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SOYBEAN [Glycine max (L.) Merr] GROWTH AND DEVELOPMENT, WHITE
MOLD [Sclerotinia sclerotiorum (Lib.) de Bary] INCIDENCE, AND YELLOW
NUTSEDGE (Cyperus esculentus L.) CONTROL AS AFFECTED BY
GLYPHOSATE AND OTHER HERBICIDES

By

Kelly Allan Nelson

A DISSERTATION
Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of
DOCTOR OF PHEOSOPHY
Department of Crop and Soil Sciences

2000

 

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ABSTRACT
SOYBEAN [Glycine max (L.) Merr] GROWTH AND DEVELOPMENT, WHITE
MOLD [Sclerotinia sclerotiorum (Lib.) de Bary] INCmENCE, AND YELLOW
NUTSEDGE (Cyperus esculentus L.) CONTROL AS AFFECTED BY
GLYPHOSATE AND OTHER HERBICmES
By

Kelly Allan Nelson

Field and greenhouse research was conducted to determine the effect of glyphosate
and other herbicides on soybean [Glycine max (L) Merr] growth and development, the
incidence of white mold [Sclerotinia sclerotiorum (Lib.) de Bary] in glyphosate-resistant
and non-resistant soybean, and the influence of glyphosate and acetolactate synthase
(ALS)-inhibiting herbicides on yellow nutsedge control and tuber production. In the
absence of white mold, soybean canopy development was reduced, reproductive
development was delayed, and yield was reduced from 130 to 270 kg ha”1 with tank
mixture treatments of bentazon/acifluorfen plus thifensulfuron plus sethoxydim and
lactofen plus bentazon plus olethodim applied to soybean at V5. Soybean tolerance to
white mold was not related to glyphosate-resistance in the cultivars evaluated. ‘8 12-49’,
‘Sl4-M7’ Roundup Ready” (RR), ‘8 19-90’, and ‘SZO-B9’ (RR) had a lower incidence of
white mold compared to ‘GL2415’, ‘GL2600’ (RR), ‘P9281’, and ‘P93B01’(RR).
Glyphosate did not affect soybean growth and development or the incidence of white mold
in glyphosate-resistant soybean. Thifensulfiiron reduced soybean leaf area and delayed
reproductive development, but did not affect the incidence of white mold. Lactofen

reduced soybean leaf area, delayed reproductive development, increased phytoalexin

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production 2 to 26 d afier treatment, and reduced the incidence of white mold compared
to untreated soybean. However, in the presence of white mold, neither lactofen nor
thifensulfuron affected soybean yield. Other protoporphyrinogen—inhibiting herbicides
such as sulfentrazone, oxyfluorfen, and oxadiazon reduced Sclerotinia sclerotiorum lesion
growth similar to lactofen. Lactofen and sulfentrazone postemergence increased leaf
phytoalexin production, but did not influence phytoalexin production in soybean stems.
Sulfentrazone preemergence also increased soybean phytoalexin production compared to
untreated soybean. Glyphosate at 840 g ae ha" suppressed yellow nutsedge 13 to 34%
greater than glufosinate at 400 g ae ha". Additional adjuvant did not increase or decrease
yellow nutsedge control with glyphosate. When glyphosate was tank mixed with
acetolactate synthase (ALS)-inhibiting herbicides aboveground visual control increased,
but tuber yield was similar to the ALS-inhibiting herbicides applied alone. Yellow
nutsedge control by the herbicides evaluated and applied at recommended use rates was
ranked halosulfiJron and chlorimuron (> 80% control and reduction in tuber density);
imazethapyr/imazapyr, imazethapyr, and glyphosate (50 to 80% control and reduction in
tuber density); and cloransulam, rimsulfuron, and imazamox (20 to 50% control and
reduction in tuber density). However, pyrithiobac controlled yellow nutsedge 48% and

reduced tuber density 60%.

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ACKNOWLEDGMENTS

A true friend is someone who shares our hopes and dreams. I want to thank
everyone who has been involved in this research for being wonderful friends. My family
and extended family deserve a special thanks for their encouragement, enthusiasm, and
assistance. A distinctive thanks goes out to my lovely wife, Teak, for her love, confidence,
help, support, and devotion throughout the last three years. This degree would not have
been possible without our family. My parents, Lloyd and Betty Nelson, for providing me
with a life time of wonderful experiences. Ken and Barbara Nelsen for their support
throughout the process. Chris, Kathy, and Zachary Long have always been there for us no
matter what the occasion and kept us accountable for the important things in life.

A special thank you is extended to Dr. Karen Renner for her encouragement,
example as a teacher and parent, editing expertise, and for helping make this degree a
reality. I would like to thank Dr. Donald Penner for all of his help and wisdom. I will
never forget all of the late night visits. I am also grateful for the motivation and friendship
of Dr. Ray Hammerschmidt and Dr. Irvin Widders. Amy Guza, Pat O’Boyle, Mike
Partika, Rob Herron, and Gary Powell deserve a huge thanks for their technical assistance.
I would also like to thank Chad Lee, Kyle Poling, Nate Kemp, Sherry White, Jason
Fausey, Luis Velasquez, Tiffiny Murrell, Brent Tharp, Christy Sprague, Joe Simmons,
Caleb Dalley, Matt Rinella, Andy Chomas, John Boyse, Dr. Jim Kells, Kyle Fiebig, Julie
Root, and Stephanie Eickholt for their assistance and support through the various aspects

of graduate school.

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TABLE OF CONTENTS

LIST OF TABLES .................................................. ix
LIST OF FIGURES ................................................. xiii
INTRODUCTION .................................................... 1
CHAPTER 1

GLYPHOSATE-RESISTAN T AND NON-RESISTAN T SOYBEAN (Glycine max)
GROWTH AND DEVELOPMENT AS AFFECTED BY GLYPHOSATE AND

POSTEMERGENCE HERBICIDE TANK MIXTURES ..................... 3
ABSTRACT ................................................... 3
INTRODUCTION ............................................... 4
MATERIALS AND METHODS .................................... 6
RESULTS ..................................................... 9
DISCUSSION ................................................. 12
IMPLICATIONS AND CONCLUSIONS ............................ 15
LITERATURE CITED .......................................... 17

CHAPTER 2

WHITE MOLD IN SOYBEAN LITERATURE REVIEW ................... 27
INTRODUCTION AND DISEASE CYCLE OF WHITE MOLD .......... 27
EVALUATING WHITE MOLD: DISEASE SEVERITY INDEX .......... 32
EFFECTS OF WHITE MOLD ON SOYBEAN YIELD ................. 33
SOYBEAN RESPONSE TO S. SCLEROTIORW AND

PHYTOALEXIN PRODUCTION ............................ 36

METHODS TO CONTROL WHITE MOLD .......................... 39

EFFECTS OF HERBICIDES ON DISEASES AND WHITE MOLD ....... 42

LITERATURE CITED .......................................... 49
CHAPTER 3

SOYBEAN (Glycine max L) CULTIVAR AND HERBICIDE SELECTION
AFFECTS SOYBEAN DEVELOPMENT AND THE INCIDENCE OF WHITE

MOLD [Sclerotinia sclerotiorum (Lib.) de Bary] ........................... 73
ABSTRACT .................................................. 73
INTRODUCTION .............................................. 75
MATERIALS AND METHODS ................................... 79
RESULTS AND DISCUSSION ................................... 83

Soybean Response ........................................ 83
Reproductive and Canopy Development ........................ 84

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Detached Leaf and TLC Phytoalexin Bioassays ................... 87
White Mold Disease Severity ................................ 88
Soybean Yield ........................................... 89
SUMMARY .................................................. 9O
LITERATURE CITED .......................................... 93

CHAPTER 4

EFFECTS OF PROTOPORPHYRINOGEN OXH)ASE INHIBITORS ON
SOYBEAN (Glycine max L.) GROWTH RESPONSE, Sclerotinia sclerotiorum
DISEASE DEVELOPMENT, AND PHYTOALEXIN PRODUCTION BY

SOYBEAN ........................................................ 115
ABSTRACT ................................................. 115
INTRODUCTION ............................................. 116
MATERIALS AND METHODS .................................. 120

Greenhouse methods ..................................... 120
Soybean tolerance ....................................... 120
Postemergence protox-inhibiting herbicide rate screen ....... 120

Necrosis of the first trifoliate of S 19-90 and S20-B9 treated
postemergence with protox-inhibiting herbicides ..... 121

S. sclerotiorum grth on protox-inhibiting herbicide treated and

untreated leaves ................................... 121
Phytoalexin production in the leaf and stem .................... 122
Phytoalexin production with sulfentrazone PRE ................. 124
Statistical Protocol ....................................... 124
RESULTS AND DISCUSSION .................................. 125
Soybean Tolerance ....................................... 125
Postemergence protox-inhibiting herbicide rate screen ....... 125

Necrosis of the first trifoliate of S 19-90 and SZO-B9 treated
postemergence with protox-inhibiting herbicides ..... 125

S. sclerotiorum growth on protox-inhibiting herbicide treated and

untreated leaves ................................... 126
Phytoalexin production in the leaf and stem ................... 128
Phytoalexin production with sulfentrazone PRE ................. 130
LITERATURE CITED ......................................... 132

CHAPTER 5

YELLOW NUTSEDGE LITERATURE REVIEW ........................ 143
MORPHOLOGY, GROWTH, AND DEVELOPMENT ................. 144

Leaves and Shoots ....................................... 144
Basal Bulbs ............................................ 145
Rhizomes .............................................. 146
Inflorescence and Flowers ................................. 147

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Tubers ................................................ 148
Breaking Dormancy ................................ 157

Sprouting and Longevity ............................. 159
COMPETITION WITH CROPS .................................. 161
Corn .................................................. 166
Soybean ............................................... 167

Cotton ................................................ 167
YELLOW NUTSEDGE RESPONSE TO HERBICIDES AND HORMONES 168
Glyphosate ............................................. 170
ALS-inhibitors .......................................... 174

Other Herbicides ........................................ 17 9
Hormones .............................................. 182
BIOLOGICAL CONTROL ...................................... 183
LITERATURE CITED ......................................... 184

CHAPTER 6

YELLOW NUTSEDGE (Cyperus esculentus) CONTROL AND TUBER
PRODUCTION WITH GLYPHOSATE AND ALS—INHIBITING

HERBICIDES ..................................................... 205
ABSTRACT ................................................. 205
INTRODUCTION ............................................. 207
MATERIALS AND METHODS .................................. 211

Greenhouse Methods ..................................... 211
Field Methods .......................................... 213
RESULTS ................................................... 215
Greenhouse Studies ...................................... 215
Field Studies ............................................ 216
Yellow Nutsedge Growth .................................. 219
DISCUSSION ................................................ 219
LITERATURE CITED ......................................... 223

CHAPTER 7

YELLOW NUTSEDGE (Cyperus esculentus) CONTROL AND TUBER YIELD

WITH GLYPHOSATE AND GLUFOSINATE ........................... 235
ABSTRACT ................................................. 235
INTRODUCTION ............................................. 236
MATERIALS AND METHODS .................................. 239

Greenhouse Methods ..................................... 239

Effect of Spray Volume on Yellow Nutsedge Control with
Glyphosate and Glufosinate with and without AMS . . . 239
Herbicide Injection Experiment ........................ 240

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Production, and Tuber Sprout ................... 240
Dose-Response of Glyphosate and Glufosinate ............ 242
Field Methods .......................................... 242
RESULTS AND DISCUSSION .................................. 244
Effect of Spray Volume and AMS on Yellow Nutsedge Control with
Glyphosate and Glufosinate ........................... 244
Herbicide Injection Experiment .............................. 245
Effect of Glyphosate and Glufosinate on Yellow Nutsedge Control, Tuber
Production, and Sprouting ........................... 246
Effect of Additional Adjuvant with Glyphosate on Yellow Nutsedge
Control, Tuber Production, and Tuber Sprout ............. 247
Dose-Response of Glyphosate and Glufosinate .................. 248
LITERATURE CITED ......................................... 251
SUMMARY ....................................................... 261

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LIST OF TABLES

CHAPTER 1
GLYPHOSATE-RESISTANT AND NON-RESISTAN T SOYBEAN (Glycine max)
GROWTH AND DEVELOPMENT AS AFFECTED BY GLYPHOSATE AND
POSTEMERGEN CE HERBICIDE TANK MIXTURES

Table 1: Soybean injury 21 d after treatment, and redzfar red light at the soil

surface 14 and 28 d after treatment in 1997 and 1998. ................... 22

Table 2: Soybean vegetative stage 7 d after treatment, reproductive stage 20 and

80 d after treatment, and yield in 1997 and 1998. ...................... 23
CHAPTER 2

WHITE MOLD IN SOYBEAN LITERATURE REVIEW
Table 1: The effect of the environment and soybean growth characteristics on the
infection and incidence of white mold in the field (F) and greenhouse or laboratory

(G). ......................................................... 58
Table 2: Class designations for white mold disease severity calculations. ..... 61
Table 3: Phytoalexins isolated from Glycine spp. ....................... 62

Table 4: Effects of disease or stimulus on soybean phytoalexins and the associated
cultivar. ...................................................... 63

Table 5: Soybean cultivars with reported tolerance to white mold in the field or

greenhouse ..................................................... 67

Table 6: Fungicides for Sclerotinia spp. control. ....................... 68

Table 7: Effect of herbicides and adjuvants on Sclerotinia spp.. ........... 69
CHAPTER 3

SOYBEAN (Glycine max L.) CULTIVAR AND HERBICIDE SELECTION
AFFECTS SOYBEAN DEVELOPMENT AND THE INCIDENCE OF WHITE
MOLD [Sclerotinia sclerotiorum (Lib.) de Bary]
Table 1: Soybean grth characteristics, environmental conditions, and cultural
practices that influence the infection and incidence of S. sclerotiorum. ..... 100

Table 2: Soybean injury from postemergence herbicides 3 and 7 d after treatment
(DAT) in 1998 and 1999. ........................................ 101

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herbicide in 1998 and 1999 14 and 21 d after herbicide treatment (DAT) ..... 102

Table 4: Inhibited area of the fourth (treated) trifoliolate 12 d after treatment
(DAT) and the eighth trifoliolate 26 DAT for glyphosate-resistant and non-
resistant soybean cultivars averaged over herbicide treatment using a
Cladosporium cucumerinum TLC phytoalexin bioassay in 1998 and 1999. . . . 103

Table 5: White mold disease severity index and soybean yield for glyphosate-
resistant and non-resistant soybean isolines treated with postemergence herbicides
in 1998 and 1999. ............................................. 104

Table 6: Summary of the effects of thifensulfuron and lactofen on soybean
development, physiology, incidence of white mold and yield compared to untreated
soybean. .................................................... 105

CHAPTER 4
EFFECTS OF PROTOPORPHYRINOGEN OXIDASE INHIBITORS ON
SOYBEAN (Glycine max L.) GROWTH RESPONSE, Sclerotinia sclerotiorum
DISEASE DEVELOPMENT, AND PHYTOALEXIN PRODUCTION BY
SOYBEAN

Table 1: Effect of herbicides and adjuvants on Sclerotinia spp. ........... 136

Table 2: Necrosis of the first trifoliolate of S 19-90 and $20-89 treated with a
postemergence application of protox-inhibiting herbicides in the greenhouse. . 138

Table 3: Diameter of the S. sclerotiorum lesion on leaves of S 19-90, 820-89,
GL2415, GL2600, P9381, and P93B01 treated with protox-inhibiting
herbicides. ................................................... 1 3 9

Table 4: Fresh weight reduction and TLC phytoalexin bioassay of phytoalexin
production in the leaf and stem of soybean treated with lactofen and sulfentrazone
averaged over S 19-90, 820-B9, GL2415, GL2600, P9281, P93B01 cultivars. 140

Table 5: Stern and leaf weight reduction and phytoalexin production of S 19-90,
820-B9, GL2415, GL2600, P9281, and P93BOl soybean cultivars averaged over

untreated, lactofen, and sulfentrazone herbicide treatments. .............. 141

Table 6: Fresh weight, fresh weight reduction, and TLC phytoalexin bioassay for
phytoalexin production of sulfentrazone applied preemergence on soybean
cultivars ..................................................... 142

  
  
 

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YELLOW NUTSEDGE LITERATURE REVIEW
Table 1: Taxonomic key from Shippers et a1. (1995) for Cyperus esculentus
varieties based primarily on flower characteristics of plants grown in the field and
over different fertility, temerature, and day length conditions in the greenhouse. 193

Table 2: Environmental stimulus and the effect on yellow nutsedge tuber
production and sprouting. ....................................... 194

Table 3: Yellow nutsedge sprouting methods and environmental conditions for
yellow nutsedge sprouted in controlled environments. .................. 196

Table 4: Herbicide efficacy or yellow nutsedge control in the field or greenhouse
and the effect on yellow nutsedge tubers. ............................ 198

Table 5: Yellow nutsedge control and the effect on tubers with glyphosate. . . 202

Table 6: Absorption and translocation of glyphosate, ALS-inhibitors, and other
herbicides in yellow nutsedge. .................................... 204

CHAPTER 6

YELLOW NUTSEDGE (Cyperus esculentus) CONTROL AND TUBER

PRODUCTION WITH GLYPHOSATE AND ALS-INHIBITING HERBICIDES
Table 1: Yellow nutsedge visual control, height, panicle number, root fresh
weight, shoot dry weight reduction, tuber number and weight reduction in the
greenhouse 14 weeks after treatment with glyphosate plus AMS, ALS-inhibiting
herbicides, and tank mixtures of glyphosate and the ALS-inhibiting herbicides. 229

Table 2: Yellow nutsedge control 8 weeks after treatment (WAT), shoot density 8
WAT, and dry weight reduction 8 WAT, height reduction 6 WAT, and tuber
sprout 42 WAT following treatments with glyphosate plus AMS, ALS-inhibiting
herbicides, and tank mixtures of glyphosate and the ALS-inhibiting herbicides in
the field in 1997 and 1998. ....................................... 230

CHAPTER 7
YELLOW NUTSEDGE (Cyperus esculentus) CONTROL AND TUBER YIELD
WITH GLYPHOSATE AND GLUFOSINATE
Table 1: The effect of ammonium sulfate and spray volume on yellow nutsedge dry
weight reduction with glyphosate and glufosinate 28 d after treatment ...... 256

Table 2: Height and control of yellow nutsedge by glyphosate and glufosinate
injected into plants grown in the greenhouse 28 d after treatment. ......... 257

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following glyphosate and glufosinate in the greenhouse and field
in 1997 and 1998. ............................................. 258

Table 4: Yellow nutsedge control, height, dry weight, tuber density, tuber fi'esh

weight, and tuber sprouting with glyphosate plus AMS alone and with NIS, COC,
and MSO in the greenhouse and field in 1997 and 1998 .................. 259

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CHAPTER 1
GLYPHOSATE-RESISTAN T AND NON-RESISTANT SOYBEAN (Glycine max)
GROWTH AND DEVELOPMENT AS AFFECTED BY GLYPHOSATE AND
POSTEMERGENCE HERBICIDE TANK MIXTURES
Figure 1: Leaf area index for each soybean cultivar in the untreated control,
bentazon/acifluorfen at 1030 g ha" plus thifensulfuron at 2.2 g ha'l plus
sethoxydim at 240 g ha'l plus UAN and COC at 1.5% v/v, and lactofen at 105 g
ha'l plus bentazon at 1120 g ha‘l plus olethodim at 140 g ha'l plus UAN and COC
at 1.5% v/v treatments combined over 1997 and 1998. ................... 24

Figure 2: Height of each soybean cultivar in the untreated control,
bentazon/acifluorfen at 1030 g ha‘l plus thifensulfuron at 2.2 g ha’l plus
sethoxydim at 240 g ha'l plus UAN and COC at 1.5% v/v, and lactofen at 105 g
ha‘l plus bentazon at 1120 g ha‘1 plus olethodim at 140 g ha‘1 plus UAN and COC
at 1.5% v/v treatments combined over 1997 and 1998. ................... 25

Figure 3: Soybean above ground biomass of the untreated control,
bentazon/acifluorfen at 1030 g ha’l plus thifensulfuron at 2.2 g ha‘1 plus
sethoxydim at 240 g ha'l plus UAN and COC at 1.5% v/v, and lactofen at 105 g
ha'l plus bentazon at 1120 g ha‘l plus olethodim at 140 g ha“1 plus UAN and COC
at 1.5% v/v treatments combined over 1997 and 1998. ................... 26

CHAPTER 2
WHITE MOLD IN SOYBEAN LITERATURE REVIEW
Figure 1: Sclerotinia sclerotiorum (Lib.) de Bary disease cycle in soybean in the

fidd. ........................................................ 71

Figure 2: Rf-values of phytoalexins separated with thin-layer chromatography

using chloroformzacetonezNH40H (50:50: 1) (Ingham et a1. 1981). .......... 72
CHAPTER 3

SOYBEAN (Glycine max L.) CULTIVAR AND HERBICIDE SELECTION
AFFECTS SOYBEAN DEVELOPMENT AND THE INCIDENCE OF WHITE
MOLD [Sclerotinia sclerotiorum (Lib.) de Bary]
Figure 1: Soybean leaf area index for cultivar combined over herbicide treatment
in 1998 and 1999.. ............................................ 106

xiii

   

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4.5 g ha‘l plus 0.25% v/v nonionic surfactant (NIS) plus 28% urea ammonium
nitrate (UAN) at 2.3 L ha", and lactofen at 70 g ai ha'l plus 0.25% v/v nonionic
surfactant plus UAN at 2.3 L ha“1 combined over cultivar in 1998 and 1999. . 107

Figure 3: Soybean leaf area index for the untreated control, thifensulfuron at 4.5 g
ha" plus 0.25% v/v nonionic surfactant (NIS) plus 28% urea ammonium nitrate
(UAN) at 2.3 L ha" and lactofen at 70 g ha‘l plus 0.25% v/v nonionic surfactant
plus UAN at 2.3 L ha“1 combined over cultivar in 1998 and 1999. ......... 108

Figure 4: Soybean flowers plant'1 for thifensulfuron at 4.5 g ha‘l plus 0.25% v/v
nonionic surfactant plus UAN at 2.3 L ha'1 and lactofen at 70 g ai ha'l plus 0.25%
v/v nonionic surfactant plus UAN at 2.3 L ha“l combined over glyphosate-resistant
and non-resistant isolines in 1998 and 1999. .......................... 109

Figure 5: S. sclerotiorum lesion diameter of soybean cultivars using a detached
leaf bioassay of the fourth trifoliolate (treated) ....................... 110

Figure 6: S. sclerotiorum lesion diameter using a detached leaf bioassay of the
fourth trifoliolate treated with herbicides. ............................ 1 11

Figure 7: TLC phytoalexin bioassay using Cladosporium cucmerinum. ..... 112
Figure 8: Linear regression of the detached leaf bioassay of S. sclerotiorum lesion

diameter of the fourth trifoliolate (treated) averaged over harvest date (0 to 26
DAT) and disease severity index for glyphosate-resistant (black shapes) and non-

resistant (open shapes) cultivars. .................................. 113
Figure 9: Development of S. sclerotiorum in soybean at East Lansing, MI. . . 114
CHAPTER 6

YELLOW N UTSEDGE (Cyperus esculentus) CONTROL AND TUBER

PRODUCTION WITH GLYPHOSATE AND ALS-INHIBITING HERBICIDES
Figure 1: Yellow nutsedge tuber density and fresh weight following glyphosate
plus AMS, ALS-inhibiting, and tank mixtures of glyphosate and ALS-inhibiting
herbicide treatments in 1997 and 1998 ............................... 231

Figure 2: Correlation of yellow nutsedge tuber density (x) in the soil 42 weeks
after herbicide treatment at recommended rates with observed visual control (y) of
above ground plants averaged over 1997 and 1998. .................... 232
Figure 3: Correlation of yellow nutsedge tuber fresh weight (x) with tuber density
(y) averaged over 1997 and 1998. ................................. 233

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CHAPTER 7

YELLOW NUTSEDGE (Cyperus esculentus) CONTROL AND TUBER YIELD

WITH GLYPHOSATE AND GLUFOSINATE
Figure 1: Response of yellow nutsedge shoot dry weight to glyphosate (solid log-
logistic curve) and glufosinate (clotted log-logistic curve) plus ammonium sulfate
(AMS) at 20 g L'1 4 weeks after treatment ............................ 260

 

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The use of effective, broad spectrum postemergence herbicide treatments has
encouraged the adoption of reduced tillage practices and the transformation of crop
production from wide to narrow row culture. The utilization of glyphosate-resistant
soybean technology has rapidly increased in the past few years. This technology allows a
postemergence application of glyphosate, a non-selective herbicide, to soybean with no
observed phytotoxicity.

White mold, caused by the fungus [Sclerotinia sclerotiorum (Lib.) de Bary], is a
disease on the rise and is a prominent problem in Michigan. A majority of the glyphosate-
resistant soybean have been grown in narrow-rows for early canopy closure and weed
control. However, white mold is prevalent in narrow—row soybean cultures. In 1997,
white mold was wide-spread and questions regarding the susceptibility of glyphosate-
resistant soybean and the effect of glyphosate (formulated as Roundup Ultra®) on the
incidence of white mold in soybean were asked by producers, agribusiness personnel, and
extension specialists. However, other postemergence herbicides may change soybean
growth and development and reduce the incidence of white mold [Sclerotinia
sclerotiorum (Lib.) de Bary] in soybean. An interaction between herbicide treatments and
the incidence of white mold in soybean could affect production practices and weed control

re commendations. A weed control timing that targets white mold suppression could be
attractive and profitable for producers.

Glyphosate-resistant crops have provided new opportunities for cost-effective
perennial weed management for Michigan producers. However, yellow nutsedge is a

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perennial weed that is difficult to control with glyphosate applied at a rate typically used
for annual weed management. As glyphosate-resistant crops become more prevalent,
producers may select for weeds that are not completely controlled by glyphosate like
yellow nutsedge. Tank mixture treatments of glyphosate with residual ALS-inhibiting
herbicides could increase control and reduce below ground tuber production which could
affect recommendations and the management of this weed in glyphosate-resistant crops. In
addition, treatments that reduce tuber production could also help eradicate this weed
species over time or manage this weed in a glyphosate-resistant rotational crop production
system.

Field and greenhouse research was conducted to determine the effect of glyphosate
and other herbicides on soybean growth and development, the incidence of white mold in
glyphosate-resistant and non-resistant soybean, and yellow nutsedge control and tuber

production.

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CHAPTER 1
GLYPHOSATE-RESISTANT AND NON-RESISTANT SOYBEAN (Glycine max)
GROWTH AND DEVELOPMENT AS AFFECTED BY GLYPHOSATE AND
POSTEMERGENCE HERBICIDE TANK MISTURES

Abstract: Field research was conducted to evaluate the effects of glyphosate and
postemergence herbicide tank mixtures on soybean development, canopy development,
and yield of glyphosate-resistant and non-resistant cultivars. Herbicide treatments were
applied to V5 soybean. Glyphosate did not affect growth, development, or yield of
resistant soybean compared to the untreated control. Soybean injury was 14 to 18% 21 d
after treatment (DAT) from postemergence tank mixture treatments of
bentazon/acifluorfen + thifensulfuron + sethoxydim and lactofen + bentazon + olethodim.
Red:far red light that reached the soil surface 28 DAT was dependent on the soybean
cultivar and herbicide treatment. Soybean injury caused by postemergence herbicide tank
mixtures reduced vegetative development 7 DAT, reproductive development 20 and 80
DAT, height, and dry weight. The leaf area index was reduced by the postemergence tank
mixtures up to 52 DAT, and canopy development was delayed 70 and 80 DAT depending
on the cultivar. Soybean yield in plots treated with herbicides other than glyphosate was
reduced 130 to 270 kg ha'1 compared to the untreated control. Yield of soybean cultivars
varied by year with a ranking of A1900 = A2704 = AG2701 > AG1901 in 1997, and

A2704 > AG2701 > A1900 > AG1901 in 1998.

Abbreviations: AMS, ammonium sulfate ((NH4)ZSO4); ‘Asgrow 1900’, A1900; ‘Asgrow
1901’, AG1901; ‘Asgrow 2701’, AG2701; ‘Asgrow 2704’, A2704; COC, crop oil
concentrate; DAT, days after treatment; LAI, leaf area index; UAN, 28% urea ammonium
nitrate (NH4NO3).

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INTRODUCTION

Tank mixtures of postemergence herbicides were usually necessary for
broadspectrum weed control prior to the introduction of glyphosate-resistant soybean
cultivars (Fielding and Stoller 1990, Green 1991, Hart and Roskamp 1998, Monks et a1
1993). Glyphosate applications to glyphosate-resistant soybean did not injure soybean
(Lich et al. 1997, Nelson and Renner 1999) which allowed for rapid canopy closure
(Nelson and Renner 1999). Other postemergence herbicides, such as thifensulfuron,
lactofen, and acifluorfen cause chlorosis, necrosis, or stunting of soybean (Hart and
Roskamp 1998, Kapusta et al. 1986, Wichert and Talbert 1993). Such herbicide injury
may persist in weed-free conditions up to 21 d after treatment yet result in no yield loss
(Kapusta et al. 1986). However, the use of postemergence herbicides may alter the canopy
and delay canopy closure such that late germinating weeds or diflicult-to-control weeds
may escape control (Mickelson and Renner 1997, Nelson and Renner 1999).

A change in canopy development may also influence development of white mold
[Sclerotinia sclerotiorum (Lib.) deBary], a common disease problem for producers in the
north central region. White mold is common in narrow-row soybean culture (Grau and
Radkey 1984) and crop rotations that include hosts like dry edible bean (Phaseolus
vulgaris L.) (Schwartz et al. 1978). Postemergence herbicides such as lactofen may
suppress white mold in soybean (Dann et al. 1999) and affect soybean cyst nematode
reproduction (Levene et al. 1998). The interaction between herbicide treatments and
soybean canopy closure could also affect soybean production practices and weed control

recommendations. A weed control timing that would also target white mold suppression

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could be attractive and profitable.

Soybean restricts light penetration to the soil surface by forming a dense leaf
canopy near the upper portion of a mature canopy. The upper 20% of the canopy
intercepts 90% of the total photosynthetically active radiation and produces a majority of
the plant’s photosynthate, yet has only 30% of the total LAI (leaf area index) (Hatfield and
Carlson 1978, Sakamoto and Shaw 1967). Light quality may stimulate weed seed
germination or stem elongation of a crop or weed. Far-red light penetrates soybean
canopies more readily than red light. Singh et al. (1968) reported a sharp peak of
absorption with approximately 12 to 24% intensity in the infrared radiation region once
the soybean canopy had closed. Phytochrome conversion from Pr (red) to Pfr (far-red) is
necessary for the germination of some weed seeds. For instance, red light was related to
redroot pigweed seed germination (Gallagher and Cardina 1998). A low level of red light
(3 umol m‘z) stimulated buried redroot pigweed seed germination in the laboratory
(Gallagher and Cardina 1998).

Soybean cultivars vary in their competitiveness with weeds (Burnside 1972,
Bussan et al. 1997). Bussan et a1. (1997) reported that several high yielding cultivars in
weed-free conditions were also high yielding in weed-infested conditions. Soybean
canopy area and height were evaluated 30 to 45 days afier planting, but there was no
correlation between these parameters and the competitive ability of the cultivars evaluated
(Bussan et al. 1997). Predicting the outcome of an interaction between a weed and crop
species is difficult due to the numerous yield limiting factors involved in crop production

and the weed species evaluated. Soybean cultivars may also differ in their photosynthetic

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rates (Johnston et al. 1969), leaf orientation (Blad and Baker 1972), nodulation efficiency
(Hunt et al. 1990), and growth habits (Huang et al. 1993) as a result of differential light
interception with respect to the quantity and quality of light intercepted. These differences
may ultimately increase or decrease soybean yield and help the plant gain a competitive
advantage over weeds.

Several producers have expressed concerns regarding the susceptibility of
genetically modified soybean to postemergence herbicides other than glyphosate. Other
studies have reported differential sensitivity between soybean cultivars to herbicides, but
herbicide sensitivity was not linked to other herbicide resistance in the cultivar (Burnside
1972, Connelly et al. 1988, Dayan et al. 1996 and 1997, Griffin and Habetz 1989, Wax et
al. 1974). Other research has evaluated acifluorfen and bentazon applied at the V3
(Levene et al. 1998) and V6 (Browde et al. 1994) stage of development and the
interaction with nematodes. However, no research has evaluated how the soybean canopy
would be affected when a herbicide treatment for weed control and white mold
suppression was applied and the implications on late weed seed germination. The objective
of this research was to evaluate the effects of glyphosate and postemergence herbicide
tank mixtures, timed for weed control and white mold suppresion, on soybean vegetative
and reproductive development, canopy development, and yield of glyphosate-resistant and
non-resistant cultivars in a weed- and disease-free environment.

MATERIALS AND METHODS
Field research was conducted in 1997 and 1998 at the Bean and Beet Research

Farm near Saginaw, MI (43° N, 83° W). The field was fall plowed and spring field

  
   

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cultivated in 1997. The soil was a Misteguay clay (Aeric Haplaquept, fine, mixed, mesic)
with pH 7.9 and 3.7% organic matter. In 1998, the field was fall plowed and spring field
cultivated twice. The soil was a Misteguay silty clay with pH 8 and 1.9% organic matter.
The study was arranged in a split-plot design with four replications. Soybean cultivar was
the main plot and the sub-plot was herbicide treatment. ‘Asgrow 1900’ (A1900),
‘Asgrow 1901’ (AG1901 Roundup Ready"), ‘Asgrow 2701’ (AG2701 Roundup Ready”),
and ‘Asgrow 2704’ (A2704 sulfonylurea-tolerant) soybean were planted with tool-bar
mounted International 185 (International Harvester Co., Chicago, IL) planter units in 38
cm rows at 350,000 seeds ha‘l on May 23, 1997 and May 12, 1998 in plots 2.7 by 12.2 m
that were maintained weed-free.

Herbicide treatments included an untreated control, glyphosate (N-
(phosphonomethyl)glycine) (formulated as Roundup Ultra", Monsanto Co., St. Louis,
M0) at 840 g ha‘l plus ammonium sulfate (AMS) at 20 g L", bentazon/acifluorfen (3-(1-
methylethyl)-( 1H)-2, 1,3-benzothiadiazin-4(3H)-one 2,2-dioxide)/(5-[2-chloro-4-
(trifluoromethyl)phenoxy]-2-nitrobenzoic acid) (formulated as Galaxy“, BASF, Research
Triangle Park, NC) at 1030 g ha'l plus thifensulfuron (3-[[[[(4-methoxy-6-methyl-1,3,5-
triazin-2-yl)amino]carbonyl]amino]sulfonyl]-2-thiophenecarboxylic acid) at 2.2 g ha’l plus
sethoxydim (2-[1-(ethoxyimino)butyl]-5-[2-(ethylthio)propyl]-3 -hydroxy-2-cyclohexen-1 -
one) at 240 g ha'1 plus UAN (28% urea ammonium nitrate) and COC (crop oil
concentrate) (Herbimax, paraffinic oil plus emulsifiers plus surfactants, Loveland
Industries Inc, Greeley, CO) at 1.5% v/v, and lactofen ((:)-2-ethoxy-l-methyl-Z-oxoethyl

5-[2~chloro-4-(trifluoromethyl)phenoxy]-2-nitrobenzoate) at 105 g ha'l plus bentazon at

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[2-(ethylthio)propyl]-3-hydroxy-2-cyclohexen-l-one) at 140 g ha'l plus UAN and COC at
1.5% v/v. Herbicide treatments were selected that may affect leaf area and soybean height
and had limited or no soil residual to allow weed seed germination. Herbicide treatments
were applied with a tractor-mounted compressed-air plot sprayer equipped with 8003 flat-
fan tips (Spraying Systems Co., Wheaton, IL) delivering 17 8 L ha’l at 207 kPa and 6.3 km
h". Soybean were 23 cm tall and at the V5 growth stage (Fehr and Caviness 1977) at the
time of application. Air temperature was 28 and 22 C, and relative humidity was 40 and
78% in 1997 and 1998, respectively.

Visual injury was estimated 7, 21, and 28 DAT. Injury symptoms included leaf
necrosis, chlorosis, and soybean stunting which were rated on a scale of 0 (no effect) to
100% (crop death). Five photosynthetically active radiation light measurements were
recorded in each plot with a one-m SunScan Canopy Analysis System (Dynamax Inc.,
Houston TX) perpendicular to the soybean row from the time of herbicide application
until maturity to estimate soybean LAI. Incident and diffused light measurements have
been utilized as an effective non-destructive method to measure soybean LAI (Walker et
al. 1988). Three redzfar red light (Skye-Probetech, Perkasie, PA) readings were measured
between soybean rows with a single photocell 14 and 28 DAT. All light measurements
were recorded at the soil surface at approximately 1230 h. Vegetative and reproductive
stages were recorded according to Fehr and Caviness (1977) for three randomly sampled
plants in each plot. Dry weights were measured for one m of soybean row prior to

herbicide application, 35, 56, and 77 DAT according to Hunt et al. (1987). Soybean were

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harvester and moisture adjusted to 13%.

Data were subjected to analysis of variance and means separated using Fisher’s
Protected LSD (p50.05). Data were combined over years and main effects presented
where interactions were not observed.

RESULTS

Glyphosate did not injure glyphosate-resistant soybean (Table 1). Soybean treated
with glyphosate had similar vegetative development, reproductive development, leaf area
index, dry weight, height, and yield compared with the untreated control; therefore,
glyphosate data is not presented in Table 2 or Figures 1, 2, and 3.

Soybean injury was 17% from lactofen + bentazon + clethodim and 14% from
bentazon/acifluorfen + thifensulfiiron + sethoxydim 7 DAT (data not presented). By 21
DAT, soybean injury was 14 to 18% from bentazon/acifluorfen + thifensulfuron +
sethoxydim and lactofen + bentazon + clethodim (Table 1). AG1901 and AG2701 injury
fi'om bentazon/acifluorfen + thifensulfuron + sethoxydim and lactofen + bentazon +
clethodim was greater than injury to A2704. This could be due to increased tolerance of
A2704 to components of the postemergence herbicide treatments (Sebastian et al. 1989,
Simpson and Stoller 1996).

The red2far red light was greatest at the soil surface in the absence of a crop
(glyphosate treatment of non-resistant cultivars A1900 and A2704) (Table 1). By 28
DAT, the redzfar red light at the soil surface beneath the canopy of A1900, AG1901 and

A2704 treated with bentazon/acifluorfen + thifensulfuron + sethoxydim and A2704 treated

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with lactofen + bentazon + clethodim was similar to the untreated control. There was no
difference in the red:far red light at the soil surface 14 and 28 DAT in the untreated
control of each cultivar. Such effects on light quality below the canopy were found at
high plant populations where the far red:red light was greater compared to the ratio below
the canopy of low populations (Burkey and Wells 1991). The red:far red light was similar
for all cultivars treated with bentazon/acifluorfen + thifensulfuron + sethoxydim 14 and 28
DAT. An application of lactofen + bentazon + clethodim to A1900 and AG1901 may
create an environment that is favorable for weed seed germination since the red:far red
light was greater in these treatments 28 DAT compared to the untreated control.

Soybean response to herbicides, as measured by LAI, was influenced by cultivar
(Figure 1). At 40 and 52 DAT, LAI was greatest in the untreated control followed by
bentazon/acifluorfen + thifensulfiiron + sethoxydim and lactofen + bentazon + clethodim
(data not presented). Canopy development was delayed by these herbicide treatments for
A1900 and A1901 70 DAT and for A1900, AG2701, and A2704 80 DAT. Delayed
development resulted in slower maturity which had characteristically later leaf abscission.
The untreated control of A1900 and AG1901 reached a LAI equal to the peak LAI of
soybean treated with the postemergence tank mixture treatments approximately two
weeks prior to these cultivars treated with postemergence herbicide tank mixtures. The
untreated control of AG2701 and A2704 reached a LAI equal to the peak LAI of soybean
treated with the postemergence tank mixture treatments approximately one week prior to
these cultivars treated with postemergence tank mixtures. AG2701 and A2704 had a

larger LAI compared to A1900 and AG1901 from 24 DAT until harvest. In previous

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research, light interception by soybean cultivars was affected by postemergence herbicides
(Ralston and Witt 1998).

Several researchers have evaluated light interception in the soybean canopy (Board
and Harville 1992, Burkey and Wells 1991, Egli 1994, Neeser et al 1997, Wells 1991,
Wells et a1 1993). Much of the research has evaluated the utilization of narrow-row
spacings to capture more light and thus reach the highest yield potential of soybean.
However, weeds compete with soybean for light. Rapid soybean canopy development is
important for soybean to be competitive with weed species. Rapid canopy closure reduces
the reproductive potential of weed species through shading (Neeser et al. 1997, Keeley
and Thullen 1978, Santos et al. 1997).

Soybean height was AG2701 = A2704 > AG1901 > A1900 48 DAT until harvest
in the untreated control (Figure 2). Bentazon/acifluorfen + thifensulfirron + sethoxydim
and lactofen + bentazon + clethodim applied postemergence to A1900 and AG1901
caused season-long stunting compared with the untreated control. AG2701 was also
stunted until 35 DAT.

Bentazon/acifluorfen + thifensulfuron + sethoxydim and lactofen + bentazon +
clethodim reduced soybean dry weight compared to the untreated control 35 and 56 DAT
(Figure 3). By 77 DAT, there were no longer differences in dry weights between
treatments due to earlier senescence in the untreated control plots. In other research, dry
matter production was related to the intercepted radiation (Sinclair and Horie 1989,
Shibles and Weber 1966). A linear relationship between the percent light interception and

dry matter increase per day was positively correlated (Shibles and Weber 1965). Canopy

11

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development and light interception may be affected by the morphological traits of a
soybean cultivar; however, increased yield was not always reported (Huang et al. 1993,
Wells et al. 1993).

Bentazon/acifluorfen + thifensulfiiron + sethoxydim and lactofen + bentazon +
clethodim reduced the vegetative growth stage 7 DAT (Table 2), but by 14 DAT
vegetative stage was not affected by herbicide treatment (data not shown). Herbicide
treatments had limited effects on vegetative development. However, reproductive
development was delayed from an application of bentazon/acifluorfen + thifensulfuron +
sethoxydim or lactofen + bentazon + clethodim when compared to the untreated control
during the transition between full flower and early pod (20 DAT) and physiological
maturity (80 DAT).

The yield of each soybean cultivar differed each year and data is therefore
presented for each year separately. Bentazon/acifluorfen + thifensulfuron + sethoxydim
and lactofen + bentazon + clethodim reduced soybean yield from 130 to 270 kg ha‘1
compared to the untreated control in 1997 and 1998. Soybean cultivar yield was A1900 =
A2704 = AG2701 > AG1901 in 1997, and A2704 > AG2701 > A1900 > AG1901 in
1998. Rainfall in July and August totaled 9.5 inches in 1997 and 2.7 inches in 1998, and
may be the primary factor that caused lower soybean yield in 1998.

DISCUSSION

Factors that affect canopy development and light interception may also reduce

soybean yield. Light interception is influenced by the crop species, population (Shibles and

Weber 1966), fertility (Flénet and Kiniry 1995), planting date (Board and Harville 1992),

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maturity group (Board and Harville 1992), growth stage (Luxmoore et al. 1971), insects
(Board et al. 1997), row spacing and leaf orientation (Keeley and Thullen 1978, Baker and
Meyer 1966), and leaf morphology (Egli et al. 1970, Wells et al. 1993). The effect of
these factors on canopy development depends on the soybean growth stage. For instance,
soybean that were 50% defoliated at R3 recovered 20 days later for maturity group IV
soybean (Board and Harville 1993). Light interception before R5 was considered essential
for soybean yield (Board and Harville 1993). Controversy regarding light interception
during R1 to R5 or R5 to R7 and the effect on yield has been argued. Increased soybean
yield in narrow-row soybean has been related to increased light interception in the early
reproductive grth stages compared to wide-row cultures (Shibles and Weber 1965 and
1966, Board and Harville 1992, Hicks et al. 1969). However, Egli (1994) reported that
soybean yield did not rely on increased light interception in the early reproduction stages.
In our research, postemergence herbicides applied at the V5 stage of development, prior
to flowering, affected season long canopy development and reduced yield. The degree of
this effect may depend on the row-spacing, plant population, environmental conditions at
the time of application, or the cultivar selection.

Soybean height was reduced and canopy development was reduced and delayed
from bentazon/acifluorfen + thifensulfuron + sethoxydim and lactofen + bentazon +
clethodim. Shorter beans could contribute to increased light penetration thus resulting in a
smaller LAI as indicated in this research. The reduction and delay in soybean development
was more evident for early maturing soybean (A1900 and AG1901) than late group 2

cultivars (AG2701 and A2704). Delayed soybean development may increase the risk of

13

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yield loss due to frost when producers grow late group 2 soybean in the northern latitudes.
A1900 indicated quick canopy development in the untreated control which may make it
very competitive with weeds early in the season when injury is not incurred. An altered
canopy from herbicide injury also affected the red:far red light reaching the soil surface
which may affect weed seed germination. An altered canopy could also affect the canopy
microclimate and influence white mold development. Lactofen applied postemergence is
currently labeled for white mold suppression (Anonymous 1998). A change in canopy
development, in addition to the physiological effects (Dann et a1. 1999, Levene et al.
1998), as a result of postemergence herbicides may be a factor in reducing white mold in a
non-irrigated soybean culture.

Soybean treated with bentazon/acifluorfen + thifensulfuron + sethoxydim and
lactofen + bentazon + clethodim had a more branched grth habit compared to the
untreated control (personal visual observation). Soybean appeared to counteract herbicide
injury and stunting by branching, but this did not completely compensate for the injury.
The effect of light on canopy development may be observed in the branching
characteristics of the soybean plant. For example, soybean planted in wide-row spacings
branched more which resulted in increased light interception per plant (Shibles and Weber
1966). The percent main and branched reproductive dry matter in optimal and late planted
soybean depended on the cultivar (Board et al. 1990). The lower portion of the soybean
plant, below the dense cover of the soybean canopy, receives limited light. Shaded leaves
may continue to photosynthesize when supplemental light is provided (Johnston et al.

1969). An open canopy could reduce the loss of abscised pods and flowers which could

14

   

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help offset the effect of postemergence herbicides on yield.

Soybeans have an incredible ability to recover from injury caused by
postemergence herbicides. Several soybean grth characteristics are altered by a late
application of postemergence herbicides for rescue weed control or timing for weed
control and white mold suppression. A reduction in yield may occur when tank mixture
treatments of bentazon/acifluorfen + thifensulfuron + sethoxydim or lactofen + bentazon +
clethodim are applied at the V5 stage of development. However, at the benefit of white
mold suppression and reduced weed seed production may justify such treatments.

IMPLICATIONS AND CONCLUSIONS

Soybean reproductive development was delayed by postemergence herbicide
treatments, but vegetative development was not affected 7 DAT. LAI up to 52 DAT,
height, and dry weight up to 56 DAT were reduced by postemergence herbicide tank
mixture treatments and the degree of reduction was related to cultivar. The red:far-red
light was lower in the untreated control compared to the postemergence herbicide
treatments 28 DAT for certain treatments and cultivars. Soybean treated at the V5 stage
of soybean development with bentazon/acifluorfen + thifensulfuron + sethoxydim or
bentazon + lactofen + clethodim yielded less than the untreated control. Soybean that are
not injured have more rapid canopy closure which reduces light quality and quantity at the
soil surface. This is important to reduce late germinating weeds and potential weed seed
production, and to maximize soybean yield potential. Changes in the time of maximum leaf

area and canopy development may affect the microclimate in the canopy and the potential

15

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for white mold development and infection. Future research should evaluate the effects of

postemergence herbicides on canopy development and the incidence of white mold.

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LITERATURE CITED

Anonymous. 1998. Cobraop Herbicide. Supplemental Label. Valent U.S.A. Corporation,
Walnut Creek, CA. pp 7.

Baker, D. N. and R. E. Meyer. 1966. Influence of stand geometry on light interception
and net photosynthesis in cotton. Crop Sci. 6:15-18.

Bauer, T. A., K. A. Renner, and D. Penner. 1995. ‘Olathe’ pinto bean (Phaseolus
vulgaris) tolerance to postemergence imazethapyr and bentazon. Weed Sci. 43:276-282.

Blad, B. L. and D. G. Baker. 1972. Orientation and distribution of leaves within soybean
canopies. Agron. J. 64:26-29.

Board, J. E. and B. G. Harville. 1992. Explanations for greater light interception in
narrow- vs. wide-row soybean. Crop Sci. 32: 198-202.

Board, J. E. and B. G. Harville. 1993. Soybean yield component responses to a light
interception gradient during the reproductive period. Crop Sci. 33:772-777.

Board, J. E., B. G. Harville, and A. M. Saxton. 1990. Branched dry weight in relation to
yield increases in narrow-row soybean. Agron. J. 82:540-544.

Board, I. E., A. T. Wier, and D. J. Boethel. 1997. Critical light interception during seed
filling for insecticide application and optimum soybean grain yield. Agron. J. 89:369-374.

Browde, J. A., L. P. Pedigo, M. D. K. Owen, G. L. Tylka, and B. C. Levene. 1994.
Growth of soybean stressed by nematodes, herbicides, and simulated insect defoliation.
Agron. J. 86:968-974.

Burke, K. O. and R. Wells. 1991. Response of soybean photosynthesis and chloroplast
membrane fianction to canopy development and mutual shading. Plant Physiol. 97:245-

252.

Burnside, O. C. 1972. Tolerance of soybean cultivars to weed competition and herbicides.
Weed Sci. 20:294-297.

Bussan, A. J., O. C. Burnside, J. H. Orf, E. A. Ristau, and K. J. Puettmann. 1997. Field
evaluation of soybean (Glycine max) genotypes for weed competitiveness. Weed Sci.
45:31-37.

Connelly, J. A., M. D. Johnson, J. W. Gronwald, and D. L. Wyse. 1988. Bentazon
metabolism in tolerant and susceptible soybean (Glycine max) genotypes. Weed Sci.
36:41 7-423.

17

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Dann, E. K., B. W. Diers, and R. Hammerschmidt. 1999. Suppression of Sclerotinia
stem rot of soybean by lactofen herbicide treatment. Phytopathology 89:598-602.

Egli, D. B. 1994. Mechanisms responsible for soybean yield response to equidistant
planting patterns. Agron. J. 86:1046-1049.

Egli, D. B., J. W. Pendleton, and D. B. Peters. 1970. Photosynthetic rate of three

soybean communities as related to carbon dioxide levels and solar radiation. Agron. J.
62:411-414.

Fehr, W. R. and C. E. Caviness. 1977. Stages of soybean development. Spec. Rep. 80.
Coop. Ext. Service. Iowa State Univ, Ames.

Fielding, R. J. and E. W. Stoller. 1990. Effects of additives on the efficacy, uptake, and
translocation of the methyl ester of thifensulfuron. Weed Sci. 38: 172-178.

Flénet, F. and J. R. Kiniry. 1995. Efficiency of biomass accumulation by sunflower as

affected by glucose requirement of biosynthesis and leaf nitrogen content. Field Crops
Res. 44:119-127.

Gallager, R. S. and J. Cardina. 1998. Phytochrome-mediated Amaranthus germination 1:
effect of seed burial and germination temperature. Weed Sci. 46:48-52.

Grau, C. R. and V. L. Radkey. 1984. Effects of cultivars and cultural practices on
Sclerotinia stem rot of soybean. Plant Dis. 68:56-58.

Green, J. M. 1991. Maximizing herbicide efficiency with mixtures and expert systems.
Weed Technol. 5:894-897.

Griffin, J. L. and R. J. Habetz. 1989. Soybean (Glycine max) tolerance to preemergence
and postemergence herbicides. Weed Technol. 3 2459-462.

Hart, S. E. and G. K. Roskamp. 1998. Soybean (Glycine max) response to thifensulfuron
and bentazon combinations. Weed Technol. 12: 179-184.

Hatfield, J. L. and R. E. Carlson. 1978. Photosynthetically active radiation, CO2 uptake,
and stomatal diffusive resistance profiles within soybean canopies. Agron. J. 70:592-596.

Hicks, D. R., J. W. Pendleton, R. L. Bernard, and T. J. Johnston. 1969. Response of
soybean plant types to planting pattern. Agron. J. 61:290-293.

Huang, 8., D. A. Ashley, and H. R. Boerrna. 1993. Light intensity, row spacing, and
photoperiod effects on expression of brachytic stem in soybean. Crop Sci. 33:29-37.

18

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Hunt, P. G., K. P. Bumham, and T. A. Matheny. 1987. Precision and bias of various
soybean dry matter sampling techniques. Agron. J. 79:425-428.

Hunt, P. G., M. J. Kasperbauer, and T. A. Matheny. 1990. Influence of Bradyrhizobium

japonicum strain and far-red/red canopy light ratios on nodulation of soybean. Crop Sci.
30: 1306-1308.

Johnson, T. J ., J. W. Pendleton, D. B. Peters, and D. R. Hicks. 1969. Influence of

supplemental light on apparent photosynthesis, yield, and yield components of soybeans.
Crop Sci. 9:577-581.

Kapusta, G., L. A. Jackson, and D. S. Mason. 1986. Yield response of weed-free

soybeans (Glycine max) to injury from postemergence broadleaf herbicides. Weed Sci.
34:304-307.

Keeley, P. E. and R. J. Thullen. 1978. Light requirements of yellow nutsedge (C yperus
esculentus) and light interception by crops. Weed Sci. 26:10-16.

Levene, B. C., M. D. K. Owen, and G. L. Tylka. 1998. Response of soybean cyst
nematodes and soybeans (Glycine max) to herbicides. Weed Sci. 46:264-270.

Lich, J. M., K. A. Renner, and D. Penner. 1997. Interaction of glyphosate with
postemergence soybean (Glycine max) herbicides. Weed Sci. 45: 12-21.

Lycan, D. W. and S. E. Hart. 1999. Physiological response of soybean (Glycine max)
and two weed species to thifensulfuron and bentazon combinations. Weed Sci. 47: 143-
148.

Luxmoore, R. J., R. J. Millington, and H. Marcellos. 1971. Soybean canopy structure
and some radiant energy relations. Agron. J. 63:111-114.

Marvel, J. N., C. A. Beyrouty, and E. E. Gbur. 1992. Response of soybean grth to
root and canopy competition. Crop Sci. 32:797-801.

Mickelson, J. A. and K. A. Renner. 1997. Weed control using reduced rates of
postemergence herbicides in narrow and wide row soybean. J. Prod. Agric. 10:431-437.

Monks, C. D., J. W. Wilcut, and J. S. Richburg, III. 1993. Broadleaf weed control in

soybean (Glycine max) with chlorimuron plus acifluorfen or thifensulfuron mixtures.
Weed Technol. 7 :3 17-321.

Neeser, C., R. Aguero, and C. J. Swanton. 1997. Incident photosynthetically active

radiation as a basis for integrated management of purple nutsedge (Cyperus rotundus).
Weed Sci. 45:777-783.

19

  
  

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Nelson, K. A. and K. A. Renner. 1999. Cost-effective weed management in wide- and
narrow-row glyphosate resistant soybean. J. Prod. Agric. 12:460-465.

Ralston, J. L. and W. W. Witt. 1998. Comparison of postemergence broadleaf herbicides
in glyphosate tolerant and glyphosate susceptible soybeans. Abstr. Weed Sci. Soc. Am.
38:7.

Sakamoto, C. M. and R. H. Shaw. 1967. Light distribution in field soybean canopies.
Agron. J. 5927-9.

Santos, B. M., J. P. Morales-Payan, W. M. Stall, T. A. Bewick, and D. G. Shilling. 1997.
Effects of shading on the growth of nutsedges (Cyperus spp.). Weed Sci. 45:670-673.

Schwartz, H. F., J. R. Steadman, and D. P. Coyne. 1978. Influence of Phaseolus
vulgaris blossoming characteristics and canOpy structure upon reaction to sclerotinia
sclerotiorum. Phytopathology 68:465-470.

Shibles, R. M. and C. R. Weber. 1965. Leaf area, solar radiation interception, and dry
matter production by soybeans. Crop Sci. 5:575—578.

Shibles, R. M. and C. R. Weber. 1966. Interception of solar radiation and dry matter
production by various planting patterns. Crop Sci. 6:55-59.

Simpson, D. M. and E. W. Stoller. 1996. Thifensulfuron and imazethapyr interaction and
the ALS enzyme in sulfonylurea-tolerant soybean (Glycine max). Weed Sci. 44:763-768.

Sinclair, T. R. and T. Horie. 1989. Leaf nitrogen, photosynthesis, and crop radiation use
efficiency: a review. Crop Sci. 29:90-98.

Singh, M., D. B. Peters, and J. W. Pendleton. 1968. Net and spectral radiation in
soybean canopies. Agron. J. 60:542-545. .

Walker, G. K., R. E. Blackshaw, and J. Dekker. 1988. Leaf area and competition for
light between plant species using direct sunlight transmission. Weed Technol. 22159-165.

Wax, L. M., R. L. Bernard, and R. M. Hayes. 1974. Response of soybean cultivars to
bentazon, bromoxynil, chloroxuron, and 2,4-DB. Weed Sci. 22:35-41.

Wells, R. 1991. Soybean growth response to plant density: relationships among canopy
photosynthesis, leaf area, and light interception. Crop Sci. 31:755-760.

20

 

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response to differing leaf and stem morphology. Crop Sci. 33:520-524.

Wichert, R. A. and R. E. Talbert. 1993. Soybean [Glycine max (L.)] response to lactofen.
Weed Sci. 41:23-27.

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24

Figure 1. Leaf area index for each soybean cultivar in the untreated control,
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gha" plus UAN and COC at 1.5% v/v, and lactofen at 105 g ha‘1 plus bentazon at 1120 g
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Figure 2. Height of each soybean cultivar in the untreated control, bentazon/acifluorfen at
1030 g ha’l plus thifensulfiJron at 2.2 g ha'l plus sethoxydim at 240 g ha‘1 plus UAN and
COC at 1.5% v/v, and lactofen at 105 g ha'1 plus bentazon at 1120 g ha" plus clethodim at
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25

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26

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CHAPTER 2
WHITE MOLD IN SOYBEAN LITERATURE REVIEW
Introduction and Disease Cycle of White Mold. A pathogen causes the physiological
processes of the infected plant to deviate from normal. The interaction between a
pathogen, compatible host plant, and suitable environment is essential for the infection,
growth, survival, and persistence of the disease. The disease will not be expressed if there
is an absence of or variation in one of these factors. In other instances, a vector is
necessary for a disease to infect a plant, and without the vector the disease will not persist.
The interaction between the host plant and the environment are necessary to cause stress
in the plant or provide growing conditions that facilitate the infection and subsequent
colonization of the disease. The environmental conditions must be conducive for
inoculation, infection, and the development of the pathogen in the plant. Entry of a
pathogen into a plant may take place through direct penetration through the cuticle,
natural openings like stomata or abscised plant parts, vectors, or wounds. Finally, the
pathogen must be genetically compatible to infect a plant. Particular methods to control
diseases include protection, elimination, eradication, exclusion, and sanitation. A
combination of cultural, genetic, and chemical methods may effectively suppress diseases
like white mold caused by Sclerotinia sclerotiorum (Lib.) de Bary and thus reduce yield
losses in crops like soybean [Glycine max (L.) Merr].
White mold was first reported in the United States in 1946 (Chamberlain 1951)
and is common in the North Central United States (0H, IA, IN, IL, NE, ND, MI, WI, and

MN). World-wide, white mold has been reported in China, South America (Marinelli et al.

27

 

 

 

 

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1998), and Afiica (Thompson and Van der Westhuizen 1979). Boland and Hall reported in
1982 that there was an increase of white mold occurrence in soybean in Ontario. The
popular agriculture press has also reported that white mold was increasing in the United
States. An Illinois survey in 1998 reported that 45% of the fields evaluated had white
mold present (Hartman et al. 1998).

S. sclerotiorum is a soil-borne ascomycete fungus that causes wilting and
premature death of Group 0 to Group III soybean in the regions mentioned above. S.
sclerotiorum does not cause soybean seedling blight (Chamberlain 1951); however
soybean death usually occurs prior to or during pod fill. S. sclerotiorum undergoes a
simple life cycle (Figure 1 in color). The effects and importance of this disease depend
heavily on the environmental conditions, adopted cultural methods, and cultivar selection.

The mycelium and sclerotia of S. sclerotiorum over-winter in the soil or on
decaying organic matter. Sclerotia are found in fields with a history of white mold,
surrounding fence rows, orchards, in nearby fields, and around low growing plants like
dandelion (T araxacum officinale Weber in Wiggers), white clover (T rifolium repens L.)
(Abawi and Grogan 1975), or Canada thistle [Cirsium arvense (L.) Scop] (Bronsten and
Sands 1986). Sclerotia are black aggregates of tightly packed mycelium covered by a rind
for protection (Merriman 1976). Sclerotia are similar in appearance to mouse or rat feces
and may survive several years in the soil (Coley-Smith and Cooke 1971) even after
fumigation treatment (Merriman 1976). Sclerotia have contaminated soybean seed
(Hartman et al. 1998, Hoffman et al. 1998) and can be distributed to other locations on

machinery, by animals, or in the soil.

28

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Carpogenic germination of sclerotia results in the production of tan to brown
apothecia that are 1 to 7 mm in diameter and resemble a small, frail golf-tee. A single
sclerotia may produce over two apothecia in a growing season (Schwartz and Steadman
1978). The apothecia produce several million ascospores that are released from the asci
for approximately nine d (Schwartz and Steadman 1978) and are carried by air currents to
nearby plants. Ascospores germinate and form mycelium which produces oxalic acid
(Godoy et al. 1990). Oxalic acid allows the growth and spread of mycelium in soybean
tissue (Cline and Jacobsen 1983, Radke and Grau 1986). Most inoculum originates from
the apothecia since no secondary inoculum has been reported which is unlike other
important crop diseases (Boland and Hall 1982). In general, S. sclerotiorum ascospores
infect soybean through and grow on senescent tissues like flower petals which provide a
rich nutrient source and a location for easy penetration into the plant (Boland and Hall
1982, Boland and Hall 1987, Boland and Hall 19883, Cline and Jacobsen 1983). Early
research indicated the role and importance of oxalic acid in pathogenicity. Maxwell and
Lumsden (1970) demonstrated that oxalic acid was produced by S. sclerotiorum and
increased in bean (Phaseolus vulgaris L.) after infection depending on the isolate and
culture media. Later, Marciano et al. (1983) compared strong and weakly virulent S.
sclerotiorum isolates. High oxalic acid production by the strongly virulent isolate
indicated an important role in virulence. However, the role of oxalic acid as a
pathogenicity factor was confirmed with mutants of the fungus (Godoy et al. 1990).

Mutants that did not produce oxalic acid, but were capable of producing pectinase and

29

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cellulase, were not pathogenic (Godoy et al. 1990). Oxalic acid kills plant tissue and
allows the pathogen to penetrate and overcome bean tissue.

Field variability of disease incidence has been related to the number and location of
apothecia in the field and environmental conditions that affect apothecia production and
development. There has been a strong relationship between the apothecia number and the
incidence of white mold in the field (Boland and Hall 1982, Boland and Hall 1988b,
Phipps 1983). Apothecia at a New York location were produced from April 23 to June
15, and in a controlled environment apothecia formation was greatest at 11 to 15 C
(Abawi and Grogan 1975). However, dried sclerotia produced fewer apothecia (Abawi
and Grogan 1975). Ascospore germination produced by apothecia was 95% at S to 30 C,
but the fastest germination, mycelial growth, germ tube growth, greatest sclerotia
production, and best lesion development with bean was at 25 C (Abawi and Grogan
1975). These factors may help explain the variability of white mold development in the
field.

Infection by S. sclerotiorum usually occurs after the soybean canopy has closed,
during flowering (Cline and Jacobsen 1983), and when the leaf or plant surface was wet
for 40 to 112 h (Abawi and Grogan 1975, Boland and Hall 1982, 1988a). In a growth
chamber at 20 C, a leaf wetness of 70 to 120 h was adequate for white mold infection and
growth (Boland and Hall 1988a). Infection of bean took place where an old blossom was
attached, where a cast blossom laid on the leaf, or where mechanical injury to the plant
was observed (Abawi and Grogan 1975, Boland and Hall 1982). In addition, the

microclimate in the canopy was considered an important factor influencing disease

30

 

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susceptibility of bean (Blad et al. 1978). Other conditions that were favorable for
infection included fields where corn surrounded the crop or irrigation was utilized for
growing soybean (Fuller et al. 1984).

A typical sign of S. sclerotiorum disease develops on the lower portion of the
soybean stem which becomes covered with a “white mold” that is cottony in appearance.
The fiJngus girdles the stem and may infect the petioles, branches, pods, or leaves under
wet conditions. Lesions that originate from the leaf axils at nodes 3 and 4 turned from a
grey to tannish color, and develop up and down the stem (Cline and Jacobsen 1983,
Boland and Hall 1982, Grau et al. 1982). Later, sclerotia, black and cylindrical in
appearance, are produced in the white mycelium both in and on the stem. White mold
symptoms generally develop approximately 15 to 40 cm above the soil surface in July or
August during the R1 to R2 stage of development, and from R3 to R5 the disease has
made significant grth on the plant (Boland and Hall 1988a, Grau et a1. 1982, Phipps
1983). Infected soybean plants first appear grey-green and wilted, and then turn necrotic.
Disease incidence is visible by mid-August to mid-September (Boland and Hall 1988a). In
the fall, the stems appear bleached where the mycelium was present which is characteristic
of a plant infected with white mold and sclerotia pepper the surface of the stem. Table 1
provides a summary of the environmental and soybean grth characteristics that
influence the infection and incidence of white mold in the field and greenhouse.

The white mold pathogen may infect soybean seed (Hartman et al. 1998, Hoffman
et al. 1998, Thompson and Van der Westhuizen 1979, Yang et al. 1998) or over-winter as

sclerotia in the soil which may survive for several seasons. Fields with white mold may

31

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contaminate seed with sclerotia since combines may not completely separate all of the
sclerotia from the grain. Infected seed may produce mycelium from shriveled seed
(Thompson and Van der Westhuizen 1979) and in normal appearing seed (Hartman et al.
1998). Infected seed may transfer white mold to an uninfected field (Hoffman et al. 1998,
Yang et al. 1998). It is important to plant seed that is not contaminated with sclerotia,
clean machinery between fields where white mold is known to exist, and to be aware of
the spread by animals or other environmental methods.

Evaluating White Mold: Disease Severity Index. Several researchers have developed
and defined indexes to rate the disease severity of white mold in soybean. The DSI
(disease severity index) rating characterizes the extent or type of symptom exhibited by the
plant (Table 2). DSI’s have been determined during the R6 to R8 stage of soybean
development (Boland and Hall 1988a). The classes evaluated are defined by the
researcher and may vary depending on the researcher (Table 2). A scale from O to 3

where O = no disease, 1 = lesions on lateral branches only, 2= lesions on the main stem and
no effect on pod fill, and 3 = lesions on the main stem with plant death and poor pod fill
has been utilized in several field studies to determine DSI (Dann et al. 1998, Dann et al.

1999, Grau et al. 1982, Kim et al. 1999). The following formula was used to calculate the

DS1:

W

DSI = J
3 x total number of rated plants * 100

 

Several greenhouse assays have been developed to evaluate and screen soybean

cultivar sensitivity to white mold. Cline and J acobsen (1983) evaluated carrot, ascospore,

32

 

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and limited term inoculation using celery petiole sections for inoculation. Limited term
inoculation was the most successful method for identifying differences in soybean
susceptibility (Cline and Jacobsen 1983). The measurement of lesion length by an oxalic
acid treatment and soluble stern pigment levels may be additional method to evaluate
soybean tolerance to white mold (Wegulo et al. 1998). Limitations in the greenhouse
prevent researchers from evaluating and factoring in the effects of lodging, canopy
architecture, height, flowering characteristics, and maturity which may be simple
avoidance mechanisms that help reduce the incidence of white mold. Since light is less
intense in a greenhouse, soybean stems do not grow as large which could affect cultivar
sensitivity to white mold; therefore, there were limited correlations between field and
laboratory experiments (Nelson et al. 1991b). Nonetheless, Pennypacker and Risius
(1999) argued that PAR levels may have an effect on correlations between the greenhouse
and field. Care is advised when using these methods since greenhouse and laboratory
assays don’t always indicate field resistance.
Effect of White Mold on Soybean Yield. Numerous factors, like disease, affect soybean
yield. Diseases may completely destroy a crop or reduce yield to an unprofitable margin.
S. sclerotiorum causes premature soybean death which results in the production of
shriveled pods with little or no seed. Soybean cultivars may differ in white mold
tolerance, but yield losses may not be realized under low levels of infestation due to yield
compensation by nearby soybean plants.

Several researchers have determined the relationship between white mold disease

severity or incidence and soybean yield. Grau and Radkey (1984) reported that there was

33

a negative correlation between the incidence of white mold or disease severity and
soybean yield. Chun et al. (1987) reported that for every 10% increase in disease incidence
from 0 to 52%, soybean yield was reduced 7.8% (235 kg ha" or 3.5 bu A") in Michigan.
At a maximum level of disease infestation (52%), soybean yield was reduced more than
40% (Chun et al. 1987). Hoffman et al. (1998) determined that for every 10% increase in
disease incidence in group III soybean in Illinois, yield was reduced 147 to 263 kg ha“1 (2
to 4 bu A") depending on the cultivar. Similarly, Yang et al. (1999) reported a 170 to 335
kg ha" (2.5 to 5 bu A") yield loss for every 10% increase in disease incidence in Northern
Iowa (r2 = 0.59 to 0.83). Finally, for every 10% increase in disease severity, yield loss
totaled 370 kg ha" (5.5 bu A") in Michigan (Kim et al. 1999). We can conclude that yield
reductions by white mold may range from 147 to 370 kg ha" (2 to 5.5 bu A") for every
10% increase in disease severity depending on the environment and cultivar.
Narrow-row soybean usually yield more than or equal to wide-row soybean due to
greater light interception in the absence of disease (Shibles and Weber 1965 and 1966,
Board and Harville 1992, Hicks et al. 1969). Narrow-row soybean canopy closure was
also reported earlier than wide-row soybean in other research which benefits weed control
(Burnside and Colville 1964; Mickelson and Renner 1997; Wax and Pendleton 1968).
However, greater yield reductions by white mold may occur in narrow- compared to wide-
row soybean due to an environment that encourages white mold development. For
instance, soybean yield was lower in narrow-row (25 to 38 cm) compared to wide-rows
(76 cm) due to increased disease incidence in the narrow-row soybean (Grau and Radkey

1 984). Therefore, the environment in narrow-row soybean may be more conducive to the

34

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carpogenic germination, infection, and growth of S. sclerotiorum. White mold disease is
promoted by a cool, wet environment that is typical of narrow-row soybean since air
movement and sunlight penetration is limited. However, narrow-row soybean don’t
always yield less than wide-row soybean in the presence of white mold. In a study
conducted by Buzzell et al. (1993), soybean yields were greater in 23 compared to 45 cm
soybean rows and there was no difference in disease severity between 23 and 69 cm rows.
‘Essex’ and ‘Forrest’ soybean cultivars infested with white mold planted in 25 cm rows
yielded more compared to soybean planted in 76 cm rows (Phipps 1983). Similarly,
narrow-rows (25 cm) had higher yields than wide-rows (50, 75, or 100 cm) two of three
years when infested with Rhizoctonia solani (Joye et al. 1990). Soybean cultivars may
have a higher incidence of disease in narrow-rows, but a reduction in yield is not always
observed due to the yield compensation of nearby soybean plants. The effect of row
spacing may depend on the adaptability of the cultivar to a wide- or narrow-row culture.
Several studies have reported the contamination of seed by mycelium and sclerotia
as mentioned earlier, but it is important to reiterate as contaminated seed may encourage
the spread of this disease. Contaminated soybean seed may produce white mold mycelium
from shriveled seed of infected plants (Thompson and Van der Westhuizen 1979, Hartman
et al. 1998, Hoffman et al. 1998, Nicholson et al. 1978). In addition, sclerotia were
reported in 88% of the lots evaluated by Hartman et al. (1998). Seed-home white mold
incidence ranged from 0.3 to 0.7% depending on the cultivar (Hoffman et al. 1998). Seed
quality, oil, and germination were usually reduced as the incidence of disease increased

(Hofi‘man et al. 1998). White mold may not only reduce yield, but also soybean quality

35

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and could spread the disease to non-contaminated fields via contaminated seed. This is
important for producers to consider when bin-run seed is planted the following year.
Soybean Response to S. sclerotiorum and Phytoalexin Production. A plant must
detect the presence of a pathogen before it can initiate an active defense response. A plant
may undergo a hypersensitive response after it has detected the presence of a pathogen
(Baker and Orlandi 1995, Goodman and Novacky 1994, Mehdy 1994, Sutherland 1991).
This results in the rapid localized death of the host plant tissue around the infected area.
A hypersensitive response is often coordinated with the activation of other defense
mechanisms in a plant (Baker and Orlandi 1995, Goodman and Novacky 1994, Mehdy
1994, Sutherland 1991). For instance, the plant cell membranes at the infected area may
break down and release of toxic substances fiom the vacuole (Goodman and Novacky
1994, Mehdy 1994, Sutherland 1991). In addition, phenolic oxidases are produced and
utilized to synthesize quinones from phenols (Nicholson and Hammerschmidt 1992).
Phenols are precursors of lignin which is utilized to seal off an infection. In surrounding
cells, PR-proteins (pathogenesis related proteins) and phytoalexins are synthesized
(Goodman and Novaky 1994, Nicholson and Hammerschmidt 1992, Mehdy 1994).
Phytoalexins are antimicrobial compounds produced by plants after disease infection or
treatment with biotic or abiotic elicitors (Hammerschmidt 1999). The role of phytoalexins
and phenolic compounds in defense have been reviewed (Nicholson and Hammerschmidt
1992, VanEtten et al. 1989) and have been identified in numerous plant species. The
interaction between the hypersensitive response and PR-proteins, peroxidase, and

phytoalexin production helps to contain and combat an invading pathogen.

36

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Oxalic acid is produced by white mold as a pathogenicity factor (Godoy et al.
1990). After soybean was inoculated with S. sclerotiorum, browning of the epidermal
cells within 24 h resembled a hypersensitive response (Sutton and Deverall 1984). Several
phytoalexins have been identified in Glycine spp. which are good indicators of disease
expression (Table 3). Ingham (1982) reported that Glycine max produced up to twelve
phytoalexins including four isomers of glyceollin. Soybean phytoalexins were produced in
response to diseases, nematodes, herbicides, or external stimuli to whole plants, excised
plants, and cell suspensions (Table 4). Methods to isolate, purify, separate, and detect
glyceollin and other phytoalexins were reviewed by Ingham (1982). Phytoalexins have
been isolated from soybean tissue using several techniques, but the separation of
phytoalexins using thin-layer chromatography (TLC) and a chloroform:acetone:NH4OH
(50:50: 1) mobile phase was summarized in Figure 2 (Ingham et al. 1981). An antifungal
bioassay in situ with Cladosporium cucumerinum was also utilized to identify
phytoalexins on the TLC plates (Keen et al. 1971).

Glyceollin is an isoflavonoid pterocarpan phytoalexin. Soybean foliage (200 g)
may produce 5 to 10 mg of glyceollin I and 2 to 5 mg of glyceollin III (Giannini et al.
1991). Near-isogenic lines of ‘Harosoy’ (susceptible) and ‘Harosoy 63’ (resistant) have
been evaluated for glyceollin production in the hypersensitive response to Phytophthora
megasperma infection (Yoshikawa et al. 197 8) and several other studies (Table 4).
However, when other cultivars were evaluated, glyceollin was not considered the reason
for differential tolerance to Phytophthora megasperma(01ah et al. 1985). Nonetheless,

several correlative studies have reported the importance of glyceollin in the resistance of

37

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‘Harosoy 63’ to Phytophthora megasperma (Table 4). Similarly, glyceollin I accumulated
in a nematode resistant soybean cultivar 8 h after penetration while the susceptible cultivar
had minimal glyceollin production (Huang and Barker 1991). These studies have
indicated that soybean phytoalexins like glyceollin may play an important role in disease
resistance.

The mode of action of glyceollin (isomers I, II, and III) was evaluated by Kaplan et
al. (1980). Glyceollin inhibited electron transport, but did not affect oxidative
phosphorylation in isolated soybean mitochondria (Boydson et al. 1983, Kaplan et al.
1980). The inhibition of electron transport was associated with the inner membrane of the
mitochondria (Boydson et al. 1983). Glyceollin inhibited proton transport by the
plasmalemma (EDso of 50 uM) at a lower rate than the plasma membrane (EDso of 80
uM) of red beet (Beta vulgaris L.) (Giannini et al. 1988). The effects of glyceollin on
ATPase may affect the transport of solutes across the cell membrane (Giannini et al.
1988). The isomers of glyceollin were compared to rotenone (Kaplan et al. 1980), a
common insecticide and piscicide produced from the roots of the bean family, and mode of
action similarities were reported by Boydson et al. (1983).

Favaron et al. (1988) reported that glyceollin was produced as a result of
polygalacturonase (PG) activity by S. sclerotiorum. Three endogenous and one exogenous
PG were produced by S. sclerotiorum (F avaron et al. 1988). PG cleaves pectin molecules
hydrologically. Both PG II and IV have been isolated during S. sclerotiorum infection
(F avaron et al. 1993). This interaction could be significant since glyceollin inhibits white

mold ascospore germination and hyphae growth (Sutton and Deverall 1984). If there was

38

3.113 arcane
muse.<mt
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a high accumulation of glyceollin, S. sclerotiorum mycelial growth could be inhibited.
However, as much as 200 ug glyceollin ml" was required to inhibit white mold mycelium
grth in vitro (Sutton and Deverall 1984).
Methods to Control White Mold. Integrated methods to manage white mold are
currently the most practical methods to reduce the prevalence of this disease. Crop
rotation, plant population, tillage, chemicals, cultivar selection, row spacing, and effective
weed control may help reduce the incidence of white mold (Table l) and yield loss caused
by white mold. Planting white mold tolerant or resistant soybean cultivars is the
recommended strategy to reduce yield loss. In addition, avoiding cultivars with parentage
from ‘Williams’ or ‘Asgrow A3127’, which are both susceptible to white mold and had
higher DSI than cultivars that did not have these genotypes in their parentage, may help
also reduce the incidence of this disease (Kim et al. 1999). Table 5 summarizes resistant
soybean cultivars that have been reported in the literature in the field or greenhouse.
Disease escape means like canopy architecture, maturity, plant height, flowering,
row spacing, and plant population may affect the incidence of white mold for a given
cultivar in the field (Table 1). For instance, taller plants may lodge more due to excessive
vegetative growth which was correlated with an increase in the incidence of white mold
(Buzzell et al. 1993). Buzzell et al. (1993) recommended planting early maturing cultivars
that are tolerant to lodging to reduce white mold infestations. Lodging has been correlated
with disease severity (Kim et a1. 1999), but white mold can be the cause of increased
lodging rather than the result. In addition, cultural practices may help escape white mold

problems in the field. Irrigation timing is critical to reduce white mold. Soybeans that

39

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were irrigated before and after flowering had higher yields than soybean irrigated season
long (Grau and Radkey 1984). In addition, wide-row soybean may reduce or prevent
extended leaf wetness periods needed for white mold growth and development.

Several studies have evaluated canopy development in dry bean that may be useful
for soybean producers and researchers. Canopy development influenced the density of
apothecia which resulted in a larger number of apothecia produced in the row compared to
between the row (Schwartz and Steadman 1978). A dense canopy was cooler, wetter,
and had more disease than an open canopy (Blad et al. 1978). Compact bean plants
produced by TIBA (2,3,5-triiodobenzoic acid) treatment were more susceptible to S.
sclerotiorum than trellis-grown plants (Coyne et al. 1974). Flowering characteristics
(determinate vs. indeterminate) also influenced disease severity in bean (Schwartz et al.
1978). Deterrninate beans and wide- (76 cm) rows were less susceptible to white mold
than a near-isogenic indeterminate line and a narrow-row (25 cm) bean culture (Steadman
et al. 1973). Fuller et al. (1984) reported that the incidence of white mold on bean
cultivars was greatly affected by the adjacent row and the growth characteristics of the
bean in that row. Thus, avoidance mechanisms for white mold may not be effectively
evaluated when single rows are evaluated in dry edible bean cultivar comparisons. We can
conclude that such changes in the canopy development may be useful in reducing the
incidence of this disease in soybean.

A combination of tillage and crop rotation may reduce the incidence of white
mold. Abawi and Grogan (1975) reported that plowed fields had fewer sclerotia;

however, the sclerotia buried below the soil were still viable. Limited research on the

40

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effects of tillage on white mold development has been conducted or reported; however,
sclerotia generally germinate near the soil surface. S. sclerotiorum has a wide crop host
range (Table 6) and is often greater when susceptible or host species such as soybean,
lettuce (Lactuca sativa L.), alfalfa (Medicago sativa L.), red clover (T rifolium pratense
L.), celery (Apium graveolens L.), crown vetch (Corom'lla varia L.) (Marinelli et al. 1998,
Maxwell and Lumsden 1970), bean (Abawi and Grogan 1975), peanut (Arachis hypogaea
L.) (Marinelli et al. 1998, Phipps 1982), or sunflower (Helianthus annuus L.) (Kohler and
Friedt 1999, Marciano et al.1983, Marinelli et al. 1998) are grown in a crop rotation with
soybean. In a rotational year, planting a non-host crop in no-till while the use of deep
tillage to bury sclerotia prior to planting soybean may help reduce infestations. However, a
three-year crop rotation in bean had no effect on the sclerotia in the field even though
germinated apothecia were observed in sugar beet in the rotation year (Schwartz and
Steadman 1978). Some weeds such as pigweed spp. (Amaranthus spp.), common
ragweed (Ambrosia artemisiifolia L.), mustard (Brassica spp.), velvetleaf (A butilon
theophrasti Medicus), common lambsquarters (Chenopodium album L.), and Canada
thistle (Bronsten and Sands 1986) may also serve as hosts for white mold when a non-host
crop is planted in the rotation. Research has evaluated white mold as a biocontrol agent
for Canada thistle in Montana (Bronsten and Sands 1986), but the use in crop production
is limited due to the susceptibility of several crops to the disease. Nonetheless, the control
of host weeds in a rotational crop may be important to help reduce the sclerotia in the soil
for years When soybean are grown. A reduction in diseased soybean plants in the field has

reduced the number of sclerotia m‘2 (Buzzell et a1. 1993). The management of white mold

41

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sclerotia is similar to managing the weed seed bank in grain crops.

The application of fungicides for white mold control is costly and a single
application has met with limited success. The use of a fungicide like thiophanate methyl
(Topsin"")l may control white mold in soybean (Crop Protection Chemicals Reference
1997). Topsin should be applied from R1 to R2 and a subsequent application may be
made one to two weeks later (Crop Protection Chemicals Reference 1998). Similarly, an
application of benomyl (Benlate‘m)2 has been utilized in white bean to control white mold
(Natti 1971). In other crops, several fungicides have been utilized to control Sclerotinia
spp. (Table 4). Due to the cost of these products, their use has not been justified on a
large scale in soybean. In addition, several applications of the systemic induced resistance
activator 2,4—dichloroisonicotinic acid or benzothiadiazole reduced the disease severity of
white mold in susceptible (‘Williams 82’) cultivars compared to tolerant (‘5 19-90’)
cultivars (Dann et al. 1998).

Effects of Herbicides on Diseases and White Mold. A herbicide may increase,
decrease, or have no effect on a disease depending on the herbicide mode of action,
application timing, and application rate. Most of the research conducted and reported has
focused on soil-applied herbicides. In Altman and Campbell’s (1977) review of the effect

of herbicides on plant disease, only seven postemergence herbicides were reported. Since

1977, numerous postemergence herbicides have been discovered, introduced, and utilized

 

1Topsin, Elf Atochem North America, Inc., Agrichemicals Group, 2000 Market St., 21“
Floor Philadelphia, PA 19103-3222.

2Benlate, E. I. du Pont de Nemours and Company, Agricultural Products, Wilmington, DE
19898.

42

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for weed control in soybean and other agronomic crops, and the recent introduction of
herbicide-resistant crops has allowed new opportunities and options for weed control in
these crops. The effects of glyphosate on several plant fungi and the interaction between
the plant and fungus were reported and variable responses were observed (Black et al.
1996, Johal and Rahe 1984, Levesque and Rahe 1992, Rahe et al. 1990). Levesque and
Rahe (1992) recently reviewed the interaction between glyphosate and plant pathogens.
Glyphosate prevented phaseollin production by bean (Johal and Rahe 1990) and glyceollin
production by soybean at sub-lethal rates, but did not affect hypersensitive cell death
(Holliday and Keen 1982). Similarly, glyphosate inhibited the synthesis of medicarpin of
lucerne (Medicago sativa L.) when challenged with Verticillium albo-atrum (Latunde-
Dada and Lucas 1985), and enhanced mycelia growth of Alternaria cassiae on leaves of
of sicklepod (Cassia obtusifolia L.) (Sharon et al. 1993). Variable results on the
incidence of disease have been reported when glyphosate was applied to glyphosate-
resistant (Lee and Penner 1999, Penner et al. 1997, Sanoto et al. 2000) and non-resistant
soybean (Holliday and Keen 1982, Keen et al. 1982, Sanogo et al. 2000) in controlled
environments. Several studies with glyphosate involved non-glyphosate resistant soybean
cultivars and no research has reported the effects of glyphosate on glyceollin production
with glyphosate-resistant cultivars. However, no effect on S. sclerotiorum was reported in
the field or greenhouse when glyphosate was applied to ‘Asgrow 2701’ or ‘GL 2600’
soybean (Lee and Penner 1999, Penner et al. 1997).

The diphenyl ether herbicides have directly affected the incidence of white mold

(Dann et a1. 1999) and phytoalexin production by soybean (Dann et al. 1999, Komives and

43

 

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Cassida 1983, Levene et al. 1998). Acifluorfen increased phenylalanine amonia-lyase
activity in soybean which was followed by tissue necrosis in spinach (Komives and Cassida
1982). The aromatic amino acids, phenylalanine or tyrosine, are precursors of secondary
metabolites that were related to the observed phytotoxicity (Komives and Cassida 1982).
Phytotoxicity caused by the herbicide was needed to produce these secondary metabolites
in soybean (Komives and Cassida 1983). Acifluorfen at 5 ppm induced production of
glyceollin (79 ppm) and glyceofuran (16 ppm) in soybean (Komives and Cassida 1983).
The quantities of glyceollin I, II, and III in soybean seedlings following treatment with 5
ppm acifluorfen were 7, 19, and 38 pg g" fresh weight, respectively (Komives 1983). In
other research, V4 soybean treated with lactofen increased glyceollin production by 8.3 to
10.2 ug g" in ‘Williams 82’ and ‘S 19-90’ 4 DAT as the rate of lactofen increased from
0.04 to 0.11 kg ha" (Dann et al. 1999). ‘S 19-90’ had higher levels of glyceollin when
treated with lactofen compared to ‘Williams 82’ (Dann et al. 1999). An application of
lactofen to soybean not resistant to glyphosate at R1 reduced white mold disease severity
and increased yield under high infestations; however, under low infestations yield was
reduced by lactofen (Dann et al. 1999). Soybean leaves in this study were harvested 3
DAT, 4 DAT, and 6 WAT and therefore researchers could not determine if glyceollin
levels peaked and were subsequently reduced or remained high during white mold
infection. Similarly, when acifluorfen, lactofen, bentazon, COC, or NIS (X-77) were
applied at the V3 stage of development soybean cyst nematode (Heterodera glycines
Ichonohe) egg production was reduced 50 to 60% 8 weeks after application, and

acifluorfen, bentazon, COC and NIS increased glyceollin levels 1.3 to 1.9 fold in the root

44

 

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(Levene et al. 1998). F luazifop, sethoxydim, and imazethapyr did not affect egg
production, and lactofen had no effect on glyceollin levels in the root (Levene et al. 1998).

Herbicides may act directly or indirectly on the growth and infectious ability of a
disease on a plant species. F oliar blight of soybean by Rhizoctonia solani Kuhn AG—l IA
was reduced with alachlor, paraquat, and glufosinate more than with pendimethalin,
acifluorfen, or glyphosate (Black et al.1996). However, Duncan and Paxton (1981)
reported a synergistic effect between Phytophthora root rot and trifluralin on soybean.
Similarly, Millhollon and Koike (1985) reported reduced sugarcane (Saccharum
officinarum L.) yield when plants were injured by hexazinone and sugarcane mosaic virus
or ratoon stunting disease (Clavibacter xyli). The presence of a pathogen in the soil may
aid the herbicide in controlling certain plant species by predisposing the plant to injury
which allows the invasion of the disease. Lévesque and Rahe (1992) reported a ten-fold
difference in LDso values for glyphosate applied to plants grown in raw compared with
autoclaved soils. Higher rates of glyphosate were needed on soils with low fungal levels
(Levesque and Rahe 1992).

The physiological effects of herbicides on white mold are not completely
understood. The effects of several herbicides on the incidence of Sclerorim’a spp. has been
summarized (Table 7). Radke and Grau (1986) discussed the possible effects of the
herbicide mode of action of several herbicides including the triazines on the carpogenic
germination of white mold. Simazine and atrazine affected apothecia development, but

metribuzin had no effect (Radke and Grau 1986). However, cultural methods like

45

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moldboard plowing reduced Phytopthora root rot more than herbicide use (chloramben,
sethoxydim, and metribuzin) (Levesque and Rahe 1992).

Other reviews have summarized the effect of herbicides on diseases (Altman and
Cambell 1977, Katan and Eshel 1973, Levesque and Rahe 1992). Disease resistance and
herbicide resistance is desirable in crop species due to the advantages for crop protection
and improved yields. Herbicides may stimulate resistance to disease by preventing disease
infection (Uchimiya et al. 1993), or a herbicide may stimulate phytoalexin production after
the herbicide is applied (Dann et a1. 1999, Levene et al. 1998). Similarly, weeds that were
atrazine resistant biotypes of Epilobioum ciliatum and Senecio vulgaris were also resistant
to powdery mildew (Sphaerotheca epilobii [Wallr.] Sacc.) and Epilobium cilatum var.

fischeri [Blumer] Braun (Clay et al. 1991). However, herbicide resistant Lolium rigidum
also had the toxicant Clavibacter toxicus present in the grass (Riley and Gill 1994).
Linkage between herbicide resistance and disease susceptibility in released crop cultivars
could have detrimental effects on crop yield.

Postemergence herbicides, such as thifensulfuron, lactofen, and acifluorfen, cause
chlorosis, necrosis, or stunting of soybean (Hart and Roskamp 1998, Kapusta et al. 1986,
Wichert and Talbert 1993). Such herbicide injury may persist in weed-free conditions up
to 21 d afier treatment yet result in no yield loss (Kapusta et a1. 1986). However, at low
levels of white mold, lactofen reduced soybean yield when applied for white mold
suppression (Dann et al. 1999). The use of postemergence herbicides may alter the
canopy and delay canopy closure such that late germinating weeds or difficult to control

weeds may escape control (Mickelson and Renner 1997, Nelson and Renner 1999).

46

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The leaf area index is the ratio of the canopy area to the ground area and could be
used to determine the openness and the length of the disease free period available for
producers. Rapid light interception is essential to maintain yield, suppress late germinating
weeds, and quickly reach critical photosynthetic rates (Hatfield and Carlson 1978,
Sakamoto and Shaw 1967). Interactions between diseases or insects and herbicides can
reduce yields more than either factor acting alone (Browde et al. 1994). Browde et al.
(1994) reported that yield reductions were primarily attributed to a reduction in PAR
interception by soybean at R4 to R5 (Browde et al. 1994). Postemergence applications of
acifluorfen and bentazon reduced soybean LAI and yield in two of the three years
evaluated (Browde et al. 1994). Similarly, postemergence tank mixture treatments of
bentazon/acifluorfen + thifensulfuron + sethoxydim and lactofen + bentazon + clethodim
applied to V5 soybean injured soybean 14 to 21%, and reduced vegetative development 7
DAT, reproductive development 20 and 80 DAT, height, and dry weight (Nelson and
Renner 1998). The leaf area index was also reduced by these postemergence tank mixtures
up to 52 DAT, and canopy development was delayed 70 and 80 DAT depending on the
cultivar (Nelson and Renner 1998). Soybean yield with herbicides other than glyphosate
was reduced 130 to 270 kg ha" compared to the untreated control (Nelson and Renner
1998). Soybean that are not injured have more rapid canopy closure which reduces light
quality and quantity at the soil surface. This is important to reduce late germinating weeds
and potential weed seed production, and to maximize soybean yield potential. However,

changes in the time of maximum leaf area and canopy development may affect the

47

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microclimate in the soybean canopy and the potential for white mold development and

infection.

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LITERATURE CITED

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temperature and moisture on infection of beans by Whetzelinia sclerotiorum.
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Altman, J. and C. L. Campbell. 1977. Effect of herbicides on plant diseases. Annu. Rev.
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Baker, C. J. and E. W. Orlandi. 1995. Active oxygen in plant pathogenesis. Annu. Rev.
Phytopathol. 33 2299-321 .

Bhattacharyya, M. K. and E.W.B. Ward. 1986. Expression of gene-specific and age-
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Bhattacharyya, M. K. and E.W.B. Ward. 1987. Biosynthesis and metabolism of
glyceollin I in soybean hypocotyls following wounding or inoculation with Phytophthora
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Black, D. B., J. S. Russin, J. L. Griffin, and J. P. Snow. 1996. Herbicide effects on
Rhizoctonia solani in vitro and Rhizoctonia foliar blight of soybean (Glycine max). Weed
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Blad, B. L., J. R. Steadman, and A. Weiss. 1978. Canopy structure and irrigation

influence white mold disease and microclimate of dry edible beans. Phytopathology
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Boland, G. J. and R. Hall. 1982. White mold of soybeans. Can. J. Plant Pathol. 4:304.

Boland, G. J. and R. Hall. 1987. Evaluating soybean cultivars for resistance to
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Boland, G. J .and R. Hall. 1986. Growthroom evaluation of soybean cultivars for
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Boland, G. J. and R. Hall. 1988a. Epidemiology of sclerotinia stem rot of soybean in
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Boland, G. J. and R. Hall. 1988b. Relationships between the spatial pattern and number

of apothecia of Sclerotinia sclerotiorum and stem rot of soybean. Plant Pathol. 37:329-
336.

49

qudston. R.

of electron ”31

Bresten. BS 4
(made thistle

Brande. l A.
we Tatum”

J 85 ail-313

53332.16. 0 C

13:22:03. rou

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(55c. 11' L a
3377:3333 an.

.L f5»

. 3 .434

‘
‘
‘-

Boydston, R., J. D. Paxton, and D. E. Koeppe. 1983. Glyceollin: A site-specific inhibitor
of electron transport in isolated soybean mitochondria. Plant Physiol. 72: 151-155.

Brosten, BS. and D. C. Sands. 1986. Field trials of Sclerotinia sclerotiorum to control
Canada thistle (Cirsium arvense). Weed Sci. 34:377-380.

Browde, J. A., L. P. Pedigo, M. D. K. Owen, and G. L. Tylka. 1994. Soybean yield and
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Burnside, O. C. and W. L. Colville. 1964. Soybean and weed yields as affected by
irrigation, row spacing, tillage, and arniben. Weeds 12:109-112.

Buzzell, R. I., T. W. Welacky, and T. R. Anderson. 1993. Soybean cultivar reaction and
row width effect on Sclerotinia stem rot. Can. J. Plant Sci. 73 :1 169-1 175.

Casale, W. L. and L. P. Hart. 1986. Influence of four herbicides on carpogenic

germination and apothecium development of Sclerotinia sclerotiorum. Phytopathology
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Chamberlain, D. W. 1951. Sclerotinia stem rot of soybeans. Plant Dis. Rep. 35:490-491.

Chun, D., L. B. Kao, J. L. Lockwood, and T. G. Isleib. 1987. Laboratory and field
assessment of resistance in soybean to stem rot caused by Sclerotinia sclerotiorum. Plant
Dis. 71:811-815.

Clay, D. V., C. Nash, and J. A. Bailey. 1991. An association between triazine resistance

and powdery mildew resistance Epilobium ciliatum and Senecio vulgaris. Pestic. Sci.
33: 189-196.

Cline, M. N. and B. J. J acobsen. 1983. Methods for evaluating soybean cultivars for
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Coley-Smith, J. R. and R. C. Cooke. 1971. Survival and germination of fungal sclerotia.
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Coyne, D. P., J. R. Steadman, and F. N. Anderson. 1974. Effect of modified plant
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severity, and components of yield. Plant Dis. Rep. 58:379-382.

Crop Protection Chemicals Reference. 1998. 13‘“ ed. C & P Press. New York, NY. pp.
2230

50

DmE K .8
sisters ml: 2.
yields and the Re

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runes Europe

Demsee \' , l
messes in 5::
spear. 11. 533.1 .'
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2‘ ~.. '
"‘“Ci'se a:

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. 5.;e:‘ P *

ug?."'.‘.
l SC‘JK.‘~".H
!. _
M...
\u 'u?‘ k
"a‘ l 0“— .
“NLA

Dann, E. K., B. W. Diers, J. Byrunr, and R. Hammerschmidt. 1998. Effect of treating
soybean with 2,6-dichloroisonnicotinic acid (INA) and benzothiadiazole (BTH) on seed
yields and the level of disease caused by Sclerotinia sclerotiorum in field and greenhouse
studies. European J. of Plant Pathol. 104:271-278.

Dann, E. K., B. W. Diers, and R. Hammerschmidt. 1999. Suppression of Sclerotinia
stem rot of soybean by lactofen herbicide treatment. Phytopathology 892598-602.

Degousée, N., C. Triantaphylidés, and J. Montillet. 1994. Involvement of oxidative
processes in the signaling mechanisms leading to the activation of glyceollin synthesis in
soybean (Glycine max). Plant Physiol. 1042945-952.

Duncan, D. R. and J. D. Paxton. 1981. Trifluralin enhancement of phytophthora root rot
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F avaron, F., P. Alghisi, and P. Marciano. 1992. Characterization of two Sclerotinia
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Favaron, F., P. Alghisi, P. Marciano, and P. Magro. 1988. Polygalacturonase isoenzymes
and oxalic acid produced by Sclerotinia sclerotiorum in soybean hypocotyls as elicitors of
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Favaron, F., R. Peretto, P. Bonfante, and P. Alghisi. 1993. Differential absorption and
localization of two Sclerotinia sclerotiorum endo-polygalacturonases in soybean
hypocotyls. Physiol. and Mol. Plant Pathol. 43:353-364.

Fuller, P. A., J. R. Steadman, and D. P. Coyne. 1984a. Enhancement of white mold
avoidance and yield in dry beans by canopy elevation. HortScience 19:78-79.

Fuller, P. A., D. P. Coyne, J. R. Steadman, and R. F. Mumm. 1984b. Inter- and intra-row
inter-genotypic competition influences selection for avoidance of white mold disease in dry
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Giannini, J. L., D. P. Briskin, J. S. Holt, and J. D. Paxton. 1988. Inhibition of plasma
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78:1000-1003.

Giannini, J. L., J. S. Halvorson, and G. O. Spessard. 1991. High yield isolation and effect
on proton leakage of glyceollins I and III. Phytochemistry 30:3233-3236.

51

Gods}. G. J F
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Plusedus m {'51

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Godoy, G., J. R. Steadman, M. B. Dickman, and R. Dam. 1990. Use of mutants to
demonstrate the role of oxalic acid in pathogenicity of Sclerotinia sclerotiorum on
Phaseolus vulgaris. Physiol. Mol. Plant Pathol. 37: 179-191.

Goodman, R. N. and A. J. Novacky. 1994. The Hypersensitive Reaction in Plants to
Pathogens. APS Press, St. Paul, MN. pp. 244.

Grau, C. R. and V. L. Radke. 1984. Effects of cultivars and cultural practices on
Sclerotinia stem rot of soybean. Plant Dis. 68:56-58.

Grau, C. R., V. L. Radke, and F. L. Gillespie. 1982. Resistance of soybean cultivars to
Sclerotinia sclerotiorum. Plant Dis. 66:506-508.

Hart, S. E. and G. K. Roskamp. 1998. Soybean (Glycine max) response to thifensulfuron
and bentazon combinations. Weed Technol. 12:179-184.

Hartman, G. L., L. Kull, and Y. H. Huang. 1998. Occurrence of Sclerotinia sclerotiorum

in soybean fields in east-central Illinois and enumeration of inocula in soybean seed lots.
Plant Dis. 82:560-564.

Hatfield, J. L. and R. E. Carlson. 1978. Photosynthetically active radiation, CO2 uptake,
and stomatal diffusive resistance profiles within soybean canopies. Agron. J. 70:592-596.

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isoflavonoids from leaves of soybean. Phytochemistry 20:795-798.

52

 

 

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Johal, G. S. and J. E. Rahe. 1990. Role of phytoalexins in the suppression of Phaseolus
vulgaris to Colletotrichum lindemuthianum by glyphosate. Can. J. of Plant Pathol.
12:225-235.

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Biochemistry and Physiol. 18: 191-196.

53

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Kornives, T. and J. E. Casida. 1983. Acifluorfen increases the leaf content of
phytoalexins and stress metabolites in several crops. J. Agric. Food Chem. 31:751-755.

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treatment on yield of sugarcane (Saccharum officinarum). Weed Sci. 34: 137-142.

54

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Morris, P. F ., M. E. Savard, and E. W. B. Ward. 1991. Identification and accumulation
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55

 

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Phipps, P. M. and D. M. Porter. 1982. Sclerotinia blight of soybean caused by
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56

 

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to infection. Physiol. and Mol. Plant Pathol. 39:79-93.

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(Lib.) de Bary on soybean in South Africa. Phytophylactia 11:145-148.

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loss caused by Scleriotinia sclerotiorum. Plant Dis. 83:456-461.

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disease resistance in soybean: enhanced synthesis of an elicitor-releasing factor, [34,6-
endoglucanase. Physiol. and Mol. Plant Pathol. 37:367-376.

Yoshikawa, M., K. Yamauchi, and H. Masago. 1978. Glyceollin: its role in restricting

[brigal grth in resistant soybean hypocotyls infected with Phyophthroa megasperma
var. sojae. Physiol. Plant Pathol. 12:73-82.

57

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Table 1. The effect of the environment and soybean grth characteristics on the
infection and incidence of white mold in the field (F) and greenhouse or laboratory LG).

 

Characteristic and the effect on white mold infection or incidence‘I Source
Apothecia
(F) Disease severity increased as the number of apothecia increased. Boland and Hall 1988b
Fertility
(G) Increased fertility reduced lesion length. Chun et al. 1987
Flowering
(F) Lesions resulted from tissue in direct contact with dead Boland and Hall 1982
blossoms.
(F) White mold symptoms developed after flowering. Boland and Hall 198 8a
(F) White flower cultivars had a greater incidence than purple Grau et al. 1982
flower cultivars.
(F) Flower timing influenced DSI under irrigated culture. Grau and Radkey 1984
Height

(F) Correlated disease incidence to increased height over 3 years. Boland and Hall 1987
(F) Correlated disease incidence to height in 1 environment with 18 Kim et al. 1999

similar genotypes.
Injury
(G) Mechanical injury caused longer lesions to develop than lesions Chun et al. 1987
originating from leaf axil.
Inoculum
(G) Dried sclerotia reduced apothecia production. Abawi and Grogan 1975

(G) Young ascospores (5 to 8 (1 old) were more infectious than 11 d Chun et al. 1987
old ascospores.

Irrigation
(F) Irrigated soybean had a greater DSI than non-irrigated. Grau and Radkey 1984

(F) All season irrigation had greater DSI than a pre/post flower Grau and Radkey 1984
irrigation timing.

Light
(G) No effect on lesion length. Chun et al. 1987
(F) After canopy closure disease increased. Boland and Hall 1988a

(G) As PAR increased from 475 nmol ' m‘zs", soybean tolerance to Pennypacker and Risius
white mold increased. 1999

 

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Table 1 (cont ’d).

 

Lodging
(F) Correlated disease incidence to increased lodging for two years. Boland and Hall 1987

(F) Earlier maturing soybean had a lower disease incidence even Buzzell et al. 1993
under severe lodging.
(F) Increased DSI was correlated with increased lodging. Kim et al. 1999
Maturity group
(F) Correlated reduced incidence to shorter maturity. Boland and Hall 1987
(F) Earlier maturing soybean had less disease. Buzzell ct al. 1993

(F) No correlation between white mold incidence and group I, II, Chun et al. 1987
and III soybean, but group 0 had the lowest incidence of disease.

(F) Shorter maturity reduced susceptiblity. Grau et al. 1982
(F) Infrequent correlations between maturity group and DSI were Kim et al. 1999
reported.
(F) Maturity group affected disease severity and had a linear Yang et al. 1999
relationship with the incidence of disease.
Plant age
(G) Younger plants (3 week old) had larger lesions than 7 week old Chun et al. 1987
soybean.
(G) Five to six week old plants developed the largest lesions. Chun et al. 1987
Rainfall
(F) Leafwetness of 44 to 66 h was needed to insure infection. Boland and Hall 1982
(F) Leave wetness for 40 to 112 h was needed for infection. Boland and Hall 1988a
(G) A controlled environment at 20 C needed 70 to 120 h of plant Boland and Hall 1988a
wetness for infection.

(F) Soil moisture greater than -5 bars was needed for good moist Boland and Hall 1988a
conditions for white mold.

Row spacing
(F) No effect of row spacing (23 to 69 cm) on disease incidence. Buzzell et al. 1993
(F) Greater DSI in 25 to 38 cm rows than 76 cm in 2 of 3 years. Grau and Radkey 1984

(F) Some cultivars had larger differences in incidence as row width Grau and Radkey 1984
increased.

(P) Increased seeding rates had higher incidences of white mold. Phipps 1983

(F) Yield was higher in narrow- than wide-rows in the presence of Phipps 1983
white mold.

 

/

59

 

 

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Table 1 (cont ’d).

 

Temperature

(F) As the corn heat unit rating increased from 2400 to 3300 for Buzzell et al. 1993
different cultivars, there was a higher incidence of disease.

(G) Soybean incubated at 15 to 20 C had longer lesions than 25 to Chun et al. 1987
30 C.

(F) Lower air temperature was associated with increased disease Grau and Radkey 1984
incidence.

(G) Shorter lesions were observed at 15 compared to 20 and 25 C. Nelson et al. 1991a
(G) 10 to 22 C was considered an optimal temperature treatment for Pennypacker and Risius

S. sclerotiorum growth. 1999
Correlations between the Greenhouse and Field
A positive correlation between a lab assay for cultivar Chun et al. 1987
susceptibility and field susceptibility of some cultivars was
reported.
Excise stem technique had limited use. Nelson et al. 1991b

Soluble stern pigment and lesion length caused by oxalic acid Wegulo ct al. 1998
were repeatable measurements in the laboratory.

 

'Abbreviations: DSI, disease severity index; F, field; G, greenhouse or laboratory; PAR,
photosynthetically active radiation.

60

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—

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lineal If

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Table 2. Class designations for white mold disease severity calculations.

Source Class designation

 

Chun et al. 1987, Dann et al. 1998 and 1999, Grau et al. 1982, Grau and Radkey 1984,
Kim et al. 1999

0. No symptoms

1. Lesions on the lateral branches only

2. Lesions on the main stem, wilting, some pods, no effect on pod fill
3. Lesions on the main stem, plant death, poor pod fill

Boland and Hall 1988a

0. No symptoms

1. < 5 cm lesions

2. Expanding lesions on the stem or branches

3. Up to 1/2 of the branches or stem are colonized

4. > ‘/2 of the branches or stem are colonized and/or plant death

Boland and Hall 1986

0. No symptoms

Small flecks on the stem

Lesions < 2 mm in diameter

1/3 of stem encircled by lesions

2/3 of stem encircled by lesions

Complete encircling of the stem by lesions and/or plant death

Cline and Jacobsen 1983

S‘PP’NS‘

0. No symptoms

Small lesions and water soaked flowers

Water soaked petioles

Lesions on the main stem and 25% mycelium growth.
Mycelium growth on 50% of the plant

Dead plants

9:59.“?

 

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Table 3. Phytoalexins isolated from Glycine spp. (Ingham 1982).

Glycine spp.

Glycine canescens
Glycine clandestina
Glycine falcata
Glycine gracilis

Glycine latrobeana

Glycine max

Glycine soya

Glycine tabacina

Glycine tomentella

Pterocarpans

Pterocarpans
Pterocarpans

Pterocarpans

Ptercarpans

Isoflavones

Pterocarpans

Coumestans

Ptercarpans

Pterocarpans

Pterocarpans

62

Phytoalexin common name
(-)-canescacarpin
(-)-g1yceollin I
(-)-glyceollin II
(-)-clandestacarpin
(-)-glyceollin I
(-)-glyceollin II
(-)-glyceollin II
(-)-glyceollin III
(—)-glyceollin I
(-)-glyceollin II
(-)-glyceollin III
daidzein
isoformononetin
(-)-glyceocarpin
(-)-glyceofi.lran
(-)-glyceollin I
(-)-glyceollin II
(-)-glyceollin III
(-)-glyceollin IV
(-)-glycinol
(-)-9-0-methylglyceofuran
coumestrol

sojagol
(-)-glyceollin I
(-)-glyceollin III
(-)-glyceollin IV
(-)-clandestacarpin

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66

Table 5. SM
greenhouse
Cultix'ar

Ace

ism“ A25:

Clay

Corsoy
Casey 79

Dassel

E37115

Ha'din
Hoigsgn
Hofgsgn ‘8
quk

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W
Pam

EW 16‘?

Table 5. Soybean cultivars with reported tolerance to white mold in the field or

 

 

greenhouse.
Cultivar Source
Ace Boland and Hall 1987
Asgrow A2506 Kim et al. 1999
Clay Nelson et al. 1991b
Colfax Kim et al. 1999
Corsoy Chun et al. 1987, Cline and Jacobsen 1983, Grau et al. 1982, Grau
and Radkey 1984
Corsoy 79 Boland and Hall 1986, Dann et al. 1998, Kim et al. 1999, Yang et al.
1999
Dassel Wegulo et al. 1998, Yang et al. 1999
Evans Chun et al. 1987
Hardin Chun et al. 1987
Hodgson Boland and Hall 1986, Grau et al. 1982, Grau and Radkey 1984
Hodgson 78 Chun et al. 1987, Grau et al. 1982
Hawk Boland and Hall 1986
Maple Arrow Boland and Hall 1987
Maple Presto Boland and Hall 1987, Nelson et al. 1991b
McCall Boland and Hall 1987, Nelson et al. 1991b
Parker Wegulo et al. 1998

Pioneer 1677

Boland and Hall 1986

Portage Nelson et al. 1991b

S 1 9-90 Dann et al. 1998 and 1999, Kim et al. 1999, Wegulo et al. 1998,
Yang et al. 1999

Uni on Cline and Jacobsen 1983

 

67

 

 

Iablt’ 6. Full?
1098')

 

fungtldf
Beniale (DU PC

Born {60“ 3’

Elam (15K BlO
Cagec (Micro l
Pal-me (Bayer)

lloiap I’Kfiéne-l

R“? T '1 ‘ I
k l ancne-P

I‘m-L
+33! '5
,En krijf

\

T able 6. Fungicides for Sclerotinia spp. control (Crop Protection Chemicals Reference

 

 

] 9981.

Fungicide er) Application rate

Benlate (Du Pont)
Bean 24 to 64 oz A"
Brassica 32 to 96 oz A"
Carrot 4 to 48 oz A"
Dandelion 8 to 32 oz A"
Tomato 8 to 80 oz A"

Botran (Gowan)

Cucumber 1.3 lbs/A
Celery 2 to 5.3 lbs/A
Florence fennel 2 to 5.3 lbs/A
Lettuce 2 to 3.3 lbs/A
Potato 2 lbs/A
Snap bean 2 to 4 lbs/A

Bravo (ISK Biosciences)

Sweet potato

3 to 3.75 lbs/A

Peanut 2.5 pt A"
Captec (Micro Flow)
Dichondra 1 qt/ 100 gal at 1 gal 100 ft'2
F Olicure (Bayer)
Peanut 7.2 oz A"
Mocap (Rhone-Poulenc)
Peanut 2.1 lbs 1000 ft" of row
Lettuce 1.5 to 2 pt/A
Peanut 2 pt A"
Potato 2 pt A"
RO‘Vral (Rhone-Poulenc)
Bean 1.5 to 2 pt A"
Topsin (Elf Altochem)
Bean 1 to 1.5 lb A"
___ Soybean 0.75 to 1 lb A"

 

68

 

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71

a my

...‘

5‘ ’
RI”

a. Rf0.71 Methyl ester of glyceofitran.

 

b. Rf 0.56 Isoformonoetin.

HJCO

 

 

Figure 2. Rf-values of phytoalexins separated with thin-layer chromatography using
ChloroformzacetonezNH4OH (50:50:1) (Ingham et al. 1981).

72

SOY:
Al

Abstract:

L303: heme
resismt and
expefiments i
and non-resis
su'emzmmm

here of m

CHAPTER 3
SOYBEAN (Glycine max L.) CULTIVAR AND HERBICIDE SELECTION
AFFECTS SOYBEAN DEVELOPMENT AND THE INCIDENCE OF
WHITE MOLD [Sclerotinia sclerotiorum (Lib.) dc Bary]
Abstract: Postemergence herbicides can affect soybean [Glycine max (L.) Merr.]
development and the incidence of white mold [Sclerotinia sclerotiorum (Lib.) de Bary] in
glyphosate-resistant and non-resistant soybean. The effects of glyphosate, lactofen, and
thifensulfiaron on soybean canopy development, flower number plant", S. sclerotiorum
lesion diameter, phytoalexin production, white mold incidence, and yield of glyphosate-
resistant and near isogenic non-resistant soybean cultivars were evaluated in field
experiments in 1998 and 1999 at East Lansing, MI. Near isogenic glyphosate-resistant
and non-resistant cultivars had similar canopy development, flower number plant", S.
sclerotiorum lesion diameter, phytoalexin production, and white mold disease severity.
Yield of glyphosate-resistant and non-resistant soybean near isolines was similar except
GL2415 yielded 390 kg ha’1 greater than GL26OO Roundup Ready® (RR). Soybean
tolerance to S. sclerotiorum was not related to glyphosate-resistance. S 12-49, Sl4-M7
(RR), S 1990, and SZO-B9 (RR) had a lower incidence of white mold than GL2415,
GL26OO (RR), P9281, and P93B01 (RR). S 12-49, Sl4-M7 (RR), S 19-90, and SZO-B9
(RR) had peak flowering one week before apothecia were observed while GL2415,
GL2600 (RR), P9281, and P93B01 (RR) had peak flowering one week afler apothecia
were observed in the field. Glyphosate at 840 g ae ha'l plus 28% urea ammonium nitrate
(UAN) at 2.3 L ha’1 did not injure soybean, and relative humidity or temperature at the

soil surface 19 d after treatment (DAT), reproductive development, canopy development,

73

flower nun
sex-em}; or
P, 1:0

.. 0V \'

Zeszon diam

' Y I
I0 untreated
I . t

I
deseggpmen

flower number plant", S. sclerotiorum lesion size, phytoalexin production, disease
severity, or yield did not differ from untreated soybean. Lactofen at 7 0 g ai ha" plus
0.25% v/v nonionic surfactant (NIS) plus UAN at 2.3 L ha" reduced S. sclerotiorum
lesion diameter 2 to 26 DAT, increased phytoalexin production 2 to 26 DAT, and reduced
white mold disease severity by 50% compared to thifensulfuron, glyphosate, or untreated
soybean. Lactofen and thifensulfuron at 4.5 g ai ha" plus 0.25% v/v NIS plus UAN at 2.3
L ha" injured soybean 3 to 17% 7 DAT, reduced canopy development 3 to 28 DAT, and
delayed reproductive development, but neither herbicide affected soybean yield compared
to untreated soybean. The reduction in white mold incidence following an application of
lactofen may be attributed to increased phytoalexin production, reduced canopy
development, and delayed reproductive development.

Nomenclature: glyphosate, N-(phosphonomethyl)glycine; lactofen, (i)-2-ethoxy-1-
methyl-Z-oxoethyl 5-[2-chloro-4-(trifluoromethyl)phenoxy]-2-nitorbenzoate;
thifensulfuron, 3-[[[[(4-methoxy-6-methyl-1,3,5-triazin-2-
yl)amino]carbonyl]amino]sulfonyl]-2-thiophenecarboxylic acid, soybean, Glycine max (L.)
Merr.; white mold, Sclerotinia sclerotiorum (Lib.) de Bary.

Additional index words: acetolactate synthase-inhibitor, canopy development, cultivar,
diphenyl ether, disease severity, herbicide resistant, isolines, leaf area index, phytotoxicity,
postemergence, protoporphyrinogen inhibitor, Roundup Ready“, white mold.
Abbreviations: AMS, ammonium sulfate; ‘Novartis S 19-90’, ‘8 19-90’; ‘Novartis 820-
B9’, ‘SZO-B9’; ‘Novartis Sl4-M7’, ‘Sl4-M7’; ‘Novartis S 12-49’, ‘8 12-49’, ‘Great
Lakes 2415’, ‘GL2415’; ‘Great Lakes 2600’, ‘GL2600’; ‘Pioneer 9281’, ‘P9281’;
‘Pioneer 93801 ’,‘P93B01 ’; DAT, d after treatment; DSI, disease severity index; LAI, leaf

area index; NIS, nonionic surfactant; PDA, potato dextrose agar; UAN, 28% urea
ammOnium nitrate.

74

 

 

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INTRODUCTION

White mold or Sclerotinia stem rot [Sclerotinia sclerotiorum (Lib.) de Bary], a
common disease in the north central region of the United States as well as other locations
in the world, is caused by a soil—borne ascomycete firngus, S. sclerotiorum (Marinelli et al.
1998, Thompson and Van der Westhuizen 1979). A recent Illinois survey reported S.
sclerotiorum in 45% of the sampled soybean fields (Hartman et al. 1998). Symptoms of
white mold include wilting and premature death of Group 0 to Group III soybean. S.
sclerotiorum has a broad crop host range (Abawi and Grogan 1975, Kohler and Friedt
1999, Marciano et al.1983, Marinelli et al. 1998, Maxwell and Lumsden 1970) and is
more prevalent in soybean when other host species are grown as a rotational crop. Several
soybean growth characteristics, environmental conditions, and cultural practices that
influence the infection and incidence of S. sclerotiorum have been evaluated in field and
controlled environments (Table 1). These studies have shown that the management of this
disease is complicated and depends heavily on the environmental conditions of a given
year, adopted cultural methods, and cultivar selection.

S. sclerotiorum causes premature soybean death which results in the production of
shriveled soybean pods with little or no seed. Soybean cultivars differ in white mold
tolerance, but yield losses may not always occur under low levels of disease due to yield
compensation of nearby soybean plants (Hart 1998). A negative correlation between

White mold disease severity or incidence and soybean yield in Illinois (Hoffman et al.
1998), Iowa (Yang et al. 1999), Michigan (Chun et al. 1987, Kim et al. 1999), and

Wisconsin (Grau and Radkey 1984) has been reported. Yield reductions caused by S.

75

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sclerotiorum may range from 147 to 370 kg ha" (2 to 5.5 bu A") for every 10% increase
in disease severity depending on the environment and cultivar. Planting soybean cultivars
that are tolerant to white mold is strongly recommended to reduce yield loss (Boland and
Hall 1986, Boland and Hall 1987, Chun et al. 1987, Cline and Jacobsen 1983, Dann et al.
1998 and 1999, Grau et al. 1982, Grau and Radkey 1984, Kim et al. 1999, Nelson et al.
1991b, Wegulo et al. 1998, Yang et al. 1999). Avoiding cultivars with parentage from
white mold sensitive ‘Williams’ or ‘Asgrow A3127’ cultivars may reduce the incidence of
this disease (Kim et al. 1999). Limited research has evaluated S. sclerotiorum tolerance in
glyphosate-resistant soybean, and no published research has evaluated the effect of
glyphosate and other postemergence herbicides on the incidence of white mold in
glyphosate-resistant soybean.

A herbicide treatment may prevent disease infection (Uchimiya et al. 1993) or
stimulate phytoalexin production (Dann et al. 1999, Levene et al. 1998) which may help
reduce the incidence of disease. Phytoalexins are antimicrobial compounds produced by
plants afler disease infection or treatment with biotic or abiotic elicitors (Hammerschmidt
1999). Glycine spp. produce twelve known phytoalexins (Ingham 1982). The amount
and type of these compounds produced may vary depending on the cultivar, disease, or
stimulus evaluated. The production of soybean phytoalexins in response to herbicides

(Dann et al. 1999, Holliday and Keen 1982, Keen et al. 1982, KOmives and Cassida 1983,
Levene et al. 1998), other chemicals (Degouse'e et al. 1994, Favaron et al. 1988, Ingham
et al- 1981, Yoshikawa et al. 1978), or diseases (Huang and Barker 1991, Ingham et al.

1981 , Morris et al. 1991, Olah et al. 1985, Yoshikawa et al. 1978), and the effect of

76

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soybean phytoalexins on S. sclerotiorum (Sutton and Deverall 1984), Phytopthora
megasperma f. sp. glycinea (Bhattacharyya and Ward 1986, Yoshikawa et al. 197 8),
nematodes (Heterodera glycines Ichonohe) (Kaplan et al. 1980, Levene et al. 1998) have
been reported. These studies suggest that phytoalexins are an important component of
soybean disease tolerance.

The recent introduction of herbicide-resistant crops including glyphosate-resistant
soybean, has raised concerns regarding the susceptibility of glyphosate-resistant soybean
to disease when glyphosate is applied for weed control (Lee and Penner 1999, Sanogo et
al. 2000). Levesque and Rahe (1992) reviewed the interaction between glyphosate and
plant diseases. Glyphosate reduced or prevented phytoalexin production in bean
(Phaseolus vulgaris L.) (J ohal and Rahe 1990), lucerne (Medicago saliva L.) (Latunde-
Dada and Lucas 1985), soybean (Holliday and Keen 1982), and enhanced mycelia growth
of Alternaria cassiae, a biocontrol agent of sicklepod (Cassia obtusifolia L.) (Sharon et
a1. 1993). In addition, glyphosate increased the severity and fi'equency of Fusarium solani
f. sp. glycines in the roots of glyphosate-resistant and non-resistant soybean cultivars
(Sanogo et al. 2000). The research to date with glyphosate and the effects on
phytoalexins has involved non-glyphosate resistant soybean cultivars; however, no
research has reported the effects of glyphosate on phytoalexin production with glyphosate-

resistant cultivars. A linkage between herbicide resistance and increased disease
susceptibility in commercial crop cultivars could have detrimental effects on crop yield and

the acceptance of a cultivar in the market. The interaction between herbicide treatments

77

 

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and the incidence of white mold could significantly affect soybean production practices
and weed control recommendations.

Postemergence herbicides, such as thifensulfuron, lactofen, and acifluorfen, cause
chlorosis, necrosis, and/or stunting of soybean (Hart and Roskamp 1998, Kapusta et al.
1986, Wichert and Talbert 1993). Postemergence herbicides may alter the canopy and
delay canopy closure such that late germinating weeds or difficult to control weeds may
not be controlled (Mickelson and Renner 1997, Nelson and Renner 1998). Postemergence
tank mixture treatments of bentazon/acifluorfen plus thifensulfuron plus sethoxydim and
lactofen plus bentazon plus clethodim applied to V5 soybean reduced reproductive
development 20 and 80 DAT and canopy development up to 52 DAT (Nelson and Renner
1998). Changes in soybean morphology, the time of maximum leaf area, and canopy
development may affect the microclimate in the soybean canopy and the potential for S.
sclerotiorum infection and white mold development.

The objectives of this research were to determine if: 1) glyphosate-resistant
cultivars differed in reproductive development, canopy development, flowering
characteristics, phytoalexin production, white mold disease severity, and soybean yield
compared to non-resistant near isolines and 2) if postemergence herbicides including
glyphosate, lactofen, and thifensulfirron influenced soybean response, reproductive

development, canopy development, flowering characteristics, phytoalexin production,

white mold disease severity, and soybean yield compared to untreated soybean.

78

MATERIALS AND METHODS

Field research was conducted in 1998 and 1999 at the Michigan State University
Research Farm S. sclerotiorum nursery at East Lansing (42° 43' N, 84° 33' W). The soil
was a Capac sandy loam (fine-loamy, mixed mesic Aeric Ochraqualf) with 1.5% organic
matter and pH 6.5 in 1998. The 1998 site was fall chisel plowed, soil finished three times
in the spring, and fertilized with 170 kg ha" of 0-0-60 prior to planting. In 1999, the soil
was a Capac loam with pH 6.8 and 1.7% organic matter. The field was soil finished twice
in the spring and fertilized with 170 kg ha" of 6-24-24 prior to planting. Plots were
maintained weed-free throughout the season.

Research was arranged in a split-plot design with three replications. Cultivar was
the main plot and herbicide treatment was the sub-plot. Plots were 1.5 by 6.1 m and
trimmed back to 1.5 by 4.3 m prior to harvest. Near isolines of glyphosate-resistant and
non-resistant soybean, ‘S 12-49’, ‘S14-M7’ Roundup Ready" (RR), ‘8 19-90’, ‘S20—B9’
(RR), ‘GL2415’, ‘GL2600’ (RR), ‘P9281’, and ‘P93BOI’ (RR), were planted May 13,
1998 and May 11, 1999 in 38 cm rows at 543,000 seeds ha'l with a final stand of 474,000
plants ha". Herbicide treatments included an untreated control, thifensulfuron at 4.5 g ai
ha" plus 0.25% v/v nonionic surfactant’ (NIS) plus 28% urea ammonium nitrate (UAN) at

2.3 L ha", lactofen at 70 g ai ha" plus 0.25% v/v nonionic surfactant plus UAN at 2.3 L

k

 

3Nornonic surfactant was Activator-90, a mixture of alkyl polyoxyethylene ether and free
fatty acids, Loveland Industries Inc, PO. Box 1289, Greeley, CO 80632.

79

ha", and glyphosate (formulated as Roundup Ultra‘")4 at 840 g ae ha’l plus UAN at 2.3 L
ha". Glyphosate was only applied to glyphosate-resistant soybean.

Herbicide treatments were applied on June 25, 1998 and June 20, 1999 to soybean
20 to 25 cm tall, in the V5 to V6 stage of development, and prior to R1 for all cultivars
(Fehr and Caviness 1977). Treatments were applied with a C02 propelled hand-boom
calibrated to deliver 178 L ha'1 at 207 kPa, traveling 6.3 km h“, and equipped with 8003’
flat-fan nozzles spaced 51 cm apart and 48 cm above the soybean canopy. The air
temperature was 32 C with 52% relative humidity in 1998, and 28 C with 40% relative
humidity in 1999. Supplemental im'gation was provided in the evenings beginning July 5,
1998 and July 3, 1999 10 and 13 d after herbicides were applied, respectively.
Approximately 3 mm of water was provided daily during soybean flowering to encourage
white mold development.

Soybean injury from 0 to 100% (0 = no visual crop injury and 100 = complete crop
death) was evaluated 3, 7, 14, and 28 DAT based on the combined visual effects of the
herbicides on necrosis, chlorosis, and stunting. Relative humidity and temperature at the
soil surface were measured 19 DAT in all plots at solar noon since the greatest differences
in microclimate conditions were reported at this time of the day in other research (Blad et
al. 197 8). Flower number of three randomly selected plants in each plot was recorded and

averaged.

 

‘Roundup Ultra, Monsanto Co., 800 N. Lindbergh Blvd, St. Louis, MO 63167.

’Teejet flat-fan tips. Spraying Systems Co., North Ave. and Schmale Road, Wheaton, IL
60188.

80

 

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Light measurements were recorded at 7 to 14 (1 intervals from the time of herbicide
application until 85 DAT with a SunScan Canopy Analysis System? Five measurements
were recorded with a one-m light measurement device diagonal to the soybean row.
Incident and diffused light measurements have been utilized as an effective non-destructive
method to measure soybean LAI (Walker et al. 198 8).

Trifoliolate leaves located at the fifth node (fourth trifoliolate) were harvested from
three separate herbicide treated plants in each plot 0, 2, 4, 7, 12, and 26 DAT except
thifensulfuron applied to S 12-49, S14-M7, GL2415, GL2600, P9281, and P93B01
cutivars. In addition, untreated trifoliolates located at the ninth node (eighth trifoliolate)
were harvested 26 DAT from three separate plants in each plot except thifensulfiiron
applied to S 12-49, S1 4-M7, GL2415, GL2600, P9281, and P93B01 cutivars. One leaf
from each trifoliolate was evaluated in a detached leaf bioassay, and the other leaves were
stored at -20 C and evaluated for antifiingal compounds using a TLC phytoalexin
bioassay. Excised leaves used in the detached leaf bioassay were placed in a 150 by 15
mm petri dish7 with moistened filter paper to ensure a humid environment for S.
sclerotiorum growth. A potato dextrose agar (PDA) plug, 2.4 mm in diameter, from the
margin of S. sclerotiorum was placed on the leaves and the diameter of growth was
measured with an electronic digital caliper after incubating 48 h. The TLC phytoalexin
bioassay utilized Cladosporium cucumerinum to determine and quantify antifungal

compounds produced by soybean (Keen et al. 1971). This bioassay was used to evaluate

 

GDynamax Inc., 10808 Fallstone #350, Houston TX 77099.
7VWR Scientific Products, 800 E. Fabyan Pkwy, Batavia, IL 60510.

81

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the relative total antifimgal activity (phytoalexin production) between treatments.
Soybean leaves were weighed, cut into 0.5 to 1.0 cm pieces, and extracted with 40:60
ethanolzwater v/v for 4 h. The extract was evaporated8 at 40 C under reduced pressure to
approximately 1 ml and transferred to a 10 ml test tube. The flask was then rinsed with 1
ml water. The extract and water rinsate was combined and extracted three times with 1 ml
ethyl acetate. The ethyl acetate extract was dried with anhydrous magnesium sulfate and
then evaporated under a flow of N2. The residue was resuspended in ethyl acetate at 50 ul
g'l leaf fresh weight and applied to a silica-gel (250 um) thin-layer chromatography plate9
at 50 pl in 1998 and 30 ill in 1999. The plate was developed in an
acetone:chloroform:ammonia hydroxide solution at 50501 v/v. C. cucumerinum spores
were suspended in a 100 ml solution of 0.7 g KHZPO4, 0.4 g KNO3, 0.3 g Nazl-IPO4, 0.1 g
MgSO4, 0.1 g NaCl, and 5.0 g D-glucose that was adjusted to pH 6.5. A fine spray of
suspended spores was applied to the TLC plate and the plate was placed in a sealed
chamber with 100% humidity. Plates were incubated for 2 to 3 d and stored at -20 C.
Photo copies10 of the plates were made and clear areas with no C. cucumerinum growth
were quantified with a leaf area meter“. Total areas were calculated that corresponded to
Rfvalues for the methyl ester of glyceofuran (Rf 0.71); isoformonoetin and a mixture of

glyceollin I, II, and III (Rf 0.5 to 0.56); and glyceofuran and the precursor of glyceollin II

 

8Rotavapor, Buchi, Labortechnik AG, Postfach, CH-9230, Flawil.

9Analtech, Inc., P. O. Box 7558, 75 Blue Hen Dr., Newark, DE 19713.
loCannon, imageRUNNER 330S, One Cannon Plaza, Lake Success, NY 11042.
llLI-3000, LI-COR, 4421 Superior Street, Lincoln, NE 68504.

82

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and III (Rf 0.25) previously reported using chloroform:acetone:NH4OH (50:50:] v/v) as
the mobile phase and a silica-gel TLC plate as the stationary phase (Ingham et al. 1981,
Keen 1982 ).

Near physiological maturity, thirty plants per plot were evaluated for the incidence
of white mold. The disease severity index was calculated according to the scale (0 = no
symptoms, 1 = lesions on the lateral branches only, 2 = lesions on the main stem but no
effect on pod fill, and 3 = lesions on the main stem and pod fill was reduced) described by
Grau and Radkey (1984). Soybean were harvested with a Massey 10'2 small plot
harvester and moisture adjusted to 13%. An analysis of variance was conducted and
percent data for visual injury were transformed to the arcsine prior to the analysis. The
transformation did not affect the conclusions so original data is presented. Data were
combined over years and main effects presented where interactions were not observed.
TLC phytoalexin bioassay data were subjected to an F Max test for homogeneity (Kuehl
1994) and data were combined over location since variances for both locations were
homogenous. Means were separated using Fisher’s Protected LSD at p50.05.

RESULTS AND DISCUSSION
Soybean Response. Soybean response to herbicide treatments differed by cultivar (Table
2), and was characteristic of the herbicide family (Gunsolus and Curran 1992).
Glyphosate did not injure glyphosate-resistant soybean while thifensulfuron injury to

soybean was minimal (Table 2). Lactofen caused 8 to 12% injury 3 DAT and injury

 

”Kincaid Equipment Manufacturing, PO. Box 400, Haven, KS 47543.

83

 

increased to 13 to 17% 7 DAT. Injury was no longer visually evident 35 DAT from any
herbicide treatment (data not presented).

Reproductive and Canopy Development. Reproductive development of soybean is
classified according to flowering, pod development, seed development, and maturity (Fehr
and Caviness 1977). Soybean in R1 are in beginning flower, R2 are in fiill flower, and R3
are in beginning pod. Flowering has almost completely ceased by R4 (fiill pod), and most
of the flowers at this stage are concentrated near the top of the plant. The reproductive
stage of soybean 14 and 21 DAT (Table 3) as well as canopy development (Figure 1)
varied by cultivar and year. In 1998, all cultivars were at the same reproductive stage
except P93BOl 14 DAT, and all cultivars were at full bloom with some of the late Group I
or early Group II soybean producing pods by 21 DAT (Table 3). In 1999, the late Group
I and early Group II soybean were at fiill flower and P93B01 had barely begun flowering
14 DAT. One week later, S 12-49 started producing pods while the other cultivars were
primarily in fiill bloom except P93B01. Drier growing conditions two weeks prior and one
week following herbicide application (5.4 cm in 1998 and 2.3 cm in 1999) coupled with
cooler (6 to 7 C) minimum and maximum average temperatures in the week prior to
herbicide application probably delayed reproductive development in 1999 compared to
1998. Soybean cultivar LAI was similar between glyphosate-resistant and non-resistant
near isolines for all measurement dates except S 12-49 and Sl4-M7, and GL2415 and
GL2600 21 DAT in 1998 (Figure 1). In 1999, LAI was only different between P9281 and

P93B01 isolines 7 DAT. S 12-49 had one of the lowest LAI throughout the season in

84

1998. Peak canopy development for all of the cultivars was 35 DAT in 1998 and 35 to 48
DAT in 1999.

Glyphosate did not affect reproductive development when compared to the
untreated control (data not presented). However, thifensulfirron and lactofen delayed
soybean reproductive stage, flowering, pod development 3 to 35 DAT (Figure 2), and
reduced canopy development 3 to 28 DAT (Figure 3) in 1998 and 1999. These results are
similar to research conducted with postemergence tank mixtures applied to V5 soybean
(Nelson and Renner 1998). Canopy development was reduced by thifensulfuron and
lactofen more in 1999 than 1998 compared to the untreated control. These differences
were probably due to rainfall prior to and following herbicide application. Yearly
differences in herbicide effects on canopy ground cover ratings have been attributed to
rainfall differences following herbicide application (Donald 1998). In 1999, only 1.8 cm of
rain fell from 0 to 7 DAT which decreased LAI of treated soybean compared to the
untreated control. Irrigation was not initiated until 10 and 13 d after herbicides were

applied in 1998 and 1999, respectively (Figure 3).

Soybean Flowering. Postemergence herbicide treatments influenced flower number
plant’1 and peak flowering for all cultivars (Figure 4). Interactions were expected and
observed due to the maturity differences of the cultivars evaluated; therefore, data were
analyzed for the paired isolines separately. Flower number plant’1 in glyphosate-resistant
cu ltivars was no different than non-resistant soybean (data not presented). Peak flowering
was 14 DAT for S 12—49, S14-M7, S 19-90, and S20-B9 and 28 DAT for GL2415,

GL2600, P9281, and P93BOl. Thifensulfirron and lactofen reduced S 12-49, S14-M7, S

85

 

19-90, and S20-B9 flower number plant" 7 and 14 DAT, and reduced GL2415 and
GL2600 flower number plant" 14 DAT. Conversely, P9281 and P93BOl treated with
lactofen had more flowers plant" than the untreated control 28 DAT.

Soybean flowering is primarily controlled by photoperiod (J effers 1987). Lactofen
stressed soybean such that reproductive development was delayed, supporting previous
research (Nelson and Renner 1998). Lactofen reduced the flower number of Group I
soybeans (S 12-49, S14-M7, S 19-90, and 820-B9); had no effect on mid-Group II
soybean flower number (GL2415 and GL2600); and increased late Group H flower
number (P9281 and P93BOl) compared to untreated soybean. Thus, lactofen’s affect on
soybean flowering may increase when applied at or near the time of flowering since the
reproductive stage of the cultivars 14 and 21 DAT indicated some of the Group II
soybean were beginning to flower or were at full flower at this point in time (Table 3).

The first appearance of S. sclerotiorum apothecia under the soybean canopy was
observed 21 DAT when the mid- and late-Group II soybean were in firll bloom and had
peak flowering one week later. The relative humidity in the irrigated soybean canopy at
the soil surface was 73 to 76% at 1230 h 19 DAT and did not differ between cultivars,
herbicides, or years (data not presented). However, air temperature at the soil surface 19
DAT was slightly higher in the thifensulfuron (26.2 C) and lactofen (26.1 C) treatments
compared to the untreated control (25.6 C) and glyphosate (25.6 C) treatments.
Environmental conditions can affect apothecia production, ascospore germination, and
mycelial growth (Abawi and Grogan 1975). A dense canopy with a cool and wet

microclimate had a greater incidence of disease than an open, warm and dry canopy in

86

bean (Blad et al. 1978). Therefore, soybean injury caused by thifensulfuron and lactofen
that delayed reproductive development, reduced canopy development, and influenced
flower numbers plant" could influence the severity of white mold in glyphosate-resistant
and non-resistant cultivars.

Detached Leaf and TLC Phytoalexin Bioassays. No recurrent differences between
glyphosate-resistant and non-resistant isolines were observed using the S. sclerotiorum
detached leaf bioassay (data not presented). Therefore, only data for the non-glyphosate
resistant soybean cultivars are presented to demonstrate the validity of the bioassay and
differences between cultivars through the sample period (Figure 5). S. sclerotiorum lesion
diameter was greater for GL2415 and P9281 compared to 8 12-49 and S 19-90 prior to
herbicide treatments. This cultivar difference continued fi'om 2 to 12 DAT. S.
sclerotiorum lesion diameter was reduced by lactofen from 2 to 26 DAT compared to
thifensulfuron (data not shown), the untreated control, or glyphosate treatments (Figure
6). The effect of thifensulfiiron on lesion diameter was only evaluated on S 19-90 and
S20-B9. None of the herbicide treatments affected the S. sclerotiorum lesion diameter of
the eighth trifoliolate (untreated leaf) 26 DAT (data not shown).

The TLC phytoalexin bioassay indicated the presence of antifungal compounds
that corresponded to Rf values previously reported for the methyl ester of glyceofiiran (Rf
0.71); isoformonoetin and a mixture of glyceollin I, II, and III (Rf 0.5 to 0.56); and
glyceofuran and the precursor of glyceollin II and ID (Rf 0.25) (Ingham et al. 1981, Keen
1982 ). Soybean phytoalexin production did not differ due to cultivar from 0 to 7 DAT

(data not presented). However, S 12-49 inhibited C. cucumerinum growth more than

87

S14-M7 while P9281 inhibited C. cucumerinum grth more than P93B01 12 DAT
(Table 4). There was no difference in total antifungal compound production between
cultivars for the fourth trifoliolate (herbicide treated) 26 DAT (data not presented).
However, S 12-49 and S20-B9 produced more phytoalexins in the eighth trifoliolate
(untreated leaves) than GL2415, GL2600, P9281, or P93B01 (Table 4).

Extracts from lactofen-treated plants inhibited C. cucumerinum growth more than
the untreated control 2 to 26 DAT thus indicating increased phytoalexin production with
this treatment (Figure 7). Glyphosate and thifensulfiiron did not affect phytoalexin
production compared to the untreated control. The diphenyl ether herbicides like lactofen
inhibit protoporphyrinogen oxidase which causes the accumulation of protoporphryin IX
(Scalla and Matringe 1994). The exposure to light causes the peroxidative destruction of
membrane fatty acids and subsequent cell death (Scalla and Matringe 1994). Similar
observations have been reported when plants undergo a hypersensitive response to disease
infection (Baker and Orlandi 1995, Bhattacharyya and Ward 1986, Mehdy 1994, Sutton
and Deverall 1984, Yoshikawa et al. 1978) or abiotic factors (Degousée et al. 1994,
Ingham et al.1981) which have resulted in phytoalexin production. Phytoalexins reduced
S. sclerotiorum growth in vitro (Sutton and Deverall 1984) and have been related to
increased white mold tolerance of soybean treated with lactofen in the field (Dann et al.
1999)

White Mold Disease Severity. No difference in disease severity between glyphosate
resistant and non-resistant near isoline cultivars was observed (Table 5). S 12-49, S 14-

M7, S 19-90, and S20-B9 had a lower incidence of disease than GL2415, GL2600,

88

P9281, and P93B01 (Table 5). A linear response between the detached leaf bioassay of S.
sclerotiorum lesion diameter 0 to 26 DAT and disease severity at the end of the season
was established by averaging the S. sclerotiorum lesion diameter mean for the six harvest
dates of the fourth trifoliolate. Linear regression analysis indicated soybean with small
lesion diameters had a low incidence of disease late in the season (P = 0.0001) (Figure 8).
White mold disease severity was reduced by all lactofen treatments compared to
the untreated control (Table 5). Thifensulfuron or glyphosate did not affect white mold
severity in the field compared to the untreated control (Table 5). This research does not
support research that glyphosate (Holliday and Keen 1982, Keen et al. 1982, Sanogo et a1.
2000) or ALS-inhibiting herbicides (Sanogo et al. 2000) increase the incidence of disease.
Our research supports other research showing similar disease susceptibility in glyphosate—
resistant and non-resistant cultivars (Lee and Penner 1999, Sanogo et al. 2000) and a
reduction in the incidence of disease following a lactofen application (Dann et al. 1999,
Sanogo et al. 2000).
Soybean Yield. No interaction between cultivar and herbicide treatment was observed
for soybean yield; therefore, yield by cultivar combined over herbicide treatment is
presented (Table 5). Soybean treated with lactofen, thifensulfuron, or glyphosate
treatments had yields similar to the untreated control. Yield of GL2415, S 12-49, P9281,
S14-M7, and S 19-90 was similar in the presence of white mold when combined over
herbicide treatments. GL2415 was the only cultivar that yielded more than its glyphosate-
resistant near isoline (GL2600). Seed of GL2600 was a fifth backcross derivative of

GL2415 at this point in time. When data was sorted and analyzed by herbicide treatment,

89

GL2415 yielded 480 and 720 kg ha" more than GL2600 when treated with thifensulfiiron
and lactofen, respectively, and S 19-90 yielded 400 kg ha" more than S20—B9 when
treated with lactofen.
SUMMARY

Apothecia, the fi‘uiting bodies that produce ascospores which are responsible for S.
sclerotiorum infection of soybean plants, were first observed (personal observation) in the
field approximately 21 DAT (mid-July) (Figure 9). Soybean LAI during this time was 6.5
to 8.0 in 1998 and 6.5 to 7.5 in 1999 (Figure 1). The cottony “white mold” appeared
approximately two weeks later on the lower portion of the soybean stem (Figure 9) and at
other locations on the plants during the peak LAI period of the growing season (Figure 1).

Soybean tolerance to S. sclerotiorum was not related to glyphosate-resistance.
Glyphosate-resistant and non-resistant soybean cultivars had similar canopy development
(Figure 1), flower numbers (data not presented), S. sclerotiorum lesion diameters (data
not presented), phytoalexin production (Table 4), incidence of white mold (Table 5), and
yield (Table 5) except for GL2415 which yielded 390 kg ha" more than GL2600. Yield
differences between GL2415 and GL2600 did not appear to be related to canopy
development differences between near isolines (Figure l). The yield difference between
glyphosate-resistant and non-resistant soybean averaged over herbicide treatment was
probably related to the combination of herbicide injury in the presence of white mold
(Table 5) or this difference may be due to the genetic differences between the near isoline

cultivars since GL2600 was only a fifth backcross derivative of GL241 5.

9O

In our research, cultivar selection had a large influence on reducing the incidence
of white mold in the field (Table 5). The detached leaf bioassay utilized in this and other
research (Dann et al. 1998 and 1999) may provide a simple method for predicting the
relative sensitivity of soybean cultivars to S. sclerotiorum (Figure 8). Lesion diameter
(Figure 5) and phytoalexin production (Table 4) indicated an inherent difference in cultivar
susceptibility to S. sclerotiorum; however, maturity (Table 3), canopy development
(Figure 1), and peak flowering (Figure 4) may also help to reduce disease in more tolerant
cultivars like S 12-49, S14-M7, S 19-90, and S20-B9. In our research, later maturing
cultivars had a greater incidence of white mold than earlier maturing cultivars which was
similar to other studies (Boland and Hall 1987, Buzzell et al. 1993, Chun et al. 1987, Grau
et al. 1982, Yang et a1. 1999).

This research indicated that glyphosate did not affect soybean response,
reproductive development, canopy development, flower number plant", S. sclerotiorum
lesion size, phytoalexin production, disease severity, or yield compared to untreated
soybean. Table 6 summarizes the effects of thifensulfirron, and lactofen on the parameters
evaluated in this research compared to no herbicide treatment. Thifensulfiiron and
lactofen affected soybean development, but only lactofen affected S. sclerotiorum lesion
diameter, phytoalexin production, and disease severity. Therefore, increased phytoalexin
was related to reduced disease in lactofen-treated soybean. However, lactofen affected
canopy development (LAI) more than thifensulfiiron (Figure 3). In this research, lactofen
and thifensulfuron treatments probably did not affect soybean yield compared to untreated

soybean because nearby plants are known to help compensate for yield losses (Hart 1998).

91

A reduction in disease severity would also reduce sclerotia production. Buzzel et
al. (1993) reported a reduction in the number of sclerotia present in the soil when the
incidence of white mold was reduced. Herbicide treatments and cultivars evaluated in this
research that reduced the incidence of white mold may therefore reduce sclerotia
production. The management of S. sclerotiorum sclerotia is similar to managing the weed
seed bank in grain crops.

In summary, glyphosate-resistant and non-resistant soybean were equally
susceptible to white mold; glyphosate did not affect the incidence of white mold in
glyphosate-resistant soybean; and lactofen may be used as a tool to help manage white
mold, but increased soybean yield may not always occur. This information will help
soybean growers to make informed decisions on how weed management programs impact

soybean development, incidence of white mold, and soybean yield.

92

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98

 

   

Yo

R
r

Yoshikawa, M., K. Yamauchi, and H. Masago. 1978. Glyceollin: its role in restricting
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99

Table I. Soybean growth characteristics, environmental conditions, and cultural practices
that influence the infection and incidence of S. sclerotiorum.

 

Soybean growth characteristic,

environmental condition, or cultural

practice that was evaluated. Source

Apothecia Boland and Hall 1988b

Fertility Chun et al. 1987

Flowering Boland and Hall 1982, Boland and Hall 198 8a, Grau et al.
1982, Grau and Radkey 1984

Height Boland and Hall 1987, Kim et al. 1999

Injury Chun et al. 1987

Inoculum Abawi and Grogan 1975, Chun et a1. 1987

Irrigation Grau and Radkey 1984

Light Chun et al. 1987, Boland and Hall 1988a, Pennypacker and
Risius 1999

Lodging Boland and Hall 1987, Buzzell et a1. 1993, Kim et al. 1999

Maturity group Boland and Hall 1987, Buzzell et al. 1993, Chun et al.
1987, Grau et al. 1982, Kim et al. 1999, Yang et al. 1999

Plant age Chun et al. 1987

Rainfall Boland and Hall 1982, Boland and Hall 1988a

Row spacing Buzzell et al. 1993, Grau and Radkey 1984, Phipps 1983

Temperature Buzzell et a1. 1993, Chun et al. 1987, Grau and Radkey

1984, Nelson et al. 1991a, Pennypacker and Risius 1999

100

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101

Table 3. Reproductive stage of development for soybean cultivars combined over
herbicide in 1998 and 1999 14 and 21 d after herbicide treatment (DAT).

 

 

 

 

l4 DAT 21 DAT
Cultivar” 1998 1999 1998 1999
S 12-49 1.8 1.4 2.4 2.6
Sl4-M7 1.9 1.5 2.7 2.1
S 19-90 1.7 1.2 2.4 2.1
$20-39 1.9 1.0 2.4 2.0
GL2415 1.7 0.5 2.0 1.9
GL2600 1.6 0.6 2.1 2.0
P9281 1.6 0.6 2.0 2.0
P93B01 1.0 0.2 2.0 1.4
LSD (1:50.05) 0.4 0.3 0.3 0.4

 

'Lactofen at 70 g ai ha‘1 and thifensulfiiron at 4.5 g ha'l were applied with 0.25% v/v
nonionic surfactant plus UAN (28% urea ammonium nitrate) at 2.3 L ha". Glyphosate
(formulated as Roundup Ultra“) at 840 g ha'l was applied with UAN at 2.3 L ha“.

l’Data were combined over herbicide treatments. Group I soybean included S 12-49, 814-
M7, S 19-90, and 820-B9. Group H soybean included GL2415, GL2600, P9281, and
P93B01.

102

Table 4. Inhibited area of the fourth (treated) trifoliolate 12 d after treatment (DAT) and
the eighth trifoliolate 26 DAT for glyphosate-resistant and non-resistant soybean cultivars
averaged over herbicide treatment using a Cladosporium cucumerinum TLC phytoalexin
bioassay in 1998 and 1999.‘

 

 

 

 

12 DAT 26 DAT
Soybean cultivar” fourth trifoliolate eighth trifoliolate
cm2
S 12-49 12.4 8.4
S14-M7 10.8 6.2
S 19-90 10.1 7.1
SZO-B9 10.9 8.8
GL2415 10.1 5.8
GL2600 11.4 5.8
P9281 10.8 4.2
P93B01 9.2 4.7
LSD (3,5005) 1.6 2.2

 

a'Lactofen at 70 g ai ha‘1 and thifensulfuron at 4.5 g ha’l were applied with 0.25% v/v
nonionic surfactant plus UAN (28% urea ammonium nitrate) at 2.3 L ha". Glyphosate
(formulated as Roundup Ultra‘”) at 840 g ha’l was applied with UAN at 2.3 L ha".
t’Data were combined over the untreated control, lactofen, and glyphosate treatments.

103

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104

Table 6. Summary of the effects of thifensulfuron and lactofen on soybean development,
physiology, incidence of white mold and yield compared to untreated soybean.‘

 

 

Parameterb Thifensulfuron Lactofen
1. Injury Increased Increased
2. Relative humidity 19 DATc No Effect No Effect
3. Temperature 19 DAT° Increased Increased
4. Reproductive stage Delayed Delayed
5. Canopy development (LAI) Reduced Reduced
6. Flower number plant“l Reduced or Reduced or

No Effect Increased
7. S. sclerotiorum lesion diameter No Effectd Reduced
8. Phytoalexin production No Effect" Increased
9. Disease severity index No Effect Reduced
10. Yield No Effect No Effect

 

'Lactofen at 70 g ai ha‘1 and thifensulfuron at 4.5 g ha" were applied with 0.25% v/v
nonionic surfactant plus UAN (28% urea ammonium nitrate) at 2.3 L ha".
bAbbreviations: DAT, days after treatment; and LAI, leaf area index.

‘Measured at the soil surface.

dS 19-90 and SZO-B9 were the only cultivars evaluated.

105

   

 

 

 

 

   

 

 

 

 

   

 

 

 

 

   

 

 

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Figure 1. Soybean leaf area index for cultivar combined over herbicide treatment in 1998
and 1999. Vertical lines indicate the LSD (p50.05). Comparisons between cultivars for
similar days within years are valid.

106

Amodwe 93 2: 282 3:: 35> .82 Ba 32 s 32:8 56 8:558 “.2 a

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107

 

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Y
Rainfall (cm)

 

—o— Untreated — a.— . -Thifensulfuron + NIS + UAN - - o - - Lactofen + NIS + UAN

 

 

 

F’g‘ure 3. Soybean leaf area index for the untreated control, thifensulfuron at 4.5 g ha'l
plus 0.25% v/v nonionic surfactant plus UAN at 2.3 L ha‘1 and lactofen at 70 g ai ha‘1 plus
925% v/v nonionic surfactant plus UAN at 2.3 L ha’1 treatments combined over cultivar
1n 1 998 and 1999. Vertical lines indicate the LSD (p50.05).

108

812-49 and 314-517

 

 

 

a. 7..
c
1!
a.
2
e
3
.0
II.
o s 10 15 20 25 :o 35 4o 45 so
Days after treatment (d)
3 19-00 and 320-39
b. 3;,
C
2
a.
2
0
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2
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0 5 to 15 20 25 30 35 40 45 50
Days after treatment (d)
GL2416 and GLZIOO
C. ': ,9.
E
.2!
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0 s 10 15 20 25 so 35 4o 45 so
Days after treatment (d)
P9281 and P93301
14
d. '- 12
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3 4
2 2
IL
0

 

0 5 10 l5 20 25 30 35 40 45 50

Days after treatment (6)

 

l—o— Untreated —a —Thifensulluron + NIS + UAN me.- Lactofen + NIS + UAN]

Figure 4. Soybean flowers plant‘1 for thifensulfuron at 4.5 g ha'l plus 0.25% v/v nonionic
surfactant plus UAN at 2.3 L ha’1 and lactofen at 70 g ai ha'l plus 0.25% v/v nonionic
surfactant plus UAN at 2.3 L ha'l combined over glyphosate-resistant and non-resistant
isolines in 1998 and 1999. Group I soybean included S 12-49, Sl4-M7, S 19-90, and

S20-B9. Group II soybean included GL2415, GL2600, P928], and P93B01. Vertical lines
Indicate the LSD (p50.05).

109

 

16

 

 

 

 

 

 

 

 

 

 

 

14 A8
:6: 12 - v ""0"-%
g 10 T I' ‘3‘~\ '" ,,,, :I;' ’ ._
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g ' 2L4," " I
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4 , T , r , I

O 5 10 15 20 25 30
Days after treatment (d)

—<>—-S 12-49 - A -S19-90 —O—GL2415 —D—P9281

 

 

 

Figure 5. S. sclerotiorum lesion diameter of soybean cultivars using a detached leaf
bioassay of the fourth trifoliolate (treated). Data were combined over herbicide treatments
for soybean cultivars not resistant to glyphosate in 1998 and 1999. Vertical lines indicate
the LSD (p50.05).

110

18

 

 

 

 

 

 

 

 

 

 

 

 

 

 

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. 14 — ,’ ‘\ - M
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O t 1 , l T
O 5 10 15 20 25 30
Days after treatment (d)
—O— Untreated -----<> Lactofen + NIS + UAN --El- - Glyphosate + UAN

 

Figure 6. S. sclerotiorum lesion diameter using a detached leaf bioassay of the fourth

trifoliolate treated with herbicides. Herbicide treatments included an untreated control,
glyphosate at 840 g ha’l plus 0.25% v/v nonionic surfactant plus UAN at 2.3 L ha", and
lactofen at 70 g ai ha'l plus 0.25% v/v nonionic surfactant plus UAN at 2.3 L ha“. Data
were combined over cultivars in 1998 and 1999. Vertical lines indicate the LSD (p50.05).

111

 

14

 

 

 

 

 

 

 

 

 

 

 

 

 

 

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, I; if
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s 1 El
7 2
6 , , r , I
o 5 1o 15 20 25 so
Days after treatment (d)
—o— Untreated --D-- Thifensdfuron + NIS + UAN
'"A -- Lactofen + NIS + UAN —<>-- Glyphosate + UAN

 

 

 

Figure 7. TLC phytoalexin bioassay using Cladosporium cucumerinum. The untreated
control, thifensulfuron at 4.5 g ha‘1 plus 0.25% v/v nonionic surfactant plus UAN at 2.3 L
ha", glyphosate at 840 g ha’l plus 0.25% v/v nonionic surfactant plus UAN at 2.3 L ha",
and lactofen at 70 g ai ha'l plus 0.25% v/v nonionic surfactant plus UAN at 2.3 L ha'1
treatments were combined over cultivars in 1998 and 1999. S 19-90 and SZO-B9 were the
only cultivars evaluated with thifensulfiiron. Vertical lines indicate the LSD (p$0.0S).

112

60

 

 

 

 

 

 

 

 

 

 

 

 

‘15 50 ‘GLZBOO
9. AP928 P913801
3 40 GL2415
'D
.5
1E 30
i
20
8
§ y = 7.878x - 53.707
5 10 r2=0.9559 -——
P=0.0001
0 , T , . . Y
8 9 10 11 12 13 14 15

Lesion diameter (mm)

Figure 8. Linear regression of the detached leaf bioassay of S. sclerotiorum lesion
diameter of the fourth trifoliolate (treated) averaged over harvest date (0 to 26 DAT) and
disease severity index for glyphosate-resistant (solid shapes) and non-resistant (open
shapes) cultivars. All data were combined over herbicide treatment in 1998 and 1999.

113

10 First apothecia Mycelium growth and sclerotia production

 

 

 

 

 

 

 

 

 

appe ranc A

x i t r \ \

g 8 I .

1: / First white \

E 6 mycelium

0 / appearance \

a 4 Y

“(a / Wilted soybean followed by plant deatl\
2 v

3 0,6
0 1 T I F T I i T T

 

 

 

170 180 190 200 210 220 230 240 250 260 270
Julian calendar days

Figure 9. Development of S. sclerotiorum in soybean at East Lansing, MI.

114

CHAPTER 4
EFFECTS OF PROTOPORPHYRINOGEN OXIDASE INHIBITORS ON
SOYBEAN (Glycine max L.) GROWTH RESPONSE,
Sclerotinia sclerotiorum DISEASE DEVELOPMENT,
AND PHYTOALEXIN PRODUCTION BY SOYBEAN
Abstract. Greenhouse research indicated protoporphyrinogen oxidase (protox)-inhibiting
herbicides other than lactofen may provide additional options for white mold [Sclerotinia
sclerotiorum (Lib.) de Bary] management. Research was conducted to determine soybean
growth response, Sclerotinia sclerotiorum lesion development, and phytoalexin
production in soybean treated with protox-inhibiting herbicides. Postemergence (POST)
injury to soybean from oxyfluorfen at 17.5 g ai ha", carfentrazone at 1.8 g ai ha",
sulfentrazone at 9.0 g ai ha", fomesafen at 280 g ai ha“, acifluorfen at 425 g ai ha",
flumiclorac at 30 g ai ha", CGA-248757 at 4 g ai ha", and oxadiazon at 280 g ai ha" was
equal to or less than injury from lactofen at 70 g ai ha“. Several protox-inhibiting
herbicides reduced S. sclerotiorum lesion diameter in a detached leaf bioassay depending
on the cultivar. All protox-inhibiting herbicides except acifluorfen and CGA—248757
reduced S. sclerotiorum lesion development on treated leaves compared to the untreated
control plants in a detached leaf bioassay. Phytoalexin production in treated leaves and
stems following a postemergence application of sulfentrazone and lactofen was
determined. Lactofen and sulfentrazone increased leaf phytoalexin production similarly,
but neither herbicide affected stem phytoalexin production compared to the untreated

control. Sulfentrazone applied preemergence (PRE) at 210 g ha’l reduced soybean fresh

weight 8 to 69% depending on the cultivar and increased phytoalexin production of S 19-

115

90 and SZO-B9 Roundup Ready‘ID (RR) compared to untreated S 19-90 and 820-B9 (RR)
cultivars. Sulfentrazone PRE also increased phytoalexin production of S 19-90 and 820-
B9 (RR) cultivars compared to GL2415, GL2600 (RR), P9281, and P93BOl (RR)
cultivars. This research indicated that a POST application of protox-inhibiting herbicides
other than lactofen as well as a PRE application of sulfentrazone may increase phytoalexin
production which could increase soybean tolerance to white mold.

Nomenclature: acifluorfen, 5-[2-chloro—4-(trifluoromethyl)phenoxy]-2-nitrobenzoic acid;
carfentrazone, a,2-dichloro-5-[4-(difluoromethyl)-4,5-dihydro-3 -methyl-5-oxo-1H-1,2,4-
triazol-1-yl]-4-fluorobenzenepropanoic acid; CGA—248757, [[2-chloro-4-fluoro-5-
[(tetrahydro-3-oxo-1H, 3H—[ 1 ,3 ,4]thiadiazolo[3,4-a]pyridazin-1-
yliden)amino]phenyl]thio]acetate (proposed common name fluthiacet-methyl); flumiclorac,
[2-chloro-4-fluoro-5-( 1 ,3 ,4,5 ,6,7-hexahydro-1 ,3 -dioxo-2H-isoindol-2-yl)phenoxy] acetic
acid; fomesafen, 5-[2-chloro-4-(trifluoromethyl)phenoxy]-N-(methylsulfonyl)-2-
nitrobenzamide; lactofen, (i)-2-ethoxy-1-methyl-2-oxoethyl 5-[2-chloro-4-
(trifluoromethyl)phenoxy]-2-nitorbenzoate; oxadiazon, 3-[2,4-dichloro-5-(l-
methylethoxy)phenyl]—5-( 1 , 1-dimethylethyl)-1 ,3 ,4-oxadiazol-2-(3H)-one; oxyfluorfen, 2-
chloro-l-(3 -ethoxy-4-nitrophenoxy)-4-(trifluoromethyl)benzene; sulfentrazone, N-[2,4-
dichloro-S-[4-(difluoromethyl)-4, 5-dihydro-3 -methyl-5-oxo-1H-1 ,2,4-triazol- 1-
yl]phenyl]methanesulfonamide; protox inhibitor, protoporphyrinogen oxidase inhibitor;
soybean, Glycine max (L.) Merr; white mold, Sclerotinia sclerotiorum (Lib.) de Bary.

Additional index words: aryl triazinone, cultivar, cyclic imide, diphenyl ether, herbicide
resistant, isolines, oxadiazole, phytotoxicity, postemergence, protoporphyrinogen
inhibitor, Roundup Ready", white mold.
Abbreviations: ‘Novartis S 19-90’, ‘8 19-90’; ‘Novartis $20-B9’, ‘820-B9’; ‘Great
Lakes 2415’, ‘GL2415’; ‘Great Lakes 2600’, ‘GL2600’; ‘Pioneer 9281’, ‘P9281’;
‘Pioneer 93BOl ’,‘P93BOl ’; DAT, d after treatment; NIS, nonionic surfactant; PDA,
potato dextrose agar; UAN, 28% urea ammonium nitrate.
INTRODUCTION
A herbicide may increase, decrease, or have no effect on the incidence of disease

depending on the herbicide mode of action, application timing, and application rate.

Several reviews have summarized the effect of herbicides on disease (Altman and Cambell

116

1977, Katan and Eshel 1973, Levesque and Rahe 1992). In Altman and Campbell’s
(1977) review, only seven postemergence herbicides were discussed. Since 1977,
numerous postemergence herbicides have been discovered, introduced, and utilized for
weed control in soybean and other agronomic crops. Recent studies (Table 1) have
evaluated the effect of postemergence herbicides and adjuvants on the incidence and
development of Sclerotinia spp. White mold is caused by the fungus Sclerotinia
sclerotiorum (Lib.) de Bary. This disease is commonly found in soybean produced in the
north central states and may reduce soybean yield under favorable environmental
conditions (Chun et al. 1987, Hoffman et al. 1998, Kim et al. 1999, Yang et al. 1999).
The protoporphyrinogen oxidase (protox)—inhibiting herbicides are used for
selective preplant incorporated, preemergence, post-directed, or postemergence weed
control in soybean, corn (Zea mays L.), peanut (Arachis hwogaea L.), cotton
(Goswium hirsutum L.), rice (Oryza sativa L.), and other crops (Ahrens 1994, Hatzios
1998). The inhibition of protox in the chloroplasts and mitochondria with these herbicides
causes an inhibition of heme and chlorophyll synthesis (Dayan et al. 1997, Scalla and
Matringe 1994). An accumulation of protoporphyrinogen IX leads to an overflow of
protoporphyrinogen IX into the thylakoid membrane which is oxidized to protoporphyrin
IX (Scalla and Matringe 1994). In the presence of light, protoporphyrin IX generates
singlet oxygen which can cause lipid peroxidation and membrane leakage which causes
subsequent death of susceptible plant species (Dayan et a1. 1997, Scalla and Matringe
1994). The protox-inhibiting herbicides include the cyclic imide (CGA-248757 and

flumiclorac), diphenyl ether (acifluorfen, fomesafen, lactofen, and oxyfluorfen), oxadiazole

117

(oxadiazon), and aryl triazinone (carfentrazone and sulfentrazone) chemical families
(Ahrens 1994, Hatzios 1998, Scalla and Matringe 1994). Lactofen (Hart et al. 1997,
Wichert and Talbert 1993), acifluorfen (Hart et al. 1997), fomesafen (Hart et al. 1997),
CGA-248757 (F ausey and Renner 1999), flumiclorac (F ausey and Renner 1999),
sulfentrazone (Dayan et al. 1996, Krausz et al. 1998) are used for weed control in
soybean.

The interaction between lactofen and the incidence of disease in soybean has been
reported in other research. Lactofen reduced the incidence of F usarium solani f. sp.
g1ycines(Sanogo et al. 2000) and S. sclerotiorum (Dann et al. 1999, Nelson and Renner
1999) in soybean. Lactofen currently has a supplemental label for white mold suppression
in soybean (Anonymous 1998). Acifluorfen and lactofen are known to increase
phytoalexin production in soybean which may help reduce the incidence of disease in the
field (Dann et al. 1999, Komives and Cassida 1983, Levene et al. 1998, Nelson and
Renner 1999). Phytoalexins are antimicrobial compounds produced by plants afier disease
infection or treatment with biotic or abiotic elicitors (Hammerschmidt 1999). Glyceollin
production in ‘Williams 82’ and ‘5 19—90’ increased from 8.3 to 10.2 pg g'l when treated
at the V4 stage with lactofen at 0.04 to 0.11 kg ha" 4 DAT (Dann et al. 1999).
Phytoalexin production increased as early as 2 DAT and persisted up to 26 DAT following
lactofen applied at the V5 stage in other research (Nelson and Renner 1999). Acifluorfen,
bentazon, and nonionic surfactant applied at the V3 stage of development reduced
soybean cyst nematode (Heterodera glycines Ichonohe) egg production by 50 to 60% and

increased phytoalexin production 4 DAT (Levene et al. 1998). The effects of these and

118

other herbicides on the incidence of disease could affect soybean weed management
practices in the north central states.

The susceptibility of soybean cultivars to S. sclerotiorum has been related to the
parentage of the cultivar (Kim et a1. 1999). Planting white mold tolerant or resistant
soybean cultivars is the recommended strategy to reduce yield loss to white mold (Boland
and Hall 1986, Boland and Hall 1987, Chun et al. 1987, Grau and Radkey 1984, Kim et a1.
1999, Nelson et al. 1991b, Wegulo et al. 1998, Yang et al. 1999). Avoiding cultivars with
parentage from ‘Williams’ or ‘Asgrow A3127’ may reduce the incidence of white mold
(Kim et al. 1999). The effect of the protox-inhibiting herbicides on the incidence of white
mold in glyphosate-resistant and non-resistant cultivars has been restricted to lactofen
(Nelson and Renner 1999, Sanogo et al. 2000). The effects of many of the other protox-
inhibiting herbicides on the incidence of disease has not been studied.

This research extends the evaluation of protox-inhibiting herbicides beyond.
lactofen to other diphenyl ether herbicides and the cyclic imide, oxadiazole, and aryl
triazinone chemical families using a challenge inoculation of S. sclerotiorum. This
research reports on the relative phytoalexin production between glyphosate-resistant and
non-resistant cultivars, and the implications regarding the systemic effect of lactofen and
sulfentrazone on phytoalexin production. Our objectives were to determine: 1) soybean
tolerance to the protox-inhibiting herbicides, 2) the effect of protox-inhibiting herbicides
treatments to a challenge inoculation of S. sclerotiorum, 3) the effect of sulfentrazone and
lactofen on phytoalexin production in the stem and leaf, and 4) the effect of sulfentrazone

applied preemergence on whole plant phytoalexin production.

119

MATERIALS AND METHODS
Greenhouse methods. Soybean were planted in 910 m] plastic pots filled with BACCTO
potting soil”. Soybean were grown in a greenhouse with a 16-h photoperiod of natural
and supplemental sodium vapor lighting which provided an additional photosynthetic
photon flux density” of 120 HE m‘2 ' s". Herbicides were applied using a traveling-belt
sprayer traveling at 1.5 km h", delivering 234 L ha'l at 193 kPa, and equipped with an
8001 even flat-fan nozzle". All postemergence herbicide treatments included nonionic
surfactant“ (NIS) at 0.25% and 28% urea ammonium nitrate (UAN) at 2.3 L ha".
Soybean tolerance.
Postemergence protox-inhibiting herbicide rate screen. ‘5 19-90’ soybean were planted
and grown in the conditions previously described. Oxyfluorfen, carfentrazone,
sulfentrazone, lactofen, fomesafen, acifluorfen, flumiclorac, CGA-248757, and oxadiazon
were applied at several application rates with NIS and UAN. Herbicide treatments were
applied at 1430 to 1600 h. Soybean were 13 cm tall and at the V2 stage of development
(F ehr and Caviness 1977) at the time of application. Whole plant visual injury was rated
on a scale from 0 (no visual injury, necrosis, chlorosis, or stunting) to 100 (complete plant

death), and soybean height was measured and the percent height reduction

 

13BACCTO is a product of Michigan Peat Co. Houston, TX 77098.
l4LI-COR. 4421 Superior Street, Lincoln, NE 68504.

lsTeejet flat-fan tips. Spraying Systems Co., North Ave. and Schmale Road, Wheaton, IL
601 88.

l"Nonionic surfactant was Activator-90, a mixture of alkyl polyoxyethylene ether and free
fatty acids, Loveland Industries Inc., PO. Box 1289, Greeley, CO 80632.

120

calculated 14 DAT. The application rate of each protox-inhibiting herbicide was selected
based on injury that was similar or less than lactofen applied at 70 g ai ha“. The study had
four replications and was repeated four times.

Necrosis of the first trifoliolate of S 19—90 and S20-B9 treated postemergence with
protox-inhibiting herbicides. Growing conditions and herbicide applications methods in
this experiment were described previously. Soybean were 13 cm tall and at the V2 growth
stage of development at the time of application. Herbicide application rates (Table 2) that
resulted in soybean injury equal to or less than lactofen were determined from the prior
experiment. The whole plant was covered with a plastic bag with only the first trifoliolate
exposed to the herbicide treatments. Visual necrosis to the treated trifoliolate was
determined 7 DAT on a scale from 0 (no necrosis) to 100 (complete leaf necrosis). This
experiment was a two factor factorial with three replications and was repeated in time.
Factors included soybean cultivar and herbicide treatment.

S. sclerotiorum growth on protox-inhibiting herbicide treated and untreated leaves.
‘8 19-90’, ‘820-B9’, ‘GL2415’, ‘GL2600’, ‘P9281’, and ‘P93BOl’ soybean cultivars
were planted as previously described. Herbicide treatments were applied as previously
described at the rates listed in Table 3. The whole plant was covered with a plastic bag
with only the first trifoliolate exposed to the herbicide treatments. Soybean were at the
V2 stage of development with the second trifoliolate beginning to open at the time of
application. The first (treated) and second (untreated) trifoliolates were harvested from
each treatment 7 DAT to determine if plant defense was localized or translocated to

actively growing tissue in the soybean. The excised leaves were placed in a 150 by 15 mm

121

petri dish17 with a moistened filter paper to ensure a humid environment for white mold
growth. A PDA (potato dextrose agar) plug, 2.4 mm in diameter, from the margin of S.
sclerotiorum was placed on the leaves and the horizontal, vertical, and tangential diameter
of grth was measured with an electronic digital caliper after incubating 48 h. The three
diameter measurements were averaged prior to an analysis of variance. This experiment
was a three factor factorial with three replications and was repeated in time. Factors
included cultivar, herbicide treatment, and harvested leaf.

Phytoalexin production in the leaf and stem. Herbicide treatments were applied as
previously described. Soybean were 16 cm tall and at the V3 grth stage at the time of
application. Treated leaves (first and second trifoliolate) and the stem plus petioles
between the treated leaves were analyzed for phytoalexins using a TLC (thin-layer
chromatography) phytoalexin bioassay. Leaves attached to the plant were harvested and
evaluated in the phytoalexin bioassay.

The TLC phytoalexin bioassay utilized Cladosporium cucumerinum to determine
and quantify antifungal compounds produced by soybean (Keen et a1. 1971). This
bioassay was used to evaluate the relative total antifungal activity (phytoalexin
production) between treatments. Soybean leaves were weighed, cut into 0.5 to 1.0 cm
pieces, and extracted with 40:60 ethanolzwater v/v for 4 h. The extract was evaporated18
at 40 C under reduced pressure to approximately 1 ml and transferred to 10 ml test tube.

The flask was then rinsed with 1 ml water. The extract and water rinsate was combined

 

l7VWR Scientific Products, 800 E. F abyan Pkwy, Batavia, IL 60510.
"Rotavapor, Buchi, Labortechnik AG, Postfach, CH—9230, Flawil.

122

and extracted three times with 1 ml ethyl acetate. The ethyl acetate extract was dried with
anhydrous magnesium sulfate and evaporated under a flow of N2. The residue was
resuspended in ethyl acetate at 50 ul g'1 leaf fresh weight and applied to a silica-gel (250
um) thin-layer chromatography plate19 at 30 ul g'1 fresh weight. The plate was developed
in an acetone:chloroform:ammonia hydroxide solution at 5025021 v/v. C. cucumerinum
spores were suspended in a 100 ml solution of 0.7 g KH2P04, 0.4 g KNO3, 0.3 g
Nazi-IP04, 0.1 g MgSO,, 0.1 g NaCl, and 5.0 g D-glucose that was adjusted to pH 6.5. A
fine spray of suspended spores was applied to the TLC plate and the plate was placed in a
sealed chamber with 100% humidity. Plates were incubated for 2 to 3 d and stored at -20
C. Photo copies20 of the plates were made and clear areas with no C. cucumerinum
growth were quantified with a leaf area meter“. Total areas were calculated that
corresponded to Rfvalues for the methyl ester of glyceofuran (Rf 0.71); isoformonoetin
and a mixture of glyceollin I, II, and III (Rf 0.5 to 0.56); and glyceofiiran and the
precursor of glyceollin II and III (Rf 0.25) previously reported using
chloroforrnzacetonezNH4OH (5025021 v/v) as the mobile phase and a silica-gel TLC plate
as the stationary phase (Ingham et a1. 1981, Keen 1982 ).

The sum of the total inhibited area was calculated and the change in phytoalexin
production was equal to the difference in phytoalexin production between the untreated

cultivar and the same cultivar treated with sulfentrazone or lactofen. The experiment was

 

19Analtech, Inc., P. O. Box 7558, 75 Blue Hen Dr., Newark, DE 19713.
20Cannon, imageRUNNER 330S, One Cannon Plaza, Lake Success, NY 11042.
21LI-3000, LI-COR, 4421 Superior Street, Lincoln, NE 68504.

123

a three factor factorial with three replications and was repeated in time. The factors
included herbicide treatment, soybean cultivar, and leaf or stem sample.

Phytoalexin production with sulfentrazone PRE. Two ‘S 19-90’, ‘S20-B9’,
‘GL2415’, ‘GL2600’, ‘P9281’, and ‘P93BOI ’ soybean seed were planted three cm deep in
910 ml plastic pots of Spinks loamy sand (sand, mixed, mesic Psammentic Hapludalfs)
with pH 5.2 and 1.1% organic matter. Sulfentrazone at 210 g ha'l was applied
preemergence as previously described and pots were watered with 50 ml water to simulate
0.6 cm of rainfall for herbicide activation. Pots were misted lightly to ensure adequate
moisture for germination. The whole plant was harvested 28 d after application, fresh
weights were measured, and fresh weight reduction calculated for each cultivar. The
whole plants were evaluated for phytoalexin production using the methods previously
described. This study was a two factor factorial design with three pots for each of the
four replications and was repeated in time. The factors were herbicide treatment and
cultivar.

Statistical protocol. All research was arranged as a randomized complete block design.
Data were subjected to an analysis of variance according to the factorial description in
each section. Percent data for visual injury or necrosis were transformed to the arcsine
prior to the analysis. The transformation did not affect the conclusions so original data is
presented. Visual necrosis rating data for the experiment evaluating S. sclerotiorum
growth on protox-inhibiting herbicide treated and untreated leaves were subjected to an F

Max test for homogeneity (Kuehl 1994) and data were combined over run since variances

124

for both runs were homogenous. Data were combined over time and means separated
using Fisher’s Protected LSD at p_<_0.05.

RESULTS AND DISCUSSION
Soybean Tolerance.
Postemergence protox-inhibiting herbicide rate screen. Lactofen injured soybean 8% and
reduced height -2% 14 DAT (data not presented). Visual injury from oxyfluorfen at 17.5
g ha", carfentrazone at 1.8 g ha", fomesafen at 280 g ha", acifluorfen at 425 g ha",
flumiclorac at 30 g ha", CGA—248757 at 4 g ha", and oxadiazon at 280 g ha‘l was from 6
to 14% and was no different from lactofen at 70 g ha'l (data not presented).
Sulfentrazone at 8.8, 17.5, and 32 g ha'l reduced soybean height 18, 23, and 27%,
respectively (data not presented). Acifluorfen also reduced soybean height by 15% (data
not presented). Some herbicide treatments stimulated soybean height, but such
observations have been reported with sub-lethal doses of herbicides which exhibit
hormetic grth increases (Brain and Cousens 1989). Injury to soybean from
sulfentrazone at 4.4 g ha'l was 14%. Therefore, the rate was reduced in the following
study. The accumulation of protoporphyrin IX in the presence of light results in the
generation of singlet oxygen which causes the peroxidation of membranes and subsequent
cell destruction (Dayan et al 1997b, Scalla and Matringe 1994). This results in the
bronzing of soybean foliage which causes necrotic lesions characteristic of the injury
symptoms observed.
Necrosis of the first trifoliolate of S 19-90 and S20-B9 treated postemergence with

protox-inhibiting herbicides. NIS plus UAN did not cause soybean necrosis (Table 2).

125

Oxyfluorfen, acifluorfen, oxadiazon, and lactofen caused leaf necrosis that did not differ
between herbicides and cultivars (Table 2). Soybean treated with fomesafen had necrosis
equal to that of the other protox-inhibitors when applied to 820-B9, but not S 19-90.
Sulfentrazone at 1.1 g ha'l did not injure soybean. Therefore, fiirther preliminary research
determined the sulfentrazone rate (9 g ha") which caused leaf necrosis equal to lactofen
(data not presented).

S. sclerotiorum growth on protox-inhibiting herbicide treated and untreated
soybean leaves. Soybean response to challenge inoculations of S. sclerotiorum was
dependent on the cultivar and herbicide treatment (Table 3). In the absence of herbicides,
GL2415 and GL2600 had larger lesions than S 19-90 and 820-B9. A correlation between
lesion diameter and disease severity in the field indicated cultivars with smaller lesion
diameters had a lower incidence of white mold in the field (Nelson and Renner 1999).
There was no difference in lesion diameters between untreated controls of glyphosate-
resistant and non-resistant near isolines 7 DAT (Table 3). These results were similar to
field research using the same near isogenic cultivars (Nelson and Renner 1999). NIS plus
UAN did not affect S. sclerotiorum lesion diameter compared to untreated soybean;
therefore, the adjuvant and fertilizer portion of the herbicide treatment may not affect the
incidence of white mold. Similarly, adjuvants did not affect the incidence of white mold in
field research (Lee and Penner 1999), and the addition of adjuvants to protox-inhibiting
herbicides (Levene et al. 1998) did not have an additive effect on reducing soybean cyst

nematode reproduction. However, adjuvants are known to increase phytotoxicity (Dayan

126

et al. 1996, Fausey and Renner 1999) and foliar absorption (Dayan et al. 1996) of the
protox-inhibiting herbicides.

The S. sclerotiorum lesion diameter differed by cultivar and herbicide treatment
(Table 3). Oxifluorfen and lactofen reduced S. sclerotiorum lesion diameter of five of the
cultivars evaluated compared to the untreated controls. All treatments averaged over
cultivar reduced S. sclerotiorum lesion diameter compared to the untreated control (Table
3). Oxifluorfen, sulfentrazone, and oxadiazon reduced lesion size similar to lactofen;
however, sulfentrazone and lactofen are the only herbicides labeled for weed control in
soybean (crop Protection Chemicals Reference 1998).

Treated and untreated soybean leaves responded differently to challenges of S.
sclerotiorum depending on the herbicide treatment (Table 3). The second trifoliolate
(untreated) of untreated, NIS plus UAN, oxyfluorfen, carfentrazone, sulfentrazone,
lactofen, fomesafen, treatments had larger S. sclerotiorum lesion diameters than the first
trifoliolate (treated). This may be due to a thinner cuticle and younger tissue which may
allow easier disease penetration by a disease. Similarly, older soybean leaves were more
resistant to a Phytophthora megasperma challenge on both susceptible or resistant
cultivars compared to less mature leaves (Bhattaaryya and Ward 1986). NIS and UAN
did not afi'ect S. sclerotiorum lesion diameter of the first (treated) or second (untreated)
trifoliolate compared to untreated soybean. All protox-inhibiting herbicides reduced S.
sclerotiorum lesion diameter compared to untreated plants when averaged over cultivar
except the first trifoliolate treated with acifluorfen and CGA-248757. All protox-

inhibiting herbicides reduced S. sclerotiorum lesion diameter on the second trifoliate.

127

A reduction in lesion diameter in the second trifoliate was expected since acifluorfen plus
NIS increased root phytoalexin levels 4 DAT (Levene et al. 1998). The protox-inhibiting
herbicide treatments evaluated induced a systemic induced resistance response that caused
a reduction in the lesion diameter of S. sclerotiorum in the untreated trifoliolate.

Oxyfluorfen and lactofen provided consistent S. sclerotiorum suppression.
Oxyfluorfen caused 48 to 87% necrosis while lactofen caused 24 to 48% necrosis (data
not presented). Oxadiazon and acifluorfen caused injury similar or greater than that fiom
oxyfluorfen or lactofen (data not presented), but did not consistently reduce S.
sclerotiorum lesion diameter. After soybean was inoculated with S. sclerotiorum,
browning of the epidermal cells within 24 h resembled a hypersensitive response (Sutton
and Deverall 1984). The protox-inhibiting herbicides may mimic such a hypersensitive
response that results in the production of phytoalexins.

Several of the protox-inhibiting herbicide treatments evaluated suppressed S.
sclerotiorum lesion diameter growth. However, since sulfentrazone is labeled for weed
control in soybean, a reduced rate of sulfentrazone (96% of the preemergence application
rate) could be a cost-effective treatment for white mold suppression. Other researchers
have reported severe postemergence injury from sulfentrazone (35 to 55%) at 34 to 56 g
ha“1 (Dayan et al. 1996). Further research should evaluate the postemergence tolerance of
soybean to sulfentrazone in the field.

Phytoalexin production in the leaf and stem. Lactofen was selected in this experiment
as a standard because it consistently reduced S. sclerotiorum diameter in the previous

experiment (Table 3) and its effect on phytoalexin production has been previously

128

reported (Dann et al. 1999, Nelson and Renner 1999). Sulfentrazone was selected
because this herbicide reduced S. sclerotiorum lesion size nearly as much as lactofen in the
previous study (Table 3) and a reduced rate of this herbicide could provide cost-effective
white mold suppression. Leaf and stem fresh weight and phytoalexin production in this
study differed by herbicide treatment (Table 4). Lactofen and sulfentrazone reduced
soybean leaf fresh weight more than stem fresh weight which could be related to abscised
leaves and necrosis caused by the herbicide treatment. Lactofen and sulfentrazone reduced
stem fresh weight 8 to 12%, but had no effect on total phytoalexin production and had no
net effect on the relative phytoalexin production in the stem. Total phytoalexin production
of the untreated control was similar in the leaf and stem when averaged over cultivar.
Lactofen and sulfentrazone did not induce phytoalexin production in the stem, but
increased phytoalexin production in the leaf compared to untreated soybean.

Total antifungal production was greater in S 19-90 and 820-39 compared to
GL2415 or GL2600 (Table 5). 820-89 produced more total leaf phytoalexins than S 19-
90 while P93BOl produced more total leaf phytoalexins than P9281. All cultivar leaves
had more total and relative phytoalexin production compared to stems regardless of
cultivar.

We did not observe an increase in total phytoalexin production in the stem when
plants were treated with lactofen or sulfentrazone (Table 4). S. sclerotiorum lesions
usually originate from the leaf axils of the lower soybean nodes and develop along the
length of the stem which causes the premature death of the entire soybean plant (Cline and

Jacobsen 1983, Boland and Hall 1982, Grau et al. 1982). S. sclerotiorum mycelial grth

129

might be inhibited if there was an accumulation of phytoalexins in the stem (Sutton and
Deverall 1984) which could help explain a reduction in disease severity by lactofen (Dann
et al. 1999, Nelson and Renner 1999).

Phytoalexin production is a good indicator of defense expression (Dann et al.
1999, Hammerschmidt 1999, Levene et al. 1998). However, the protox-inhibiting
herbicides generate active oxygen that may activate other soybean defense mechanisms.
Active oxygen species produced in response to a pathogen have been considered elicitors
in a hypersensitive response and responsible for cell wall lignification and lipid
peroxidation (Baker and Orlandi 1995, Mehdy 1994, Sutherland 1991). The mechanical
and diffusion barriers formed as a result of an oxidative, burst may also explain reduced S.
sclerotiorum lesion growth. A combination of defense mechanisms other than phytoalexin
production alone may contribute to a reduction in S. sclerotiorum lesion growth.
Phytoalexin production with sulfentrazone PRE. Sulfentrazone is applied PRE at 210
g ha‘1 in Michigan and other states for weed control in soybean (Crop Protection
Chemicals Reference 1998). Sulfentrazone PRE reduced fresh weight of all soybean
cultivars except GL2415 compared to untreated cultivars (Table 6). Fresh weight was
reduced 8 to 69% compared to the untreated control depending on the cultivar.
Sulfentrazone reduced GL2600 fresh weight 43% more than GL2415. Differences in
soybean cultivar tolerance to sulfentrazone have been reported (Dayan et al. 1996 and
1997b, Swantek et al. 1998). Herbicide tolerance for sensitive and tolerant varieties was
related to the rapid metabolism, but differences in sensitivity were also related to the

tolerance of the cultivars to the stresses caused by peroxidation (Dayan et al 1997b).

130

Phytoalexin production was greater in S 19-90 and SZO—B9 treated with sulfentrazone
than untreated soybean and GL2415, GL2600, P9281, or P93B01 treated with
sulfentrazone. Whole plant phytoalexin analysis ranked phytoalexin production S 19-90 =
$20-B9 > GL2415, GL2600, P9281, and P93BOl.

Soybean tolerance to the protox-inhibiting herbicides varies and is rate dependent.
Several of the protox-inhibiting herbicides reduced S. sclerotiorum lesion diameter. The
effect of the protox-inhibiting herbicides appeared to be systemic and increased in leaves
that were actively growing at the time of application, yet not in the stem tissue. However,
a reduction of S. sclerotiorum lesion diameter was not always related to the degree of leaf
tissue necrosis following a herbicide treatment. Sulfentrazone at 9 g ha'1 increased leaf
phytoalexin production similar to lactofen, but neither treatment affected phytoalexin
production in the stem. Sulfentrazone PRE may suppress white mold if soybean
phytoalexin production remained high from nine to ten weeks following planting and
application. However, soybean cultivar tolerance would be important to consider if this
treatment was used for white mold suppression. Other postemergence herbicides with a
mode of action similar to lactofen may provide white mold suppression. Further
considerations of the cost-effectiveness, phytotoxicity, and efficacy of these treatments are

needed.

131

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'6801'17801 :19 [(Holoqtedotfiqd "an/0s 'JeA vuuadsvSaw vxoqzqdopfqd
Aq slfitooodfiq ueoqfios ut poonput [eonuoqo [efiurgnue ue :utuooseqdfixorpfiH
-v9 'IL6I 'efiwmd '3 i put; ‘eoru 'a ‘uwa o 'a ‘SWIS 'r 'r ‘1 ‘N ‘uaex

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Table 2. Necrosis of the first trifoliate of S 19-90 and SZO-B9 7 DAT treated with a
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Necrosis

Herbicide treatment Rate S 19-90 S20-B9

g ai ha'1 %
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Fomesafen + NIS + UAN 280 33 57
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139

Table 4. Fresh weight reduction and TLC phytoalexin bioassay of phytoalexin production
m the leaf and stem of soybean treated with lactofen and sulfentrazone averaged over S

l9-90, SZO-B9, GL2415, GL2600, P9281, and P93BOl cultivars.

 

 

 

 

 

 

 

 

 

 

TLC phytoalexin bioassay

Fresh weight reduction Inhibition area A inhibition area'

Treatmentb Rate Stem Leaf Stem Leaf Stem Leaf
g ai ha" % cm2

Untreated 0 0 2. 1 3 .0 O O
Lactofen 70 12 49 2.0 7.6 -O.2 4.6
Sulfentrazone 9 8 41 1.5 7.5 -O.6 4.4

7 —- 1.0 —— -— 0.9 ——

LSD (950.05)

‘A inhibition area on the TLC plate was the difference between the treated plants and the
untreated plants for each cultivar to calculate a relative phytoalexin production for each

 

cultivar
l’All herbicide treatments included nonionic surfactant at 0. 25% v/v and 28% urea

ammonium nitrate at 2 3 L ha".

140

Table 5. Stern and leaf weight reduction and phytoalexin production of S 19-90, 820-39,
GL2415, GL2600, P928], and P93B01 soybean cultivars averaged over untreated,
lactofen, and sulfentrazone herbicide treatments.

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

TLC phytoalexin bioassay
Fresh weight reduction Inhibition area A inhibition area
Cultivar Stem Leaf Stem Leaf Stem Leaf
% cm2 cm2
S 19-90 6 22 3.0 5.9 -O.5 3.5
820-39 5 28 2.7 7.8 -O.5 4.2
GL2415 9 35 1.2 4.6 0.2 1.9
GL2600 1 24 1.0 4.6 -O.4 2.0
P928] 10 39 1.5 5.5 0.2 3.0
P93BOI 9 32 1.9 7.9 -0.7 3.3
LSD (950.05) —— NS --— 1.4 1.3

 

‘A inhibitition area on the TLC plate was the difference between the treated plants and the
untreated plants for each cultivar to calculate a relative phytoalexin production for each

cultivar.

141

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142

CHAPTER 5
YELLOW N UTSEDGE LITERATURE REVIEW

Yellow nutsedge (C yperus esculentus L.) is a problematic perennial weed found
throughout the world (Bendixen and Nandihalli 1987, Lapham 1985, Schippers et al.
1995). The anatomy, morphology, chemical composition, and grth of yellow nutsedge
were reviewed by Doll (1983) and Wills (1987). Yellow nutsedge is a prolific weed
primarily due to its ability to propagate through rhizomes and tubers. Several buds on a
tuber allow for easy reinfestation after tillage or selective herbicide treatments. In reviews
by Glaze (1987) and William and Bendixen (1987), infestation prevention, crop selection
and rotation, row spacing, tillage, fumigation, and herbicide treatments were some of the
options available to help manage yellow nutsedge. The long-term control of this weed
depends on killing the parent tuber and preventing daughter tuber production.

Yellow nutsedge has four varieties described as: var. esculentus found in Africa,
India, southern Europe, and northern North America; var. Ieptostachyus found in North
America, South America, and Europe; var. macrostachyus found in Central America,
southern United States, and the Netherlands; and var. heermannii found in southeastern
United States and the Netherlands (Schippers et al. 1995). Varietal differences were
based primarily upon the size and shape of the floral structures (Schippers et al. 1995). In
a morphological description of Illinois Cyperacea, two perennial varieties of C yperus
esculentus were prevalent (Mohlenbrock 1960). Cyperus esculentus var. esculentus was
described as a typical weed with a wide distribution throughout the state while var.

leptostachyus was less prevalent and had longer spikelets and achenes than C. esculentus

143

(Mohlenbrock 1960). Additionally, var. leptostachyus had more leaves per shoot, wider
leaves, shorter leaf length, more rhizomes, lighter tubers, and a greater number of
flowering plants compared to var. esculentus (Costa and Appleby 1976). There were also
differences in the susceptibility of the yellow nutsedge varieties to various herbicide
treatments; therefore, it was important to identify the variety that was researched (Costa
and Appleby 1976). A taxonomic key (Table 1) was described to determine differences
between varieties based on floral characteristics of C yperus esculentus grown over
different growing conditions (Schippers et al. 1995).

MORPHOLOGY, GROWTH, AND DEVELOPMENT
Leaves and Shoots. Yellow nutsedge may produce over 139 shoots m'2 (Keeley et a1.
1983), grow up to 40 cm tall, and produce a leaf area over 2500 cm2 (Holt and Orcutt
1991). The leaves of yellow nutsedge are green with a mix of yellow and have parallel
venation characteristic of a Kranz anatomy found in C4 or monocot plants (Cafial et al.
1990, Wills et al. 1980). Axillary buds originate at the prophyllar which is the first
structure that emerges from the basal bulb (Jansen 1971). Leaves grow from the basal
bulb and the intercalary meristem located at the base of each leaf (Stoller and Woolley
1983, Wills et al. 1980). Individual leaves grow for 24 to 40 d at a sigmoid growth rate of
2.2 to 4.5 cm (1’1 (Jansen 1971). New leaves emerged every 4 to 5 d (Jansen 1971), and
leaves develop and green under low light conditions (3 11E of2 ' s") (Stoller and Woolley
1983). A thick waxy cuticle (1 to 1.3 pm) covers a layer of epidermal cells with large
vacuoles on the adaxial leaf surface (Wills et al. 1980). The cuticle on the lower leaf

surface is half the thickness of the adaxial surface (Wills et al. 1980). On a cellular level,

144

the xylem is adaxial to the phloem in the vascular bundles (Wills et a1. 1980). The
photosynthetic tissue was concentrated in two cell layers that surround the vascular
bundles in leaves (Wills et al. 1980). Stomates are concentrated at approximately 400
mrn‘2 on the abaxial leaf surface in parallel rows (Wills et al. 1980).

Basal Bulbs. Basal bulbs are similar to a tuber, but the leaf primordia in the basal bulb
produce leaves instead of dormant leaf primordia that surround the tuber (Gifford and
Bayer 1995, Wills et al. 1980). Basal bulbs are the origination of the primary vegetative
growth structure (Jansen 1971). Basal bulbs develop from a leaf associated with the
rhizome and are the result of stem swelling from the rhizomes that originate fi’om a tuber
or adventitious rhizomes (Bendixen 1970b, Gifford and Bayer 1995, Stoller and Woolley
1983, Wills et al. 1980). The meristematic activity in the basal bulb produces above
ground leaf material, and roots and rhizomes which may develop into secondary shoots or
tubers (Bendixen 1970b, Gifford and Bayer 1995, Stoller and Woolley 1983). Basal bulbs
may produce approximately 15 rhizomes that develop into secondary and tertiary plants or
tubers (Jansen 1971).

Basal bulb development has been related to light exposure, growth medium, and
temperature. Basal bulb development was stimulated by exposure to light for 10 to 15
min (Stoller et al. 1972). In the absence of a growth medium (moist cotton), basal bulb
development may be hindered (Stoller and Woolley 1983). When no medium was present,
light exposure had no effect, but a temperature treatment greater than 10 C stimulated
basal bulb development (Stoller and Woolley 1983). However, both light and temperature

stimulated basal bulb development in a medium (expanded mica) (Stoller and Woolley

145

1983). Once temperatures were reduced below 10 C, light was the primary stimulus for
basal bulb development (Stoller and Woolley 1983). Light quality did not affect basal bulb
development since red and far-red light had similar effects on basal bulb initiation as white
light (Stoller and Woolley 1983). The most favorable conditions for basal bulb
development were achieved at high temperatures (20/30 C and 25/35 C) (Stoller and
Woolley 1983).

Rhizomes. Yellow nutsedge contributes up to 74% of its dry weight to root and rhizome
material (Holt and Orcutt 1991). Rhizomes are underground stems that are important for
the spread and vegetative propagation of yellow nutsedge. The rhizome length between
the basal bulb and tuber may be long or short depending on the stimulus discussed above.
Rhizomes develop from axillary buds in the basal bulb (Gifford and Bayer 1995) which
produce secondary shoots, additional rhizomes, and tubers (Bendixen 1970b).

Rhizomes usually do not grow longer than 40 cm long (Tumbleson and
Kommedahl 1961); however, Jansen (1971) observed rhizomes that were up to 60 cm in
length and had over 33 intemodes before developing into a basal bud or tuber. Extensive
rhizome growth was observed when plants were grown under long photoperiods (Jansen
1971). Rhizomes also grow from the tuber during sprouting (Stoller et al. 1972) and are
covered by scale leaves that protect the apical meristem (Wills et al. 1980). These small
(0.5 to 1.0 cm long), scale-like leaves called cladophylls or dormant leaf primordia are
produced along the length of the rhizome (Jansen 1971, Wills et al. 1980). Rhizomes may
develop into basal bulbs or tubers (Bendixen 1970b, Wills et al. 1980). At a 16 h

photoperiod, Jansen (1971) observed that rhizomes had their greatest development into

146

basal buds, but when the photoperiod was less than 14 h, no new shoots were produced
(Jansen 1971).

Rhizomes are produced from July to October in Illinois (Jordan-Molero and Stoller
1978). The rhizome consists of an endo and epidermal layer of cells that are separated by
cortical cells (Wills et al. 1980). The vascular bundles in the rhizome have an endodermal
layer that develops around them, and as the rhizome matures the vascular system is
encompassed with lignified cells (Gifford and Bayer 1995, Wills et al. 1980). The
lignification of the cells in the rhizome results in rhizomes that appear “wiry”. Usually 5 to
6 vascular bundles are found in a rhizome cross-section with the xylem toward the center
and phloem toward the outside (Gifford and Bayer 1995). Veins in the rhizomes are
directly connected with the basal bulb, tubers, roots, and leaves (Bendixen 1973, Wills et
al. 1980).
Inflorescence and Flowers. Yellow nutsedge inflorescence are present as early as July
and flowering continues to late August (Hill et al. 1963, Jordan-Molero and Stoller 1978).
Yellow nutsedge may produce up to 600,000 inflorescence ha‘l (Hill et al. 1963). The
inflorescence grow from meristematic tissue located at the base of the involucral leaves
(Jansen 1971). The flowering characteristics and seed biology of yellow nutsedge were
reviewed (Doll 1983, Stoller and Sweet 1987). The inflorescence size and shape helps
taxonomists differentiate between different varieties (Table 1) of yellow nutsedge
(Schippers et al. 1995). Yellow nutsedge produces a compound umbel inflorescence on a
stem or rachis that is triangular in shape. Spikelets are usually 1 to 3 cm long (Salzman et

al. 1997). Yellow nutsedge requires a 12 to 14 h photoperiod once the plants reach six

147

weeks old (Jansen 1971) to induce flowering. The inflorescence may produce from 1227
to 6685 flowers per inflorescence, and 17% of the flowers produced seed that was viable
approximately two weeks after anthesis (Thullen and Keeley 1979). However, some
populations produced infertile inflorescence (Holt 1994).

Seed. Seed production is essential for the survival of several plant species; however, seed
production is not considered a major factor in the spread of yellow nutsedge. Several
researchers have reported viable seed production by yellow nutsedge (Hill et al. 1963,
Justice and Whitehead 1946, Thullen and Keeley 1979). Yellow nutsedge produced over
90,000 seed plant", and over half sprouted in a study by Hill et al. (1963). Seed
germination may range from 1 to 7 8% and is best when a temperature greater than 21 C
was achieved (Thullen and Keeley 1979). Seed was germinated by Hill et al. (1963) at
20/35 C temperature and with a 16/8 h photoperiod cycle. Yellow nutsedge seed weighed
fiom 0.15 to 0.21 mg (Hill et al. 1963) or 0.13 to 0.31 mg (Thullen and Keeley 1979). As
the seed weight increased, the percent germination also increased (Thullen and Keeley
1979)

Tubers. Yellow nutsedge tubers are enlargements produced at the terminal tip of a
rhizome and are characterized by a cessation of rhizome growth and subsequent swelling
of the rhizome (Bendixen 1970b, Bendixen 197 3, Gifford and Bayer 1995, Tumbleson and
Kommedahl 1961, Wills et al. 1980). Tubers consist of compressed nodes that are densely
arranged at the terminal end of the rhizome with a lignified epidermis and leaves that are
tightly packed against the tuber surface (Bendixen 1973). Scale leaves encompass the

enlargement of a rhizome as the tuber develops from an immature white tuber to a mature

148

brown, tan, or black tuber (Gifford and Bayer 1995). The scale leaves encompass the
tuber once it is mature for protection (Jansen 1971). Mature tubers have a lignified inner
cortex and endodermis (Wills et al. 1980). During sprouting, roots are produced from the
tubers (Wills et al. 1980) and act as part of the vascular system (Gifford and Bayer 1995).
A primary thickening meristem that undergoes periclinal cell division is responsible for the
enlargement of the tuber and establishing roots (Gifford and Bayer 1995).

Several studies have evaluated the chemical composition of yellow nutsedge
tubers. Tubers have a diverse chemical composition which includes hormones and
phenolic compounds like p-coumaric, ferulic, p-hydroxybenzoic, syringic, vanillic,
salicylic, protocatechuic, eugenol, and caffeic acids. The total dry weight composition of
yellow nutsedge includes 15% oil, 12% starch, 9% carbohydrate, 7.5% protein, 0.06% K,
and 0.02% P (Stoller et al. 1972). Most of the stored reserves (60%) are consumed when
a tuber sprouts the first time (Stoller et al. 1972). Several factors like light, fertility, and
hormones influence the carbohydrate level in yellow nutsedge tubers. For instance,
carbohydrate yield in the tubers under a 12 h photoperiod was less than that under a 15.5
h photoperiod (Garg et al. 1967). As temperature, gibberellic acid (GA), or nitrogen
levels increased, tuber carbohydrate levels were reduced regardless of the photoperiod.

Tuber composition also depends on the specific ecotype. Stoller and Weber
(1975) evaluated two yellow nutsedge ecotypes, one from Illinois and one from Georgia,
for tuber composition differences. They determined that the tubers were composed of
47% starch, 3% sugar, 7% protein, and 5% fatty acids (Stoller and Weber 1975). Across

six states, carbohydrate levels in yellow nutsedge varied from 45 to 75% of the total dry

149

weight (Matthiesen and Stoller 1978). Starch was the prevalent carbohydrate and
triglycerides composed approximately 80% of the lipids (Matthiesen and Stoller 1978).
The tuber lipid content was 5 to 7% on a dry weight basis (Stoller and Weber 1975).
Fructose and sucrose were the sugars detected in tubers in the greatest quantity (Garg et
al. 1967, Thullen and Keeley 1978). Bendixen (1973), Wills et a1. (1980), and Tumbleson
and Kommedahl (1961) reported that tuber parenchyma (Gifford and Bayer 1995) cells
were also high in starch content.

Allelopathic compounds are the products of one plant that inhibit the growth of
another plant. Tubers contain a sprouting inhibitor that affects nearby tubers and several
crop species (Drost and Doll 1980, Jangaard et al. 1971, Mufiiz and Tames 1982,
Tumbelson and Kommedahl 1962). The extract was water soluble and nonvolatile
(Tumbleson and Kommedahl 1962).

The identification of viable tubers is useful for evaluating tuber production when
control mechanisms are implemented. Taylorson (1967) classified tubers into four groups:
1) black skinned and flacid which were usually dead; 2) black skinned and turgid which
were usually older tubers; 3) brown and turgid; 4) white to tan colored. In other research,
dead tubers were described as spongy, soft, and hollow (Thullen and Keeley 1975).
Viable tubers were hard and fleshy with a white interior (Stoller and Wax 1973, Stoller et
a1. 1979). Young tubers are small and white colored with parenchyma cells that serve as
an epidermal layer (Thullen and Keeley 1978, Wills et al. 1980) while mature tubers are
large and brown to black colored (Gifford and Bayer 1995, Jansen 1971, Jordan-Molero

and Stoller 1978). Other research has classified tubers based on size (Ghafar and Watson

150

1983a and 1983b, Keeley and Thullen 1970). Small tubers (157 mg) have a shorter life
span compared to large (662 mg) tubers which is important for evaluating the longevity of
tubers in the soil (Thullen and Keeley 1975); however, no differences in sprouting between
tubers with different sizes have been reported (Thullen and Keeley 1975, Stoller et al.
1972)

Keeley et al. (1983) reported initial tuber counts of 449 to 553 0.1 m3, but
maximum tuber densities were reported by Lapham (1985) which totaled 15,344 m’2 with
10,411 viable tubers m‘2 in field research. Tumbleson and Kommedahl (1961) reported
that over 99% of the tubers produced by yellow nutsedge were located in the top 25 cm of
soil. Yellow nutsedge tubers may constitute up to 0.5% (w/w) of the top 15 cm of soil in
the field (Drost and Doll 1980). Up to 10% of the total dry weight produced by yellow
nutsedge was contributed to tubers (Holt and Orcutt 1991). Stoller and Sweet (1987),
Bhowmik (1997), and Doll (1983) reviewed tuber formation, sprouting, longevity,
dormancy, and the variability between biotypes.

Tubers are classified as the noxious perennial structure that allow the persistence
of this troublesome weed, but in areas of Egypt, Spain, Burkina Faso, and the United
States a cultivated variety of yellow nutsedge called chufa (Cyperus esculentus L. cv.
Chufa) was raised (DeVries 1991). In the United States, chufa was described as a food
source for turkeys (Daniel’s Sons 1997). The primary morphological differences between
the weed and chufa were tuber color, rhizome abundance, and the failure to flower
(DeVries 1991). DeVries (1991) reviewed the characteristic differences between the

yellow nutsedge weed and crop. Chufa is also called tiger nut and earth almond in Egypt

151

or patemoster in Spanish countries (DeVries 1991). The edible tubers are Mediterranean
in origin with a hazelnut taste, and have a higher oil content than the weedy yellow
nutsedge (DeVries 1991). The chufa tuber is grey orange, but the weed tuber is grey
brown (DeVries 1991). Chufa tubers do not have the cold tolerance found in the weed
and are more palatable due to reduced fiber coating the tuber surface (DeVries 1991).

For the weedy species, one tuber planted in the spring may produce greater than
1900 shoots and up to 6900 tubers in one year (Tumbleson and Kommedahl 1961). In
other research, one plant produced over 100 tubers (Salzman et al. 1997) and with no
crop one plant produced over 170 tubers (Lotz et al. 1991). Keeley and Thullen (1983)
reported that yellow nutsedge tuber number (y) was directly related to the number of
shoots (x) with the equation y = 15.68x - 57.1.

Tubers are usually less than 10 mm long (Bendixen 1973), but may range from 3
to 11 mm in length (Tumbleson and Kommedahl 1961). Larger tubers have a longer life
span and produced uniform sprouts with a greater weight than smaller tubers (Thullen and
Keeley 1975). Variability of tuber size has been reported by several researchers. For
instance, tuber weight ranged from 157 to 662 mg tuber" dry weight (Thullen and Keeley
1975), 400 to 600 mg tuber‘l fresh weight (Wilen et al. 1996a, 1996b), 50 to 120 mg
tuber" dry weight (Stoller and Wax 1973), 500 mg tuber" (Keeley and Thullen 1974), and
160 to 285 mg tuber" fresh weight with 40 to 60% moisture (Tumbleson and Kommedahl
1961). Tuber size has been related to the ecotype of nutsedge. For instance, populations
in California differed in tuber weight from 6 to 25 mg tuber" (Holt 1994), and tubers from

six states in the Midwest varied fi'om 70 to 710 mg tuber" (Matthiesen and Stoller 1978).

152

Tuber fresh and dry weights for Illinois tubers were 255 and 135 mg tuber", respectively,
but a Georgian ecotype fresh and dry weights were 862 and 538 mg tuber", respectively
(Stoller and Weber 1975). Finally, Costa and Appleby (1976) reported that C. esculentus
var. esculentus produced tubers with a fresh weight of 490 mg from the field and 330 mg
from the greenhouse, while the variety C. esculentus var. leptostachjms yielded tubers
with a fresh weight of 290 mg from the field an 90 mg from the greenhouse. Such
differences in tuber size might affect early growth, the reproductive potential, and the
control of this weed depending on the climate or latitude.

Several environmental factors affect yellow nutsedge tuber production and sprout.
These factors were summarized in Table 2. In the field, temperature, relative humidity,
soil depth, and light are the major environmental factors that affect tuber sprout. The
production of tubers is related to day length. Short days, less than a 12 h photoperiod,
stimulated tuber production. Tubers were observed as early as 7 d afier exposure to short
days, but do not reach maturity until 28 d afier the short d treatment (Bendixen 1973).
Temperature affects tuber survivability and sprouting. If tubers are subjected to freezing
temperatures, winter kill may result. Warm (30 C) and moist conditions are most
conducive for tuber sprout. Most tubers need to be stored in moist conditions because
dessication reduces tuber sprout. Finally, the effect of hormones on tuber production and
sprouting is dependent on the hormone. Several studies indicate that hormones have a
large impact on yellow nutsedge tubers.

The survival of this perennial can result from the ability to produce more than one

sprout per tuber even after mechanical removal of a shoot from the tuber (Stoller et al.

153

1972, Wills et al. 1980, Thullen and Keeley 1975). Stoller et al. (1972) produced plants
from tubers that were sprouted up to three times; however, tuber sprout dropped from 80
to 52 to 27% for tubers that were sprouted one, two, and three times, respectively. Buds
are concentrated at the terminal end of the tuber and range from 1 to 7 per tuber
(Bendixen 1973, Thullen and Keeley 1975, Tumbleson and Kommedahl 1961), but usually
number 4 to 5 (Thullen and Keeley 1975). The oldest and largest buds of a tuber usually
sprout from the terminal end of the tuber cone first (Bendixen 1973). Tubers have
survived up to 64 weeks even if the sprout was removed from the tuber (Thullen and
Keeley 1975).

Tuber production may vary depending on the time of the year or location. In
Georgia, tuber production was greatest in May and June and fell off in the summer months
(Taylorson 1967), but in Minnesota tubers were produced from August to September
(Tumbleson and Kommedahl 1961). In Illinois, tuber production began in August and
yellow nutsedge needed to be controlled by August 1 to avoid tuber production (Jordan-
Molero and Stoller 1978).

Yellow nutsedge varieties and their geographic origin were related according to
Schippers et al. (1995). Horak and Holt (1986) determined that yellow nutsedge had
limited genetic diversity which was characteristic of plants that have insignificant sexual
reproduction. This is not surprising since underground tubers are the primary means of
reproduction and propagation. Some genetic variation may occur among populations, but
less variation occurs within populations (Horak et al. 1987). Holt (1994) evaluated the

variability of yellow nutsedge phenotypic traits and reported high variability between

154

populations in California rather than within the populations. Quantitative differences such
as emergence, flowering, height, dry weight production, rachis number and length, and
spikelet characteristics were greater than isozyme differences, and isozyme differences
provided a limited resource for identifying variability (Holt 1994). Thus, the possibilities
for herbicide resistance by yellow nutsedge may be low due to the limited number of
genotypes and the lack of viable seed. If a biotype was resistant to a herbicide, then the
resistance should be widespread for that biotype due to the limited variability within
populations.

Some field research has evaluated yellow nutsedge tuber production. Researchers
have reported that tuber samples were taken from a 15 cm diameter by 30 cm deep
(Keeley and Thullen 1983), two 10 cm deep by 1820 cm2 samples (Keeley and Thullen
197 5), and Neeser et al. (1997) sampled 13 cm diameter by 30 cm deep soil cores for
purple nutsedge (Cyperus rotundus L.). Banks (1983) evaluated tuber production from
herbicide treatments and samples were removed from the top 15 cm of soil. Gutman and
Watson (1980) described plans for a soil sampler and washing device for tuber separation.
The soil sampler was 15 cm3 and tubers were separated with a 25 mm seed sieve (Ghafar
and Watson 1983a, Gutman and Watson 1980). Since the tubers are concentrated in the
top 20 cm of soil, samples that are taken to this depth would provide a good estimate of
the tubers present in the soil.

Several management decisions in crop production may affect tuber production.
When yellow nutsedge control is implemented, tuber production usually depends on the

competitive nature of the crop. For example, an integrated approach to control yellow

155

nutsedge with a combination of cropping system, mechanical, and chemical control for at
least two years reduced tuber production 97 to 99% in cotton (Gossypium hirsutum L.)
(Keeley et al. 1983); however, tuber production may rapidly increase from 3 to 12 fold if
not controlled in cotton (Keeley and Thullen 1975, 1983). In addition, Hauser et al.
(1974) utilized cotton, corn and peanut rotations to reduce yellow nutsedge tuber
production. In corn, herbicide treatments reduced tuber density more than the untreated
control within one year after treatment (Stoller et al. 1979). By two and three years after
similar treatments were applied, tuber density in the plots was reduced to 20 and 15% of
the original density, respectively (Stoller et al. 1979). However, eradication was not
achieved after 3 years of intensive management in corn (Stoller et al. 1979).

In other research, several herbicide treatments reduced tuber yield after two years
in cotton or soybean (Banks 1983). Stoller et al. (1975) reported that bentazon killed the
parent tubers of treated plants and no regrth was observed with this treatment.
Cultivation reduced purple nutsedge tuber density in the presence and absence of
herbicides (Webster and Coble 1997). Cultivation caused tubers to be exposed and
susceptible to dessication (Keeley et al. 1983). Soil fertility not only increases crop yield
but may affect weed growth. At high nitrogen levels, tuber production was less, but shoot
production increased especially under long photoperiods (Garg et al. 1967). Fertility
levels may affect tuber yield since nitrogen applications to the C. esculentus var.
leptostachyus increased tuber dry weight (Appleby and Paller 1978). Adjusting fertility

and fertility placement may reduce the grth and depletion of nutrients by this weed.

156

Table 3 summarizes sprouting methods and environmental conditions that have
been documented or employed for yellow nutsedge sprouting in controlled environments.
Tuber collection for these experiments depended upon the tuber dormancy characteristics,
resistance to dessication, and dormancy considerations. Keeley and Thullen (1974) and
Banks (1983) collected tubers in March for use in greenhouse experiments to evaluate
sprouting. For other evaluations, Jansen (1971) reported that leaf length, fresh weight of
parent and peripheral shoots, shoot number, rhizome fresh weight, number of rhizome tip
types, mature height, and number of flowering plants were suitable and repeatable
measurements for yellow nutsedge research.

Breaking Dormancy. Tubers are dormant from summer to fall as low levels of
carbohydrates were available for sprouting (Taylorson 1967). Thomas (1969) broke tuber
dormancy by storing tubers at 4 C for 21 d while Stoller and Wax (1973) exposed the
tubers to 2 C to break dormancy. Appleby and Paller (1978) stored tubers in plastic bags
to maintain a humid environment at 5 C for 3 weeks to break dormancy. Research has
been contradictory with respect to whether washing tubers affects sprouting (Taylorson
1967, Tumbleson and Kommedahl 1962). However, washed tubers in Minnesota research
had 70 to 81% greater sprout than unwashed tubers (Tumbleson and Kommedahl 1961).
Table 3 summarizes some of the sprouting methods and environmental conditions needed
to effectively store and sprout yellow nutsedge tubers. Tubers sprouted at 25 to 30 C with
or without a diurnal cycle has been utilized as an effective sprouting method. A petri dish
with two filter papers surrounding the tubers and wrapped with parafilm has been used to

make sure that the tuber with plenty of moisture available. A combination of washing and

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cold storage in humid environments helps break tuber dormancy and ensure viable tubers
for research purposes. Preventing the exposure of tubers to light or diurnal cycles do not
appear to have a large influence on tuber sprout although several researchers have
included this procedure in their experiments. These conditions will insure a dependable
and repeatable environment that will encourage tuber sprout.

Internal tuber chemical composition appears to affect and regulate tuber sprouting.
Hormone treatments may increase or prevent tuber sprouting. A treatment of 100 ppm N-
6 benzyl adenine stimulated purple nutsedge sprouting (Zandstra and Nishimoto 1977).
Also, GA promoted tuber sprouting within one week after planting, but had no effect 2 or
5 weeks after the tubers were planted (Tumbleson and Kommedahl 1962). J angaard et al.
(1971) inhibited tuber sprouting with high levels of phenols (eugenol and salicylic acid);
however, limited salicylic acid was found in the tubers. Phenolic levels in the tubers were
low in the spring and fall (Mufiiz and Tames 1982). Jangaard et al. (1971) and Mufiiz and
Tames (1982) concluded that phenolic compounds had almost no effect upon tuber
dormancy. Abscisic acid inhibited and delayed tuber sprouting depending on the rate
applied to the tubers (Jangaard et al. 1971). Abscisic acid (ABA) levels were highest in
November (6.81 ug g" fresh weight), but during sprouting (April) no ABA was detected
(Mufiiz and Tames 1982). Due to ABA fluctuations and its effect on sprouting, ABA
could be a tuber dormancy regulating compound. Finally, 2,4-D reduced shoot growth 12
d after treatment (DAT) and was translocated to actively growing tissue, but was not
metabolized (Bhan et al. 1970). Table 2 summarizes some of the effects of hormones on

yellow nutsedge.

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Sprouting and Longevity. Yellow nutsedge sprouts from the distal or terminal portion of
the tuber (Tumbleson and Kommedahl 1962). When yellow nutsedge tubers sprout, one
or two buds appeared as white protrusions on the tuber. Yellow nutsedge was considered
sprouted when a bud protruded the scales of the tuber (Holt and Orcutt 1996). The
subsequent sprout of other buds was observed when the growing bud was removed
(Bendixen 1973). Taylorson (1967) reported that tuber sprouting was greatest from
tubers harvested from February to March. However, yellow nutsedge usually emerged
from April to July in Illinois fields (Jordan-Molero and Stoller 1978, Stoller and Wax
1973). Tubers may remain viable in the soil for at least two years (Stoller and Wax 1973).
Tuber sprout was not affected by the tuber weight, but increased shoot biomass was
produced from heavier tubers (Stoller et al. 1972).

Tuber ecotype differences were noted by Stoller and Weber (1975) with respect to
winter survivability. An Illinois ecotype had excellent sprouting (100%) after a cold
treatment, over winter (87%), or fiesh afler harvest in October (91 to 94%); however, the
Georgia ecotype had only 8% sprout after overwintering and 67 to 7 8% after cold
treatment or fresh out of the field (Stoller and Weber 1975). Stoller and Weber (1975)
reported that starch, sugar, and lipid levels increased in the tuber during cold treatment;
however, they concluded that differences in several tuber constituents may be related to
tuber cold tolerance. The adaption of an ecotype and environmental factors may
significantly affect yellow nutsedge tuber survival. Table 2 summarizes some of the
environmental effects on tubers which may directly or indirectly affect tuber production or

sprouting. Mechanical soil disturbance may increase tuber sprouting in the field

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(Taylorson 1967); however, tubers are susceptible to desiccation and low temperatures
(Thomas 1969).

The temperature threshold for tuber sprouting was 6 C for yellow nutsedge while
for purple nutsedge it was 11.2 C (Holt and Orcutt 1996). This would imply that yellow
nutsedge has an excellent chance for early sprouting and establishment, and would help
explain why purple nutsedge was not found in northern latitudes. Yellow nutsedge also
had the broadest sprouting temperature range compared to purple nutsedge and
johnsongrass (Holt and Orcutt 1996). The LDso for temperature was -7 C, but tubers
could easily survive temperatures of -4 C (Stoller and Wax 1973). Thomas (1969)
reported that tubers had the greatest survivability at 90% relative humidity and room
temperature. Under alternating temperature regimes tubers readily sprouted and
stimulated quick basal bulb development in contrast to constant temperatures (Stoller and
Woolley 1983), and as storage time increased, the number of sprouts that emerged from a
tuber increased (Thullen and Keeley 1975).

The depth of tubers in the soil has a large impact upon sprouting. Tubers at 2.5
cm depth had 1% sprouting the following year while sprouting from 10 to 20 cm deep was
55 and 84%, respectively (Stoller and Wax 1973). No shoots emerged from tubers
planted 30 cm deep in Minnesota (Tumbleson and Kommedahl 1961). Tuber sprout
ranged from 49 to 57% 3 to 9 weeks after planting (Dixon and Stoller 1982), 80 to 100%
for tubers in Illinois (Stoller et al. 1972, Stoller et al. 1979, Stoller and Wax 1973) and

100% for tubers in California (Holt and Orcutt 1996, Keeley et al. 1979). Black skinned

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turgid tubers usually had 15% or less sprouting compared with brown tubers (Taylorson
1967)

Recently, researchers have tried to predict yellow nutsedge emergence using
environmental parameters in models to help predict cultivations (W ilen et al. 1996a).
However, emergence patterns for California and Arizona genotypes were different and
under dry conditions these models had difficulty predicting emergence (W ilen et al. 1996a,
1996b). For the California genotype, emergence was reported 47 to 61 d after planting in
dry soil (-0.06 MPa) and 33 to 49 d after planting in wet soil (-0.03 MPa); however,
emergence for the Arizona genotype emerged 51 to 76 d after planting in dry soil (-0.06
MPa) and 43 to 61 d afier planting in wet soil (-0.03 MPa) (Wilen et al. 1996a). Both
genotypes emerged in California earlier than in Arizona even under similar growing
conditions (Wilen et al. 1996a). Predictions of yellow nutsedge emergence could be made
with a 2 d accuracy under adequate rainfall conditions (Wilen et al. 1996b). These
methods may help producers make economical decisions for cultivation timings to control
this weed.

COMPETITION WITH CROPS

Yellow nutsedge is competitive with several crop species early in the growing
season (Doll 1983, Ghafar and Watson 1983a, Keeley 1987). Yellow nutsedge densities
from 40 to 1000 plants m'2 have reduced agronomic and horticultural crop yields (Keeley
1987). Based on the leaf area index (LAI), yellow nutsedge was not considered a big
competitor for light late in the season (Ghafar and Watson 1983a). However, yellow

nutsedge does compete for moisture, nutrients and light, and may exert allelopathic effects

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on crop species early in the season (Keeley 1987). Holt and Orcutt (1991) reported that
yellow nutsedge was more competitive than johnsongrass, purple nutsedge, and cotton
when the above-ground yield and aggressiveness was measured and compared.
Nonetheless, tuber production may be reduced up to 96% depending on the cropping
system and the adopted herbicide management program (Keeley et al. 1979). Although
yellow nutsedge plants rarely grow taller that 20 to 60 cm (Salzman et al. 1997), a dense
stand may intercept light and deplete soil moisture and nutrients before the crop is able to
grow above this aggressive perennial.

Crop selection has an important effect on the ability of yellow nutsedge to
compete. Canopy development differences between crops caused bush bean to be more
competitive than corn, followed by bell pepper (Capiscum annum L.) with purple
nutsedge for incident PAR (Neeser et al. 1997). Competition for water and nutrients is
usually greater than for light due to the low growth habit of nutsedge. However, crops
may require 8 to 9 weeks before they reach 90% light interception (LI) (Keeley and
Thullen 1978). Shading yellow nutsedge effectively reduced shoot number, height, and
tuber and shoot biomass in field and greenhouse research (Keeley and Thullen 1978,
Santos et al. 1997). Flowering was also reduced when yellow nutsedge was in 30 to 47%
shade (Keeley and Thullen 1978). A competitive crop like hemp (Cannabis sativa L.)
reduced tuber and shoot production by yellow nutsedge from 99 to 100% (Lotz et al.
1991). However, tuber production with barley or rye was only reduced 40% (Lotz et al.
1991). Fallow treatments reduced tuber sprout 44% and tuber yield 99% (Tumbleson and

Kommedahl 1961) and glyphosate treatments on fallow areas reduced tubers by 98%

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(Keeley et al. 1979). In one year, viable tubers m‘2 were reduced by about 7 5% with one
year of control (Lapham 1985).

Keeley and Thullen (197 8) reported that there was a strong correlation between
yellow nutsedge growth, tuber production, and shade levels. The amount of light available
to yellow nutsedge decreased as corn population increased (Ghafar and Watson 1983 a).
Yellow nutsedge compensates for low light by increasing shoot grth and reducing root
and tuber production (Santos et al. 1997). Lotz et al. (1991) reported that adequate light
was important for tuber production. Yellow nutsedge may survive under reduced light
intensities because of its low light compensation point (where the CO2 fixed equals CO2
respired) (Santos et al. 1997). Tuber production was reduced under 30 and 70% shade by
approximately 20 and 75%, respectively, when grown under shade cloth (J ordan-Molero
and Stoller 1978). However, yellow nutsedge aboveground growth at 30% shade was
similar to plants grown in full sunlight (Jordan-Molero and Stoller 1978). Similarly,
reducing the light intensity by 43 and 18% reduced tuber production 51 and 3% as well as
tuber weight 56 and 30%, respectively, in the greenhouse 80 d after planting (Lotz et al.
1991). Tuber production was reduced by 95% under 80% shade, and with no shade each
plant averaged 62 tubers (Santos et al. 1997). Under 0% shade, yellow nutsedge produced
from 400 to 1100 tubers, but under 73% shading only 100 to 300 tubers were produced
(J ordan-Molero and Stoller 1978). Keeley and Thullen (1978) reported that 94%
continuous shade reduced tuber production by 94%, but under 80 to 94% continuous
shade there was still a net increase in tubers. Tuber dry weight (y) was reduced linearly in

controlled environments by shading treatments (x): y = 12.46 - 0.16x.

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Several cultural methods like crop planting date, density, selection, rotations,
cultivation, fertility, and weed-free periods may be adopted to manage yellow nutsedge
(Ghafar and Watson 1983a and 1983b, Glaze 1987, Jordan-Molero and Stoller 1978,
Keeley 1987). Crop selection and crop density are key factors that help suppress yellow
nutsedge (Neeser et al. 1997, Keeley 1987, Keeley and Thullen 1978, Keeley et al. 1979).
For instance, alfalfa reached 90% L1 in 2 to 3 weeks, but onion required 26 weeks before
light interception reached 95% (Keeley and Thullen 1978). Based on light interception
data, alfalfa, barley, corn, potatoes, and safflower were the best competitors with yellow
nutsedge (Keeley and Thullen 1978). At high com populations (133,300 plants ha"), the
number of tubers and tuber size was reduced compared to lower corn populations (33,300
plants ha") (Ghafar and Watson 1983a). Tuber production was reduced 97 to 99% when
a combination of cropping systems, mechanical and chemical control methods were used
in cotton (Keeley et al. 1983). This was also evident when herbicides were applied in a
cotton, corn, and peanut rotation in Georgia which killed 78 to 99% of the tubers
depending upon the herbicide use intensity (Hauser et al. 1974). Several herbicide
treatments in cotton and soybean reduced tuber yield after two years of use (Banks 1983).
Intense cultivation alone reduced tuber production from 97 to 99% (Hauser et al. 1974).
Cultivation helps remove early flushes of yellow nutsedge in cotton and corn, but does not
always provide excellent yellow (Keeley and Thullen 197 8) or purple nutsedge control
(Webster and Coble 1997). By delaying planting time, yellow nutsedge growth, dry
matter production, tuber number and weight were reduced (J ordan-Molero and Stoller

1978). Late planted corn had smaller tubers produced in the corn row while large ones

164

were produced between the corn rows (Ghafar and Watson 1983b). The planting date
with the highest corn yield and the lowest yellow nutsedge growth in Canada was the third
week of May (Ghafar and Watson 1983b).

Holt and Orcutt (1991) measured yellow nutsedge competition as the relative
“agressiveness” (A) in terms of dependent variables [A=1.86 - 1.5(relative growth rate) +
200.96(unit leaf rate) + 0.71(ln height) - 0.28(initial propagule weight)]. Yellow nutsedge
was more aggressive than cotton (Holt and Orcutt 1991). The unit leaf rate (mg cm'z' d")
was the best variable that described the relative competitive difference between yellow
nutsedge, cotton, johnsongrass, and purple nutsedge (Holt and Orcutt 1991).

Allelopathic effects of yellow nutsedge residue reduced corn and soybean growth
in greenhouse research, but the effect did not interact with soil texture (Doll 1983, Drost
and Doll 1980). The placement of yellow nutsedge residue beside the seed reduced corn
and soybean growth more than the absence of the residue (Drost and Doll 1980).

Soybean were more susceptible than corn to increased rates (0 to 0.675%) of yellow
nutsedge residue (Drost and Doll 1980). Similarly, Tumbelson and Kommedahl (1962)
reported that yellow nutsedge tuber extract inhibited or delayed red clover, barley, oats,
pea, alfalfa, and soybean gemiination and inhibited root development of some species.
However, Tumbelson and Kommedahl (1962) reported that tuber extracts had no effect
on the germination of wheat or corn unless high rates were present with the corn seeds.
Tumbelson and Kommedahl (1962) developed an alfalfa germination bioassay that could

detect the tuber extract.

165

Corn. Yellow nutsedge densities in corn have been recorded up to 2580 to 3875 plants
m’2 (Tumbleson and Kommedahl 1961), and untreated yellow nutsedge in corn produced
over 1200 tubers m'2 (Stoller et al. 1979). Yellow nutsedge reduced corn biomass 7% in
container-grown plants 6 weeks after planting (Campbell and Hartwig 1982). In the field,
yellow nutsedge densities of 300 and 1200 tubers rn'2 reduced corn yields by 17 and 41%,
respectively (Stoller et al. 1979). However, silage corn reduced tuber production from 4
to 15% (Lotz et al. 1991). Corn intercepted 70% of the incident photosynthetically active
radiation (PAR) at the ground level by June 12, and at July 18, 95% of the incident PAR
was intercepted by com (Lotz et al. 1991). Tuber residue was evaluated from 0 to
0.075% (w/w) which reduced shoot growth of corn and soybean more than foliage residue
(Drost and Doll 1980). However, corn shoot and root growth were reduced by tuber
residue by 46 and 45%, respectively (Drost and Doll 1980). Finally, corn yield was
reduced 8% for every 100 shoots m‘2 and tubers were produced under the corn canopy
(Stoller et al. 1979).

Herbicides may reduce yield loss from yellow nutsedge in corn depending on the
level of yellow nutsedge infestation. For instance, alachlor preplant incorporated (PPI)
prevented yield loss from yellow nutsedge (Stoller et al. 1979). Yield of corn treated with
no postemergence herbicides equaled yield of corn treated with postemergence bentazon
or cultivated, and tuber production in cultivated corn was similar to tuber production in
corn treated with a postemergence application of bentazon (Stoller et al. 1979). Corn
treated with postemergence and postemergence-directed herbicide applications had from 7

to 8% yield loss when a PPI weed control program was not applied (Stoller et al. 1979).

166

Moderate herbicide use controlled yellow nutsedge from 70 to 76% in corn which was
similar to the 70% control with cultivation alone, while intense herbicide use provided
91% control (Hauser et al. 1974).

Soybean. Soybean had greater susceptibility to tuber residue than corn (Drost and Doll
1980). Soybean shoot and root growth was reduced by tuber residues 42 and 55%, and
shoot residues 35 and 61%, respectively (Drost and Doll 1980). However, the leachate
reduced shoot growth by only 16% (Drost and D01] 1980). Soybean seed yield was
reduced by densities from 200 to 300 shoots rn’2 when poor yellow nutsedge control was
attained with herbicides (Banks 1983). Untreated soybean had 379 viable tubers m’2
(Banks 1983). Limited research has been conducted on yellow nutsedge control in
soybean.

Cotton. Variable effects of yellow nutsedge on cotton yield have been reported. Holt and
Orcutt (1991) reported that yellow nutsedge had superior growth characteristics as
measured by several growth parameters compared to cotton. Yellow nutsedge reduced
cotton seed yield 41% and stem diameter 20% two of three years, but had no effect one
year when it was allowed to compete for 25 weeks (Patterson et al. 1980). Cotton seed
and lint yield were reduced by densities from 200 to 300 shoots rn'2 when poor yellow
nutsedge control was reported (Banks 1983). Untreated cotton had 307 viable tubers rn'2
after two years (Banks 1983). Yellow nutsedge reduced cotton seed yield approximately
18 kg ha" for each plant rn'2 (Patterson et al. 1980). Patterson et al. (1980) recommended
that yellow nutsedge should be controlled between 10 and 25 weeks after planting, but

after 25 weeks it was too late. However, four weeks of competition reduced cotton seed

167

yields, delayed maturity, and reduced cotton height in research by Keeley and Thullen
(1975). Keeley and Thullen (1983) reported that a 2 to 6 week weed free period reduced
tuber production and had no effect on cotton yield. Cotton seed yield was reduced 0 to
50% by yellow nutsedge in the untreated control, but had no effect on the quality or
quantity of lint produced (Keeley and Thullen 1983). However, a more drastic effect was
reported where cotton seed yield was reduced 34% in irrigated soil by yellow nutsedge,
and quality was reduced during one year (Keeley and Thullen 1975).
A combination of cropping system, and mechanical and chemical control for at

least two years reduced tuber production 97 to 99% in cotton (Keeley et al. 1983).
Similarly, in a com-cotton rotation, cultivation plus halosulfuron and ametryn reduced
purple nutsedge tuber populations (Webster and Coble 1997). Moderate herbicide use in
cotton controlled yellow nutsedge 37% and cultivation provided 97% control, while
intense herbicide usage provided 92% control (Hauser et al. 1974). Cotton germination
threshold was from 11 to 44 C compared to yellow nutsedge which sprouted from 3 to 45
C (Holt and Orcutt 1996).

YELLOW NUTSEDGE RESPONSE TO HERBICIDES AND HORMONES

Herbicides are utilized as a primary yellow nutsedge control mechanism in crop

production. In 1968, Hauser et al. pointed out in a review that there was a dire need for
the development of herbicides that effectively control yellow nutsedge. In 1987, Pereira et
al. reviewed the efficacy of several herbicides used to control yellow nutsedge. Herbicides
included in the review were 2,4-D, atrazine, bromacil, linuron, bentazon, amitrol,

fluoridone, norflurazon, paraquat, dichlobenil, alachlor, metolachlor, nitrofen, oxyfluorfen,

168

fluorodifen, EPTC, vemolate, butylate, arsenicals, firmigants, and glyphosate (Pereira et
al. 1987). Additives that increased yellow nutsedge control with some of the herbicides
included oils and nitrogen sources like ammonium sulfate (Pereira et al. 1987). Much of
the research evaluating yellow nutsedge control was conducted before the 1980's.
Numerous papers evaluated yellow nutsedge control in agronomic crops prior to the
introduction of bentazon. The decline in yellow nutsedge research was strongly related to
the introduction of this effective postemergence herbicide used in corn and soybean. The
only thing bentazon lacks is residual control; therefore, late sprouting tubers may still
produce enough tubers under the low light intensities of the crop canopy to maintain a
competitive population under stressful environmental conditions.

Several herbicides suppress the growth and development of yellow nutsedge.
Table 4 reviews the effects of several herbicides and control methods on yellow nutsedge
tubers. Herbicides have reduced yellow nutsedge shoot density 77 to 97% (Keeley et al.
1979). For instance, EPTC applied in alfalfa reduced nutsedge shoot counts fi'om 5.5 to
6.4 m". Butylate applied in corn reduced nutsedge shoot number 80 to 85%, but yellow
nutsedge regrew late in the season; and MSMA applied in cotton reduced shoot number
80% (Keeley et al. 1979). Again, yellow nutsedge regrew late in the season (Keeley et al.
1979). Herbicide applications of imazethapyr, metolachlor, or imazethapyr plus
metolachlor reduced yield loss in peanuts to yellow nutsedge (Grichar et al. 1992).
However, one important goal is to reduce the reproductive potential of this weed which
means a reduction in tuber production. A lack of residual herbicide activity is a common

theme for producers trying to control yellow nutsedge since tubers may sprout afier a

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herbicide application or shoots may arise from underground rhizomes. The exposure of
tubers to herbicide treated soil can reduce growth and daughter tuber production (Banks
1983). A combination of cropping system management and herbicide selection can
effectively reduced tuber production to 5 to 17% of the original population and reduce
tuber viability to 3 to 9% (Keeley et al. 1979). The recent introduction of Roundup
Ready® crops and the acetolactate synthase-inhibiting (ALS, EC. 4.1.3.18) herbicides has
made more options for yellow nutsedge control in agronomic crops.
Glyphosate. Glyphosate has been utilized as an effective non-selective herbicide for the
control of perennial weeds like yellow nutsedge. Table 5 summarizes the effect of
glyphosate on yellow nutsedge. Pereira et al. (1987) reported that glyphosate was a
superior herbicide for preventing yellow nutsedge regrowth and tuber production.
Glyphosate symptoms exhibited by yellow nutsedge included leaf apex, basal bulb, and
vein tissue necrosis and chlorosis (Keeley et al. 1979), height reduction, reduced fresh
weight, and rhizome, tuber, and secondary shoot grth inhibition (Canal et al. 1990,
Villaneuva et al. 1985, Stoller et al. 1975). Glyphosate also reduced chlorophyll content 6
DAT when it was applied to yellow nutsedge (Pereira and Crabtree 1986). Similarly,
Villaneuva et al. (1985) reported that chlorophyll pigments were as susceptible to
glyphosate as carotenoid pigments. This may account for the observed symptoms
exhibited by yellow nutsedge to glyphosate.

Variable yellow nutsedge control fi'om glyphosate was usually related to the
application rate. The yellow nutsedge shoot number following an application of glyphosate

to fallow ground remained high through the fallow period, but the shoot number

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decreased by 95% by the next year (Keeley et al. 1979). However, yellow nutsedge plants
treated with 1100 or 2200 g ha" of glyphosate regrew from secondary shoots (Pereira and
Crabtree 1986) and 0 to 3% of the parent tubers were not killed (Stoller et al. 1975).
Appleby and Paller (1978) reported 10 to 12% tuber sprout and up to 42% nonviable
tuber production from 1100 and 2200 g ha" glyphosate. Regrth was less when plants
were treated at 70 compared with 40 d after emergence (Pereira and Crabtree 1986).
Young plants (30 d old) absorbed more l‘lC-glyphosate than 60 (1 old plants, but
translocation to underground plant parts was greater in the 60 (1 old plants (Pereira and
Crabtree 1986) which could account for differential control. In other research, glyphosate
translocation to the tubers decreased as the plant aged (Keeley et a1. 1985). Appleby and
Paller (1978) reduced the number of tubers produced per plant and increased the number
of nonviable tubers as the rate of glyphosate increased from 300 to 2200 g ha".

Careful selection of herbicide tank mixtures with glyphosate should identify
combinations that synergistically control yellow nutsedge since antagonistic combinations
with glyphosate have been reported on difficult-to-control weeds like velvetleaf (A butilon
theophrasti Medicus) (Lich et al. 1997), ivyleaf momingglory [Ipomoea hederacea (L.)
Jacq.] (Lich et al. 1997), purple nutsedge (Rao and Reddy 1999), and sicklepod (Cassia
obtusifolia L.) (Rao and Reddy 1999). Imazaquin plus glyphosate was the most
antagonistic tank mixture for purple nutsedge control (Rao and Reddy 1999). Interactions
could be due to reduced absorption or translocation of glyphosate in the presence of the
ALS-herbicides. The ALS-herbicides may be translocated more readily than glyphosate.

However, Starke and Oliver (1998) reported that chlorimuron did not affect the

171

absorption and translocation of l“C-glyphosate by velvetleaf or pitted morningglory, but
glyphosate increased the absorption of l‘C-chlorimuron or 1“C-imazethapyr by velvetleaf.

An application of naptalam followed by glyphosate synergistically controlled
yellow nutsedge due to the stimulation of shoot grth and inhibited tuber formation from
the naptalam (Appleby and Paller 1978). Such treatments with hormones might provide
greater yellow nutsedge control and should be explored. Pereira and Crabtree (1986)
synergistically controlled 30 d old yellow nutsedge when 1500 g ha" glyphosate was tank
mixed with 1500 g ha" oxyfluorfen due to increased absorption and translocation in young
plants. Similarly, 2,4-D and amitrol increased purple nutsedge control with glyphosate
(Suwunnamek and Parker 1975). However, photosynthesis inhibitors (diuron, atrazine,
and terbacil) antagonized purple nutsedge control with glyphosate (Suwunnamek and
Parker 1975). The addition of ammonium sulfate from 1.25 to 10 kg ha" and other
ammonium salts to glyphosate increased purple nutsedge control four fold (Suwunnamek
and Parker 197 5) while monovalent inorganic salts (NH,, K, and Na) increased purple
nutsedge control compared to untreated, divalent (Zn), and trivalent (Fe) cations (Wills
and McWorter 1985.). However, an organosilicone surfactant plus glyphosate (formulated
as Roundup Ultra")22 did not affect control, absorption, or translocation of 14C-glyphosate
(Bariuan et al. 1999).

Glyphosate is readily absorbed by and translocated in yellow nutsedge. From one
to nine (1 after yellow nutsedge was spotted with l‘lC-glyphosate, absorption and

translocation increased with time (Pereira and Crabtree 1986). In non-resistant soybean,

 

22Roundup Ultra, Monsanto Co., 800 N. Lindbergh Blvd, St. Louis, MO 63167.

172

glyphosate caused severe injury to soybean at 0.3 kg ha", while 2.2 kg ha" was required
for yellow nutsedge control (Stoller et al. 1975). Postemergence applications of 600 g
ha" glyphosate to 2 to 4 week old plants reduced parent tuber resprouting 14 to 32% with
a visual control of 43 to 45% (Keeley et al. 1985). Similarly, Appleby and Paller (197 8)
reported control from 300, 600, 1100, and 2200 g ha" was 27, 52, 97, and 100%,
respectively. One could estimated from this research that 600 g ha" would probably
provide 50% yellow nutsedge growth reduction in greenhouse research (Appleby and
Paller 1978). When a single twin shoot was treated with glyphosate the grth of the
other shoot was reduced (Keeley et al. 1985). Table 6 summarizes research on the
absorption and translocation of glyphosate by yellow nutsedge.

At high rates of glyphosate (10mM), vacuolar phenol levels were reduced, vascular
bundles developed necrotic areas, and sclerenchyma cells were formed between the
cortical parenchyma cells that separate the root and rhizome of yellow nutsedge plants
(Canal et al. 1990). Glyphosate may act as a sprouting inhibitor since 300 to 600 g ha"
increased the number of dormant tubers produced in greenhouse research (Appleby and
Paller 1978). Catalase activity in purple nutsedge was reduced 70% by glyphosate (Abu-
Irrnaileh and Jordan 1978). Glyphosate at 4000 g ha" killed the tubers of young purple
nutsedge (Zandstra and Nishimoto 1977). Young purple nutsedge tubers were a strong
sink for l‘lC-glyphosate, but tubers from older plants were not a strong sink (Zandstra and
Nishimoto 1977). Glyphosate reduced respiration in purple nutsedge tubers as denoted by

the pink color from the tetrazolium test (Zandstra and Nishimoto 1977), and glyphosate

173

was not metabolized by purple nutsedge (Zandstra and Nishimoto 1977). One of the
major problems with glyphosate is consistently killing the parent tubers.

Control of purple nutsedge with glyphosate has been evaluated by several
researchers. Glyphosate at 4000 g ha" reduced purple nutsedge fresh weight and sprouts
per tuber in the greenhouse, and in the field purple nutsedge stands were reduced by 26
and 67% with 2 and 4 kg ha" glyphosate, respectively (Zandstra et al. 1974). Glyphosate
at 2.24 kg ha" and 4.48 kg ha" controlled 17 d and 10 week old plants (Bariuan et al.
1999). In the top 13 cm of soil, tuber production was reduced 92% with 2 to 4 kg ha"
glyphosate and 89% with MSMA, but neither dicamba nor paraquat reduced tuber
production (Zandstra et al. 1974). Tuber sprouting was reduced by 49 to 52% with
glyphosate at 2.24 and 4.48 kg ha" (Bariuan et al. 1999), and 55% with MSMA (Zandstra
et al. 1974). At high humidity (90%) and low water stress (-2 bars), glyphosate at 2000 g
ha" controlled purple nutsedge 73 to 77% due to increased translocation at these
environmental conditions (Chase and Appleby 1979). Bariuan et al. (1999) reported that
the most glyphosate was translocated in the shoot 168 h after treatment. However,
translocation of glyphosate to the underground growth, was less than 1% (Chase and
Appleby 1979). The addition of additives to glyphosate may affect yellow nutsedge
control. Limited research has evaluated other adjuvants commonly used for weed control
with other herbicides. Tank mixture combinations of glyphosate with other herbicides
may be related to increased control due to the addition of additional adjuvant.
ALS-inhibitors. The ALS-inhibiting herbicides inhibit acetolactate synthase which is

essential for the production of the plant essential amino acids leucine, valine, and

174

isoleucine (Claus 1987). ALS-inhibiting herbicides are translocated to the actively
growing meristematic tissue in plants (Claus 1987). The ALS-inhibiting herbicides include
the sulfonylurea, imidazolinone, triazolopyn'midine sulfonanilide, and pyrimidinyl
thiobenzoate herbicide families. These herbicides may cause yellow nutsedge necrosis and
death, suppression, or no control depending upon the herbicide. The use of these
herbicides encompasses a wide spectrum of agronomic and horticultural crops. Several
studies have evaluated the sight of uptake for the ALS-inhibiting herbicides. Many of
these herbicides have the best yellow nutsedge control when applied postemergence which
allows for both foliar and soil uptake. Several of these herbicides provide residual control
of yellow nutsedge unlike glyphosate. Table 6 summarizes the absorption and
translocation research conducted to date with these herbicides.

Ackley et al. (1996) evaluated yellow nutsedge control with several ALS-inhibiting
herbicides and reported that halosulfilron provided 88 to 94% control, chlorimuron 78 to
91%, imazethapyr 28 to 54%, nicosulfuron 50 to 58%, primisulfuron 60 to 73%,
rimsulfuron 51 to 89%, and pyrithiobac 41 to 74%. CGA—152005 (proposed prosulfuron)
or thifensulfuron did not control yellow nutsedge (Ackley et al. 1996). In greenhouse
research, halosulfuron controlled yellow nutsedge greater than chlorimuron, while
imazethapyr had increased activity in the greenhouse compared with other herbicides in
the field (Ackley et al. 1996). Richburg III et al. (1993a) reported that nicosulfilron
provided less than 43% yellow nutsedge control, but nicosulfuron tank-mixed with
bentazon provided greater control than either herbicide applied alone. The interaction

between glyphosate, chlorimuron, irnazamox, imazaquin, and halosulfuron was evaluated

175

for purple nutsedge control (Rao and Reddy 1999). At 50% the recommended rate,
control was greatest with chlorimuron followed by halosulfuron > imazaquin > imazamox
> pyrithiobac of three week old yellow nutsedge and control was at least 27% less
compared with the full rate (Rao and Reddy 1999).

Postemergence applications of halosulfuron at 30 to 280 g ha" provided 34 to
89% yellow nutsedge control 63 DAT (Derr et al. 1996). Yellow nutsedge control with
halosulfuron was linearly related to the rate applied preemergence (PRE) (Derr et al.
1996). Chlorimuron at 10 g ha" provided 33% control 63 DAT which was similar to
3360 g ha" glyphosate (Derr et al. 1996). Control of yellow nutsedge with halosulfuron
was greater than control with 1120 g ha" bentazon (Derr et al. 1996). Similarly,
halosulfuron provided greater yellow nutsedge control than bentazon or imazaquin in
turfgrass (Czamota and Bingham 1997). Below tuber placement treatments of
halosulfuron reduced yellow nutsedge regrowth by 95% in the greenhouse, while control
decreased when halosulfuron was placed at other locations in the soil (Vencill et al. 1995).
A postemergence foliar, soil, or foliar plus soil treatment of halosulfuron reduced yellow
nutsedge dry weight 54 to 65% 30 DAT, and regrth dry weight was reduced by more
than 96% (Vencill et al. 1995). Halosulfuron reduced purple nutsedge tuber population
and shoot density in a corn followed by com rotation in the presence and absence of
cultivation (Webster and Coble 1997). In turfgrass, halosulfuron had no effect on yellow
nutsedge tuber density or weight, but reduced tuber viability with two applications (Molin
et al. 1999). As the rate of halosulfuron increased, tuber viability was reduced (Molin et

al. 1999). Similarly, yellow nutsedge control 6 weeks afier treatment (WAT) was 77 to

176

100% with halosulfuron at 0.14 kg ha" and limited regrth (Czamota and Bingham
1997). No research has evaluated the absorption or translocation of halosulfirron to
underground tubers.

Imazethapyr, an irnidazolinone herbicide, at 60 and 120 g ha" reduced yellow
nutsedge shoot fresh weight 31 to 83% PRE and 70 to 72% postemergence with a 0 to
90% reduction in regrth which was dependent on the application rate and timing (Derr
and Wilcut 1993). Imazethapyr postemergence at 70 g ha" had variable control (40 to
66%) 46 to 142 DAT, but control with a PPI application was 55 to 99% (Grichar et al.
1992). Richburg III et al. (1993b) reported that soil and foliar plus soil applied
imazethapyr provided the best yellow nutsedge control. Imazethapyr at 70 g ha" applied
above or below the tuber stimulated grth 28 DAT and stimulated regrowth when
applied below the tuber 42 DAT (Richburg III et al. 1993b). Above and below tuber
treatments with imazethapyr reduced shoot number, dry weight 28 DAT, and regrth 42
DAT (Richburg III et al. 1993b). A foliar postemergence application of imazethapyr plus
NIS provided similar control at between an early postemergence and postemergence
timing 28 DAT; however, 42 DAT regrowth was reduced by the postemergence greater
than the early postemergence timing, but no difference between timings was reported for
below ground root and tuber control (Richburg III et al. 1993b).

Imazaquin at 250 and 500 g ha" reduced yellow nutsedge fresh weight by 83 to
89% PRE and by 72 to 77% postemergence with an 85 to 95% reduction in regrth
(Derr and Wilcut 1993). Imazaquin at 0.1 to 0.5 ppmw prevented yellow nutsedge shoot

growth and provided the best control when soil-applied (N andihalli and Bendixen 1988).

177

Imazaquin may have a regulatory effect on the apical dominance of yellow nutsedge since
tuber sprouting (y) was linearly enhanced (y = 2.53 + 3.62x) by increasing imazaquin rates
(x) (Nandihalli and Bendixen 1988). Absorption was primarily by the shoot with rapid
initial absorption by the shoot compared with the roots (N adihalli and Bendixen 1988).
Shoot uptake was similar to the acetanilide herbicides and provided greater control than
root uptake (N adihalli and Bendixen 1988). Acropetal and basipetal movement of ”C-
imazaquin was observed in yellow nutsedge (N adihalli and Bendixen 1988).

Chlorimuron PRE reduced yellow nutsedge fresh weight 21 to 62%, and 81 to
83% when applied postemergence with regrowth less than 5% (Derr and Wilcut 1993).
Rates as low as 5 g ha" injured yellow nutsedge (Reddy and Bendixen 1988).
Chlorimuron controlled purple nutsedge better when COC was added as compared with a
nonionic surfactant or organosilicone adjuvant (Jordan 1996). Yellow nutsedge treated
with chlorimuron sprouted 13 to 68% by 28 DAT depending on the chlorimuron rate, and
20 to 25% of the plants sprouted when exposed to soil treated with chlorimuron (Reddy
and Bendixen 1989). A strong linear relationship between tuber sprouting (y) and
chlorimuron rate (x) was determined (y = 88.7 - 1.3x) by Reddy and Bendixen
(1989).When tubers were sprouted in soil treated with chlorimuron at 10 g ha", yellow
nutsedge dry weight was reduced by 98% 28 DAT (Reddy and Bendixen 1989). Foliar
and soil-applied chlorimuron was readily absorbed by the shoot and 53% was recovered
from the shoot within 48 h after treatment (Reddy and Bendixen 1989). When sprouting
tubers were evaluated, less than 2% of the shoot and root-applied chlorimuron was

translocated to the tubers (Reddy and Bendixen 1989). Purple nutsedge control was 20 to

178

28% greater with chlorimuron compared with imazethapyr regardless of the adjuvant and
control was greater with both herbicides when plant size was 2 to 6 cm compared to 8 to
10 cm tall (Jordan 1996). Chlorimuron was slowly degraded by yellow nutsedge (Reddy
and Bendixen 1988). Using thin-layer chromatography to evaluate metabolites, the Rf
0.57 corresponded to the chlorimuron standard, but RfO was considered a polar
metabolite (Reddy and Bendixen 1988).

Limited research has evaluated yellow nutsedge control with the triazolopyrimidine
sulfonanilide or pyrimidinyl thiobenzoate herbicides. Cloransulam-methyl, a
triazolopyrimidine sulfonanilide, did not control yellow nutsedge (Askew et al 1999).
Cloransulam-methyl, applied postemergence following trifluralin, resulted in 72% control
1 to 2 weeks after treatment (Askew et al. 1999). Chlorimuron reduced yellow nutsedge
dry weight by 94% compared to a 50% dry weight reduction with cloransulam-methyl 28
DAT in the greenhouse (Nelson and Renner 1998). Pyrithiobac, a pyrimidinyl
thiobenzoate, at 72 g ha" reduced yellow nutsedge shoot number by 29% 50 DAT (W ilcut
1999). When the placement of pyrithiobac was evaluated in the soil compared to
postemergence treatments, foliar plus soil-applied pyrithiobac provided greater yellow
nutsedge control than either placement applied alone (W ilcut 1999).

Other Herbicides. Several other herbicides have exhibited suppressive or inhibitory
capabilities on yellow nutsedge growth. A summary of the effects of these herbicides on
yellow nutsedge tubers is presented in Table 4. Differences in yellow nutsedge control
may be related to the variety treated. Costa and Appleby (1976) reported that the variety

C. esculentus var. leptostachyus was less susceptible to 2,4-D and more susceptible to

179

atrazine and metribuzin PRE and PPI than C. esculentus var. esculentus. 2,4-D stimulated
shoot initiation from the tubers and reduced rhizome development of C. esculentus var.
leptostachyus (Costa and Appleby 1976). Translocation of materials to the tubers is
greater when the tuber is young than in a mature tuber. When plants were treated with 14C
from urea or naphthaleneacetic acid young tubers accumulate the highest levels of 1“C
while parent tubers accumulate only 8% (Thullen and Keeley 1978). Holt et al. (1967)
reported the translocation of arsenic to purple nutsedge tubers. Tuber viability was not
related to the presence of arsenic, but was related to the depletion of tuber reserves (Holt
et al. 1967).

Amine methylarsonate affected the apical dominance of purple nutsedge tubers and
stimulated shoot initiation after treatment (Holt et al. 1967). Dichlobenil provided
excellent purple nutsedge control and killed tubers according to sprout tests (Waters and
Burgis 1968). PPI applications in the greenhouse of the acetanilide, thiocarbamate, and
amide herbicides delayed sprouting, but the photosynthesis inhibitors killed tubers after
emergence by causing excess consumption of stored tuber reserves in the tubers (Keeley
and Thullen 1974). Incorporated herbicides prevented tuber production and reduced tuber
sprout under normal conditions, but under cool conditions exposure to treated soil did not
affect tuber sprouting (Keeley and Thullen 1974). Soil-applied fluridone and norflurazon
controlled yellow nutsedge eight weeks after treatment and no new tubers were produced
in the greenhouse (Banks 1983). Naptalam increased shoot grth and inhibited tuber
formation (Appleby and Paller 1978). Bentazon provided good yellow nutsedge control at

1100 g ha", but plants regrew after eight weeks (Derr and Wilcut 1993). However,

180

bentazon at 0.8 and 1.7 kg ha" killed 86 and 96% of the parent tubers that were treated,
but basipetal translocation to the tuber did not occur (Stoller et al. 1975). The arsenical
herbicides (DSMA and MSMA) reduced the number of small tubers that sprouted and
killed some tubers (Keeley and Thullen 1970). Smaller and younger plants were easier to
control with the arsenicals than older larger plants (Keeley and Thullen 1970).

Sulfentrazone applied postemergence at 56 g ha" controlled yellow nutsedge 45 to
7 5% 7 DAT depending on the surfactant system utilized, but caused from 30 to 55%
injury to soybean (Dayan et al. 1996). Sulfentrazone controlled yellow nutsedge more
effectively at pH 6.2 than 4.2 when exposed to the root and sulfentrazone was
translocated from the roots to the leaf tissue (Wehtje et al. 1997). Yellow nutsedge was
more susceptible to sulfentrazone than purple nutsedge which was probably related to
increased tuber uptake of l‘lC-sulfentrazone by yellow nutsedge compared to purple
nutsedge (Wehtje et al. 1997). Postemergence applications of sulfentrazone were not
effective in controlling yellow nutsedge, but as soil applications effectively controlled
yellow nutsedge (Wehtje et al. 1997).

Alachlor was taken up in the rhizome shoot of yellow nutsedge and the best
control was observed when treated soil was placed above the tuber (Armstrong et al.
1973). Armstrong et al. (1973) reported that yellow nutsedge rapidly metabolizes alachlor
to a polar metabolite (Rf 0. 1). In the field, alachlor did not decrease tuber numbers in
soybean while other PPI treatments effectively reduced tuber yield (Banks 1983).

Metolachlor applied 20 d after emergence controlled yellow nutsedge, while

bentazon plus metolachlor applied 30 d after emergence provided 92% yellow nutsedge

181

control (Grichar et al. 1996). Metolachlor at 1.7 kg ha" gave 72 to 96% control of yellow
nutsedge depending on the application timing, and tank mixtures with imazethapyr PPI
resulted in 85 to 96% control (Girchar et al. 1992). Dixon and Stoller (1982) reported
that metolachlor had no effect on tuber sprout, 4 ppmw did not kill tubers, and less than
3% of the l‘C-metolachlor was translocated to the tuber. Metolachlor at l ppmw around
the tuber reduced growth more than placement above or below (Dixon and Stoller 1982).
l“C-metolachlor was translocated from the root to shoot when it was applied to the roots,
but basipetal movement was reported when foliar applied (Dixon and Stoller 1982).
Yellow nutsedge metabolized metolachlor more slowly than corn (Dixon and Stoller
1982). Metolachlor reduced yellow nutsedge dry weight more than alachlor (Dixon and
Stoller 1982). Yellow nutsedge metabolized metolachlor to approximately ten metabolites
that were similar to corn, but differential susceptibility between corn and yellow nutsedge
was related to slower metabolism by yellow nutsedge compared with corn (Dixon and
Stoller 1982). Dixon and Stoller (1982) also observed that l4C-metolachlor was exuded
from the yellow nutsedge roots.

Hormones. Hormones also affect the differentiation of rhizomes and the growth of
yellow nutsedge. Research has evaluated GA and auxins. GA is a plant hormone that
stimulates cell division and elongation along with increased starch degradation in cereal
grain seeds (Taiz and Zeigler 1991). Auxins are plant hormones that regulate apical
dominance, inhibit abscission, stimulate ethylene synthesis, affect flowering, stimulate fiuit
development, and stimulate root development on cuttings (Taiz and Zeigler 1991). Afier

GA was applied three times to the youngest shoot tip, yellow nutsedge rhizomes grew

182

erect under several environmental conditions (Bendixen 1970a). 1000 ppm GA reduced
tuber differentiation of plants in a 12.5 h photoperiod, but promoted tuberization under
long photoperiods (Garg et al. 1967). At low temperatures and under a short
photoperiod, GA had limited effects on rhizome suppression (Bendixen 1970a).
Indoleacetic acid (IAA) and triiodobenzoic acid (TIBA), synthetic auxins, alone or with
GA had no effect upon rhizome growth, but under high levels of naptalam or TIBA shoot
initiation was stimulated fi'om the rhizomes (Bendixen 1970a). If one could increase
rhizome differentiation into above ground growth, then treatment and control of yellow
nutsedge may be similar to an annual weed.
BIOLOGICAL CONTROL

Biological control methods were reviewed by Phatak et al. in 1987. Over 66
insects and 10 pathogens were associated with yellow nutsedge (Phatak et al. 1987).
Other organisms like insects, nematodes, bacteria, and birds had been evaluated as
biocontrol methods (Daniel’s 1997, Neeser et al. 1997, Phatak et al. 1987). The infection
of the sedge, C yperus virens, with a fungus caused flower abortion which was an effective
method of control since the sedge did not produce rhizomes. However, the success of
biocontrol methods for yellow nutsedge control has been limited. For example, Keeley et
al. (1970) reported that the infestation of the insect Bactra verutana Zeller was variable.
The insect had little effect on tuber production and showed little promise as a biocontrol

agent since the insect infested yellow nutsedge in the fall (Keeley et al. 1970).

183

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188

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Pereira, W. and G. Crabtree. 1986. Absorption, translocation, and toxicity of glyphosate
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Reddy, K. N. and L. E. Bendixen. 1989. Toxicity, absorption and translocation of soil-
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189

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Stoller, E. W., L. M. Wax, and R. L. Matthiesen. 1975. Response of yellow nutsedge
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Stoller, E. W., L. M. Wax, and F. W. Slife. 1979. Yellow nutsedge (Cyperus esculentus)
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Stoller, E. W. and J. T. Woolley. 1983. The effects of light and temperature on yellow
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190

Thomas, P.E.L. 1969. Effects of desiccation and temperature on survival of C yperus
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Thullen, R. J. and P. E. Keeley. 1975. Yellow nutsedge sprouting and resprouting
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Thullen, R. J. and P. E. Keeley. 1978. The effect of Cyperus esculentus tuber maturity on
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Thullen, R. J. and P. E. Keeley. 1979. Seed production and germination in Cyperus
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Tumbleson, M. E. and T. Kommedahl. 1961. Reproductive potential of Cyperus
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Tumbleson, M. E. and T. Kommedahl. 1962. Factors affecting dormancy in tubers of
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Vencill, W. K., J. S. Richburg, III, J. W. Wilcut, and L. R. Hawf. 1995. Effect of MON-
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Technol. 92148-152.

Villanueva, M. J. C., B. F. Mufiiz, and R. S.Tames. 1985. Effects of glyphosate on
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Waters, W. E. and D. S. Burgis. 1968. Herbicidal persistence in soil and its effect on
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Webster, T. M. and H. D. Coble. 1997. Purple nutsedge (Cyperus rotundus)
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Wehtje, G. R., R. H. Walker, T. L. Grey, and H. G. Hancock. 1997. Response of purple
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Wilcut, J. W. 1999. Influence of pyrithiobac sodium on purple (C yperus rotundus) and
yellow nutsedge (C. esculentus). Weed Sci. 46:111-115.

Wilen, C. A., J. S. Holt, and W. B. McCloskey. 1996a. Effects of soil moisture on

observed and predicted yellow nutsedge (C yperus esculentus) emergence. Weed Sci.
44: 82 1-829.

191

Wilen, C. A., J. S. Holt, and W. B. McCloskey. 1996b. Predicting yellow nutsedge
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William, R. D. and L. E. Bendixen. 1987. Year-round management of yellow nutsedge
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Wills, G. D. 1987. Description of purple and yellow nutsedge (Cyperus rotundus and C.
esculentus). Weed Technol. 1:2-9.

Wills, G. D., R. E. Hoagland, and R. N. Paul. 1980. Anatomy of yellow nutsedge
(Cyperus esculentus). Weed Sci. 28:432-437.

Wills, G. D. and C. G. McWhorter. 1985. Effect of inorganic salts on the toxicity and
translocation of glyphosate and MSMA in purple nutsedge (Cyperus rotundus). Weed
Sci. 33:755-761.

Zandstra, B. H. and R. K. Nishimoto. 1977. Movement and activity of glyphosate in
purple nutsedge. Weed Sci. 25:268-274.

Zandstra, B. H., C. K. H. Teo, and R. K. Nishimoto. 1974. Response of purple nutsedge
to repeat applications of glyphosate. Weed Sci. 22:230-232.

192

Table 1. Taxonomic key from Schippers et al. (1995) for Cyperus esculentus varieties
based primarily on flower characteristics of plants grown in the field and over different
fertility, temperature, and day length conditions in the greenhouse.

I. a) Angle between spikelet and rachis is < 35'.
b) Spikelet density is > 2 mm".
c) Ratio of ray number to bracts is < 0.75 or > 15 flowers spikelet". Variety ......... heermannii
1. Other characteristic combinations.
a) Floral scale length mean is 2 3.4 mm. Variety ........................... macrostachyus
b) Floral scale length mean is < 3.4 mm.
11. a) Floral scale length mean is _>_ 2.8 mm.
b) Height of the floral scale maximum width is _>_ 1.5 mm. Variety ............. macrostachyus
11. a) Floral scale length mean is < 2.8 mm.
b) Height of the floral scale maximum width is < 1.5 mm.
1. 3) Floral scale length mean is z 2.5 mm.
b) Floral scale reaches maximum width at z 1.4 mm.
c) Floral scale width maximum is > 1.8 mm.
d) Ratio of the top floral scale to the mean length of the spikelet floral
scales is > 0.9 or number for floral scales spikelet" is z 20. Variety . . macrostachyus
2. Other characteristic combinations.
IH. Style length with branches is z 4.2 mm. Variety ............................... esculentus
111. Style length with branches is < 4.2 mm.
a) Maximum floral scale width is 5 1.5 mm. Variety ........................ leptostachyus
b) Maximum floral scale width is > 1.5 mm.
1V. Floral scales spikelet is > 23. Variety .................................... leptostachyus
IV. Floral scales spikelet is 5 23.
a) Top floral scale lengthzfloral scale length > 0.94. Variety ................... leptostachyus
b) Top floral scale length:floral scale length 5 0.94.
V. Floral scale width is _<_ 1.8 mm and style length is _<_ 3.4 mm. Variety ............ leptostachyus
V. Floral scale width is 5 1.8 mm and style length is S 3.4 mm.
a) Floral scale width is S 1.9 mm, floral scale length mean is S 2.3 mm, and anther length is 5 1.4
mm. Variety ....................................................... leptostachyus
b) Floral scale width is > 1.9 mm, floral scale length mean is > 2.3 mm, and anther length is > 1.4
mm.
VI. a) Floral scale width is 5 2.0 mm, floral scale length mean is _>_ 2.6 mm, and style length is 5 3.5
mm or the angle between the floral scale and spikelet axis is < 17'. Variety ....... Ieptostachyus
b) Other characteristic combinations. Variety ................................. esculentus

 

193

Table 2. Environmental stimulus and the effect on yellow nutsedge tuber production and

sprouting.
Environmental stimulusa
Temperature

Increase from 21

Effect on tubers

At 12.5 h photoperiod, tuber production

Source

Garg et a1. 1967

 

 

to 27 C increased.
6 and 43 C Threshold temperatures for tuber Holt and Orcutt 1996
sprout.
Alternating Faster and improved tuber sprout. Miles et al. 1996
-7 C LD,O temperature. Stoller and Wax 1973
12 C Minimum temperature required for Stoller and Wax 1973
sprouting.
22 C vs 4 C Increased survival and weight loss at Thomas 1969
higher temperature.
Cold treatment Increased sprouting from 12% in the fall Tumbleson and
to 95% in the spring. Kommedahl 1962
Degree-days California 114 degree-day units were needed before Wilen et al. 1996b
genotype predicting emergence.
Degree-days Arizona 111 to 115 degree-day units were Wilen et al. 1996b
genotype needed before predicting emergence.
Relative Humidity
30 vs 90% Increased survival and reduced tuber Thomas 1969
weight loss at high relative humidity.
Dessication Reduced tuber sprouting. Thomas 1969
Soil Depth
2.5 to 5.1 cm Suffered winter kill. Stoller and Wax 1973

10.2 to 20.3 cm
Top 25 cm of soil

Greatest tuber emergence was observed.
Location of over 99% of the tubers.

Stoller and Wax 1973

Tumbleson and

 

 

Kommedahl 1961
Scarification
Cut tuber in half Increased tuber sprouting. Tumbleson and
Kommedahl 1962
Wash with water Increased sprouting. Tumbleson and
Kommedahl 1961,
1962
Light
Day length Long (1 had limited tuber production and Appleby and Paller
short (1 induced tuber formation. 1978

 

194

Table 2 (cont ’d).

 

Photoperiod Reduced photoperiod to 10 h induced Bendixen 1973
tuberization.

Photoperiod Reduced photoperiod from 14.5 to 12 h Costa and Appleby
to stimulate tuber production. 1976

Photoperiod Short d (12.5) stimulated greater tuber Garg et al. 1967
production than 14 and 15.5 h d.

Photoperiod 8 to 12 h had the greatest tuber Jansen 1971

development from rhizome tips and
tubers were produced by 3 months after

 

planting.

80 to 94% continuous Net gain of tubers. Keeley and Thullen

L1 1978

30% reduction in PAR Reduced tuber production 32%. Keeley and Thullen

1978
94% reduction in PAR Reduced tuber production 95%. Keeley and Thullen
1978

Crop selection Tuber production was reduced 18 to Lotz et al. 1991
43% under reduced light intensities.
Hemp had early canopy closure.

80% incident PAR Reduced tuber dry weight to 100 mg Santos et al. 1997
plant" and production to 3 plant".

Photoperiod 12 h was needed for tuber production. Santos et al. 1997

Hormones

2,4-D Shoot, rhizome, and rhizome number Bhan et al. 1970
were reduced 12 DAT.

Gibberillic acid Reduced tuberization and increased Garg et al. 1967
shoot production under short
photoperiods. Increased tuberization
and reduced shoot production under
long photoperiods.

ABA Inhibited and delayed tuber sprouting. Jangaard et al. 1971

Gibberillic acid Promoted tuber sprouting one week Tumbleson and
after planting. Kommedahl 1962

N-6 Benzyl adenine 100 ppm stimulated tuber sprouting. Zandstra and

Nishimoto 197 7

 

aAbbreviations: ABA, abscisic acid; LI, Light interception; and PAR photosynthetically
active radiation.

195

 

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203

Table 6. Absorption and translocation of glyphosate, ALS-inhibitors, and other herbicides

in yellow nutsedge.

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Glyphosate
Absorbed Translocated Source
32% of the applied 28% of the applied amount. Pereira and Crabtree
amount (1000 g ha“1 22% of the translocated herbicide was 1986
application rate). moved to below ground plant parts.
83% in shoot Keeley et al. 1985
15% to underground tissue
2% in the tubers
20% was translocated from the treated leaf Zandstra and Nishimoto
of purple nutsedge 1977
ALS-Inhibitors
Absorbed Translocated Source
Imazaquin
83% in the shoot and 1% Acropetal and basipetal movement. Nadihalli and Bendixen
from the roots after 48 h 1988
Chlorimuron
12% foliar 15% in l d, 78% remained in the leaf, Reddy and Bendixen
acropetal and basipetal, 1988
0.2% in the tuber
47% foliar in 12 h < 1% translocated to the tubers Reddy and Bendixen
1.3% root 1989
Other herbicides
Absorbed Translocated Source
2,4-D
<l9% after 24 h <1% translocated, but not metabolized Bhan et al. 1970
Metolachlor
roots absorbed 8% 53% after 13 d, acropetal Dixon and Stoller 1982
shoots basipetal movement and was exuded Dixon and Stoller 1982
Alachlor
uptake is above the tuber shoots had limited acropetal movement Armstrong et al. 1973
Sulfentrazone
43% was absorbed <1% in the tubers Wehtje et al. 1997

 

204

CHAPTER 6
YELLOW NUTSEDGE (Cyperus esculentus) CONTROL AND TUBER
PRODUCTION WITH GLYPHOSATE AND
ALS-INHIBITING HERBICIDES

Abstract: Greenhouse and field research evaluated yellow nutsedge growth, vegetative
control, and tuber production following an application of glyphosate, acetolactase
synthase (ALS)-inhibiting herbicides, and tank mixtures thereof. In the greenhouse or
field, glyphosate at 840 g ae ha‘1 did not control yellow nutsedge. Chlorimuron and
imazethapyr/imazapyr provided 90% or greater yellow nutsedge control, prevented
panicle formation, and reduced tuber density and fresh weight by 90% or more 14 weeks
after treatment (WAT) in the greenhouse. The addition of glyphosate to cloransulam or
imazethapyr increased yellow nutsedge control and reduced tuber density and fresh weight
more'than either treatment or glyphosate applied alone. Imazamox at 45 g ai ha“l applied
alone stimulated tuber production. In the field, halosulfuron and chlorimuron provided
greater than 85% yellow nutsedge control and reduced tuber density and fresh weight
more than 80 and 85%, respectively. All treatments reduced tuber fresh weight 45 to 91%
when compared to the untreated control, and tuber density was reduced 33 to 90% by all
herbicide treatments except imazamox and rimsulfuron applied alone compared to the
untreated control. Tank mixtures of glyphosate with the ALS-inhibiting herbicides
increased visual control, but did not reduce tuber production compared to the ALS-
inhibiting herbicides applied alone. Tuber sprouting was reduced 19% in plots treated

with halosulfuron and pyrithiobac compared to untreated yellow nutsedge 42 WAT.

Yellow nutsedge control by the herbicides evaluated and applied at recommended use

205

rates was ranked: halosulfuron and chlorimuron (> 80% control and reduction in tuber
density); imazethapyr/imazapyr, imazethapyr, and glyphosate (50 to 80% control and
reduction in tuber density); and cloransulam, rimsulfuron, and imazamox (20 to 50%
control and reduction in tuber density). However, pyrithiobac controlled yellow nutsedge
48% and reduced tuber density 60%. Long-term yellow nutsedge management may be

achieved with treatments that reduce tuber production.

Nomenclature: chlorimuron, 2-[[[[(4-chloro-6-methoxy-2-
pyrimidnyl)amino]carbony1]amino]sulfonyl]benzoic acid; cloransulam, 3-chloro-2-[[(5-
ethoxy-7-fluoro[ 1 ,2,4]triazolo[ 1,5-c]pyrimidin-2yl)sulfony1]amino]benzoic acid;
glyphosate, N-(phosphonomethyl)glycine; imazapyr, (i)-2-[4,5-dihydro-4-methyl-4-(1-
methylethyl)-5-oxo-1H-imidazol-2-yl]-3-pyridinecarboxy1ic acid; MON 12000 or MON
12037 (proposed halosulfuron), methyl 5-[[(4,6-dimethoxy-2-
pyrimidinyl)amino]carbony1aminosulfonyl]-3 -chloro-1 -methyl-1 -H-pyrazole-4-
carboxylate; imazethapyr, 2-[4, 5-dihydro-4-methyl-4-( 1 -methylethyl)-5-oxo- lH—imidazol-
2-yl]-5-ethyl-3-pyridinecarboxy1ic acid; imazamox, 2-[4,5-dihydro-4-methyl-4-(1-
methylethyl)-5-oxo-1H-imidazol-Z-yl]-5-(methoxymethyl)-3-pyrdinecarboxyilic acid;
pyrithiobac, 2-chloro-6-[(4,6-dimethoxy-2-pyrimidinyl)thio]benzoic acid; rimsulfuron, N-
[[(4,6-dimethoxy-2-pyrimidinyl)amino]carbonyl]-3 -(ethylsulfony1)-2-pyridinesulfonamide;
yellow nutsedge, C yperus esculentus L. #23 CYPES.

Additional index words: acetolactase synthase inhibitor, plant height, postemergence,
shoot production, tubers, tuber production.

Abbreviations: ALS, acetolactate synthase; AMS, ammonium sulfate ((NH4)ZSO,); COC,
crop oil concentrate; MSO, methylated seed oil; NIS, nonionic surfactant; WAT, weeks
afier treatment.

 

23Letters following this symbol are a WSSA-approved computer code from Composite
List of Weeds, Revised 1989. Available from WSSA, 1508 West University Ave.,
Champaign, IL 61821-3133.

206

INTRODUCTION

Yellow nutsedge is a problematic perennial weed found throughout the world
(Bendixen and Nandihalli 1987, Lapham 1985, Schippers et al. 1995). Yellow nutsedge is
prolific and aggressive due to its rapid grth (Holt and Orcutt 1991) and ability to
propagate through rhizomes and tubers. Several buds on each tuber can sprout and cause
reinfestation afier tillage or selective herbicide treatments. Yellow nutsedge competes with
crops early in the growing season (Ghafar and Watson 1983a, Keeley 1987), and densities
from 40 to 1000 plants m'2 have reduced agronomic and horticultural crop yields (Keeley
1987). In reviews by Glaze (1987) and William and Bendixen (1987), prevention, crop
selection and rotation, row spacing, tillage, fumigation, and herbicides were some of the
strategies available to manage this weed.

Yellow nutsedge tubers are enlargements produced at the terminal tip of the
rhizome (Bendixen 1970, Bendixen 1973, Gifford and Bayer 1995, Tumbleson and
Kommedahl 1961, Wills et al. 1980). Tubers consist of compressed nodes with a lignified
epidermis and scale leaves that are tightly packed against the tuber surface (Bendixen
1973). Scale leaves encompass the tuber for protection as it develops from an immature
white tuber to a mature brown, tan, or black tuber (Gifford and Bayer 1995, Jansen 1971).
Tubers are usually less than 10 mm long (Bendixen 1973), but may range from 3 to 11 mm
in length (Tumbleson and Kommedahl 1961). Larger tubers have a longer life span and
produce uniform sprouts with a greater weight than smaller tubers (Thullen and Keeley
1975). One tuber planted in the spring may produce 1900 shoots and 6900 tubers in one

year (Tumbleson and Kommedahl 1961). Up to 10% of the total dry weight produced by

207

yellow nutsedge is partitioned into the tubers (Holt and Orcutt 1991). Tumbleson and
Kommedahl (1961) reported that over 99% of the tubers produced by yellow nutsedge
were located in the top 25 cm of soil. Tubers are the primary means of propagation and
may have allelopathic effects on the crop (Drost and Doll 1980, Tumbleson and
Kommedahl 1962). The long-term control of yellow nutsedge depends on killing the
parent tuber and preventing daughter tuber production.

Crop management used in conjunction with herbicide treatments may reduce tuber
viability, germination, and yield. For instance, tuber yield, growth, and daughter tuber
production were reduced following two years of several herbicide treatments in cotton
(Gosgpium hirsutum L.) or soybean (Banks 1983). Stoller et al. (1975) reported that
bentazon killed the parent tubers of treated plants and no regrowth was observed. In other
research, cultivation reduced yellow (Stoller et al. 1979) and purple nutsedge (Cyperus
rotundus L.) (Webster and Coble 1997) tuber density in the presence and absence of
herbicides. The integration of crop rotation and herbicide selection effectively reduced
yellow nutsedge tuber production to 5 to 17% of the original population and reduced
tuber viability 3 to 9% (Keeley et a1. 1979, Keeley et a1 1983). Yellow nutsedge tubers can
resprout after a herbicide application or new shoots may arise fiom underground rhizomes
if the herbicide has no residual activity.

The introduction of glyphosate-resistant crops and the acetolactate synthase-
inhibiting (ALS, EC. 4.1.3.18) herbicides has provided more postemergence perennial
weed management options for crop producers. As glyphosate resistant crops become

more widespread, producers may select for weeds like yellow nutsedge that are not

208

completely controlled at a typical use rate of glyphosate. Variable yellow nutsedge control
with glyphosate has been reported and was related to the application rate and the size of
the plants at application. A single application of glyphosate at 430, 630 or 840 g ha’1 in
glyphosate-resistant corn did not control yellow nutsedge (Fischer and Harvey 1998).
Yellow nutsedge required at least 2200 g ha'1 of glyphosate for control, but parent tubers
were not killed (0 to 3%) with 1100 or 2200 g ha'l application regimes (Stoller et a1.
1975). However, Appleby and Paller (1978) reduced the number of tubers produced per
plant and increased the number of nonviable tubers as the rate of glyphosate increased
from 300 to 2200 g ha". Poor yellow nutsedge control with glyphosate has been related
to slow absorption and limited translocation (Periera and Crabtree 1986), plant size at the
time of application (Keeley et al 1985, Pereira and Crabtree 1986), and environmental
conditions (Chase and Appleby 1979).

Studies have evaluated yellow nutsedge control with the ALS-inhibiting herbicides
including the sulfonylurea (Ackley et al. 1996, Derr et a1. 1996, Molin et al. 1999, Reddy
and Bendixin 1988 and 1989), imidazolinone (Ackley et al. 1996, Czarnota and Bingham
1997, Derr and Wilcut 1993, Grichar et a1. 1992, Nadihalli and Bendixen 1988, Richburg
III et al. 1993), triazolopyrimidine sulfonanilide (Askew et al. 1999, Nelson and Renner
1998), and pyrimidinyl thiobenzoate (Ackley et al. 1996, Wilcut et al. 1999) herbicide
families. Several of the ALS-inhibiting herbicides controlled yellow nutsedge when there
was both foliar and soil uptake (Reddy and Bendixen 1989, Richburg IH et a1. 1993,

Vencill et al. 1995, Wilcut 1999). A postemergence application of glyphosate used in

209

 

ll

combination with an ALS-inhibiting herbicide with residual activity may provide consistent
yellow nutsedge control.

The interactions between glyphosate, chlorimuron, imazamox, imazaquin, and
halosulfuron were evaluated for purple nutsedge control in greenhouse research (Rao and
Reddy 1999). All tank mixture treatments were additive when applied to purple nutsedge
that was 6 weeks old, but 1120 g ha'1 of glyphosate alone reduced purple nutsedge fresh
weight 100% (Rao and Reddy 1999). Imazaquin and pyrithiobac applied at reduced rates
and tank mixed with glyphosate at 560 g ha‘l antagonized purple nutsedge control (Rao
and Reddy 1999). Antagonistic combinations with glyphosate plus ALS-inhibiting
herbicides have been reported on diflicult-to-control weeds like velvetleaf (A butilon
theophrasti Medicus) (Lich et al. 1997), ivyleaf morningglory [Ipomoea hederacea (L.)
Jacq.] (Lich et al. 1997), purple nutsedge (Rao and Reddy 1999), and sicklepod (Cassia
obtusifolia L.) (Rao and Reddy 1999). However, Starke and Oliver (1998) reported that
chlorimuron did not affect the absorption and translocation of l‘C-glyphosate by velvetleaf
or pitted morningglory (Ipomoea lacunosa L), but glyphosate increased the absorption of
l4C-chlorimuron or 1‘C-imazethapyr by velvetleaf. ALS-inhibiting herbicides and
glyphosate are translocated to actively growing tissue where they inhibit amino acid
synthesis. Since both herbicides are translocated, these herbicide tank mixtures could
provide greater yellow nutsedge control than either herbicide applied alone.

Additives may also increase yellow nutsedge control with glyphosate. The
addition of ammonium sulfate from 1.25 to 10 kg ha'1 and other ammonium salts to

glyphosate increased purple nutsedge control four fold (Suwunnamek and Parker 1975).

210

However, yellow and purple nutsedge control was not increased by adding ammonium
sulfate (Fischer and Harvey 1998) or organosilicone surfactants (Bariuan et al. 1999) to
glyphosate (formulated as Roundup Ultra®)z‘, respectively. Similarly, the addition of crop
oil concentrate, methylated seed oil, or nonionic surfactant to glyphosate did not affect
control (Nelson and Renner 1999).

Yellow nutsedge control depends on reducing the propagative potential of this
weed. This research was initiated to evaluate yellow nutsedge growth suppression and
tuber production following applications of glyphosate, ALS-inhibiting herbicides, and tank
mixtures of glyphosate with the ALS-inhibiting herbicides. In addition, the growth rate of
yellow nutsedge during the season was determined. Rapid growth rate of a perennial
weed like yellow nutsedge early in the season may result in a competitive advantage to this
weed, and understanding yellow nutsedge growth will improve the timing of weed
management practices.

MATERIALS AND METHODS
Greenhouse Methods.

Locally collected tubers were stored at 4 C. Tubers were germinated in a F reas
815 Incubator25 and two sprouted tubers were planted in 4-L pots of Spinks loamy sand
(mixed, mesic Psammentic Hapludalfs) with pH 5.2 and 1.1% organic matter. Plants were

grown in a 16-h photoperiod of natural and supplemental sodium vapor lighting that

 

24Roundup Ultra, Monsanto Co., 800 N. Lindbergh Blvd, St. Louis, MO 63167.
25Precision Scientific Co., 3737 West Cortland St., Chicago, IL 60647.

211

provided a photosynthetic photon flux density26 of 120 ME m'2 - s", fertilized with 0.1 g of
water soluble fertilizer (20% N, 20% P202, 20% K20) every two weeks, and surface
irrigated as needed. Environmental conditions were maintained at 27 i 5 C. Yellow
nutsedge had 6 to 8 leaves per shoot and was 13 to 18 cm tall at the time of herbicide
application. Herbicide treatments included glyphosate (formulated as Roundup Ultra) at
840 g ae ha“1 plus spray grade AMS at 20 g L‘1 applied alone and in combination with
ALS-inhibiting herbicides which were applied with recommended adjuvants (nonionic
surfactant27 (NIS) at 0.25% v/v, crop oil concentrate28 (COC) at 1.3% v/v, or methylated
seed oil29 (MSO) at 1.0% v/v) for yellow nutsedge control (Table 1). Herbicide
applications were made with a continuous link-belt sprayer traveling at 1.5 km hr'l and
equipped with an 8001 even flat-fan nozzle30 calibrated to deliver 234 L ha'1 at 193 kPa of
pressure. The soil was moist at the time of herbicide application and soil temperature was
26 i 1 C. The experiment was arranged in a randomized complete block design with four

replications and was repeated in time. Yellow nutsedge control was evaluated on a scale

 

26LI-COR. 4421 Superior Street, Lincoln, NE 68504.

27Nonionic surfactant was Activator-90, a mixture of alkylpolyoxyethylene ethers and free
fatty acids, Loveland Industries Inc., PO. Box 1289, Greeley, CO 80632.

28Crop oil concentrate was Herbimax, paraffinic oil plus emulsifiers plus surfactants,
Loveland Industries Inc., PO. Box 1289, Greeley, CO 80632.

29Methylated seed oil was Sun-It II, methylated seed oil plus surfactants, Agsco, Inc., PO.
Box 13458, Grand Forks, ND 58208-3458.

30Teejet flat-fan tips. Spraying Systems Co., North Avenue and Schmale Road, Wheaton,
IL 60188.

212

from 0 (no growth suppression) to 100 (complete weed death) 4 and 14 WAT. Height,
panicle number, shoot dry weight reduction, tuber number reduction, tuber weight
reduction, and total root fresh weight excluding tubers was evaluated 14 WAT after the
untreated control had flowered and appeared physiologically mature. Shoot dry weight,
root fresh weight, tuber number, and tuber weight reduction were calculated as the
percent reduction of the measured parameter compared to the untreated control.

Field Methods.

Research was conducted in 1997 near Alto, MI and 1998 at East Lansing, MI in
fields with a previous history of heavy, uniform yellow nutsedge infestation. The soil was
a Boyer sandy loam (coarse-loamy, mixed, mesic Typic Hapludalt) with 2.9% organic
matter and pH 6.6 in 1997 and a Marlette sandy loam (fine-loamy, mixed mesic
Glossoboric Hapludalt) with 1.7% organic matter and pH 6.7 in 1998. The field was
cultimulched 8 cm deep May 27 , 1997 and nutsedge began to emerge by June 6. In 1998,
the field was soil-finished 6 cm deep on May 13 and nutsedge emergence began on May
21.

This experiment was arranged as a randomized complete block design with four
replications each year in plots 3 by 6.1 m. Herbicide treatments were the same as
previously described. Yellow nutsedge shoot density was 1550 shoots In2 and shoots
were 10 to 20 cm tall with 4 to 8 leaves at the time of application. Herbicides were
applied on June 26, 1997 and June 23, 1998 with a tractor mounted compressed-air
sprayer traveling 6.3 km h'1 and delivering 178 L ha'1 at 207 kPa equipped with 8003 flat-

fan nozzles. In 1997, treatments were applied at 2100 h with an air temperature 25 C and

213

37% relative humidity. The treatments were applied at 1900 h with an air temperature 33
C and 30% relative humidity in 1998.

Yellow nutsedge height was recorded at frequent intervals from emergence until
flowering and the growth rate was fit to a second order polynomial using the scatter plot
option in Microsofi Excel-953‘. Visual weed control was evaluated at 2, 4, and 8 WAT,
heights at 2, 4 and 6 WAT, and two 30 by 30 cm quadrats were harvested 2, 4, and 8
WAT from each treatment to evaluate shoot density and vegetative biomass.

Initial tuber densities were measured in the field prior to herbicide application by
randomly removing soil cores from each replication. In April (42 WAT) the following
year, eight soil cores were removed with a 10 cm diameter putting green cup cutter to a
20 cm depth. Tubers were separated from the soil with a modified Kenmore 80""2 clothes
washer powered by a hydraulic orbital motor and sorted by hand into groups of small (<5
mm), medium (5-8 mm), and large (>8 mm) tuber diameter. Hard tubers were counted and
weighed for each graded category while decomposing non-viable tubers were soft, hollow,
and were discarded. Approximately thirty randomly sampled hard tubers were selected
from each plot and sprouted in 150 by 15 mm petri dishes33 between moist filter paper in
an incubator to determine the effect of field applied herbicide treatments on tuber sprout

the following year. The tuber was considered sprouted when a bud protruded from the

 

31Excel-95, Microsoft Corporation, One Microsoft Way, Redmond, Washington
98052-63 99.

32Sears, Roebuck, and Co., 3333 Beverly Road, Hoffman Estates, IL 60179.
33’VWR Scientific Products, 800 E. Fabyan Pkwy, Batavia, IL 60510.

214

scales of the tuber (Holt and Orcutt 1996). Tubers were evaluated every 7 d for 21 d for
sprouted tubers which were counted and removed upon evaluation. Percent sprout was
calculated as: sprouted tuber number/30* 100.

An analysis of variance was conducted and percent data for visual control were
transformed to the arc sine prior to the analysis. The transformation did not affect the
conclusions so original data was evaluated. Initial tuber counts were reported as a
reference and were not part of the data analysis. Means were separated using Fisher’s
Protected LSD at p50.05. Linear regression analysis was performed, and lines are
presented with symbols representing predicted values and significance at p50.01.

RESULTS
Greenhouse Studies.

Yellow nutsedge grth was suppressed by glyphosate at 840 g ha'1 14 WAT
(Table 1). There was no difference in yellow nutsedge height, panicle density, shoot dry
weight reduction, tuber number, or tuber weight following the glyphosate treatment
compared to the untreated control. Chlorimuron, imazethapyr/imazapyr, and halosulfuron
controlled yellow nutsedge 77 to 98% and reduced height, panicle density, dry weight,
tuber number, tuber weight and root fresh weight more than glyphosate applied alone.
Tank mixtures of glyphosate with these herbicides did not increase yellow nutsedge
control.

Yellow nutsedge did not form panicles following treatments of chlorimuron,
cloransulam, imazethapyr, imazethapyr/imazapyr, or halosulfirron alone or tank mixed

with glyphosate. A tank mixture of rimsulfuron plus glyphosate did not increase visual

215

control of yellow nutsedge compared with rimsulfiiron, but panicle number decreased.
Reduced panicle formation may reduce seed production although seeds are not considered
a primary mechanism of dispersal and establishment (Thullen and Keeley 1979).

Yellow nutsedge control increased 17 to 38% and tuber density and fresh weight
decreased when glyphosate was tank mixed with cloransulam or imazethapyr compared
with either treatment or glyphosate applied alone. The reduction in tuber density and fresh
weight was greatest by chlorimuron = imazethapyr/imazapyr 2 halosulfiiron 2 pyrithiobac
2 imazethapyr = rimsulfuron _>_ cloransulam = glyphosate 2 imazamox. Our results are
contrary to greenhouse research by Ackley et al. (1996) where halosulfuron suppressed
yellow nutsedge growth more than chlorimuron. We applied chlorimuron at 12 g ha’l
while Ackley et al. (1996) evaluated 9 g ha’1 of chlorimuron. Imazamox stimulated the
number of tubers produced by yellow nutsedge compared to the untreated control;
however, most of these tubers were smaller since total tuber weight was similar to the
untreated control.

Field Studies.

Glyphosate provided 53% yellow nutsedge control 56 DAT (Table 2).
Chlorimuron and halosulfuron controlled yellow nutsedge 86 and 97%, respectively.
Halosulfiiron provided 23% greater yellow nutsedge control compared to
imazethapyr/imazapyr. Tank mixtures of chlorimuron, halosulfirron, or
imazethapyr/imazapyr with glyphosate did not increase yellow nutsedge control, or reduce
shoot density, dry weight, or height as compared to the ALS-inhibiting herbicides applied

alone. Tank mixtures of glyphosate with cloransulam, imazamox, imazethapyr,

216

rimsulfuron, or pyrithiobac increased visual yellow nutsedge control by 15 to 30%
compared to either herbicide applied alone. Similarly, tank mixtures of glyphosate with
cloransulam, imazamox, imazethapyr, or rimsulfuron reduced yellow nutsedge shoot
density, dry weight, and height more than the ALS-inhibiting herbicide applied alone.
Yellow nutsedge control was greater with imazethapyr/imazapyr compared to
imazethapyr, while imazethapyr controlled yellow nutsedge and reduced shoot density, dry
weight, and height more than imazamox applied alone. Imazamox stimulated shoot
production 180% compared to the untreated control. Halosulfuron and pyrithiobac
reduced tuber sprouting by 19% compared to the untreated control. In turfgrass research,
halosulfuron had no effect on tuber density or weight, but reduced tuber viability with two
applications (Molin et al. 1999). As the rate of halosulfuron increased, tuber viability was
reduced (Molin et al. 1999). The reduction in tuber sprouting in this research could be
due to the effects of the herbicide (Molin et al. 1999, Reddy and Bendixen 1989) or the
presence of dormant tubers with reduced germination (Stoller and Wax 1973, Thullen and
Keeley 1975).

The initial tuber density and fresh weight in June prior to herbicide application was
2320 tubers rn'2 and 328 g m‘z, respectively (Figure 1a and 1b). There was no difference
in tuber density between ALS-inhibiting herbicide treatments compared to tank mixtures
of these herbicides with glyphosate except rimsulfirron plus glyphosate which reduced
tuber density more than rimsulfuron applied alone; therefore, only data for glyphosate and
the ALS-herbicides applied alone were presented. A 220% increase in tuber density was

observed in the untreated control after one year. Halosulfuron caused the greatest

217

reduction in tuber density and fresh weight of the herbicides evaluated, which was similar
to chlorimuron, imazethapyr, and imazethapyr/imazapyr, The tuber density and fresh
weight with glyphosate was the closest to the initial tuber density (23 20 tubers m'z);
therefore, comparisons with glyphosate were pertinent. Chlorimuron and halosulfiiron
reduced tuber density 64 and 79%, and fresh weight 74 and 71%, respectively, compared
to glyphosate applied alone. Tuber density was reduced by all herbicide treatments 33 to
90% except imazamox and rimsulfuron applied alone compared to the untreated control,
and tuber fresh weight was reduced 45 to 91% by all treatments compared to the
untreated control.

All treatments had small, medium, and large tubers in the plots. Small tubers (157
mg) have a shorter life span compared to large (662 mg) tubers which is important for
evaluating the longevity of tubers in the soil (Thullen and Keeley 1975); however, no
differences in germination between tubers with different sizes have been reported (Thullen
and Keeley 1975, Stoller et a1. 1972).

A linear relationship between visual control at 8 WAT and measured tuber density
the following year (42 WAT) was observed (P = 0.0001) (Figure 2). Tuber density was
reduced 625 tubers m‘2 for every 10% increase in visual control. Yellow nutsedge control
by the herbicides evaluated was ranked halosulfuron and chlorimuron (> 80% control and
reduction in tuber density); imazethapyr/imazapyr, imazethapyr, and glyphosate (50 to
80% control and reduction in tuber density); and cloransulam, rimsulfiiron, and imazamox
(20 to 50% control and reduction in tuber density). However, pyrithiobac controlled

yellow nutsedge 48% and reduced tuber density 60%.

218

The response of tuber fresh weight to the herbicide treatments was similar to the
tuber density data; therefore, tuber fresh weight and density were correlated to provide
researchers with a simple method to estimate tuber density from the tuber fresh weight.
For every tuber fresh weight g In2 measured, approximately eight tubers m‘2 were present
(Figure 3). Keeley and Thullen (1983) reported that yellow nutsedge tuber number (y)
was directly related to the number of shoots (x) at harvest with the equation y = 15.68x -
57.1. This equation overestimated the observed tuber density based on the shoot density 8
WAT in our research.

Yellow Nutsedge Growth.

Yellow nutsedge height for 1997 and 1998 was fit to a second order polynomial
curve (Figure 4). Yellow nutsedge was 15 to 20 cm tall by the third week in June which
would be approximately 4 to 5 weeks after corn and soybean planting in the Midwest. In
glyphosate-resistant corn and soybean, glyphosate is applied to weeds that are 10 and 20
cm tall, respectively, to avoid yield loss (Dalley et al. 1998, Gower et al. 1998, Horak et
a1. 1998, Levkulich, et al. 1998, Loux et al. 1998). This is usually 4 to 5 weeks after
planting for corn and soybean, respectively. The critical period for yellow nutsedge
control may be prior to 4 to 5 weeks after planting because of the rapid early growth of
this weed.

DISCUSSION

Yellow nutsedge is competitive both above and below ground, and is considered to

be a more aggressive perennial weed than purple nutsedge or johnsongrass (Holt and

Orcutt 1991). Corn yield was reduced 8% for every 100 shoots of yellow nutsedge m'2

219

(Stoller et al. 1979) while cotton seed yield was reduced 18 kg ha’1 for every shoot
measured (Patterson et al. 1980). At least a 2 to 6 week yellow nutsedge-free period
during crop establishment is needed to have no effect on cotton yield (Keeley and Thullen
1983)

Yellow nutsedge control in the greenhouse with halosulfiiron was less than the
control observed in the field. In contrast, control with imazethapyr/imazapyr was greater
in the greenhouse than in the field. Differences could be due to ample water supply and
herbicide uptake in the greenhouse for the imidazolinone herbicides. Imazamox stimulated
the number of tubers produced compared to the untreated control in the greenhouse, but
had no effect on tuber density in the field. This may be due to the longer period of growth
in the greenhouse compared to the field or plant grth regulation symptoms exhibited by
some imidazolinone herbicides (Bhalla and Shehata 1991).

Control of yellow nutsedge was observed when glyphosate was tank mixed with
several ALS-inhibiting herbicides. Reduced above ground yellow nutsedge growth is
important for limiting yellow nutsedge competition for light and moisture with a crop.
However, tuber production did not decrease compared with all of the ALS-inhibiting
herbicides except rimsulfuron applied alone. Tank mixing glyphosate with imazethapyr,
imazamox, cloransulam, or pyrithiobac would be beneficial to reduce aboveground yellow
nutsedge competition with the crop. However, for the long-term management of yellow
nutsedge, tank mixtures with glyphosate may not reduce the net tuber density in the soil

compared to the ALS-inhibiting herbicide applied alone.

220

Several cultural methods including crop selection and population, planting date,
rotations, cultivation, fertility, and weed-free periods may be adopted to manage yellow
nutsedge (Ghafar and Watson 1983a and 1983b, Glaze 1987, Jordan-Molero and Stoller
1978, Keeley 1987). Crop selection and population are key factors in suppressing yellow
nutsedge (Neeser et al. 1997, Johnson, 111 and Mullinix, Jr. 1997, Keeley 1987, Keeley
and Thullen 1978, Keeley et al. 1979). Yellow nutsedge compensates for low light by
increasing shoot growth and reducing root and tuber production (Keeley and Thullen
1978, Santos et al. 1997). A competitive crop would provide additional shading benefits
when combined with herbicide treatments that suppress yellow nutsedge and reduce tuber
production (Keeley and Thullen 1978, Lotz et al. 1991, Neeser et al. 1997).

In soybean, chlorimuron at 12 g ha’l may reduce total tuber density and fresh
weight m'2 and would be the treatment of choice followed by imazethapyr, cloransulam,
and imazamox. In glyphosate-resistant soybean, glyphosate could be tank mixed for broad
spectrum weed control without reducing yellow nutsedge control. Cloransulam and
imazamox suppress yellow nutsedge only, and tuber density was not affected over time.

In corn, halosulfirron at 35 g ha" would control yellow nutsedge and stop tuber
production. Imazethapyr or imazethapyr/imazapyr could be applied to IMI corn. These
two treatments reduced tuber density and fresh weight 42 WAT similar to halosulfiiron,
but above ground control indicated halosulfirron provided more control.

Rimsulfuron was applied at the postemergence rate for weed control in potatoes
(Renner and Powell 1998) which was one-half the rate applied by Ackley et al. (1996) and

approximately 1.3 times the amount applied in premixtures used for weed control in com.

221

Rimsulfuron did not control yellow nutsedge, but glyphosate plus rimsulfiiron reduced
yellow nutsedge tuber density and fresh weight compared to rimsulfuron applied alone.

Pyrithiobac is labeled for postemergence weed control in cotton. The addition of
pyrithiobac at 70 g ha'l to glyphosate at 840 g ha" reduced tuber fresh weight m‘2 and
improved control compared to glyphosate alone. Yellow nutsedge control was greater
with 70 g ha’l compared to 35 g ha'1 in previous research (Ackley et al. 1996, Wilcut
1999)

The use of preplant incorporated or preemergence herbicides that have yellow
nutsedge control or suppression such as sulfentrazone (Dayan et al. 1996, Wehtje et al.
1997), alachlor (Armstrong et al. 1973, Banks 1983), or metolachlor (Dixon and Stoller
1982, Grichar et al. 1996) followed by postemergence herbicide applications evaluated in
this research may increase yellow nutsedge control and reduce yellow nutsedge
competitiveness prior to a postemergence herbicide treatment. Sequential applications of
glyphosate at 840 g ha’l may improve visual yellow nutsedge control when used in
glyphosate-resistant crops (Fischer and Harvey 1998). This research allows
recommendations for yellow nutsedge control to be based not only on the vegetative
symptoms above ground, but also on tuber development and the asexual reproductive
potential of this problematic weed. Increasing our understanding of tuber dynamics will

improve our recommendations for long-term yellow nutsedge control.

222

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223

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224

 

 

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225

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226

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227

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228

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Herbicide treatment

Figure 1. Yellow nutsedge tuber density and flesh weight following glyphosate plus
AMS, ALS-inhibiting, and tank mixtures of glyphosate and ALS-inhibiting herbicide
treatments in 1997 and 1998. The dotted line indicates the initial tuber density and fresh
weight prior to herbicide applications. The solid bars represent small (<5 mm), horizontal
lines represent medium (5-8 mm), and vertical lines represent large (>8 mm) tubers in the
histogram. The vertical line indicates the LSD at pSODS. Additives included nonionic
surfactant (NIS) at 0.25% v/v, crop oil concentrate (COC) at 1.3% v/v, methylated seed
oil (MSO) at 1.0% v/v, and ammonium sulfate (AMS) at 20 g L".

231

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Tuber density (no. m'z)

Figure 2. Correlation of yellow nutsedge tuber density (x) in the soil 42 weeks after
herbicide treatment at recommended rates with observed visual control (y) of above
ground plants averaged over 1997 and 1998.

232

 

1 4000

or" 12000
E 0 0
° 10000 e
0

 

 

 

 

 

 

 

 

 

 

O
5
E. o / -7 58x +213
3 00 y— .
g .2.» r2=0.88“ _—
I... .
.8
r3
1000 1500 2000
Thher fresh weight (g m")

Figure 3. Correlation of yellow nutsedge tuber fresh weight (x) with tuber density (y)
averaged over 1997 and 1998.

233

Maximum height

 

 

   
 

 

 

 

 

50 ¢

40 --------------------------------------------------------------------------------
E
o 30 ------------------------------------------------------------------------------
V Emergence
E 20 __________________________________________________ y=-O.0033x2+l.71x-179 __
3250 r 2 = 0.77
: Tillage

10 ~I ---------------------------------------------------------------------------

O T l T I l I 1

Julian calendar day

Figure 4. Yellow nutsedge shoot height analysis in the field in 1997 and 1998.

234

CHAPTER 7

YELLOW NUTSEDGE (Cyperus esculentus) CONTROL AND TUBER
YIELD WITH GLYPHOSATE AND GLUFOSINATE

Abstract: Greenhouse and field research was conducted to evaluate yellow nutsedge
control and the effects on tuber production with glufosinate, glyphosate, and glyphosate
plus additional adjuvant. Yellow nutsedge dry weight reduction with glufosinate or
glyphosate was not affected by spray volumes ranging from 140 to 1038 L ha". The
addition of ammonium sulfate (AMS) to glyphosate and glufosinate reduced yellow
nutsedge dry weight 19% more than the herbicides alone. Glyphosate at 0.51 kg ae ha‘1
and glufosinate at 0.59 kg ae ha’l reduced yellow nutsedge grth by 50% in the
greenhouse. Glyphosate at 0.84 kg ha'1 controlled yellow nutsedge 29 to 53% and
glufosinate at 0.4 kg ha" controlled yellow nutsedge 16 to 19% in the field and
greenhouse. Glufosinate at 0.4 kg ha'1 reduced tuber density and fresh weight in the
greenhouse compared to untreated plants. Glyphosate at 0.84 kg ha'1 reduced yellow
nutsedge tuber density and sprouting in the field and tuber fresh weight in the field and
greenhouse compared to untreated plants. The addition of a nonionic surfactant,
methylated seed oil, or crop oil concentrate to glyphosate plus AMS did not increase the
effectiveness of glyphosate in the greenhouse or field.

Nomenclature: glufosinate, 2-amino-4-(hydroxymethylphosphinyl)butanoic acid;

glyphosate, N-(phosphonomethyl)glycine; yellow nutsedge, C yperus esculentus L. #34
CYPES.

 

34Letters following this symbol are a WSSA-approved computer code from Composite
List of Weeds, Revised 1989. Available from WSSA, 1508 West University Ave,
Champaign, IL 61821-3133.

235

Additional index words: ammonium sulfate, crop oil concentrate, dose-response,
methylated seed oil, nonionic surfactant, perennial weed, spray volume, and tuber
production.

Abbreviations: AMS, ammonium sulfate ((NH4)ZSO4); COC, crop oil concentrate; DAT,
d after treatment; GRSO, rate causing 50% growth reduction; and NIS, nonionic surfactant.

INTRODUCTION

Yellow nutsedge is a problematic perennial weed found throughout the world in
crop production fields (Bendixen and Nandihalli 1987, Lapham 1985, Schippers et al.
1995). Yellow nutsedge produces tubers at the terminal end of the rhizome which are
primarily responsible for the propagation and spread of this weed (Bendixen 1973, Wills et
al. 1980). The introduction of glyphosate- and glufosinate-resistant crop varieties has
provided producers with opportunities for non-selective weed control and additional
options for perennial weed management. Glyphosate has been evaluated as a non-
selective herbicide for control of numerous perennial weeds including foxtail barley
(Hordeumjubatum L.) (Donald 1988), bermudagrass [Cynodon daciylon (L.) Pers.]
(Jordan 1977), johnsongrass [Sorghum halepense (L.) Pers.] (McKinley et al. 1999,
Salisbury et al. 1991), hemp dogbane (Apocynum cannabinum L.) (Schultz and Burnside
1980, Wyrill, III and Burnside 1977), Canada thistle [Cirsium arvense (L.) Scop]
(Darwent et a1. 1994), quackgrass [Elytrigia repens (1..) Nevski] (Devine et al. 1983,
Claus and Behrens 1976), purple nutsedge (Cwerus rotundus L.) (Bariuan et al. 1999,
Villaneuva et al. 1985, Zandstra et al. 1974, Zandstra and Nishimoto 1977), and yellow
nutsedge (Cyperus esculentus L.) (Appleby and Paller 197 8, Periera and Crabtree 1986,

Stoller et al. 1975).

236

I
ll

 

 

Yellow nutsedge, however, is difficult to control with glyphosate. The glyphosate
application rate (Appleby and Paller 1978, Pereira and Crabtree 1986, Stoller et a1. 1975,
Villaneuva et al. 1985) and plant age at the time of application (Appleby and Paller 1978,
Keeley et al. 1985, Stoller et al. 1975) has influenced yellow nutsedge control. Regrth
of yellow nutsedge following an application of glyphosate at 1.1 to 2.2 kg ha'1 (Pereiera
and Crabtree 1986), and conversely, control from glyphosate at 1.1 kg ha‘1 or more (Derr
and Wilcult 1993, Stoller et al. 1975) have been reported. Glyphosate reduced tuber
production (Pereira and Crabtree 1986), germination (Stoller et al. 1975), and fresh
weight (Pereira and Crabtree 1986), and increased occurrence of nonviable tubers
(Appleby and Paller 1978). In a fallow cropping system, glyphosate at 2.24 kg ha'1 did
not reduce shoot densities during the fallow period, but shoot number decreased by 95%
the following year (Keeley et al. 1979).

Application variables such as spray volume (Buhler and Burnside 1983, Buhler and
Burnside 1987, Jordan 1977, Krausz et a1. 1996, Stahlman and Phillips 1979), the addition
of ammonium sulfate (Bruce and Kells 1990, Donald 1988, Jordan et al. 1997, Salisbury et
al. 1991), and the use of additional adjuvant (Buhler and Burnside 1983, Wyrill, HI and
Burnside 1977) increased weed control with glyphosate. However, none of these factors
have been investigated to determine their influence on yellow nutsedge control with
glyphosate. Additional adjuvant is known to increase herbicide absorption (Thompson et
al. 1996, Young and Hart 1998) and weed control with other herbicides under water-
stressed conditions (Levene and Owen 1995, Wanamarta and Penner 1989). The cuticle is

the major limiting barrier for herbicide absorption under such conditions (Hess 1985), and

237

yellow nutsedge has a thick waxy cuticle on the adaxial leaf surface (Wills et al. 1980).
Since adjuvants increase efficacy under hot, dry conditions then additional adjuvant may
help increase yellow nutsedge control with glyphosate. In addition, several premixtures of
glyphosate and other herbicides are currently marketed. Additional adjuvant for a second
herbicide may be necessary when glyphosate is tank mixed with other herbicides to
increase control of weeds like yellow nutsedge. However, additional surfactants may also
reduce the uptake of glyphosate in grass species (Gaskin and Stevens 1993). Therefore,
the effect of additional adjuvant on yellow nutsedge control with glyphosate needs to be
investigated.

Research with glufosinate has focused on annual weed control (Higgins et al.
1991, Krausz et al. 1999, Lanie et al. 1994, Steckel et al. 1997a, Steckel et al 1997b,
Tharp et al. 1999, Wilson et al. 1985). Several studies have compared annual weed
control between glufosinate and glyphosate to determine efficacy differences (Blackshaw
1989, Bruce and Kells 1990, Lanie et al. 1994, Higgins et al. 1991, Tharp et al. 1999,
Wilson et al. 1985). The relative sensitivity of bamyardgrass [Echinochloa crus-galli (L.)
Beauv.], common ragweed (Ambrosia artemisiifolia L.), fall panicum (Panicum
dichotomiflorum Michx.), giant foxtail (Setariafaberi Herrm.), and large crabgrass
[Digitaria sanguinalis (L.) Scop.] (Tharp et al. 1999) to glyphosate and glufosinate was
similar. However, research on perennial weed control with glufosinate is limited (Donn
1982), and no research has compared perennial weed control with glyphosate and

glufosinate or the effect of glufosinate on yellow nutsedge tuber production.

238

The objectives of this research were to determine: 1) the effect of spray volume
and AMS on yellow nutsedge control with glyphosate and glufosinate, 2) the effect of
glyphosate and glufosinate on yellow nutsedge control and tuber yield, 3) if additional
adjuvant with glyphosate plus AMS affected yellow nutsedge control, and 4) the
sensitivity of yellow nutsedge to glyphosate and glufosinate.

MATERIALS AND METHODS
Greenhouse Methods. Locally collected tubers were planted at in 910 ml plastic pots in
BACCTO potting soil”. A single tuber was planted in each pot and plants were fertilized
with 0.1 g of water soluble fertilizer (20% N, 20% P202, 20% K20) in 50 ml of water
every two weeks and surface irrigated as needed. Plants were grown in a greenhouse with
a 16-h photoperiod of natural and supplemental sodium vapor lighting which provided a
photosynthetic photon flux density"5 of 120 11E m’2 - s". These cultural methods were
similar for all research unless described otherwise.
Effect of Spray Volume on Yellow Nutsedge Control with Glyphosate and Glufosinate
with and without AMS. Glyphosate (formulated as Roundup Ultra”) at 0.84 kg ha'1 and
glufosinate at 0.4 kg ha"1 were applied at four spray volumes in the absence and presence
of AMS at 20 g L‘1 to yellow nutsedge 15 cm in height with six leaves. Herbicides were
applied at 1230 h with a traveling-belt sprayer traveling at 1.5 km h'1 with 152 kPa. Spray
volume was changed using 8001, 8002, 8004, and 8008 flat-fan nozzle spray nozzles at

the same pressure to attain an output of 140, 271, 542, and 1038 L ha", respectively.

 

35BACCTO is a product of Michigan Peat Co. Houston, TX 77098.
36LI-COR. 4421 Superior Street, Lincoln, NE 68504.

239

Aboveground yellow nutsedge shoot tissue was harvested 28 d after treatment (DAT),
oven-dried, and the percent dry weight reduction compared to the non-treated control was
calculated. The experiment was analyzed as a three factor randomized complete block
design with four replications repeated twice. The factors were herbicide treatment
(glyphosate and glufosinate), the presence or absence of AMS, and spray volume. Data
were subjected to an analysis of variance, means separated using Fisher’s Protected LSD
at p50.05, and main effects for each factor were presented since no interactions were
observed.

Herbicide Injection Experiment. Yellow nutsedge plants were grown in potting soil as
previously described. Herbicides were injected into plants that were 15 cm tall with six
leaves with a syringe”. A volume of 10 pl of solution injected above the basal bulb was
equivalent to the postemergence application rate previously described. These treatments
were injected above the basal bulb to overcome any effects of the cuticle on absorption.
Untreated plants injected with water were included as controls. This study was arranged
as a randomized complete block design with four replications and was repeated twice.
Height and visual control 28 DAT was recorded and analyzed using an analysis of
variance and means were separated using Fisher’s Protected LSD (pSODS).

Effect of Glufosinate, Glyphosate, and Additional Adjuvant with Glyphosate on Yellow
Nutsedge Control, Tuber Production, and Tuber Sprout. Yellow nutsedge tubers were

sprouted in a Freas 815 Incubator” between two moist filter papers for three (1 at 31 C

 

37Hamilton Company, PO. Box 10030, Reno, Nevada, 89520-0012.
38Precision Scientific Co., 3737 West Cortland St., Chicago, IL 60647.

240

 

 

prior to planting. Two moderately sized (5 to 8 mm) sprouted tubers were transferred to
four-L pots of Spinks loamy sand (sand, mixed, mesic Psammentic Hapludalfs).
Postemergence herbicide treatments included glufosinate at 0.4 kg ha" plus AMS at 20 g
L" and glyphosate (formulated as Roundup Ultra) at 0.84 kg ha" plus AMS at 20 g L"
applied alone and with nonionic surfactant” (NIS) at 0.25% v/v, crop oil concentrate40
(COC) at 1.3% v/v, methylated seed oil‘1 (MSO) at 1.0% v/v. Yellow nutsedge was 15
cm tall with six leaves at the time of application. Herbicides were applied at 1800 h with a
traveling-belt sprayer traveling at 1.5 km h", and delivering 234 L ha" at 193 kPa, and
equipped with an 8001 even flat-fan nozzle. Plants were visually rated on a scale of 0 (no
visual injury or plant suppression) to 100% (complete plant death) control four and 14
WAT. Plants were harvested after the untreated control had flowered. Plant height,
aboveground dry weight, root fresh weight, tuber density and tuber fresh weight were
measured 14 WAT. Percent dry weight reduction was calculated as 100*[1-(treated plant
dry weight/untreated plant dry weight)]. This research was arranged as a randomized
complete block design with four replications and was repeated twice. Data were subjected
to an analysis of variance and means were separated using Fisher’s Protected LSD

(1350.05).

 

39Nonionic surfactant was Activator-90, a mixture of alkyl polyoxyethylene ether and free
fatty acids, Loveland Industries Inc, PO. Box 1289, Greeley, CO 80632.

40Crop oil concentrate was Herbimax, 83% petroleum oil and 17% surfactant, Loveland
Industries Inc., PO. Box 1289, Greeley, CO 80632.

“Methylated seed oil was SunIt-II, methylated seed oil and emulsifiers, Agsco, Inc, PO.
Box 13458, Grand Forks, ND 58208-3458.

241

Dose-Response of Glyphosate and Glufosinate. A dose-response comparison of
glyphosate and glufosinate applied at 0, 0.02, 0.08, 0.35, 1.4, 5.6, and 22.4 kg ae ha" was
arranged in a completely randomized design with five replications and was repeated three
times. Glyphosate was formulated as Roundup Ultra”. All treatments included 20 g L"
spray-grade ammonium sulfate. Herbicides were applied at 1200 h with a traveling-belt
sprayer traveling at 1.5 km h", delivering 234 L ha" at 193 kPa, and equipped with an
800143 even flat-fan nozzle. Yellow nutsedge was 15 cm tall with six leaves at the time of
application. Aboveground plant shoot tissue was harvested 28 DAT, oven-dried, and the
dry weight reduction as a percent of the untreated control was calculated. Data were
subjected to a nonlinear regression analysis using a log-logistic dose-response model as
previously described by Schabenberger et al. (1999) and Tharp et al. (1999).

Field Methods. Research was conducted in 1997 near Alto, MI and 1998 at East
Lansing, MI in fields with a previous history of heavy yellow nutsedge infestation. The
soil was a Boyer sandy loam (coarse-loamy, mixed, mesic Typic Hapludalf) with 2.9%
organic matter and pH 6.6 in 1997 and a Marlette sandy loam (fine-loamy, mixed mesic
Glossoboric Hapludalf) with 1.7% organic matter and pH 6.7 in 1998. The fields were
cultimulched eight cm deep on May 27, 1997 and soil-finished six cm deep on May 13,

1998.

 

42Roundup Ultra“, isopropylamine salt of glyphosate with surfactant. Monsanto
Agricultural Company, St. Louis, MO 63167.

43Teejet flat-fan tips. Spraying Systems Co., North Avenue and Schmale Road, Wheaton,
IL 60188.

242

Herbicide treatments included glyphosate (formulated as Roundup Ultra) at 0.84
kg ha" plus AMS at 20 g L" applied alone and with NIS at 0.25% v/v, COC at 1.3% v/v,
MSO at 1.0% v/v. Glufosinate was also applied at 0.4 kg ha" with AMS at 20 g L".
Yellow nutsedge plants were 10 to 20 cm tall with four to eight leaves at the time of
application in 1997 and 1998. Herbicides were applied on June 26, 1997 and June 23,
1998 with a tractor-mounted compressed-air sprayer traveling at 6.3 km h", delivering
178 L ha" at 207 kPa, and equipped with 8003 flat-fan nozzles. In 1997, treatments were
applied at 2100 h, with an air temperature of 25 C and 37% relative humidity. The
treatments were applied at 1930 h, with an air temperature of 33 C and 30% relative
humidity in 1998.

Visual weed control was evaluated at 2, 4, and 8 WAT, heights were measured at
2, 4 and 6 WAT, and two 30 by 30 cm quadrats were harvested 8 WAT from each plot
and oven-dried. Initial tuber densities were measured fi'om each replication in the field
prior to herbicide application. Initial tuber density averaged over both years was 2320
tubers m'2 with a fresh weight of 328 g m". Eight soil cores were randomly sampled from
each plot in April (42 WAT) of the following year with a 10 cm diameter golf cup cutter
to a 20 cm depth. Tubers were separated from the soil with a custom-modified Kenmore
80‘“ clothes washer basin. Tubers were counted and weighed for each treatment. Thirty
hard tubers were randomly sampled from each treatment. The tubers were arranged

between two pieces of moist filter paper and were placed in 150 by 15 mm petri dishes45

 

“Sears, Roebuck and Co., 3333 Beverly Road, Hoffman Estates, IL 60179.
”VWR Scientific Products, 800 E. Fabyan Pkwy, Batavia, IL 60510.

243

which were transferred to an incubator set at 30 C to determine the effect of herbicide
treatments on tuber sprouting the following year. Tubers were evaluated for sprouting
weekly up to 21 d after placement in the incubator. Sprouted tubers were counted and
removed upon evaluation. A tuber was considered sprouted when a bud protruded from
the scales of the tuber (Holt and Orcutt 1996).

The experiment was arranged as a randomized complete block design with four
replications each year in plots 3 by 6.1 01. Data were subjected to an analysis of variance
and percent visual control data were transformed to the arcsine. Field and greenhouse
visual percent data were transformed to the arcsine square root; however, transformation
did not affect data interpretation and untransformed data are presented. Visual control
data were subjected to an F Max test for homogeneity (Kuehl 1994) and data were
combined over location since variances for both locations were homogenous. Means were
separated using Fisher’s Protected LSD at p50.05.

RESULTS AND DISCUSSION
Effect of Spray Volume and AMS on Yellow Nutsedge Control with Glyphosate and
Glufosinate. Glyphosate at 0.84 kg ha" reduced yellow nutsedge dry weight 64%
whereas glufosinate at 0.4 kg ha" reduced dry weight only 22% when averaged over AM S
and spray volume treatments (Table l). The addition of AMS to the herbicide containing
spray solution reduced yellow nutsedge dry weight an additional 19% compared with no
AMS. Increased control of other weed species including prickly sida (Sida spinosa L.)
(Jordan et al. 1997), entireleaf morningglory (Ipomoea hederacea var. intergriuscula)

(Jordan et al. 1997), foxtail barley (Donald 1988), johnsongrass (Salisbury et al. 1991),

244

and purple nutsedge (Suwunnamek and Parker 1975) was reported when AMS was added
to glyphosate. Additionally, there was three times less regrth of three and six week old
purple nutsedge treated with glyphosate plus AMS compared with no AMS (Suwunnamek
and Parker 1975). In contrast, yellow nutsedge control did not increase when AMS was
added to glyphosate in glyphosate-resistant corn research (Fischer and Harvey 1998).
However, AMS increased annual weed control with glufosinate applied at 0.125 to 0.25
kg ha" (Blackshaw 1989). The increased control of yellow nutsedge with AMS compared
to no AMS in our research illustrates the importance of this spray additive to increase
yellow nutsedge suppression with either glyphosate or glufosinate.

Spray volume did not affect yellow nutsedge dry weight reduction (Table 1).
Bruce and Kells (1990) similarly reported that spray volume did not influence horseweed
[Conyza canadensis (L.) Cronq.] control. In contrast, others have shown that spray
volume affects control of annual weeds (Buhler and Burnside 1987, Krausz et al. 1996).
An interaction between herbicide and spray volume was expected in our research since
lower spray volumes increased control of annual grasses with glyphosate in other research
(Buhler and Burnside 1987) and higher spray volumes may increase coverage and control
with glufosinate since it is a contact herbicide. However, the leaf morphology of yellow
nutsedge may allow the applied herbicide spray solution to accumulate in the whorl of the
plants when applied as spray volume increases so that there is little affect of spray volume
on dry weight reduction.
Herbicide Injection Experiment. A thick waxy cuticle (1 to 1.3 pm) covers a layer of

epidermal cells on the adaxial leaf surface of yellow nutsedge (Wills et al. 1980).

245

Glyphosate and glufosinate were injected into yellow nutsedge plants above the basal bulb
to reduce the potential effects of the cuticle on absorption. Glufosinate and glyphosate
decreased yellow nutsedge height similarly 28 DAT (Table 2). Glyphosate at 0.84 kg ha"
controlled yellow nutsedge 88% while glufosinate at 0.4 kg ha" provided 68% control.
The increased control with this technique supports our hypothesis that reduced absorption
across the cuticle may be a major limitation for yellow nutsedge control with glufosinate.
Common lambsquarters was the most tolerant weed species to glufosinate in the field
(Steckel et al. 1997b) and greenhouse (Steckel et al. 1997a), and had the lowest
absorption compared to giant foxtail, bamyardgrass, and velvetleaf (Steckel et al. 1997a).
Effect of Glyphosate and Glufosinate on Yellow Nutsedge Control, Tuber
Production, and Sprouting. Glyphosate at 0.84 kg ha" provided greater suppression of
yellow nutsedge compared to glufosinate at 0.4 kg ha", but neither herbicide controlled
yellow nutsedge greater than 53% in the greenhouse or field (Table 3). Glyphosate
control of yellow nutsedge was 13 to 34% greater than glufosinate in the greenhouse and
field. Glufosinate reduced tuber density 19%, and glufosinate and glyphosate reduced
tuber fresh weight 22 and 24%, respectively, compared to untreated plants in the
greenhouse. Glyphosate controlled yellow nutsedge 53% 8 WAT which resulted in a 51%
reduction in tuber density, 59% reduction in tuber fresh weight, and a 17% reduction in
tuber sprouting 42 WAT compared to untreated plants (Tables 3). Tubers that did not
sprout remained hard and turgid (data not presented). Other researchers reported a 43 to
45% reduction in shoot fresh weight 10 DAT and a 27 to 59% reduction in parent tuber

sprouting from a postemergence application of glyphosate at 0.6 kg ha'1 to 2 to 4 week

246

old plants (Keeley et al. 1985). Similarly, the tubers that did not sprout remained firm
(Keeley et al. 1985). A reduction in purple nutsedge viability has also been reported with
glyphosate (Zandstra et al. 1974). However, glyphosate at 1.1 and 2.2 kg ha" did not
affect parent tuber viability in other research (Stoller et al. 1975). The translocation of
glyphosate to yellow nutsedge tubers (Keeley et al. 1985) may explain a reduction in tuber
sprout reported in this research. Glufosinate controlled yellow nutsedge 19% in the field
and had no effect on tuber density, fi'esh weight, or tuber sprouting (Table 3) 42 WAT
compared to untreated plants.

Effect of Additional Adj uvant with Glyphosate on Yellow N utsedge Control, Tuber
Production, and Tuber Sprout. There was no effect of additional adjuvant on yellow
nutsedge control, height, dry weight, tuber density or fresh weight in the greenhouse or
field (Table 4). Yellow nutsedge leaves appeared bleached and chlorotic at the base of the
leaves one week after glyphosate was applied, and new sprouts emerging from the basal
bulb and rhizomes allowed the propagation of this weed in the greenhouse and field.
Buhler and Burnside (1987) increased annual weed control at 190 L ha" with additional
adjuvant compared to lower spray volumes; therefore, an increase in control may be
expected at higher application volumes as used in our research. Similarly, Bariuan et al.
(1999) found that organosilicone surfactants did not affect purple nutsedge control with
glyphosate formulated as Roundup Ultra". Adding an organosilicone surfactant to
glyphosate did not affect absorption or translocation of l4C-glyphosate by purple nutsedge

(Bariuan et al. 1999).

247

All glyphosate treatments had similar tuber sprouting (Table 4) and rotten tubers
present (data not presented), but glyphosate plus AMS had more hard, turgid tubers than
the treatments that included additional adjuvant (data not presented). Glyphosate may act
as a sprout inhibitor since 0.3 to 0.6 kg ha" increased the number of dormant tubers
produced in greenhouse research (Appleby and Paller 1978) and glyphosate was
translocated to parent tubers in other research (Keeley et al. 1985). The adjuvants
evaluated (Table 4) did not increase or decrease herbicide activity of glyphosate. These
adjuvants may not have been the appropriate adjuvants for use with glyphosate, but if used
with glyphosate in various tank mixtures these adjuvants will not have a negative effect on
yellow nutsedge control.

Dose-Response of Glyphosate and Glufosinate. Yellow nutsedge dry weight decreased
as the glyphosate and glufosinate rate increased (Figure 1). GR,O values of 0.51 and 0.59
kg ha" for glyphosate and glufosinate, respectively, were statistically similar between these
non-selective herbicides. Appleby and Paller (1978) reported 52, 97, and 100% yellow
nutsedge control from glyphosate at 0.6, 1.1, and 2.2 kg ha", respectively. Research with
annual weeds has indicated GRso values from 0.063 to 0.235 kg ha" for glufosinate
(Steckel et al. 1997, Tharp et al. 1999) and 0.064 to 0.16 kg ha" for glyphosate (Tharp et
al. 1999). The dose-response curve indicated glufosinate had a narrow rate range (0.27 to
1.05 kg ha") between 90% and 10% of the control while glyphosate had a broad rate
range (0.09 to 2.10 kg ha") (Figure 1). Our GR50 values for yellow nutsedge were equal
to 61% of the glyphosate (0.84 kg ha") (Jordan et al. 1997) and 148% of the glufosinate

(0.4 kg ha") (Steckel et al 1997b) rate applied for annual weed control in glyphosate- or

248

 

 

 

glufosinate-resistant crops. Therefore, yellow nutsedge control in the field would be
greater with glyphosate than glufosinate at rates applied for annual weed control which
confirmed previously conducted field and greenhouse research.

Herbicide rate and growth stage affect annual weed control with glufosinate
(Steckel et al. 1997a and 1997b, Tharp et al. 1999) and yellow nutsedge control with
glyphosate (Jordan et al. 1997, Pereira and Crabtree 1986, Stoller et al. 1975). Our
research only evaluated control of yellow nutsedge at one grth stage. Yellow nutsedge
grows rapidly early in the season (Holt and Orcutt 1996) and control by glufosinate and
glyphosate may differ at other grth stages.

In conclusion, neither glufosinate at 0.4 kg ha" nor glyphosate at 0.84 kg ha"
controlled yellow nutsedge greater than 65%. Similarly, a single application of glyphosate
at 0.43, 0.63 or 0.84 kg ha" in glyphosate-resistant corn did not control yellow nutsedge
(Fischer and Harvey 1998); Glyphosate provided greater aboveground yellow nutsedge
control compared to glufosinate, but treatments had similar tuber production. Dose-
response data (Tharp et al. 1999) and annual weed control comparisons between
glyphosate and glufosinate (Higgins et al. 1991, Lanie et al. 1994) indicate higher rates of
glufosinate may provide more consistent control of annual weeds. An increase in
glufosinate rate may increase yellow nutsedge control. The addition of AMS increased
yellow nutsedge control with glyphosate and glufosinate, but spray volume with either
herbicide or additional adjuvant tested with glyphosate did not affect yellow nutsedge
control. Split applications of these herbicides (Fischer and Harvey 1998, Krausz et al.

1999), tank mixture combinations with other postemergence herbicides that control yellow

249

nutsedge(Ack1ey et al. 1996, Derr et a1. 1996, Derr and Wilcut 1993, Reddy and
Bendixen 198 8), or the use of preplant incorporated or preemergence herbicides
(Armstrong et al. 1973, Dixon and Stoller 1982, Grichar et a1. 1996, Reddy and Bendixen
1989, Wehtje et al. 1997) for yellow nutsedge suppression or control are options that will
need to be utilized to increase yellow nutsedge control and reduce tuber production in
glyphosate or glufosinate-resistant crops. In addition, crop canopy closure may increase
yellow nutsedge suppression by competing for light and other resources and altering
yellow nutsedge growth. Yellow nutsedge compensates for low light levels by increasing
shoot grth and reducing root and tuber production (J ordan-Molero and Stoller 197 8,
Keeley and Thullen 1978, Lotz et al. 1991, Santos et al. 1997) which reduces the
propagative and reproductive potential of this perennial weed. Yellow nutsedge must be
managed in glyphosate- and glufosinate-resistant crops by practices other than a single
application of these herbicides or this weed species could become highly problematic in

these weed management systems.

250

LITERATURE CITED

Ackley, J. A., H. P. Wilson, and T. E. Hines. 1996. Yellow nutsedge (Cyperus
esculentus) control post with acetolactate synthase-inhibiting herbicides. Weed Technol.
102576-580.

Armstrong, T. F., W. F. Meggitt, and D. Penner. 1973. Absorption, translocation and
metabolism of alachlor by yellow nutsedge. Weed Sci. 21:357-360.

Bariuan, J. V., K. N. Reddy, and G. D. Wills. 1999. Glyphosate injury, rainfastness,
absorption, and translocation in purple nutsedge (Cwerus rotundus). Weed Technol.
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255

Table 1. The effect of ammonium sulfate and spray volume on yellow nutsedge dry
weight reduction with glyphosate and glufosinate 28 d after treatment.

 

 

 

 

 

 

 

 

 

Dry weight
reduction
Herbicide treatment %
Glufosinate 0.4 kg ac ha" 22
Glyphosate 0.84 kg ae ha" 64
LSD (1,500,, — 10 —
Ammonium sulfate
0 g L" 34
20 g L" 53
LSD (1,500,) --— 10 —-
Spray volume
140 L ha" 41
271 L ha" 48
542 L ha" 40
1038 L ha" 46
LSD (9:00,) — NS —

 

256

Table 2. Height and control of yellow nutsedge by glyphosate and glufosinate injected
into plants grown in the greenhouse 28 d after treatment.

 

 

Treatment Rate Height Control
kg ae ha" ---— cm —— —— %
Glyphosate 0. 84 10 88
Glufosinate 0 . 4 1 0 68
Untreated 0 25 0
LSD 0,503,) —— 3 -——— 8

 

 

257

Table 3. Yellow nutsedge control, tuber number, fresh weight, and sprouting following
glyphosate and glufosinate in the greenhouse and field in 1997 and 1998.

 

 

 

 

Herbicide treatment Tuber
Greenhouse Researcha Rate Control Shoot density Fresh weight Sprout
kgaeha" —%— —no.m'2— —gm’2-— -%-
Glyphosate + AMSb 0.84 + 20 29 6990 700 —
Glufosinate + AMS 0.4 + 20 16 6320 680 —
Untreated 0 0 7830 900 —
LSD 0,500,) — 8 — — 980 — — 150 - —
Field Research“ Rate Control Shoot density Fresh weight Sproutd
kgaeha" -%— —no.m’2— —gm’2— -%-
Glyphosate + AMS 0.84 + 20 53 2540 320 65
Glufosinate + AMS 0.4 + 20 19 3450 430 83
Untreated 0 0 5 190 770 82
LSD (1,500,, — 3 —- — 2180 - — 370 — — 14 —

 

aGreenhouse research evaluated control, tuber density, and tuber fresh weight 14 WAT
and were repeated in time. AMS was applied at 20 g L".
I’Abbreviations: AMS, ammonium sulfate; WAT, weeks after treatment.
cField research evaluated control 8 WAT, tuber density, and tuber fresh weight 42 WAT.
dData were the percentage of thirty tubers per plot that were hard, rotten, or sprouted
upon visually examination for three weeks after placement in an incubator at 30 C. Tubers
were randomly selected from soil cores sampled to a 20 cm depth 42 WAT.

258

Table 4. Yellow nutsedge control, height, dry weight, tuber density, tuber fresh weight,
and tuber sprouting with glyphosate plus AMS alone and with NIS, COC, and MSO in the
greenhouse and field in 1997 and 1998.

 

 

 

 

 

Tuber
Adjuvant Dry Shoot Fresh
Herbicide treatment‘ rate Control Height weight density weight Sprout
Greenhouse Researchb % v/v — % — - cm - -g m'z- no. rn'2 —g m’2 - - % -
Glyphosate + AMSc 0 29 41 312 6985 695 —
Glyphosate + AMS 0.25 42 35 242 5827 530 —
+ NIS
Glyphosate + AMS 1.3 32 39 316 7863 772 _—
+ COC
Glyphosate + AMS 1.0 35 39 295 7582 786 —
+ MSO
Lsomim -NS- —NS- —NS- -NS- -NS- —
Field Research‘I % v/v — % — - cm - -g m‘z- no. m‘2 -g m'2 - - % -
Glyphosate + AMS 0 53 28 387 2545 314 65
Glyphosate + AMS 0.25 58 26 323 3047 331 81
+ NIS
Glyphosate + AMS 1.3 56 27 258 2474 306 76
+ COC
Glyphosate + AMS 1.0 57 28 301 2192 264 74
+ MSO

 

'All glyphosate treatments were applied at 0.84 kg ae ha" with AMS at 20 g L".
bGreenhouse research evaluated control, height, dry weight, tuber density, and tuber fresh

weight 14 WAT and were repeated in time.
cAbbreviations: AMS, ammonium sulfate; NIS, nonionic surfactant; COC, crop oil

concentrate; MSO, methylated seed oil; and WAT, weeks after treatment.
dField research evaluated control 8 WAT, height 6 WAT, dry weight 8 WAT, tuber
density, and tuber fiesh weight 42 WAT.

259

 

 
    

A100 1 ‘ .
9 90 - .
'E 80 fl ' — Glyphosate+AMS
8 """ Glufosinate+AMS
,_ 7o -
O ..................................... .
60 ‘
\°
3; 50 " I
I
g, 40 ... ........................................ ' ...
'- - I
0;: 30 I
—I ........................................ ru-
z~ 2° .
D 10 ‘ I
o r I I

 

 

 

 

-4 -3 -2 -1 o 1 2 3 4
Herbicide rate (In kg aelha)

Figure 1. Response of yellow nutsedge shoot dry weight to glyphosate (solid log-logistic
curve) and glufosinate (dotted log-logistic curve) plus ammonium sulfate (AMS) at 20 g
L" 4 weeks after treatment. GRso values for glyphosate and glufosinate were 0.51 and
0.59 kg ha", respectively. Vertical lines with arrows indicate the predicted control with
glyphosate at 0.84 kg ha" and glufosinate at 0.4 kg ha".

260

SUMMARY

A postemergence application of bentazon/acifluorfen plus thifensulfirron plus
sethoxydim and lactofen plus bentazon plus clethodim applied at the V5 stage of soybean
development reduced canopy development, delayed reproductive development, and
reduced soybean yield 130 to 270 kg ha" compared to untreated soybean in the absence of
white mold. Yield of soybean cultivars varied by year with a ranking of ‘A1900’ =
‘A2704’ = ‘AG2701’ > ‘AG1901’ in 1997, and ‘A2704’ > ‘AGZ701’ > ‘A1900’ >
‘AG1901 ’ in 1998. In the presence of white mold, near isogenic glyphosate-resistant
(‘S14-M7’, ‘S20—B9’, ‘GL2600’, ‘P93B01’) and non-resistant (‘8 12-49’, ‘S 19-90’,
‘GL2415’, ‘P9281’) soybean cultivars were equally susceptible to white mold. However,
cultivar selection was important for white mold management since ‘S 12-49’, ‘S14-M7’,
‘S 19-90’, and ‘S20-B9’ had a lower incidence of white mold than ‘GL2415’,
‘GL2600’,‘P928 1 ’, ‘P93B01’. Glyphosate did not affect the incidence of white mold in
glyphosate-resistant soybean. A reduction in white mold incidence following an
application of lactofen may be attributed to increased phytoalexin production, reduced
canopy development, and delayed reproductive development. Lactofen may be used as a
tool to help manage white mold, but increased soybean yield may not always occur. Other
protoporphyrinogen oxidase-inhibiting herbicides such as oxyfluorfen, sulfentrazone, and
oxadiazon suppressed Sclerotinia sclerotiorum lesion grth similar to lactofen.
Sulfentrazone and lactofen increased phytoalexin production similarly.

Yellow nutsedge was suppressed by glyphosate at 840 g ae ha" 13 to 34% greater

than glufosinate at 400 g ae ha". Ammonium sulfate increased yellow nutsedge control

261

 

with glyphosate and glufosinate, but the addition of a nonionic surfactant, methylated seed
oil, or crop oil concentrate applied with glyphosate at 840 g ha" plus ammonium sulfate at
20 g L" did not increase yellow nutsedge control. Yellow nutsedge must be managed in
glyphosate- and glufosinate-resistant crops by increased use rates or more than a single
application of these herbicides or this weed species could become highly problematic in
these weed management systems. Tank mixtures of glyphosate with the ALS-inhibiting
herbicides increased visual control of yellow nutsedge, but did not reduce tuber
production compared to the ALS-inhibiting herbicides applied alone. Yellow nutsedge
control by the herbicides evaluated and applied at recommended use rates was ranked
halosulfiiron and chlorimuron (> 80% control and reduction in tuber density);
imazethapyr/imazapyr, imazethapyr, and glyphosate (50 to 80% control and reduction in
tuber density); and cloransulam, rimsulfuron, and imazamox (20 to 50% control and
reduction in tuber density). However, pyrithiobac controlled yellow nutsedge 48% and
reduced tuber density 60%. Long-term yellow nutsedge management may be achieved
with treatments that reduce tuber production. This information will help producers in the
Midwest and Michigan make informed decisions on how weed management programs
impact soybean development, incidence of white mold, soybean yield, and yellow nutsedge

tuber yield.

262