand, is stimulated to flower in short photoperiods of six to eight hours, with the time from emergence to flowering ranging from three to eight weeks (Holm et al. 1991a). Photosynthesis Purple and yellow nutsedge are C-4 plants possessing the dicarboxylic acid photosynthetic pathway (Wills 1987). In addition to being able to convert CO 2 into carbohydrates by the C-3 Calvin cycle common to all photosynthetically active plants, C-4 plants also possess the phosphoenolpyruvate pathway, through which CO 2 is efficiently collected (Wills 1987). C-4 plants are efficient both in obtaining CO 2 from the atmosphere and in recapturing CO 2 that has been expelled through plant respiration. In addition, C-4 plants can assimilate CO 2 at higher temperatures and light intensities than plants possessing only the C-3 pathway (Wills 1987). This phenomenon is demonstrated by the fact that C-4 species tend to exhibit their best growth rates at higher temperatures than do C-3 species. Furthermore, most C-3 species light saturate at 20 to 30% of full sunlight, while C-4 plants usually saturate from 50% to greater than full sunlight (Wills 1987). Perennial species, such as purple and yellow nutsedge, that fix CO 2 at high rates under elevated temperatures and light intensities, and that possess the ability to spread 19

20 by rhizomes, have the potential to become serious weed problems. As C-4 plants, purple and yellow nutsedge are sensitive to shading. While shading reduces tuber production in both species (Keeley and Thullen 1978; Stoller and Sweet 1987), it is debatable whether it reduces tuber numbers significantly. For instance, Patterson (1982) and Wills (1975) found that both species were able to efficiently divert dry matter into tuber production even when grown under 90% shade. When nutsedge tubers sprout, one or more of the many buds on the tuber begin to grow. Yellow nutsedge buds are concentrated at the apical end of the tuber (Bendixen 1973), while purple nutsedge buds gather at nodes along the entire length of the tuber (Stoller and Sweet 1987). Typically, several buds sprout simultaneously while others remain dormant for subsequent sprouting (Stoller et al. 1972). Purple nutsedge tubers exhibit apical dominance, since the sprouting of the most apical buds inhibits sprouting of the more basal buds (Bayer 1987; Stoller and Sweet 1987). Yellow nutsedge tubers, however, break dormancy in acropetal order, beginning with the oldest, most basipetal bud (Bayer 1987; Stoller and Sweet 1987). Yellow nutsedge tubers are capable of sprouting at least three times, with 60% of the total tuber dry weight, carbohydrate, starch, oil, and protein being expended on the first sprouting, while only 10% of these constituents are depleted during each of the next two sproutings (Stoller et al. 1972). In addition to the preceding constituents, organic acids are apparently consumed during the sprouting of purple nutsedge tubers (Stoller and Sweet 1987). Carbohydrate levels in tubers and rhizomes of Cyperus species are dependent upon their function. Rhizomes of Cyperus species are not storage organs, but rather serve to transport carbohydrates from older to actively growing tubers and young sprouts. Hence, they contain 20

21 more sugars than starch (Diethelm and Bocion 1993). Conversely, tubers contain more starch than sugars since they are storage organs. However, not all tubers possess the same starch-tosugar ratio. The younger the tubers, the lower the ratio of starch to sucrose and reducing sugars. This is a consequence of their higher metabolic rates and net import of carbohydrates (Diethelm and Bocion 1993). Even young, actively growing tubers possess sufficient carbohydrates for sprouting, as long as dormancy is broken (Diethelm and Bocion 1993). Cyperus tubers store much greater amounts of carbohydrates than what is required for sprouting. Therefore, some essential step in carbohydrate metabolism must be almost completely inhibited in order for a strong herbicide effect to occur. An exception would be toxic effects caused by inhibition of an essential enzyme; however, at this time it is unknown which enzyme might be involved. The study of essential enzymes in transgenic plants has not yielded conclusive results. AGPase (ADP-glucose pyrophosphorylase), an essential gene for starch synthesis, is an example. Diethelm and Bocion (1993) showed that potato tubers possessing the antisense gene for AGPase were found to have much lower starch content, but a significantly greater sugar concentration. Inhibition of starch synthesis in tubers led to a much higher number of tubers per potato plant, with the tubers being considerably smaller than those in the wild type. Although the antisense gene converted the tubers from starch to sugar-storage organs, tubers displayed normal sprouting and subsequent plant development. These researchers concluded that the results from genetically altered plants indicate that a reasonable method of eradicating perennial plants such as Cyperus species through carbohydrate starvation does not seem feasible (Diethelm and Bocion 1993). Cultural Control An integrated program that utilizes several methods is usually most effective in controlling purple and yellow nutsedge (Wax 1975). Care should be taken not to transport tubers 21

22 by cultivation and harvesting implements, and by other means stated earlier (Holm et al. 1991a). Purple and yellow nutsedge have also spread as production inputs have increased (Hauser et al. 1973). The augmentation of nutsedge densities is largely a consequence of reduced competition due to better annual weed control and ameliorated growing conditions due to the increased use of soil amendments (Hauser et al. 1973). Rotating crops, and subsequently the herbicides applied to a given field, is known to aid in nutsedge control (Wax 1975). Crop rotation may also increase a crop s competitiveness against nutsedge by reducing insect, disease, and nematode damage ( and Brecke 1997). Crop selection should include fast-growing, competitive crops that quickly form a canopy over the soil to shade nutsedge, thus reducing its growth and reproductive potential (Glaze 1987; Keeley and Thullen 1978). Crop selection also determines the planting date, and thus the timing and frequency of tillage possible, as well as the duration of weed control required to produce economical yields (Glaze 1987). Delayed planting dates make additional preplant cultivations possible (William and Bendixen 1987). For most crops, the first four to eight weeks are the most critical in terms of competition from nutsedge. The time required for a crop to produce a canopy sufficiently large to shade nutsedge varies with each crop, but is generally between four and sixteen weeks (Glaze 1987). Crops can become more competitive by using the narrowest practical row spacing, increasing seeding densities, applying fertilizers as indicated by soil tests, and by controlling insects and diseases (Brecke and Stephenson, 2006a; Doll 1983a; Glaze 1987). Besler et al. (2008) found both imazapic and diclosulam applied to peanut grown in a twin-row pattern provided better yellow nutsedge control than herbicide applications to peanut grown in singlerow spacing. 22

23 Mechanical Control The keys to successful mechanical control of nutsedge species are timeliness and frequency (Doll 1983a). Different theories exist as to the preferred timing of cultivation. Doll (1983a) found tillage to be most effective when performed prior to nutsedge being well established, whereas Stoller and Wax (1973), Williams (1982), and Glaze (1987) observed maximum benefits in nutsedge control when cultivation was delayed until a substantial number of tubers had germinated. Disadvantages to mechanical weed control measures do exist. Preplant cultivation can promote tuber germination. The first tillage operation may kill many of the shoots, but dormant buds on tubers of both purple and yellow nutsedge are capable of sprouting another two or three times (Doll 1983a). Furthermore, subsequent cultivation often helps place non-sprouted tubers in soil conditions conducive to sprouting (Doll 1983a). Soil disturbance, however, can also move tubers closer to the soil surface where they are more susceptible to desiccation and cold temperatures (Glaze 1987). Glaze (1987) claims purple nutsedge can be reduced to manageable populations by plowing or disking at intervals of three weeks or less for a minimum of two years, and by planting a winter grain or hay crop. An obvious disadvantage to this is that fields must be fallowed. It is also costly in terms of both time and energy. Doll (1983a), however, claims that two to four cultivations prior to planting are sufficient to provide crop species a competitive advantage over nutsedge. Glaze (1987) concluded that in addition to practicing cultural methods such as high plant density and narrow row spacing, cultivation during the growing season, in conjunction with the use of herbicides, will eventually provide adequate pressure to maintain nutsedge populations at manageable levels. 23

24 Research has shown that eradication of nutsedge is possible. Smith and Mayton (1942) disked 11 fields infested with nutsedge every two, three, or four weeks for two years, and almost always achieved eradication. This frequency of cultivation is obviously not practical from a farmer's perspective. Irrigation in these studies was shown to reduce the number of tillage operations needed to achieve eradication by inducing tubers to sprout more readily than if left under natural rainfall conditions. Herbicides Nonchemical control measures alone do not provide satisfactory control of nutsedge species in most cases (Wax 1975). Attempts to chemically eradicate purple nutsedge began in 1925 in India when Ranade and Burns applied a 2% solution of table salt and copper sulfate at two-week intervals to a dense stand. One hundred applications of table salt significantly reduced, but failed to eradicate the tubers. Copper sulfate merely caused leaf chlorosis, and had no effect on the tuber population (Doll 1983a). Banks (1983) conducted field and greenhouse experiments to determine the effects of soilapplied herbicides on initial yellow nutsedge control, as well as their effects on shoot regrowth and tuber production. Herbicides evaluated included alachlor, fluridone, diethatyl, metolachlor, acetochlor, norflurazon, and vernolate (Banks 1983). Herbicides were mixed in a sandy loam soil with a ph of 6.3 and 1.1% organic matter and transferred to plastic pots. Four yellow nutsedge tubers were placed in each pot. Shoot counts were taken at two-week intervals to determine initial control for different periods after treatment. Also at two-week intervals, original tubers from four pots in each replication were removed, washed, and transplanted in untreated soil. After eight weeks, counts on new shoots and tubers were taken for each pot (Banks 1983). Only fluridone and norflurazon provided 100% control of yellow nutsedge when visually rated eight weeks after planting in treated soil in a greenhouse. Fluridone, norflurazon, and 24

25 acetochlor were most effective in preventing new tuber production (Banks 1983). Soybeans were more competitive against yellow nutsedge in the field. Fluridone was most effective, and norflurazon the least effective in reducing tuber and shoot populations of yellow nutsedge in cotton. However, tubers in plots treated with fluridone tended to produce more shoots per tuber than the other treatments. All treatments except alachlor provided good control of yellow nutsedge in soybeans (Banks 1983). The acid amide herbicide metolachlor has been used to provide partial control of yellow nutsedge in several crops including corn and peanut, but it has only slight activity on purple nutsedge (Webster and Coble 1997). The photosystem II inhibitor herbicide bentazon is applied POST in corn, soybean and peanut to augment yellow nutsedge control, but as with metolachlor, bentazon lacks adequate activity on purple nutsedge (Stroller et al. 1975). Diclosulam is a triazolopyrimidine sulfonanilide herbicide used in soybean and peanut (Dotray et al. 1998). Grichar et al. (2008) reported greater than 80% control of yellow nutsedge in peanut when diclosulam was applied PRE at or kg ha -1 followed by (fb) S- metolachlor applied POST at 0.56, 1.12, or 1.46 kg ha -1. The sulfonylurea halosulfuron controls both purple and yellow nutsedge in corn, but has limited activity on most grasses and certain broad leaf weeds (Webster and Coble 1997). Mesotrione, a triketone herbicide, can be applied PRE and POST in corn to improve broadleaf weed control, while also providing added control of yellow nutsedge (Armel et al. 2008). Trader et al. (2008) reported greater than 83% yellow nutsedge control in yellow summer squash with POST applications of halosulfuron at 18 and 27 g ha -1 in combination with clomazone plus ethalfluralin PRE. Norsworthy et al. (2007) reported 66 % suppression of purple nutsedge in chile pepper (Capsicum annuum L.) from POST-directed applications of 25

26 halosulfuron at rates less than or equal to 36 g ha -1. Halosulfuron can also be used POST to control both purple and yellow nutsedge in turfgrass (Czaronta 2004) Applying herbicides at rates below those recommended by the manufacturer has been considered a possible means of minimizing the risk of carryover, provided broadspectrum weed control is not compromised (Troxler et al. 2001). For example, lower use rates alleviate carryover concerns with cotton. The manufacturer indicates cotton should not be planted within 17 months after imazapic application to peanut at the labeled rate (Anonymous, 2007). Grichar (2002) reported imazapic provided 90% yellow nutsedge control in peanut when applied at the rate of 40 g ha -1, well below the labeled rate of 70 g ha -1. Single applications and mixtures of imazapic, diclosulam, and flumioxazin provide residual control of yellow nutsedge, as well as many broadleaf weeds in peanut when applied at labeled rates (Grichar 2002, Main et al. 2005). Reduced rates of these herbicides can also be effective. Willingham et al. (2008) reported greater than or equal to 80% control of yellow nutsedge, Florida beggarweed (Desmodium tortuosum L.), hairy indigo (Indigofera hirsuta L.), and sicklepod (Senna obtusifolia L.) in peanut resulting from diclosulam at 6 g ha -1 (1/4 X) (1/4 the labeled use rate) plus flumioxazin at 20 g ha -1 (1/4 X) applied PRE, fb imazapic POST at 17 g ha -1 (1/4 X). Trifloxysulfuron is a POST herbicide developed for use in cotton, sugarcane, tomato and turfgrass. In cotton, trifloxysulfuron provides control of perennial nutsedges as well as several difficult to control broadleaf weeds (Brecke and Stephenson 2006b). Most recently, the development of glyphosate-resistant cotton and soybean cultivars has provided growers another option in nutsedge management. POST applications of this otherwise 26

27 non-selective herbicide may provide an alternative to more conventional chemical control measures (Edenfield et al. 2005). History of the Imidazolinone Family of Herbicides The imidazolinone herbicides were discovered in the mid 1970s and developed in the 1980s by scientists at American Cyanamid Company in Princeton, New Jersey. Their discovery began via random screening tests in which a compound known as phthalamide was found to exhibit herbicidal activity at 4 kg ha -1. This compound subsequently served as the template, or herbicide lead, from which chemicals with stronger activity were derived. Due to their lack of crop selectivity, the first imidazolinone herbicides to be synthesized were tested as total vegetation control agents. Modifications of these initial imidazolinones yielded compounds possessing greater selectivity, as well as higher levels of activity (Shaner and O Connor 1991). All members of the imidazolinone family of herbicides contain an imidazole ring. They differ, however, in the type of ring structure substitution at the 2 (R) position of the imidazole ring. Three possible substitutions at this position include benzene, pyridine, and quinoline. Being weak acids, the water solubility of the imidazolinones is greatly affected by soil ph (Shaner and O Connor 1991). Solubility increases significantly as the soil ph rises from 5 to 7. Adsorption, conversely, is greater in low ph (acidic) soils. Since dissipation of the imidazolinones is due primarily to microbial degradation, leaching and carryover injury to rotational crops are more of a concern in high ph (basic) soils in which microbial activity is typically reduced. Persistence also tends to be greatest in soils high in clay content and organic matter. Risks of leaching and carryover injury, however, are less of a concern in Florida than in northern regions of the country, where cool temperatures often decelerate microbial activity. Losses of these herbicides due to volatilization and photo-degradation are negligible (Shaner and 27

28 O Connor 1991). Rotational crops sensitive to carryover from imadazolinone herbicides include cotton, field corn, potato, canola (Brassica napus L. and B. campestris L.), and sugarbeet (Moyer and Esau 1996). Imidazolinone herbicides inhibit the enzyme acetolactate synthase (ALS), also known as acetohydroxyacid synthase (AHAS) (Moberg and Cross 1990; Schloss 1990; Stidham and Shaner 1990). This is the same mechanism of action as that targeted by the sulfonylurea and triazolopyrimidine sulfonanilide families of herbicides, as well as the herbicide pyrithiobac (Kleschick et al. 1990; Moberg and Cross 1990). ALS is required by plants for the synthesis of the branched chain amino acids leucine, isoleucine, and valine. Secondary effects include reduced levels of RNA and DNA synthesis, and respiration. Activity is first seen in the growing points of susceptible plants, where amino acid demands are greatest. Most tolerant plant species are able to metabolize imidazolinones more quickly than susceptible ones. There is evidence, however, that some tolerant plants possess an altered form of the ALS enzyme that is not subject to inhibition by these herbicides (Shaner and O Connor 1991). Since the ALS enzyme occurs only in plants, imidazolinones pose little threat to humans or the environment. All but one has oral LD 50 values of 5,000 mg kg -1 in rats. By comparison, the LD 50 value of table salt for rats is 3,000 mg kg -1 (Shaner and O Connor 1991; Tables 1.1 and 1.2). Imidazolinone herbicides are readily absorbed by roots and foliage, and are highly mobile once inside plants. They are translocated in both the xylem and phloem, and accumulate in meristematic tissues located in young, actively growing plant parts. Injury symptoms are characterized by cessation in growth, shortened internodes, and chlorosis followed by necrosis in the growing tips of roots and shoots, progressing to older plant parts. Treated plants may become purplish in color in response to a weakened root system. Although movement of these herbicides 28

29 within plants is rapid, death usually does not occur for approximately two weeks (Shaner and O Connor 1991). Imidazolinone herbicides can be tank mixed with several different herbicides, including members of the dinitroaniline family of herbicides. Postemergence applications require the use of a nonionic surfactant (Shaner and O Connor 1991). As a result of their versatility, low mammalian toxicity, environmental safety, and low use rates, the imidazolinones currently play a vital role in food production throughout the world (Shaner and O Connor 1991; Tables 1.1 and 1.2). Imazapic, (±)-2-(4-isopropyl-4-methyl-5-oxo-2-imidazolin-2-yl)-5-methylnicotinic acid, and imazethapyr, (±)-2-(4-isopropyl-4-methyl-5-oxo-2-imidazolin-2-yl)-5-ethylnicotinic acid, are the first herbicides to provide effective postemergence purple nutsedge control in peanuts. Imazapic was registered for use in peanut in the spring of 1996 under the trade name Cadre. Imazethapyr was registered for use in peanut in the spring of 1991 under the trade name Pursuit (Wilcut et al. 1996). Imazapic is not labeled for use in any crop other than peanut, while imazethapyr may also be used for weed control in soybean, dry beans (Phaseolus vulgaris L.), field peas (Pisum sativum L.), and alfalfa (Medicago sativa L.) (Moyer and Esau 1996). Imazapic is labeled for application in peanut postemergence, while imazethapyr may be applied PPI, preemergence, at-cracking, and postemergence. The rate for both imazapic and imazethpyr in peanut is 71 g ha -1. At-cracking and early postemergence treatments provide the best results for both herbicides (Ferrell et al. 2009). Optimum weed control occurs when weeds are 2-4 in height. At-cracking and postemergence treatments require a non-ionic surfactant at 0.25% v/v. 29

30 Imazapic provides enhanced purple and yellow nutsedge control relative to imazethapyr (Richburg et. al 1994; Richburg et. al 1993). In addition, imazapic controls Florida beggarweed and sicklepod, the two most common and troublesome weeds in peanut, whereas imazethapyr provides poor control of these weeds (Wilcut et. al 1996). Imazapic provides a broader spectrum of grass control than imazethapyr. Grass weeds controlled by imazapic include: broadleaf signalgrass (Brachiaria platyphylla L.), large and smooth crabgrass (Digitaria sanguinalis L. and D. ischaemum L.), goosegrass (Eleusine indica L.), seedling johnsongrass (Sorghum halepense L.), southern sandbur (Cenchrus echinatus L.), and Texas panicum (Panicum texanum L.). In addition, imazapic controls the following broadleaf weeds: bristly starbur (Acanthospermum hispidum L.), common cocklebur (Xanthium strumarium L.), coffee senna (Cassia occidentalis L.), Florida pusley (Richardia scabra L.), hairy indigo, common lambsquarters (Chenopodium album L.), morningglory sp. (Ipomea sp. L.), pigweed sp (Amaranthus sp. L.), prickly sida (Sida spinosa L.), wild poinsettia (Euphorbia heterophylla L.), and wild radish (Raphanis raphanistrum L.). Imazethapyr provides good control of seedling johnsongrass, and fair control of crabgrass and goosegrass. Broadleaf weeds controlled by imazethapyr include: bristly starbur, common cocklebur, coffee senna, Florida pusley, morningglory sp., pigweed sp., prickly sida, wild poinsettia, and wild radish. Imazethapyr provides better wild poinsettia control than imazapic, whereas imazapic provides better control of hairy indigo than imazethapyr (Colvin and Brecke 1997). Similar to other members of the imidazolinone class of herbicides, persistence of imazapic and imazethapyr in soil is dependent upon degradation by soil microbes. Therefore, persistence is influenced by soil moisture content, soil temperature, and degree of herbicide 30

31 adsorption to soil (Moyer and Esau 1998). Abundant rainfall and warm temperatures in the southeastern United States, particularly in Florida, are conducive to substantial microbial breakdown of these herbicides. Imazapic and imazethapyr persistence increases with increasing organic matter and clay content, and as soil ph decreases below 6.0. This is due to increased herbicide adsorption to soil under these conditions, thereby protecting them from degradation via microbes (Shaner and O Connor 1991; Loux and Reese 1993). The importance of purple and yellow nutsedge control to southeastern peanut growers has been well documented (Dowler 1992). Both species reduce yield and quality by competing for light, water, mineral nutrients, and interfere with pesticide applications (Wilcut 1994a). Furthermore, tubers and rhizomes may cause problems in peanut harvesting and processing (Richburg et al. 1996; York and Wilcut 1995; Wilcut et al. 1994a). In addition, nutsedge rhizomes can pierce peanuts, thereby predisposing them to secondary infection by pathogens (Ramirez 1982). Tuber reduction is essential to any successful management system targeting these two perennial weeds due to the important role tubers play in the reproduction and dissemination of purple and yellow nutsedge in southeastern peanut fields. Little is known about the impact of imazapic and imazethapyr on purple nutsedge tuber populations and tuber germination. Tillage alone, or in conjunction with herbicide treatments, is a method growers may employ to manage purple and yellow nutsedge. No research has explored the effects a wide variety of tillage timings and frequencies may have on purple nutsedge tuber populations. Furthermore, no research has been conducted to examine the effects of repeated glyphosate applications and tillage over multiple years on purple and yellow nutsedge tuber numbers. 31