Investigations into Multiple–Herbicide-Resistant Ambrosia artemisiifolia (Common Ragweed) in Ohio and Glyphosate-Resistance Mechanisms

Dissertation

Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy in the Graduate School of The Ohio State University

By

Jason Thomas Parrish

Graduate Program in Horticulture and Crop Science

The Ohio State University

2015

Dissertation Committee:

Dr. Mark M. Loux, Advisor

Dr. S. Kent Harrison

Dr. James D. Metzger

Dr. David M. Mackey

Dr. Anne E. Dorrance

Copyrighted by

Jason Thomas Parrish

2015

Abstract

Common ragweed (Ambrosia artemisiifolia) is a weed problem in many places throughout the world. Though it seldom dominates the landscape, common ragweed seems to be able to exploit diverse habitats. Common ragweed is primarily outcrossing and has a high rate of gene polymorphisms, leading to high genetic diversity. This high level of genetic diversity likely plays a major role in the evolution of herbicide-resistant biotypes. Whole-plant bioassays of herbicide dose-response in the greenhouse were used to characterize resistance levels to glyphosate, cloransulam-methyl, and fomesafen herbicides. Additional studies were conducted to provide insight into potential mechanisms that may contribute to the development of resistance to glyphosate in an

Ohio ragweed biotype, including 5-enolpyruvylshikimate-3-phosphate synthase (EPSPS) gene sequencing, quantitative PCR of the EPSPS gene, EPSPS enzyme immunoblot and activity/inhibition assays, 31P-nuclear magnetic resonance (NMR) studies of glyphosate- treated tissues, and whole-plant absorption and translocation studies using 14C-labeled glyphosate. A single common ragweed population from Clinton County, Ohio exhibited multiple resistance to herbicides at dosages that exceeded the rate required to kill herbicide-sensitive common ragweed biotypes from 4- to 30-fold for glyphosate,

> 1000-fold for cloransulam-methyl, and 14- to > 100-fold for fomesafen. This is the first report of a common ragweed biotype with multiple resistance to herbicides from three site-of-action (SOA) groups. Sequencing data indicated the gene coding for EPSPS has a

ii high mutation rate in all studied common ragweed biotypes, but it typically does not code for an altered amino acid sequence in the glyphosate binding area. Additional studies identified alleles of EPSPS coding for proline-to-serine and proline-to-threonine substitutions at amino acid number 106 (based upon the mature maize EPSPS numbering scheme). Previous studies by other authors have found these amino acid substitutions to confer glyphosate resistance in numerous other species. The alleles containing these mutations were not detected in previous studies of Ohio ragweed populations, and it is not known whether these alleles are translated into a functional EPSPS protein. Direct sequence analysis also suggested that there are six-to-eight or more partial- or full-length copies of the EPSPS gene in a typical diploid (2n) common ragweed plant. An immunoblot assay with common ragweed total soluble protein, as well as Palmer amaranth (Amaranthus palmeri) glyphosate-sensitive and EPSPS overexpressing glyphosate-resistant controls, showed a single plant from the glyphosate-resistant biotype with increased EPSPS expression. Quantitative PCR also showed an increased relative

EPSPS gene copy number in the same plant. 31P-NMR data showed similar uptake of glyphosate into the leaf cells and no vacuolar sequestration in all common ragweed biotypes, with lower sugar-phosphate (including shikimate-3-phosphate) accumulation relative to glyphosate-susceptible common ragweed plants. Similarly, absorption and translocation of 14C-labeled-glyphosate over 48 hours did not differ between resistant and susceptible biotypes. More research will be required to unequivocally determine the molecular basis of glyphosate resistance in common ragweed, but accumulated evidence supports the hypothesis that multiple mechanisms of glyphosate resistance are possible within a common ragweed population.

iii Acknowledgments

This work was completed thanks to the many forms of assistance and support from hundreds of people at OSU, Colorado State University, Monsanto Company,

Washington University in St. Louis, and the University of Illinois, and the love and encouragement of my friends and family outside of these institutions. You know who you are, and I could not have done this without you. Thank you for the financial support and your patience. You shared your time, knowledge, and experience, your workspaces and equipment, and even your food. We spent many hours talking about plants, the weather, sports, and international foods. Thank you for the long lunches where we shared laughter and frustrations. I enjoyed the drives across the state looking for plants or any other adventure we might encounter. Thank you for helping me when I had questions and last-minute requests. I have learned so much from so many of you. As I complete my dissertation, I am left with many debts and many memories.

iv Vita

2002...... Firelands High School

2007...... B.S. Agriculture, The Ohio State University

2008 to 2014 ...... Graduate Research/Teaching Associate,

Department of Horticulture and Crop

Science, The Ohio State University

Fields of Study

Major Field: Horticulture and Crop Science

v Table of Contents

Abstract ...... ii

Acknowledgments...... iv

Vita ...... v

List of Tables ...... x

List of Figures ...... xii

Chapter 1 : Introduction ...... 1

1.1 Common Ragweed ...... 1

1.2 Glyphosate Resistance ...... 4

1.3 Common Ragweed Resistance to Other Herbicides...... 13

1.4 Objectives ...... 15

Chapter 1 References ...... 17

Chapter 2 : Characterization of Common Ragweed Resistance to Glyphosate, Cloransulam-Methyl, and Fomesafen Herbicides...... 29

2.1 Materials and Methods: ...... 29

2.1.1 Development of a sample population ...... 29

2.1.2 Growing Conditions for Dose-Response ...... 31

vi 2.1.3 Treatments ...... 31

2.1.4 Data Collection and Analysis ...... 34

2.2 Results and Discussion ...... 37

2.2.1 Glyphosate dose-response results ...... 37

2.2.2 Cloransulam-methyl dose-response results ...... 38

2.2.3 Fomesafen dose-response results ...... 39

2.2.4 Discussion...... 40

Chapter 2 References ...... 41

Chapter 3 : Common Ragweed Target-Site Glyphosate-Resistance Mechanisms ...... 54

3.1 Materials and Methods ...... 54

3.1.1 Plant Materials ...... 54

3.1.2 Genomic DNA extraction ...... 54

3.1.3 RNA extraction and complementary DNA synthesis ...... 55

3.1.4 PCR primer design ...... 56

3.1.5 Gene sequencing of EPSPS, acetolactate synthase (ALS), and fructan 1-exohydrolase IIa (FEH) ...... 58

3.1.6 EPSPS Enzyme Activity...... 60

3.1.7 EPSPS Enzyme Quantification ...... 63

3.1.8 EPSPS relative genomic copy number determination ...... 65

3.2 Results and Discussion ...... 68

vii 3.2.1 EPSPS gene sequencing ...... 68

3.2.2 EPSPS Enzyme Activity...... 71

3.2.3 EPSPS Enzyme Quantification ...... 72

3.2.4 Real-Time Quantitative PCR ...... 72

Chapter 3 References ...... 74

Chapter 4 : Common Ragweed Non–Target-Site Glyphosate-Resistance Mechanisms . 83

4.1 Materials and Methods ...... 83

4.1.1 Development of a sample population ...... 83

4.1.2 14C-glyphosate uptake and translocation: ...... 85

4.1.3 Statistical analyses ...... 87

4.1.4 In vivo 31P-NMR investigation ...... 87

4.2 Results and Discussion ...... 88

4.2.1 14C-glyphosate uptake and translocation: ...... 88

4.2.2 Discussion...... 90

Chapter 4 References ...... 92

Chapter 5 : Conclusions ...... 97

Chapter 5 References ...... 104

References ...... 105

Appendix A : In vivo 31P-NMR Results ...... 118

viii A.1 31P-NMR observations for source tissue following glyphosate infusion ...... 118

A.2 31P-NMR observations of source tissue following glyphosate spray treatment .. 119

A.3 31P-NMR observations for sink tissue following glyphosate spray treatment .... 120

ix List of Tables

Table 2.1. Estimated glyphosate doses resulting in 50% (GR50) or 90% (GR90) reduction in fresh weight relative to untreated controls and resistance ratios of R1, R2, S1, and S2 common ragweed biotypes in greenhouse dose-response studies...... 42

Table 2.2. Estimated glyphosate doses lethal for 50% (LD50) or 90% (LD90) of R1, R2, S1, and S2 common ragweed biotypes in greenhouse dose-response studies...... 43

Table 2.3. Estimated clorasulam doses resulting in 50% (GR50) or 90% (GR90) reduction in fresh weight relative to untreated controls of R and S2 common ragweed biotypes in a greenhouse dose-response study...... 44

Table 2.4. Estimated cloransulam doses lethal for 50% (LD50) or 90% (LD90) of R and S2 common ragweed biotypes in a greenhouse dose-response study...... 45

Table 2.5. Estimated fomesafen doses resulting in 50% (GR50) or 90% (GR90) reduction in fresh weight relative to untreated controls and resistance ratios of R and S common ragweed biotypes in greenhouse dose-response studies...... 46

Table 2.6. Estimated fomesafen doses lethal for 50% (LD50) or 90% (LD90) of R and S common ragweed biotypes in greenhouse dose-response studies...... 47

Table 3.1. PCR primer pairs used for sequencing and real-time quantitative PCR of common ragweed...... 76

Table 3.2. EPSPS glyphosate-dose–response enzyme activity I50 values and R:S ratios. 77

Table 4.1. Translocation of 14C-glyphosate in common ragweed plants receiving low or high doses of glyphosate...... 94

x Table 4.2. Translocation of 14C-glyphosate in common ragweed plants harvested 8-, 24-, or 48-HAT...... 95

xi List of Figures

Figure 1.1. Schematic of the shikimate (chorismate) pathway and products...... 28

Figure 2.1. Fresh weight response curves of R1, R2, S1, and S2 common ragweed biotypes to varied doses of glyphosate in greenhouse studies...... 48

Figure 2.2. Mortality response curves of R1, R2, S1, and S2 common ragweed biotypes to varied doses of glyphosate in greenhouse studies...... 49

Figure 2.3. Fresh weight response curves of R and S2 common ragweed biotypes to varied doses of cloransulam in a greenhouse study...... 50

Figure 2.4. Mortality response curves of R and S2 common ragweed biotypes to varied doses of cloransulam in a greenhouse study...... 51

Figure 2.5. Fresh weight response curves of R and S common ragweed biotypes to varied doses of fomesafen in greenhouse studies...... 52

Figure 2.6. Mortality response curves of R and S common ragweed biotypes to varied doses of fomesafen in greenhouse studies...... 53

Figure 3.1. Alignment of predicted 5-enolpyruvylshikimate-3-phosphate synthase (EPSPS) amino acid sequences...... 78

Figure 3.2. EPSPS glyphosate-dose–response enzyme activity assay...... 80

Figure 3.3. Relative EPSPS protein abundance determined by a western immunoblot. . 81

Figure 3.4. EPSPS:FEH relative genomic copy number...... 82

Figure 4.1. Absorption of 14C-glyphosate in common ragweed plants over 48-hours. ... 96 xii Figure A.1. 31P-NMR spectra of perfused S and R common ragweed mature leaves following an 8-hour treatment with 10-mM glyphosate...... 124

Figure A.2. 31P-NMR spectra of common ragweed mature (source) leaves 24-hours after spray treatment with 3.36 kg ae ha−1 glyphosate...... 125

Figure A.3. 31P-NMR spectra of common ragweed shielded immature (sink) leaves 24 hours after spray treatment of mature leaves with 3.36 kg ae ha−1 glyphosate...... 126

Figure A.4. 31P-NMR measured cytosolic glyphosate relative to reference (MDP) 24 hours after treatment in common ragweed leaf tissues...... 127

Figure A.5. Photo of mature R4 and S1 common ragweed plants 12-days after spray treatment with 3.36 kg ae ha-1 glyphosate...... 128

xiii Chapter 1: Introduction

1.1 Common Ragweed

Ambrosia artemisiifolia (common ragweed) is a monoecious flowering plant in the tribe Heliantheae of the dicotyledonous plant family Asteraceae (Karis, 1995; Payne et al., 1964). Common ragweed is a diploid, and like most other species in the subtribe

Ambrosiinae, it has a chromosome number based on n = 18; however, common ragweed has reached a larger worldwide distribution than other species in the Ambrosiinae

(Mulligan, 1957; Payne et al., 1964). The center of origin for the Ambrosia genus is reported to be southwestern North America, but common ragweed is more prevalent in eastern North America, and fossilized pollen has been identified in Ontario, Canada, in deposits older than 60,000 years (Bassett and Crompton, 1975; Bassett and Terasmae,

1962; Eom et al., 2013). The range of common ragweed has expanded rapidly with human disturbance to the extent that the time of European settlement can be easily correlated with sharp increases in ragweed pollen observed in sediment cores

(McAndrews, 1988; Munoz and Gajewski, 2010). Common ragweed has now been found throughout North America, in every state except Alaska and a few northern provinces of Canada, and has invaded every continent except Antarctica (Bass et al.,

2000b; Chauvel et al., 2006; Csontos et al., 2010; Gaudeul et al., 2011; Hodgins and

Rieseberg, 2011; Joly et al., 2011; Kil et al., 2004; Kiss and Béres, 2006; Lavoie et al.,

2007; Maryushkina, 1991; Tokarska-Guzik et al., 2011).

1 Common ragweed is capable of germinating over a wide range of temperatures through an extended period into summer (Baskin and Baskin, 1980; Coble et al., 1981;

Dickerson and Sweet, 1971; Simard and Benoit, 2010). However, several studies found that common ragweed was one of the earliest germinating summer annual weeds, with emergence occurring over a relatively short period in early spring (Myers et al., 2004;

Stoller and Wax, 1973; Werle et al., 2014), and control of this early flush in a timely manner was the most important factor in limiting interference with soybean yields (Coble et al., 1981). Germination was shown to occur at a maximum depth of 5 cm (Stoller and

Wax, 1973), and is favored by light during low spring temperatures and darkness during higher temperatures, but seeds can enter a secondary dormancy after extended high temperatures (Baskin and Baskin, 1980; Willemsen, 1975). Common ragweed plants that emerged in mid-May in Ithaca, New York produced over 32,000 seeds per plant, whereas later plantings through July produced successively fewer seeds per plant (Dickerson and

Sweet, 1971). Plant growth and phenological development can occur over a wide range of temperatures, with leaf development rates increasing with increasing temperatures

(Deen et al., 1998).

Common ragweed becomes sensitive to photoperiod soon after emergence and reproduction is initiated earlier under short days, but the critical day length varies by provenance (Deen et al., 1998; Dickerson and Sweet, 1971). Biotypes from southern latitudes, planted in late May in Ithaca, New York (15-hour day length), initiated flowering nearly two months later while producing more vegetative growth than those from northern locations—although the reproductive development of the southern

2 biotypes was too late for the production of much viable seed in the northern United States

(Dickerson and Sweet, 1971).

As is often the case with wind-pollinated plant species, common ragweed is highly outcrossing, with greater than 90% cross-pollination in plants separated by as much as 9-meters (Friedman and Barrett, 2008). Although some studies describe common ragweed as self-compatible (Bassett and Crompton, 1975; Jones, 1936), a more recent study demonstrated self-incompatibility due to inhibition of pollen germination and pollen tube entry to the stigma (Friedman and Barrett, 2008). Like many other weeds with wide geographic distribution, common ragweed exhibits a large amount of local adaptation and phenotypic plasticity across differing geographies and climates, and within localized populations (Dickerson and Sweet, 1971; DiTommaso, 2004; Eom et al.,

2013; Leskovšek et al., 2012; Payne, 1963; Traveset, 1992).

Common ragweed’s wide range of adaptation may be aided not only by the high rate of outcrossing through wind-blown pollen, but also by intrinsic genetic variation.

Common cocklebur (Xanthium stumarium) is a closely related summer annual species

(Asteraceae, subtribe Ambrosiinae), but is largely self-pollinated. A comparison of acetolactate-synthase (ALS) gene nucleotide sequences from multiple accessions of common cocklebur versus common ragweed showed 12.5% polymorphic nucleotide positions in common ragweed and no polymorphisms in common cocklebur (Tranel et al., 2004). Even if this overestimates total genetic variation in common ragweed, it is indicative of high genetic diversity, making it difficult to differentiate heritable phenotype from environmental responses.

3 Dickerson and Sweet (1971) reported that season-long common ragweed control is important, as even small plants originating from July-planted seeds produced more than 3,000 seeds per plant. Studies have demonstrated the competitiveness, yield losses, and reproductive potential of common ragweed in corn and soybean (Coble et al., 1981;

Cowbrough et al., 2003; Simard and Benoit, 2012; Weaver, 2001), white bean (Chikoye et al., 1995), and peanut (Clewis et al., 2001), and several have attempted to calculate economic thresholds for control. Economic weed thresholds have usually disregarded additions to the soil seed bank by uncontrolled common ragweed plants, and the impact they can have on future crops (Simard et al., 2009). In addition to its impact on crop production, ragweed is now the major source of allergenic pollen in several countries

(Boulet et al., 1997; Burton et al., 2001; Léonard et al., 2010; Simard and Benoit, 2012;

Wopfner et al., 2005; Ziska et al., 2009). As common ragweed has expanded its distribution, its competitiveness with crops and its detrimental effects on human health have become a major concern worldwide (Bass et al., 2000; Burbach et al., 2009; Genton et al., 2005; Kiss and Béres, 2006; Makra et al., 2005; Tokarska-Guzik et al., 2011; Týr et al., 2009).

1.2 Glyphosate Resistance

Glyphosate (N-[phosphonomethyl]-glycine) inhibits the enzyme

5-enolpyruvylshikimate-3-phosphate synthase (EPSP synthase or EPSPS) (Steinrücken and Amrhein, 1980), and has been described as the world’s most important herbicide

(Duke and Powles, 2008a; Duke and Powles, 2008b). After glyphosate application to a susceptible plant and subsequent blockage of a critical step in the synthesis of aromatic

4 amino acids and many phenolic secondary metabolites (Figure 1.1), plants are affected by a variety of changes, including reduced photosynthetic activity and reduced stomatal conductance (Fuchs et al., 2002). In addition, glyphosate has been shown to increase a plant’s susceptibility to pathogens, including to races that are not normally pathogenic to that plant species (Brammall and Higgins, 1988; Liu et al., 1997; Rosenbaum et al., 2014;

Schafer et al., 2012; Schafer et al., 2013). Increased disease susceptibility in glyphosate- treated plants may occur by a variety of mechanisms, but perhaps most importantly by perturbations in the plants basal and systemic defense responses (Duke and Powles,

2008b; Liu et al., 1997). Defense suppression can occur with otherwise sub-lethal glyphosate concentrations, and may be an important component of glyphosate’s efficacy

(Brammall and Higgins, 1988; Sharon et al., 1992; Smith and Hallett, 2006). Seemingly contradictory to defense suppression, hormetic effects (growth stimulation at low doses) of glyphosate have also been observed in various species, but the mechanisms are unclear

(Belz and Duke, 2014; Carvalho et al., 2013; Velini et al., 2008). More research is needed to understand these paradoxical observations.

In 1996, a biotype of rigid ryegrass (Lolium rigidum) was found in Victoria,

Australia that displayed the first known case of naturally occurring glyphosate resistance in a weed species (Powles et al., 1998). Rigid ryegrass has shown a propensity for developing different mechanisms of herbicide resistance, with resistant biotypes confirmed for 12 herbicide site-of-action (SOA) groups worldwide (Heap, 2014; Preston et al., 2009). In early experiments, the Victoria glyphosate-resistant biotype displayed no differences in 14C-glyphosate uptake, translocation, or metabolism compared to glyphosate-sensitive biotypes (Feng et al., 1999). Biotype lines with the highest level of 5 glyphosate resistance were found to have slightly greater EPSPS transcript quantities than glyphosate-sensitive lines, but no differences in EPSPS enzyme sensitivity to glyphosate

(Baerson et al., 2002a).

Studies of other glyphosate-resistant rigid ryegrass biotypes have found altered translocation of glyphosate, with less 14C-glyphosate in the shoot meristems than sensitive biotypes (Wakelin et al., 2004; Yu et al., 2009). Further investigations using

31phosphorus nuclear magnetic resonance (31P-NMR) spectroscopy revealed greater accumulation of glyphosate within the vacuole of the resistant biotype (Ge et al., 2012b).

As of 2006, there were 54 confirmed glyphosate-resistant biotypes of rigid ryegrass in

Australia alone, and one was shown to have a point mutation coding for a proline-to- threonine substitution at position-106 (P106T) of the EPSPS enzyme (Wakelin and

Preston, 2006). Rigid ryegrass biotypes have since been identified with proline-to- alanine (P106A; Yu et al., 2007), proline-to-serine (P106S; Bostamam et al., 2012;

Collavo and Sattin, 2012; Simarmata and Penner, 2008), and proline-to-leucine (P106L;

Collavo and Sattin, 2012; Kaundun et al., 2011) substitutions in EPSPS.

Goosegrass (Eleusine indica) glyphosate resistance was first reported in 1997 for a biotype from Malaysia. This was the first naturally occurring biotype in which a mutation in the EPSPS gene was found to confer glyphosate resistance (Baerson et al.,

2002b). Within a goosegrass population segregating for a P106S substitution, individuals homozygous for this mutation displayed two-fold higher glyphosate resistance than individuals from the same population lacking the mutation (Kaundun et al., 2008).

Malaysian goosegrass biotypes have also been found that contain a P106T substitution

6 (Ng et al., 2003). A recently identified biotype containing alleles coding for a threonine- to-isoleucine substitution at position 102 (T102I) and the P106S substitution of the

EPSPS enzyme (combined substitutions known as “TIPS”) exhibited a high level of glyphosate resistance (Jalaludin et al., 2013). The TIPS substitutions were used in the engineered GA21 enzyme for the first generation of Roundup Ready® maize, where the addition of the T102I substitution decreased the EPSPS affinity for glyphosate and increased its affinity for the substrate phosphoenolpyruvate (PEP) compared to the P106S substitution alone (Sammons and Gaines, 2014). The T102I substitution in goosegrass is presumed to have evolved from an allele already containing the P106S substitution, because other experiments have shown that the T102I change alone greatly decreases the binding of EPSPS with both glyphosate and PEP, making it a detrimental mutation without the presence of the P106S (Jalaludin et al., 2013; Sammons and Gaines, 2014).

The first dicot species reported to have glyphosate resistance was horseweed

(Conyza canadensis) from a Delaware population in 2000 (VanGessel, 2001). In greenhouse experiments, the glyphosate rate required to reduce growth of the resistant biotype by 50% (GR50) was 8- to 13-fold higher than for a glyphosate-sensitive biotype.

Three years later, Koger et al. (2004) reported three horseweed biotypes collected from cotton and soybean fields in Mississippi and Tennessee showed 8- to 12-fold resistance to glyphosate, and a fourth biotype from Mississippi was 2- to 4-fold more resistant that the first three resistant biotypes. Other dose-response experiments with putative or confirmed glyphosate-resistant horseweed biotypes from Arkansas, Delaware, Ohio, and

Virginia found two-leaf seedlings to have GR50 values similar to sensitive biotypes, but three-fold higher resistance than sensitive biotypes at the five-leaf rosette stage (Dinelli et 7 al., 2006). Later growth stages of both biotypes required higher glyphosate doses to reached 50% growth reduction, but the resistant : susceptible GR50 ratio did not change at higher doses.

Subsequent studies of eleven glyphosate-resistant horseweed biotypes reported similar foliar retention and uptake of 14C-glyphosate compared to sensitive biotypes, but reduced translocation and reduced loading of glyphosate into the apoplast and phloem in resistant biotypes. No metabolic deactivation of glyphosate was detected in any of these biotypes (Feng et al., 2004). Another study reported that a glyphosate-resistant horseweed biotype had reduced glyphosate translocation, but semi-quantitative PCR of cDNA also revealed that it produced two- to three-fold higher EPSPS mRNA transcript levels than glyphosate-sensitive plants (Dinelli et al., 2006). Further investigations into the reduced glyphosate translocation mechanism in the resistant versus susceptible horseweed biotypes were conducted using 31phosphorus nuclear magnetic resonance

(31P-NMR) spectroscopy, and showed that a significantly greater amount of glyphosate accumulated within the vacuole of the resistant biotype (Ge et al., 2010). Glyphosate- resistant horseweed plants acclimated to 12˚C showed little accumulation of glyphosate in the vacuole, and were killed by a labeled field use rate of 1.68 kg ae ha−1 glyphosate

(Ge et al., 2011).

Transcriptome analysis and a whole-genome mouse-ear cress (Arabidopsis thaliana) microarray assay of glyphosate-treated and untreated plants of glyphosate- sensitive and glyphosate-resistant horseweed were used to identify multiple candidate glyphosate-resistance genes. Those in the ATP-binding cassette (ABC) transporter

8 family had high levels of transcript abundance, thus making this family a high priority for further analyses for the vacuolar sequestration of glyphosate as a potential resistance mechanism (Yuan et al., 2010). Three glyphosate-resistant biotypes of horseweed from

Crete, mainland Greece, and Delaware, USA were found not to have EPSPS amino acid substitutions or increased expression of EPSPS, but did have greatly increased expression of two ABC-transporter genes in leaf tissue following a glyphosate application (Nol et al., 2012). Glyphosate-resistant biotypes of two close relatives of horseweed have been found, including hairy fleabane (C. bonariensis) in Africa, Australia, Europe, and North and South America, and Sumatran fleabane (C. sumatrensis) in Europe and South

America (Dinelli et al., 2008; Heap, 2014; Mylonas et al., 2014; Okada and Jasieniuk,

2014; Santos et al., 2014; Urbano et al., 2007; Walker et al., 2011). Investigations into the glyphosate-resistance mechanism in Sumatran fleabane from Spain found decreased translocation of 14C-glyphosate and the mutation which would translate to the P106T substitution, based on cDNA sequencing of ESPS (reported as P182T based upon different numbering scheme, González-Torralva et al., 2014).

An Italian ryegrass (Lolium multiflorum; syn. Lolium perenne ssp. multiflorum) biotype resistant to glyphosate was first discovered in Chile in 2001 (Michitte et al.,

2007). Plants from this Chilean biotype exhibited 35% lower glyphosate spray retention and 40% lower uptake of 14C-glyphosate from the abaxial leaf surface than a sensitive biotype from the same region. Similar to some biotypes of rigid ryegrass, resistant Italian ryegrass plants also had increased translocation to leaf tips and decreased translocation to the shoot meristems (Lorraine-Colwill et al., 2002; Michitte et al., 2007). Experiments utilizing 31P-NMR showed accumulation of glyphosate in the cell vacuoles of glyphosate- 9 resistant rigid ryegrass and Italian ryegrass plants in all biotypes with reduced glyphosate translocation (Ge et al., 2012b). Another glyphosate-resistant Italian ryegrass biotype from Chile had the previously mentioned P106S amino acid substitution, with no change in translocation (Perez-Jones et al., 2007). Plants from an Arkansas biotype exhibiting 7-

to 13-fold glyphosate resistance were found to contain no EPSPS mutations believed to confer resistance, but did contain up to 25 times as many genomic copies of EPSPS as susceptible plants, which correlated with the observed levels of glyphosate resistance

(Salas et al., 2012). In both rigid and Italian ryegrass, decreased-translocation mechanisms appear to confer a higher level of glyphosate resistance than do EPSPS mutations, and biotypes with the highest levels of resistance most likely contain multiple resistance mechanisms (Preston et al., 2009).

Common ragweed glyphosate resistance was found in 2004 in Missouri (Pollard as cited in Brewer and Oliver, 2009) and Arkansas (Brewer and Oliver, 2009). In the same year, glyphosate-resistant biotypes of two other species in the subtribe

Ambrosiinae, including giant ragweed (Ambrosia trifida) and ragweed parthenium

(Parthenium hysterophorus), were identified in Ohio and Columbia, respectively (Heap,

2014; Stachler, 2008). The resistant biotype from Arkansas and another biotype found the next year were 10- to 21-fold more resistant to glyphosate than a glyphosate-sensitive biotype, but no differences were detected in 14C-glyphosate absorption or translocation between the two biotypes. These common ragweed biotypes were presumed to lack an altered EPSPS enzyme, because no differences in shikimate accumulation were detected

(Brewer and Oliver, 2009). Glyphosate-resistant common ragweed biotypes were reported from Ohio (also resistant to acetolactate synthase (ALS)-inhibitors), Kentucky, 10 and North Carolina in 2006; Indiana, Kansas, North Dakota and South Dakota in 2007;

Minnesota and Pennsylvania in 2008; Ontario, Canada in 2012; Alabama, Nebraska, and

New Jersey in 2013; and Mississippi in 2014, for a total of 17 glyphosate-resistant common ragweed biotypes reported to the International Survey of Herbicide Resistant

Weeds (Heap, 2014).

Glyphosate-resistant Palmer amaranth (Amaranthus palmeri) was found in North

Carolina and central Georgia in 2005 (Culpepper et al., 2006; Culpepper et al., 2008), and a biotype of waterhemp (Amaranthus tuberculatus) from Missouri exhibited multiple resistance to glyphosate, ALS-inhibiting herbicides, and protoporphyrinogen IX oxidase

(PPO or protox)-inhibiting herbicides (Legleiter and Bradley, 2008). The Palmer amaranth biotype from Georgia was six- to eight-fold more resistant to glyphosate than a sensitive biotype, based on shoot fresh weight and visual control GR50 values in a greenhouse dose response experiment (Culpepper et al., 2006). A glyphosate-resistant

Palmer amaranth biotype from Arkansas had 79- to 115-fold higher resistance than three susceptible biotypes from South Carolina (Norsworthy et al., 2008). Initial experiments with the Georgia biotype showed reduced shikimate accumulation in response to glyphosate treatment in the resistant biotype, but there were no differences in absorption, translocation, or chromosome number between the resistant and sensitive biotypes

(Culpepper et al., 2006). Mutations and reduced activity of the EPSPS enzyme were not found, but target-site duplication left 5- to 160-fold more copies of the EPSPS gene distributed throughout the genome of glyphosate-resistant Palmer amaranth plants, as evidenced by quantitative PCR of genomic DNA and fluorescent in-situ hybridization

(FISH) analysis (Gaines et al., 2009). These results were verified by heritability studies 11 and quantitative reverse-transcriptase PCR and immunoblot analyses showing higher

EPSPS expression, making this the first weed species in which EPSPS duplication has been found to confer resistance to glyphosate (Gaines et al., 2009).

Other glyphosate-resistance mechanisms notwithstanding, Palmer amaranth plants seem to require 30-to-50 genomic copies of EPSPS to survive normal field rates of glyphosate, based upon dose-response experiments with glyphosate-resistant and F2 plants of known EPSPS copy numbers (Gaines et al., 2011). 31P-NMR studies of waterhemp and Palmer amaranth biotypes with higher glyphosate resistance than suggested by their EPSPS expression levels, showed decreased cellular uptake of glyphosate (Ge et al., 2013). A glyphosate-resistant biotype of waterhemp from

Mississippi without increased EPSPS expression was found to have both decreased absorption of 14C-glyphosate and an EPSPS mutation translating to a P106S amino acid substitution, the latter being the first reported such mutation in a dicot weed species

(Nandula et al., 2013). Additional studies demonstrated that Palmer amaranth has the ability to transfer glyphosate resistance via pollen to spiny amaranth (Amaranthus spinosus), smooth pigweed (Amaranthus hybridus), and waterhemp, with hybridization observed at rates of < 0.01% to 1.4% (Gaines et al., 2012). Numerous other biotypes of waterhemp and Palmer amaranth have been reported since 2005, as well as a Mississippi biotype of spiny amaranth in 2012, and a biotype of mucronate pigweed (A. quitensis) from Argentina in 2013 (Heap, 2014).

Fourteen weed species with glyphosate-resistant biotypes have been reported thus far in the United States (Heap, 2014). In addition to 10 of those mentioned previously,

12 glyphosate-resistant biotypes of johnsongrass (Sorghum halepense) (Riar et al., 2011), junglerice (Echinochloa colona) (Alarcón-Reverte et al., 2012), kochia (Kochia scoparia)

(Waite et al., 2012; Wiersma, 2012), and annual bluegrass (Poa annua) (Binkholder et al., 2011) have been confirmed. Worldwide, glyphosate-resistant biotypes have been reported for 31 species (Heap, 2014).

1.3 Common Ragweed Resistance to Other Herbicides

Herbicide resistance has developed in many populations of Ambrosia artemisiifolia. Common ragweed was one of the earliest species in which biotypes were identified with resistance to photosystem II (PSII)-inhibitors. A triazine-resistant biotype was found in 1976 in Ontario, Canada and a phenylurea-resistant biotype was discovered in 1999 in Québec, Canada (Heap, 2014; Saint-Louis et al., 2005; Stephenson et al.,

1990). Only two additional cases of triazine-resistant common ragweed biotypes have been officially documented in the International Survey of Herbicide Resistant Weeds

(Heap, 2014). While the decline in new records could represent changes in agronomic practices and decreased reliance on PSII-inhibitors, other accounts portraying triazine- resistant common ragweed as widespread in Ontario and Hungary suggests that there is merely a lack of formal reports (Cseh et al., 2009; Stephenson et al., 1990). Triazine resistance is typically the result of a point mutation in the chloroplast gene encoding the

D1-subunit of the PSII reaction-center protein, resulting in a serine-to-glycine substitution at amino acid 264 (S264G), and a new polymerase chain reaction (PCR)- based method of detecting this polymorphism could increase the number of reported triazine-resistant biotypes (Mátyás et al., 2011).

13 Common ragweed biotypes resistant to cloransulam-methyl, an acetolactate synthase (ALS) [acetohydroxyacid synthase (AHAS)]-inhibitor in the triazolopyrimidine sulfonanilide chemical family, were confirmed in the first year of cloransulam commercialization (Patzoldt et al., 2001). A biotype from Indiana was also found to be cross-resistant to both chlorimuron-ethyl and imazaquin—ALS-inhibitors of the sulfonylurea and imidizolinone chemical families, respectively—so selection pressure for cloransulam-methyl resistance probably occurred over a number of years from applications of imidizolinone herbicides (Patzoldt et al., 2001). ALS-inhibitor resistant biotypes were found in 64 species of plants prior to common ragweed (Heap, 2014), and numerous amino acid substitutions with the potential to confer resistance had been documented (Falco et al., 1989; Tranel and Wright, 2002; Wright et al., 1998). DNA sequencing identified a nucleotide substitution that would translate as a tryptophan-to- leucine substitution at amino acid position 574 (W574L), which was previously found to confer cross-resistance among many ALS-inhibiting herbicides (Patzoldt et al., 2001).

Other studies have demonstrated that the W574L substitution is a common source of resistance to cloransulam-methyl and other ALS-inhibitors, in both common ragweed and other species (Patzoldt and Tranel, 2002; Rousonelos et al., 2012; Tranel and Wright,

2002; Zheng et al., 2005).

Biotypes of common ragweed with resistance to protoporphyrinogen IX oxidase

(PPO)-inhibiting herbicides have been found in both Delaware (2005) and Ohio (2006)

(Heap, 2014; Rousonelos et al., 2012). Resistance to PPO-inhibitors has only been found in five other species to date, and both of the common ragweed biotypes were also resistant to ALS-inhibitors (Heap, 2014). Molecular analysis of the Delaware common 14 ragweed biotype demonstrated that an amino acid substitution of arginine-to-leucine at position 98 (R98L) of the PPO enzyme conferred resistance to PPO-inhibitors, while the previously described W574L substitution in the ALS enzyme provided resistance to

ALS-inhibitors (Rousonelos et al., 2012).

1.4 Objectives

A continuous soybean field in Clinton County, Ohio had been treated with ALS- inhibitors over several years. The grower was beginning to experience poor control of common ragweed. Upon the introduction of glyphosate-tolerant soybeans in 1996, this common ragweed population was initially well controlled by a single application of glyphosate, but by 2004, the grower was again having consistent failures for common ragweed control, and a large population had built up in the field. After several years of acceptable control following a switch to PPO-inhibitors, control was again declining.

Seed was collected from plants surviving a glyphosate application in 2009 and stored for further research.

The objectives of the studies presented in the following chapters are as follows:

(1) to confirm and characterize levels of resistance to glyphosate (EPSPS-inhibitor), cloransulam-methyl (ALS-inhibitor), and fomesafen (PPO-inhibitor) in suspected multiple-resistant common ragweed biotypes using dose response assays; (2) to investigate the target site of glyphosate (EPSPS) using molecular genetic and enzymology approaches to look for evidence of mutations, amino acid substitutions, or increases in enzyme expression due to gene duplication; and (3) to use 14C-glyphosate to examine absorption and translocation and 31P-NMR spectroscopy to investigate

15 subcellular localization as a possible non–target-site mechanism regulating glyphosate uptake or translocation in R versus S biotypes.

16 Chapter 1 References

Alarcón-Reverte R, García A, Urzúa J, Fischer AJ (2012) Resistance to glyphosate in junglerice (Echinochloa colona) from California. Weed Science 61:48-54

Baerson SR, Rodriguez DJ, Biest NA, Tran M, You J, Kreuger RW, Dill GM, Pratley JE, Gruys KJ (2002a) Investigating the mechanism of glyphosate resistance in rigid ryegrass (Lolium ridigum). Weed Science 50:721-730

Baerson SR, Rodriguez DJ, Tran M, Feng Y, Biest NA, Dill GM (2002b) Glyphosate- resistant goosegrass. Identification of a mutation in the target enzyme 5- enolpyruvylshikimate-3-phosphate synthase. Plant Physiol 129:1265-75

Baskin JM, Baskin CC (1980) Ecophysiology of secondary dormancy in seeds of Ambrosia artemisiifolia. Ecology 61:475-480

Bass DJ, Delpech V, Beard J, Bass P, Walls RS (2000a) Late summer and fall (March- May) pollen allergy and respiratory disease in Northern New South Wales, Australia. Annals of allergy, asthma & immunology : official publication of the American College of Allergy, Asthma, & Immunology 85:374-381

Bass DJ, Delpech V, Beard J, Bass P, Walls RS (2000b) Ragweed in Australia. Aerobiologia 16:107-111

Bassett IJ, Crompton CW (1975) The biology of Canadian weeds.: 11. Ambrosia artemisiifolia L. and A. psilostachya DC. Canadian Journal of Plant Science 55:463-476

Bassett IJ, Terasmae J (1962) Ragweeds, Ambrosia species, in Canada and their history in postglacial time. Canadian Journal of Botany 40:141-150

Belz RG, Duke SO (2014) Herbicides and plant hormesis. Pest Management Science 70:698-707

Binkholder KM, Fresenburg BS, Teuton TC, Xiong X, Smeda RJ (2011) Selection of glyphosate-resistant annual bluegrass (Poa annua) on a golf course. Weed Science 59:286-289

Bostamam Y, Malone JM, Dolman FC, Boutsalis P, Preston C (2012) Rigid ryegrass (Lolium rigidum) populations containing a target site mutation in EPSPS and reduced glyphosate translocation are more resistant to glyphosate. Weed Science 60:474-479

Boulet LP, Turcotte H, Laprise C, Lavertu C, Bedard PM, Lavoie A, Hébert J (1997) Comparative degree and type of sensitization to common indoor and outdoor

17 allergens in subjects with allergic rhinitis and/or asthma. Clinical & Experimental Allergy 27:52-59

Brammall RA, Higgins VJ (1988) The effect of glyphosate on resistance of tomato to Fusarium crown and root rot disease and on the formation of host structural defensive barriers. Canadian Journal of Botany 66:1547-1555

Brewer CE, Oliver LR (2009) Confirmation and resistance mechanisms in glyphosate- resistant common ragweed (Ambrosia artemisiifolia) in Arkansas. Weed Science 57:567-573

Burbach GJ, Heinzerling LM, Röhnelt C, Bergmann KC, Behrendt H, Zuberbier T (2009) Ragweed sensitization in Europe – GA2LEN study suggests increasing prevalence. Allergy 64:664-665

Burton WN, Conti DJ, Chen C-Y, Schultz AB, Edington DW (2001) The Impact of Allergies and Allergy Treatment on Worker Productivity. Journal of Occupational and Environmental Medicine 43:64-71

Carvalho LB, Alves PL, Duke SO (2013) Hormesis with glyphosate depends on coffee growth stage. Anais da Academia Brasileira de Ciências 85:813-822

Chauvel B, Dessaint F, Cardinal-Legrand C, Bretagnolle F (2006) The historical spread of Ambrosia artemisiifolia L. in France from herbarium records. Journal of Biogeography 33:665-673

Chikoye D, Weise SF, Swanton CJ (1995) Influence of common ragweed (Ambrosia artemisiifolia) time of emergence and density on white bean (Phaseolus vulgaris). Weed Science 43:375-380

Clewis SB, Askew SD, Wilcut JW (2001) Common ragweed interference in peanut. Weed Science 49:768-772

Coble HD, Williams FM, Ritter RL (1981) Common ragweed (Ambrosia artemisiifolia) interference in soybeans (Glycine max). Weed Science 29:339-342

Collavo A, Sattin M (2012) Resistance to glyphosate in Lolium rigidum selected in Italian perennial crops: bioevaluation, management and molecular bases of target-site resistance. Weed Research 52:16-24

Cowbrough MJ, Brown RB, Tardif FJ (2003) Impact of common ragweed (Ambrosia artemisiifolia) aggregation on economic thresholds in soybean. Weed Science 51:947-954

Cseh A, Cernák I, Taller J (2009) Molecular characterization of atrazine resistance in common ragweed (Ambrosia artemisiifolia L.). Journal of Applied Genetics 50:321-327 18 Csontos P, Vitalos M, Barina Z, Kiss L (2010) Early distribution and spread of Ambrosia artemisiifolia in Central and Eastern Europe. Bot. Helv. 120:75-78

Culpepper AS, Grey TL, Vencill WK, Kichler JM, Webster TM, Brown SM, York AC, Davis JW, Hanna WW (2006) Glyphosate-resistant Palmer amaranth (Amaranthus palmeri ) confirmed in Georgia. Weed Science 54:620-626

Culpepper AS, Whitaker JR, MacRae AW, York AC (2008) Distribution of glyphosate- resistant Palmer amaranth (Amaranthus palmeri) in Georgia and North Carolina during 2005 and 2006. J. Cotton Sci 12:306-310

Deen W, Hunt T, Swanton CJ (1998) Influence of temperature, photoperiod, and irradiance on the phenological development of common ragweed (Ambrosia artemisiifolia). Weed Science 46:555-560

Dickerson CT, Jr, Sweet RD (1971) Common ragweed ecotypes. Weed Science 19:64-66

Dinelli G, Marotti I, Bonetti A, Catizone P, Urbano JM, Barnes J (2008) Physiological and molecular bases of glyphosate resistance in Conyza bonariensis biotypes from Spain. Weed Research 48:257-265

Dinelli G, Marotti I, Bonetti A, Minelli M, Catizone P, Barnes J (2006) Physiological and molecular insight on the mechanisms of resistance to glyphosate in Conyza canadensis (L.) Cronq. biotypes. Pesticide Biochemistry and Physiology 86:30-41

DiTommaso A (2004) Germination behavior of common ragweed (Ambrosia artemisiifolia) populations across a range of salinities. Weed Science 52:1002- 1009

Duke SO, Powles SB (2008a) Glyphosate-Resistant Weeds and Crops. Pest Management Science 64:317-318

Duke SO, Powles SB (2008b) Glyphosate: a once-in-a-century herbicide. Pest Management Science 64:319-325

Eom SH, DiTommaso A, Weston LA (2013) Effects of soil salinity in the growth of Ambrosia artemisiifolia biotypes collected from roadside and agricultural field. Journal of Plant Nutrition 36:2191-2204

Falco SC, McDevitt RE, Chui CF, Hartnett ME, Knowlton S, Mauvais CJ, Smith JK, Mazur BJ (1989) Engineering herbicide-resistant acetolactate synthase. Dev Ind Microbiol 30:187-194

Feng PCC, Pratley JE, Bohn JA (1999) Resistance to glyphosate in Lolium rigidum. II. Uptake, translocation, and metabolism. Weed Science 47:412-415

19 Feng PCC, Tran M, Chiu T, Douglas Sammons R, Heck GR, CaJacob CA (2004) Investigations into glyphosate-resistant horseweed (Conyza canadensis): retention, uptake, translocation, and metabolism. Weed Science 52:498-505

Friedman J, Barrett SCH (2008) High outcrossing in the annual colonizing species Ambrosia artemisiifolia (Asteraceae). Annals of Botany 101:1303-1309

Fuchs MA, Geiger DR, Reynolds TL, Bourque JE (2002) Mechanisms of glyphosate toxicity in velvetleaf (Abutilon theophrasti medikus). Pesticide Biochemistry and Physiology 74:27-39

Gaines TA, Shaner DL, Ward SM, Leach JE, Preston C, Westra P (2011) Mechanism of resistance of evolved glyphosate-resistant Palmer amaranth (Amaranthus palmeri). Journal of Agricultural and Food Chemistry 59:5886-5889

Gaines TA, Ward SM, Bukun B, Preston C, Leach JE, Westra P (2012) Interspecific hybridization transfers a previously unknown glyphosate resistance mechanism in Amaranthus species. Evolutionary Applications 5:29-38

Gaines TA, Zhang W, Wang D, Bukun B, Chisholm ST, Shaner DL, Nissen SJ, Patzoldt WL, Tranel PJ, Culpepper AS, Grey TL, Webster TM, Vencill WK, Sammons RD, Jiang J, Preston C, Leach JE, Westra P (2009) Gene amplification confers glyphosate resistance in Amaranthus palmeri. Proceedings of the National Academy of Sciences 107:1029-1034

Gaudeul M, Giraud T, Kiss L, Shykoff JA (2011) Nuclear and chloroplast microsatellites show multiple introductions in the worldwide invasion history of common ragweed, Ambrosia artemisiifolia. PloS one 6:e17658

Ge X, d'Avignon DA, Ackerman JJH, Duncan B, Spaur MB, Sammons RD (2011) Glyphosate-resistant horseweed made sensitive to glyphosate: low-temperature suppression of glyphosate vacuolar sequestration revealed by 31P NMR. Pest Management Science 67:1215-1221

Ge X, d'Avignon DA, Ackerman JJH, Ostrander E, Sammons RD (2013) Application of 31P-NMR spectroscopy to glyphosate studies in plants: Insights into cellular uptake and vacuole sequestration correlated to herbicide resistance. Pages 55-83 Handbook on Herbicides : Biological Activity, Classification and Health and Environmental Implications. Hauppauge, NY, USA: Nova Science Publishers, Inc.

Ge X, d'Avignon DA, Ackerman JJH, Sammons RD (2010) Rapid vacuolar sequestration: the horseweed glyphosate resistance mechanism. Pest Management Science:n/a-n/a

Ge X, d’Avignon DA, Ackerman JJH, Collavo A, Sattin M, Ostrander EL, Hall EL, Sammons RD, Preston C (2012b) Vacuolar glyphosate-sequestration correlates with glyphosate resistance in ryegrass (Lolium spp.) from Australia, South 20 America, and Europe: A 31P NMR investigation. Journal of Agricultural and Food Chemistry 60:1243-1250

Genton BJ, Shykoff JA, Giraud T (2005) High genetic diversity in French invasive populations of common ragweed, Ambrosia artemisiifolia, as a result of multiple sources of introduction. Molecular Ecology 14:4275-4285

González-Torralva F, Gil-Humanes J, Barro F, Domínguez-Valenzuela J, De Prado R (2014) First evidence for a target site mutation in the EPSPS2 gene in glyphosate- resistant Sumatran fleabane from citrus orchards. Agron. Sustain. Dev. 34:553- 560

Heap I (2014) The international survey of herbicide resistant weeds. http://www.weedscience.com. Accessed November 5, 2014

Hodgins KA, Rieseberg L (2011) Genetic differentiation in life-history traits of introduced and native common ragweed (Ambrosia artemisiifolia) populations. Journal of Evolutionary Biology 24:2731-2749

Jalaludin A, Han H, Powles S (2013) Evolution in action: a double amino acid substitution in the EPSPS gene endows high-level glyphosate resistance. Pages 35-35 in Proceedings of the Global Herbicide Resistance Challenge. Fremantle, Australia: Australian Herbicide Resistance Initiative

Joly M, Bertrand P, Gbangou RY, White M-C, Dubé J, Lavoie C (2011) Paving the way for invasive species: road type and the spread of common ragweed (Ambrosia artemisiifolia). Environmental Management 48:514-522

Jones KL (1936) Studies on Ambrosia, I. The inheritance of floral types in the ragweed, Ambrosia elatior L. American Midland Naturalist 17:673-699

Karis PO (1995) Cladistics of the subtribe Ambrosiinae (Asteraceae: Heliantheae). Systematic Botany 20:40-54

Kaundun SS, Dale RP, Zelaya IA, Dinelli G, Marotti I, McIndoe E, Cairns A (2011) A novel P106L mutation in EPSPS and an unknown mechanism(s) act additively to confer resistance to glyphosate in a South African Lolium rigidum population. Journal of Agricultural and Food Chemistry 59:3227-3233

Kaundun SS, Zelaya IA, Dale RP, Lycett AJ, Carter P, Sharples KR, McIndoe E (2008) Importance of the P106S target-site mutation in conferring resistance to glyphosate in a goosegrass (Eleusine indica) population from the Philippines. Weed Science 56:637-646

Kil JH, Shim KC, Park SH, Koh KS, Suh MH, Ku YB, Suh SU, Oh HK, Kong HY (2004) Distributions of naturalized alien plants in South Korea. Weed Technology 18:1493-1495 21 Kiss L, Béres I (2006) Anthropogenic factors behind the recent population expansion of common ragweed (Ambrosia artemisiifolia L.) in Eastern Europe: is there a correlation with political transitions? Journal of Biogeography 33:2156-2157

Lavoie C, Jodoin Y, De Merlis AG (2007) How did common ragweed (Ambrosia artemisiifolia L.) spread in Québec? A historical analysis using herbarium records. Journal of Biogeography 34:1751-1761

Legleiter TR, Bradley KW (2008) Glyphosate and multiple herbicide resistance in common waterhemp (Amaranthus rudis) populations from Missouri. Weed Science 56:582-587

Léonard R, Wopfner N, Pabst M, Stadlmann J, Petersen BO, Duus J, Himly M, Radauer C, Gadermaier G, Razzazi-Fazeli E, Ferreira F, Altmann F (2010) A New Allergen from Ragweed (Ambrosia artemisiifolia) with Homology to Art v 1 from Mugwort. Journal of Biological Chemistry 285:27192-27200

Leskovšek R, Datta A, Knezevic SZ, Simončič A (2012) Common ragweed (Ambrosia artemisiifolia) dry matter allocation and partitioning under different nitrogen and density levels. Weed Biology and Management 12:98-108

Liu L, Punja ZK, Rahe JE (1997) Altered root exudation and suppression of induced lignification as mechanisms of predisposition by glyphosate of bean roots (Phaseolus vulgaris L.) to colonization by Pythium spp. Physiological and Molecular Plant Pathology 51:111-127

Lorraine-Colwill DF, Powles SB, Hawkes TR, Hollinshead PH, Warner SAJ, Preston C (2002) Investigations into the mechanism of glyphosate resistance in Lolium rigidum. Pesticide Biochemistry and Physiology 74:62-72

Makra L, Juhász M, Béczi R, Borsos Ek (2005) The history and impacts of airborne Ambrosia (Asteraceae) pollen in Hungary. Grana 44:57-64

Maryushkina VY (1991) Peculiarities of common ragweed (Ambrosia artemisiifolia L.) strategy. Agriculture, Ecosystems & Environment 36:207-216

Mátyás K, Taller J, Cseh A, Poczai P, Cernák I (2011) Development of a simple PCR- based assay for the identification of triazine resistance in the noxious plant common ragweed (Ambrosia artemisiifolia) and its applicability in higher plants. Biotechnology Letters 33:2509-2515

McAndrews JH (1988) Human disturbance of North American forests and grasslands: The fossil pollen record. Pages 673-697 in Huntley B, Webb T, III, eds. Vegetation history: Springer Netherlands

22 Michitte P, De Prado R, Espinoza N, Pedro Ruiz-Santaella J, Gauvrit C (2007) Mechanisms of resistance to glyphosate in a ryegrass (Lolium multiflorum) biotype from Chile. Weed Science 55:435-440

Mulligan GA (1957) Chromosome numbers of Canadian weeds. I. Canadian Journal of Botany 35:779-789

Munoz SE, Gajewski K (2010) Distinguishing prehistoric human influence on late- Holocene forests in southern Ontario, Canada. The Holocene 20:967-981

Myers MW, Curran WS, VanGessel MJ, Calvin DD, Mortensen DA, Majek BA, Karsten HD, Roth GW (2004) Predicting weed emergence for eight annual species in the northeastern United States. Weed Science 52:913-919

Mylonas PN, Giannopolitis CN, Efthimiadis PG, Menexes GC, Madesis PB, Eleftherohorinos IG (2014) Glyphosate resistance of molecularly identified Conyza albida and Conyza bonariensis populations. Crop Protection 65:207-215

Nandula VK, Ray JD, Ribeiro DN, Pan Z, Reddy KN (2013) Glyphosate resistance in tall waterhemp (Amaranthus tuberculatus) from Mississippi is due to both altered target-site and nontarget-site mechanisms. Weed Science 61:374-383

Ng CH, Wickneswari R, Salmijah S, Teng YT, Ismail BS (2003) Gene polymorphisms in glyphosate-resistant and -susceptible biotypes of Eleusine indica from Malaysia. Weed Research 43:108-115

Nol N, Tsikou D, Eid M, Livieratos IC, Giannopolitis CN (2012) Shikimate leaf disc assay for early detection of glyphosate resistance in Conyza canadensis and relative transcript levels of EPSPS and ABC transporter genes. Weed Research 52:233-241

Norsworthy JK, Griffith GM, Scott RC, Smith KL, Oliver LR (2008) Confirmation and control of glyphosate-resistant Palmer amaranth (Amaranthus palmeri) in Arkansas. Weed Technology 22:108-113

Okada M, Jasieniuk M (2014) Inheritance of glyphosate resistance in hairy fleabane (Conyza bonariensis) from California. Weed Science 62:258-266

Patzoldt WL, Tranel PJ (2002) Molecular analysis of cloransulam resistance in a population of giant ragweed. Weed Science 50:299-305

Patzoldt WL, Tranel PJ, Alexander AL, Schmitzer PR (2001) A common ragweed population resistant to cloransulam-methyl. Weed Science 49:485-490

Payne WW (1963) The morphology of the inflorescence of ragweeds (Ambrosia- Franseria: Compositae). American Journal of Botany 50:872-880

23 Payne WW, Raven PH, Kyhos DW (1964) Chromosome numbers in Compositae. IV. Ambrosieae. American Journal of Botany:419-424

Perez-Jones A, Park K-W, Polge N, Colquhoun J, Mallory-Smith CA (2007) Investigating the mechanisms of glyphosate resistance in Lolium multiflorum. Planta 226:395-404

Powles SB, Lorraine-Colwill DF, Dellow JJ, Preston C (1998) Evolved resistance to glyphosate in rigid ryegrass (Lolium rigidum) in Australia. Weed Science 46:604- 607

Preston C, Wakelin AM, Dolman FC, Bostamam Y, Boutsalis P (2009) A decade of glyphosate-resistant Lolium around the world: Mechanisms, genes, fitness, and agronomic management. Weed Science 57:435-441

Riar DS, Norsworthy JK, Johnson DB, Scott RC, Bagavathiannan M (2011) Glyphosate resistance in a johnsongrass (Sorghum halepense) biotype from Arkansas. Weed Science 59:299-304

Rosenbaum KK, Miller GL, Kremer RJ, Bradley KW (2014) Interactions between glyphosate, Fusarium infection of common waterhemp (Amaranthus rudis), and soil microbial abundance and diversity in soil collections from Missouri. Weed Science 62:71-82

Rousonelos SL, Lee RM, Moreira MS, VanGessel MJ, Tranel PJ (2012) Characterization of a common ragweed (Ambrosia artemisiifolia) population resistant to ALS- and PPO-inhibiting herbicides. Weed Science 60:335-344

Saint-Louis S, DiTommaso A, Watson AK (2005) A common ragweed (Ambrosia artemisiifolia) biotype in southwestern Québec resistant to linuron. Weed Technology 19:737-743

Salas RA, Dayan FE, Pan Z, Watson SB, Dickson JW, Scott RC, Burgos NR (2012) EPSPS gene amplification in glyphosate-resistant Italian ryegrass (Lolium perenne ssp. multiflorum) from Arkansas. Pest Management Science 68:1223- 1230

Sammons RD, Gaines TA (2014) Glyphosate resistance: state of knowledge. Pest Management Science

Santos G, Oliveira RS, Constantin J, Constantin Francischini A, Machado MFPS, Mangolin CA, Nakajima JN (2014) Conyza sumatrensis: A new weed species resistant to glyphosate in the Americas. Weed Biology and Management 14:106- 114

Schafer JR, Hallett SG, Johnson WG (2012) Response of giant ragweed (Ambrosia trifida), horseweed (Conyza canadensis), and common lambsquarters 24 (Chenopodium album) biotypes to glyphosate in the presence and absence of soil microorganisms. Weed Science 60:641-649

Schafer JR, Hallett SG, Johnson WG (2013) Soil microbial root colonization of glyphosate-treated giant ragweed (Ambrosia trifida), horseweed (Conyza canadensis), and common lambsquarters (Chenopodium album) biotypes. Weed Science 61:289-295

Sharon A, Amsellem Z, Gressel J (1992) Glyphosate suppression of an elicited defense response: Increased susceptibility of Cassia obtusifolia to a mycoherbicide. Plant Physiology 98:654-659

Simard M-J, Benoit DL (2012) Potential pollen and seed production from early- and late- emerging common ragweed in corn and soybean. Weed Technology 26:510-516

Simard M-J, Panneton B, Longchamps L, Lemieux C, Légère A, Leroux GD (2009) Validation of a management program based on a weed cover threshold model: Effects on herbicide use and weed populations. Weed Science 57:187-193

Simard MJ, Benoit DL (2010) Distribution and abundance of an allergenic weed, common ragweed (Ambrosia artemisiifolia L.), in rural settings of southern Quebec, Canada. Canadian Journal of Plant Science 90:549-557

Simarmata M, Penner D (2008) The basis for glyphosate resistance in rigid ryegrass (Lolium Rigidum) from California. Weed Science 56:181-188

Smith DA, Hallett SG (2006) Interactions between chemical herbicides and the candidate bioherbicide Microsphaeropsis amaranthi. Weed Science 54:532-537

Stachler JM (2008) Characterization and management of glyphosate-resistant giant ragweed (Ambrosia trifida L.) and horseweed [Conyza canadensis (L.) Cronq.]. Ph.D Dissertation. Columbus, OH: The Ohio State University

Steinrücken HC, Amrhein N (1980) The herbicide glyphosate is a potent inhibitor of 5- enolpyruvylshikimic acid-3-phosphate synthase. Biochemical and Biophysical Research Communications 94:1207-1212

Stephenson GR, Dykstra MD, McLaren RD, Hamill AS (1990) Agronomic practices influencing triazine-resistant weed distribution in Ontario. Weed Technology 4:199-207

Stoller EW, Wax LM (1973) Periodicity of germination and emergence of some annual weeds. Weed Science 21:574-580

Tokarska-Guzik B, Bzdęga K, Koszela K, Żabińska I, Krzuś B, Sajan M, Sendek A (2011) Allergenic invasive plant Ambrosia artemisiifolia L. in Poland: threat and selected aspects of biology. Biodiversity: Research and Conservation 21:39-48 25 Tranel PJ, Jiang W, Patzoldt WL, Wright TR (2004) Intraspecific variability of the acetolactate synthase gene. Weed Science 52:236-241

Tranel PJ, Wright TR (2002) Resistance of weeds to ALS-inhibiting herbicides: what have we learned? Weed Science 50:700-712

Traveset A (1992) Sex expression in a natural population of the monoecious annual, Ambrosia artemisiifolia (Asteraceae). American Midland Naturalist 127:309-315

Týr Š, Vereš T, Lacko-Bartošová M (2009) Occurrence of common ragweed (Ambrosia artemisiifolia L.) in field crops in the Slovak Republic. Herbologia 10:1-9

Urbano JM, Borrego A, Torres V, Leon JM, Jimenez C, Dinelli G, Barnes J (2007) Glyphosate-resistant hairy fleabane (Conyza bonariensis) in Spain. Weed Technology 21:396-401

VanGessel MJ (2001) Glyphosate-resistant horseweed from Delaware. Weed Science 49:703-705

Velini ED, Alves E, Godoy MC, Meschede DK, Souza RT, Duke SO (2008) Glyphosate applied at low doses can stimulate plant growth. Pest Management Science 64:489-496

Waite J, Thompson CR, Peterson DE, Currie RS, Olson BLS, Stahlman PW, Al-Khatib K (2012) Differential kochia (Kochia scoparia) populations response to glyphosate. Weed Science 61:193-200

Wakelin AM, Lorraine-Colwill DF, Preston C (2004) Glyphosate resistance in four different populations of Lolium rigidum is associated with reduced translocation of glyphosate to meristematic zones. Weed Research 44:453-459

Wakelin AM, Preston C (2006) A target-site mutation is present in a glyphosate-resistant Lolium rigidum population. Weed Research 46:432-440

Walker S, Bell K, Robinson G, Widderick M (2011) Flaxleaf fleabane (Conyza bonariensis) populations have developed glyphosate resistance in north-east Australian cropping fields. Crop Protection 30:311-317

Weaver SE (2001) Impact of lamb’s-quarters, common ragweed and green foxtail on yield of corn and soybean in Ontario. Canadian Journal of Plant Science 81:821- 828

Werle R, Sandell LD, Buhler DD, Hartzler RG, Lindquist JL (2014) Predicting emergence of 23 summer annual weed species. Weed Science 62:267-279

Wiersma AT (2012) Regional whole plant and molecular response of Kochia scoparia to glyphosate. MS Thesis. Fort Collins, CO: Colorado State University 26 Willemsen RW (1975) Effect of stratification temperature and germination temperature on germination and the induction of secondary dormancy in common ragweed seeds. American Journal of Botany 62:1-5

Wopfner N, Gadermaier G, Egger M, Asero R, Ebner C, Jahn-Schmid B, Ferreira F (2005) The spectrum of allergens in ragweed and mugwort pollen. International archives of allergy and immunology 138:337-346

Wright TR, Bascomb NF, Sturner SF, Penner D (1998) Biochemical mechanism and molecular basis for ALS-inhibiting herbicide resistance in sugarbeet (Beta vulgaris) somatic cell selections. Weed Science:13-23

Yu Q, Abdallah I, Han H, Owen M, Powles S (2009) Distinct non-target site mechanisms endow resistance to glyphosate, ACCase and ALS-inhibiting herbicides in multiple herbicide-resistant Lolium rigidum. Planta 230:713-723

Yu Q, Cairns A, Powles S (2007) Glyphosate, paraquat and ACCase multiple herbicide resistance evolved in a Lolium rigidum biotype. Planta 225:499-513

Yuan JS, Abercrombie LLG, Cao Y, Halfhill MD, Zhou X, Peng Y, Hu J, Rao MR, Heck GR, Larosa TJ, Sammons RD, Wang X, Ranjan P, Johnson DH, Wadl PA, Scheffler BE, Rinehart TA, Trigiano RN, Stewart CN (2010) Functional genomics analysis of horseweed (Conyza canadensis) with special reference to the evolution of non-target-site glyphosate resistance. Weed Science 58:109-117

Zheng D, Patzoldt WL, Tranel PJ (2005) Association of the W574L ALS substitution with resistance to cloransulam and imazamox in common ragweed (Ambrosia artemisiifolia). Weed Science 53:424-430

Ziska LH, Schlesinger WH, Epstein PR (2009) Rising CO2, climate change, and public health: Exploring the links to plant biology. Environmental Health Perspectives 117:155-158

27 Figure 1.1. The shikimate (chorismate) pathway and products.

Adapted from Duke and Powles (2008b). The site of inhibition by glyphosate is indicated with a red arrow and the dotted arrow indicates regulatory feedback inhibition.

28

Chapter 2: Characterization of Common Ragweed Resistance to Glyphosate, Cloransulam-Methyl, and Fomesafen Herbicides

2.1 Materials and Methods:

2.1.1 Development of a sample population

In Autumn 2009, seed was collected from common ragweed plants that survived multiple glyphosate applications in a soybean field in Clinton County, Ohio, where a large population of glyphosate-resistant plants had developed over time. Seed from a presumably glyphosate-sensitive common ragweed biotype was purchased from Azlin

Seeds in Leland, Mississippi in 2010, and collected from a non-crop area in Columbus,

Ohio in 2011.

Prior to stratification, a sample of seeds from each common ragweed biotype was placed in mesh packets and submerged in a 1000-mL beaker of tap water containing

0.1% v/v non-ionic surfactant to improve permeation of water through the seed hulls.

The packets were removed from the water/surfactant solution and rinsed twice, and then covered in damp sand in a closed chest cooler (for moisture retention) at a temperature of

4˚C for five weeks. The stratified seeds were planted directly into 1.9-L greenhouse containers filled with potting media (Canadian sphagnum peat moss, perlite, dolomitic limestone, composted bark, vermiculite, wetting agent, and starter nutrient charge with gypsum; Fafard® 3B, Sun Gro; Agawam, Massachusetts), and supplemented with 700 g of controlled release fertilizer (Osmocote® Plus Lo-Start® 12–14 Month, 15-9-11, 29 Everris NA; Dublin, OH) blended into each 79.3-L bag of media. The containers were placed into a research greenhouse under 16-hour photoperiod controlled by 1000-watt metal halide lamps that provided approximately 200 µmol m-2 s-1 of supplemental lighting. Air temperatures in the greenhouse were maintained near 27˚C daytime and

20˚C nighttime by a computerized climate management system (EnviroSTEP;

Wadsworth Control Systems; Arvada, Colorado). An automated overhead irrigation system (JetRain PolyRail; Dramm Co.; Manitowoc, Wisconsin) was set to water three times daily, with supplemental hand watering as needed to maintain adequate soil moisture and prevent wilting. Emerged plants were thinned to one seedling per container. Seedlings were approximately 16 cm tall with three-to-five pairs of true leaves four weeks after planting. At this time, 30 plants from the glyphosate-resistant Clinton

County biotype and 10 plants from the presumed glyphosate-sensitive Leyland,

Mississippi biotype were treated with glyphosate. The potassium salt of glyphosate

(Roundup WeatherMAX®, 540 g ai L−1; Monsanto Company; St. Louis, Missouri) was applied at a rate of 8.4 kg ae ha−1 (10 field use rate) with ammonium sulfate (AMS) solution (N-Pak® AMS Liquid, 407 g L−1; Winfield Solutions, LLC; St. Paul, Minnesota) at 5% v/v using a pneumatic track sprayer equipped with an even flat spray tip (Teejet

8001EVS; Spraying Systems Co.; Carol Stream, Illinois) calibrated to apply 140 L ha−1 of spray solution at a speed of 3.5 km hr−1. An additional 10 plants from the glyphosate- sensitive biotype were left untreated. Plant survival was assessed four weeks after treatment. Surviving plants from the glyphosate-resistant biotype were divided into two groups of 10- and 12-plants (R1 parents and R2 parents, respectively) based upon degree of stem dieback, and placed into two separate greenhouse compartments for pollination

30 within groups. The 10 untreated plants from the glyphosate-sensitive biotype were placed into a third greenhouse chamber for pollination (S1 parents). Greenhouse chambers were maintained under conditions similar to those mentioned previously. At senescence, entire plants with seeds were harvested, placed into paper bags, and allowed to dry for one month in the greenhouse. Seeds were subsequently removed from the plants by hand, and cleaned with sieves and an air-powered seed separator. The

Columbus, Ohio biotype (S2) was collected later and not subjected to this greenhouse seed-generation process. Dry seeds were stored in plastic bottles at 4˚C until needed.

2.1.2 Growing Conditions for Dose-Response

Common ragweed seeds from the R1, R2, S1, and S2 progeny populations were placed in mesh bags and stratified in damp sand for eight weeks, after which they were rinsed with water and a 500-ppm solution of peroxyacetic acid disinfectant (X3™;

Phyton Corp; New Hope, Minnesota) and sown in germination towels at 25˚C. Seeds were checked daily, and once radicals emerged, were planted into 0.8-L greenhouse containers (Dillen 5AZATW; Myers Industries; Akron, Ohio) with potting media, fertility, and greenhouse conditions as described previously.

2.1.3 Treatments

To mimic the microbial environment of field soil, and supplement herbicidal activity, soil samples were collected from several field locations where common ragweed and giant ragweed were found growing (Bethel, 2013). The field soil was periodically blended by hand with composted and fresh plant material, including common and giant ragweed, to create a stockpile of inoculum for subsequent experiments. A slurry was 31 created by blending a 1 to 5 ratio of the non-sterile soil and tepid water in a powered concrete mixer (Model No. 817, The J.B. Foote Coundry Company, Fredericktown,

Ohio) for one hour, making about 20 L of slurry. Continuing agitation, aliquots of 150- mL slurry were applied to the surface of media around each plant approximately 24 hours before glyphosate application, and watered into the media with overhead irrigation.

Seedlings were 10-to-15 cm tall with three-to-five pairs of mature true leaves four weeks after planting. At this time, the plants were treated with the potassium salt of glyphosate

(MON 78623, 474 g ae kg−1; Monsanto Company). The glyphosate was applied at rates of 0, 0.012, 0.06, 0.12, 0.3, 0.6, 1.2, 2.4, 4.8, 12, 24, and 60 kg ae ha−1 with 0.25% v/v surfactant (MON 56151; Monsanto Company) and 5% v/v N-Pak AMS liquid. Herbicide was applied with a pneumatic track sprayer as described previously. The lowest rate

(0.012 kg ae ha−1) was only applied to the S1 and S2 biotypes, and the highest rate

(60 kg ae ha−1) was only applied to R1 and R2 biotypes. Treatments were arranged as a randomized complete block design with six replicates, and maintained under greenhouse conditions similar to those described previously, except no irrigation was applied for

24 hours following glyphosate treatment.

A second run was conducted similarly, except glyphosate rates were shifted to 0-,

0.012, 0.06, 0.12, 0.24, 0.48, 0.75, 1.2, 2.4, 4.8, 12, and 24 kg ae ha−1 to give better confidence in the response curve around lower doses. The lowest rate (0.012 kg ae ha−1) was only applied to the S1 and S2 biotypes, and the highest rate (24 kg ae ha−1) was only applied to R1 and R2 biotypes. Four replicates were used for the R1, R2, and S2 biotypes, and three replicates were used for the S1 biotype.

32 Similar dose-response assays were conducted for the herbicides cloransulam- methyl and fomesafen. Cloransulam, an acetolactate synthase (ALS)-inhibitor, and fomesafen, a protoporphyrinogen IX oxidase (PPO)-inhibitor, are two alternatives to glyphosate that can be effective for common ragweed control. An initial fomesafen dose- response assay (results not shown) and the assay for cloransulam were conducted along with the second glyphosate dose-response run. The original Clinton County common ragweed population (R) was known to have little or no control with ALS-inhibitors and declining control with PPO-inhibitors (Loux and Stachler, personal communication).

Selection for the R1 and R2 biotypes in the greenhouse had not involved ALS- or

PPO-inhibitors, so they were presumed to be equivalent in their response to ALS- and

PPO-inhibitors. Therefore, plants of uniform size (8-to-10 cm tall with two-to-three pairs of mature true leaves) were selected from both R-biotypes for these experiments. Only the S2-biotype was used as a presumed-sensitive control for the cloransulam study. Rates of cloransulam-methyl (FirstRate® 84DF, 84% w/w; Dow AgroSciences; Indianapolis,

Indiana) were 0, 0.007, 0.07, 0.71, 3.6, 7.1, and 14 g ai ha−1 for the S2 biotype, and 0, 7.1,

14, 71, 710, and 7100 g ai ha−1 for the R biotype. The cloransulam spray mixtures contained 0.25% v/v nonionic surfactant (NIS) (Activator 90; Loveland Products;

Greenley, Colorado) and 5% v/v N-Pak® AMS Liquid. Five replicates of each biotype were used in this study.

The fomesafen dose-response assays reported here were conducted later, and both the S1- and S2-biotypes were used as controls. The sodium salt of fomesafen

(Flexstar® 1.88L, 225 g ai L−1; Syngenta Crop Protection, LLC; Greensboro, North

Carolina) was applied at rates of 0, 0.11, 0.56, 2.8, 14, 70, 175, 350, 700, and 1750 33 g ai ha−1, with Activator 90 NIS at 0.5% v/v and N-Pak® AMS Liquid at 5% v/v. The

175, 700, and 1750 g ai ha−1 rates of fomesafen were only applied to the R biotype. The fomesafen experiment was repeated with slightly larger plants (10 to 12 cm tall; 4 to 5 pairs of true leaves). In the second fomesafen experiment, the 0.11-g ai ha−1 rate was not used, the rates of 0.56 to 700 g ai ha−1 from the first run were applied to all biotypes, and rates of 1750 and 8750 g ai ha−1 were applied to the R biotype. Additionally, crop oil concentrate (Prime Oil®; Winfield Solutions, LLC) was used at 0.5% v/v in place of

NIS, to compensate for the larger plant size. Spray application parameters for the cloransulam and fomesafen experiments were identical to those described previously.

Five replicates of the R biotype and four replicates of each sensitive biotype were used in each run of the fomesafen study.

2.1.4 Data Collection and Analysis

Injury relative to the untreated controls and mortality was visually estimated, and shoot fresh weight was measured, 21 days after treatment (DAT) in the glyphosate experiments and 28 DAT in the cloransulam-methyl and fomesafen experiments. Fresh weight data were converted to percentages of the mean fresh weight of untreated control plants in each respective biotype for presentation.

Data from the glyphosate and fomesafen experiments were analyzed using the

PROC MIXED procedure of SAS (SAS Institute, Cary, North Carolina) with biotype, run, and treatment as class (independent) variables, and replicate as a random effect in a three-way combined ANOVA. The run by treatment interactions were significant for the visual injury estimation (data not shown), adjusted fresh weight, and mortality data (p <

34 0.05), likely due to the changes in rates between runs—but more importantly for these analyses, the run by biotype and run by biotype by treatment interactions were not significant (p >> 0.05). Therefore, data from both runs were pooled for regression analyses of glyphosate and fomesafen dose-responses.

Fresh weight data from the glyphosate study were subjected to regression analysis with three-parameter Weibull-2 functions (Equation [2.1]):

�(�) = � (1 − exp− exp�(log(�) − log(�))) [2.1] where ( x ) is the rate of herbicide, ( d ) is the upper limit, ( e ) is the inflection point, about which the function is asymmetric and not equal to the GR50 (estimated herbicide dose required to reduce fresh weight by 50%), and ( b ) is the slope around ( e ). Euler’s constant is expressed as ( exp ) here to differentiate from the variable ( e ).

Asymmetry was less-clearly defined by plant responses to the chosen doses of cloransulam and fomesafen, so these curves were estimated using three-parameter log- logistic functions (Equation [2.2]):

d f (x) = [2.2] 1+ exp(b(log(x)- log(e))) where parameters ( x, d, b, and exp ) are as described for Equation [2.1], and the function is symmetric around ( e ), which is equal to the GR50.

35 Mortality data were best described by a two-parameter binary log-logistic function (Equation [2.3]):

1 f (x) = [2.3] 1+ exp(b(log(x)- log(e))) where parameters (x, b, and exp) are as described for Equation [2.1], parameter (e) is as described for Equation [2.2], and parameter (d) from Equation [2.2] is equal to 1. The drc argument type=”binomial” was used for mortality data (Ritz and Strebig, 2013).

Regression was analyzed using the drc package in R (R Core Team, 2014; Ritz and Streibig, 2005), including estimation of herbicide doses required to reduce fresh weight by 50% (GR50) and 90% (GR90), and estimation of doses required to kill 50% of the biotype (LD50) and 90% of the biotype (LD90) values. The relative magnitude of herbicide resistance in the resistant biotypes was estimated by calculating R:S ratios

(GR50 of resistant biotype/GR50 of susceptible biotype). Tests of the same population regression model were made using an F-test (Equation [2.4]) for comparing the regression equations (Harrison et al., 2001; Zar, 1996):

SSt - SSp (m +1)(k -1) F = [2.4] SSp

DFp

where (SSt) is the total residual sums of squares from regression of the combined biotypes or runs, (SSp) is the pooled residual sums of squares, equal to the sum of the residual sums of squares from the individual runs, (m) is the number of independent variables, (k)

36 is the number of regressions being compared, and (DFp) is the pooled residual degrees of freedom, equal to the sum of residual degrees of freedom from compared regressions.

Combined regression analysis data are presented across susceptible biotypes and runs for analyses in which the null hypotheses for the F-test were accepted.

2.2 Results and Discussion

2.2.1 Glyphosate dose-response results

The S1 and S2 biotypes were confirmed to be susceptible to glyphosate, with

-1 -1 GR50 estimates of 0.05 and 0.03 kg ae ha and GR90 estimates of 0.4 and 0.3 kg ae ha , respectively (Table 2.1; Figure 2.1). The R1 and R2 biotypes both were confirmed resistant to glyphosate, with GR50 estimates of 0.35 and 0.20 and GR90 estimates of 4.8

-1 and 1.7 kg ae ha , respectively. The R:S ratios of GR50 estimates range from 4:1 (R2:S1) to 11:1 (R1:S2), and GR90 R:S ratios range from 4:1 (R2:S1) to 15:1 (R1:S2).

The GR90 estimates for the R1 and R2 biotypes were six- and two-fold higher,

-1 respectively, than the standard 0.84 kg ae ha field use rate of glyphosate, while the GR90 estimates for the S1 and S2 biotypes were below labeled field use rates (50% and 40% of the standard rate, respectively). To provide a frame of reference, it is notable that the maximum rate of glyphosate currently allowable in single application for soybeans is

-1 1.7 kg ae ha (two times the “normal” use rate), which was the GR90 estimate for the R2 sample of the Clinton County biotype. This demonstrates that some resistant plants are clearly not immune to field rates of glyphosate, but instead exhibit elevated levels of tolerance to the herbicide. Shifting the focus to plant survival, the LD50 estimates for the

R1 and R2 biotypes were 6.8 and 2.7 kg ae ha-1, respectively (Table 2.2; Figure 2.2). 37 These are eight-fold (R1) and three-fold (R2) higher than the typical use rate of

-1 0.84 kg ae ha , and 5-fold (R2:S1) to 19-fold (R1:S2) higher than the LD50 estimates for the sensitive biotypes. LD90 estimates and R:S ratios are also shown in Table 2.2. The

R:S ratio was even larger for estimates of 90% mortality (LD90), ranging from 9:1

(R2:S1) to 30:1 (R1:S2). The LD50 range we observed is similar to that of glyphosate- susceptible and glyphosate-resistant common ragweed biotypes reported by Brewer and

Oliver (2009). This could be an indication that similar resistance mechanisms are involved in the Ohio and Arkansas biotypes.

2.2.2 Cloransulam-methyl dose-response results

Resistance to cloransulam-methyl was also confirmed in the Clinton County,

Ohio (R) biotype of common ragweed (Figure 2.3; Figure 2.4). Only one individual in this biotype was affected enough by the highest dose of 7100 g ai ha-1 (1000-fold higher than the field use rate of 7.1 g ai ha-1) to be evaluated as dead 28 DAT. One plant seemed nearly unaffected at the highest dose, and mean growth reduction only reached 65%. In contrast, the highest tested dose at which any replicates from the S2 biotype survived was

-1 0.71 g ai ha —one tenth of the field use rate. The GR50 and GR90 estimates for the S2

-1 biotype were 0.1 and 1.2 g ai ha , respectively, and the LD50 and LD90 estimates were 0.6 and 0.8 g ai ha-1 (Table 2.3 and Table 2.4). For the S2 biotype, all estimates were well below the recommended field use rate of cloransulam-methyl. Estimates of effective doses for the R biotype are difficult to calculate without a measured upper limit, but the

-1 GR50 can be estimated at approximately 4800 g ai ha , for an R:S2 ratio in the vicinity of

50,000-to-1.

38 2.2.3 Fomesafen dose-response results

Data from the S1 and S2 biotypes of common ragweed were pooled for comparison to the Clinton County, Ohio (R) biotype in the fomesafen studies, and will be referred to collectively as the “S” biotype. As suspected, the R biotype was confirmed resistant to fomesafen. Fomesafen was effective for control of the S biotypes, with an

-1 -1 estimated GR50 of 3.5 g ai ha , only one-percent of the 1 rate of 350 g ai ha (Table 2.5;

Figure 2.5). The S plants were uniformly sensitive, progressing quickly to an estimated

-1 GR90 of 15 g ai ha . Conversely, the R biotype did not appear to be fully segregated for resistance, with some individual plants appearing to respond as sensitive or intermediate in phenotype. All plants suffered at least transient damage from fomesafen rates of 2.8 g

-1 -1 ai ha and higher. The GR50 estimate for the R biotype was 48 g ai ha , which was 14- fold higher than the GR50 of the S biotype. The estimated GR90 for the R biotype was

540 g ai ha-1, a 36-fold resistance ratio.

Analysis of plant survival showed a clearer difference between the R and S common ragweed biotypes (Table 2.6; Figure 2.6). Nearly one-third of the S plants treated with fomesafen at a rate of 14 g ai ha-1 were killed, and 15 out of 16 S plants

-1 succumbed to the 70 g ai ha rate, leading to LD50 and LD90 estimates of 20 and 50 g ai ha-1, respectively. The mixed nature of the R biotype was evident from mortality of fewer than 30% of the individuals treated with each rate of 70, 175, 350, and 700 g fomesafen ha-1, and vigorous regrowth of other individuals at the same rates. The

-1 estimated LD50 for the R biotype was 1400 g ai ha , a 70-fold resistance ratio. Five R plants were treated with fomesafen at 8750 g ai ha-1, and one plant was surviving after 28

39 days. The lack of a dose providing 90% to 100% control did not allow for precise estimation of an LD90.

2.2.4 Discussion

This is the first report of a common ragweed biotype with multiple resistance to herbicides from three site of action (SOA) groups (Heap, 2014). Few herbicide options currently exist for postemergence control of common ragweed in soybeans, outside of these three SOA groups. Aggressive management techniques integrating varied cultural practices, such as crop rotation, herbicide rotation, and tillage, should be employed to prevent development of resistance to other herbicides. Resistance to PPO-inhibitors has been relatively slow to develop, with only six species worldwide having reported resistant biotypes, compared to 145 species with biotypes resistant to ALS-inhibitors

(Heap, 2014). However, the outcrossing nature of common ragweed increases the possibility of gene flow transferring resistance from outside populations, if not already present within a population. This enhances the ability of common ragweed to evolve rapidly, particularly under heavy selection pressure from a single herbicide SOA.

Herbicide combinations were not used in these studies, but it can be assumed that resistances to all three SOA groups are present in most individuals from the Clinton

County, Ohio (R) biotype, based on observations that nearly 100% of the individual plants treated with glyphosate or cloransulam-methyl showed resistance. Combinations of these herbicides could provide some growth suppression, and the extent of cross- resistance to other chemical families within these SOA groups is unknown.

40 Chapter 2 References

Bethel JD (2013) Evaluation of glyphosate resistant giant ragweed (Ambrosia trifida) in Ohio soybean (Glycine max) fields. MS Thesis. Columbus, OH: The Ohio State University

Brewer CE, Oliver LR (2009) Confirmation and resistance mechanisms in glyphosate- resistant common ragweed (Ambrosia artemisiifolia) in Arkansas. Weed Science 57:567-573

Harrison SK, Regnier EE, Schmoll JT, Webb JE (2001) Competition and fecundity of giant ragweed in corn. Weed Science 49:224-229

Heap I (2014) The international survey of herbicide resistant weeds. http://www.weedscience.com. Accessed November 5, 2014

R Core Team (2014) R: A Language and Environment for Statistical Computing. R Foundation for Statistical Computing. Vienna, Austria. http://www.R-project.org/.

Ritz C, Strebig J (2013) Package ‘drc’.

Ritz C, Streibig JC (2005) Bioassay analysis using R. Journal of Statistical Software 12:1-22

Zar JH (1996) Biostatistical Analysis. 3rd edn. Upper Saddle River, NJ: Prentice-Hall

41 Table 2.1. Estimated glyphosate doses resulting in 50% (GR50) or 90% (GR90) reduction in fresh weight relative to untreated controls and resistance ratios of R1, R2, S1, and S2 common ragweed biotypes in greenhouse dose-response studies.

Estimate GR ± SE Biotype xx R:S1 p R:S2 p Type (kg ae ha−1) R1 0.35 ± 0.11 6.8 0.046 11.4 0.034 R2 0.20 ± 0.05 3.8 NS 6.4 0.036 GR50 S1 0.051 ± 0.015 – – – – S2 0.031 ± 0.009 – – – – R1 4.8 ± 1.1 12 0.004 15 0.004 R2 1.7 ± 0.4 4.1 0.011 5.3 0.008 GR90 S1 0.42 ± 0.09 – – – – S2 0.33 ± 0.07 – – – – Glyphosate doses are presented as kg ae ha−1 ± standard errors of the estimates. Fresh weights were measured 21 days after treatment (DAT) and data from two experimental runs were pooled. GR50 and GR90 estimates and resistance ratios (R:S) for glyphosate-resistant (R1 and R2) and glyphosate-sensitive (S1 and S2) common ragweed biotypes were calculated from curves fit using three-parameter Weibull-2 functions (Figure 2.1; Equation [2.1]). Recommended field use rates of glyphosate range from 0.84 to 1.7 kg ae ha-1.

42 Table 2.2. Estimated glyphosate doses lethal for 50% (LD50) or 90% (LD90) of R1, R2, S1, and S2 common ragweed biotypes in greenhouse dose-response studies.

Estimate LD ± SE Biotype xx R:S1 p R:S2 p Type (kg ae ha−1) R1 6.8 ± 1.60 13 0.001 19 0.001 R2 2.7 ± 0.56 5.3 0.002 7.4 0.002 LD50 S1 0.5 ± 0.08 – – – – S2 0.4 ± 0.06 – – – – R1 30 ± 12 26 0.049 31 0.050 R2 10 ± 3.6 8.8 0.043 10 0.042 LD90 S1 1.1 ± 0.3 – – – – S2 1.0 ± 0.3 – – – – Glyphosate doses are presented as kg ae ha−1 ± standard errors of the estimates. Plants were visually scored live or dead 21 days after treatment (DAT) and data from two experimental runs were pooled. LD50 and LD90 estimates and resistance ratios (R:S) for glyphosate-resistant (R1 and R2) and glyphosate-sensitive (S1 and S2) common ragweed biotypes were calculated from curves fit using two-parameter log-logistic functions (Figure 2.2; Equation [2.3]). Recommended field use rates of glyphosate range from 0.84 to 1.7 kg ae ha-1.

43 Table 2.3. Estimated clorasulam doses resulting in 50% (GR50) or 90% (GR90) reduction in fresh weight relative to untreated controls of R and S2 common ragweed biotypes in a greenhouse dose-response study.

Estimate GR ± SE 95% Biotype xx Type (g ai ha−1) Confidence Interval R 4800 ± 3000 −1400–11000 GR50 S2 0.079 ± 0.03 0.009–0.148 R > 7100 – GR 90 S2 1.1 ± 0.3 0.51–1.76 Cloransulam-methyl doses are presented as g ai ha−1 ± standard errors of the estimates. Fresh weights were measured 28 days after treatment (DAT) in one experimental run. GR50 and GR90 estimates and 95% confidence intervals for resistant (R) and sensitive (S2) common ragweed biotypes were calculated from curves fit using three-parameter log-logistic functions (Figure 2.3; Equation [2.2]). The recommended field use rate of cloransulam-methyl is 7.1 g ai ha-1.

44 Table 2.4. Estimated cloransulam doses lethal for 50% (LD50) or 90% (LD90) of R and S2 common ragweed biotypes in a greenhouse dose-response study.

Estimate LD ± SE 95% Biotype xx Type (g ai ha−1) Confidence Interval R > 7100 – LD50 S2 0.57 ± 1.18 −1.74–2.87 R > 7100 – LD90 S2 0.82 ± 1.00 −1.15–2.78 Cloransulam-methyl doses are presented as g ai ha−1 ± standard errors of the estimates. Plants were visually scored live or dead 28 days after treatment (DAT). LD50 and LD90 estimates and 95% confidence intervals for the herbicide-sensitive (S2) common ragweed biotype were calculated from curves fit using two-parameter log-logistic functions (Figure 2.4; Equation [2.3]). The herbicide-resistant Clinton County biotype did not approach 50% mortality 28 DAT at the highest tested rate of cloransulam. The recommended field use rate of cloransulam-methyl is 7.1 g ai ha-1.

45 Table 2.5. Estimated fomesafen doses resulting in 50% (GR50) or 90% (GR90) reduction in fresh weight relative to untreated controls and resistance ratios of R and S common ragweed biotypes in greenhouse dose-response studies.

Estimate GR ± SE Biotype xx R:S p Type (g ai ha−1) R 47.6 ± 12.8 13.6 0.004 GR50 S 3.49 ± 0.60 – – R 539 ± 103 35.7 < 0.001 GR90 S 15.1 ± 2.47 – – Fomesafen doses are presented as g ai ha−1 ± standard errors of the estimates. Fresh weights were measured 28 days after treatment (DAT) and data from two experimental runs were pooled. GR50 and GR90 estimates and resistance ratios (R:S) for resistant (R) and sensitive (pooled S1 and S2) common ragweed biotypes were calculated from curves fit using three-parameter log-logistic functions (Figure 2.5; Equation [2.2]). Recommended field use rates of fomesafen range from 350 to 420 g ai ha-1.

46 Table 2.6. Estimated fomesafen doses lethal for 50% (LD50) or 90% (LD90) of R and S common ragweed biotypes in greenhouse dose-response studies.

Estimate LD ± SE Biotype xx R:S p Type (g ai ha−1) R 1400 ± 629 69.3 0.046 LD50 S 20.2 ± 4.1 – – R 19 000 ± 19 000 > 100 NS LD90 S 50.1 ± 16.9 – – Fomesafen doses are presented as g ai ha−1 ± standard errors of the estimates. Plants were visually scored live or dead 28 days after treatment (DAT) and data from two experimental runs were pooled. LD50 and LD90 estimates and resistance ratios (R:S) for resistant (R) and sensitive (pooled S1 and S2) common ragweed biotypes were calculated from curves fit using two- parameter log-logistic functions (Figure 2.6; Equation [2.3]). Recommended field use rates of fomesafen range from 350 to 420 g ai ha-1.

47

Figure 2.1. Fresh weight response curves of R1, R2, S1, and S2 common ragweed biotypes to varied doses of glyphosate in greenhouse studies. ) l 100 R1 o r t R2 n o

c S1

80 f

o S2

% (

t 60 h g i e 48 W

40 h s e r F

t 20 o o h S 0

0 0.0084 0.084 0.84 8.4 84 Glyphosate Dose (kg glyphosate ae ha-1)

Glyphosate dose-response curves of glyphosate-resistant (R1 and R2) and glyphosate-sensitive (S1 and S2) common ragweed biotypes. Fresh weights were measured 21 days after treatment (DAT) and data from two experimental runs were pooled. Each point represents the mean fresh weight of three to ten replicates as a percentage of the respective control mean fresh weight. Curves were fit using a three-parameter Weibull-2 function (Equation [2.1]). Recommended field use rates of glyphosate range from 0.84 to 1.7 kg ae ha-1.

Figure 2.2. Mortality response curves of R1, R2, S1, and S2 common ragweed biotypes to varied doses of glyphosate in greenhouse studies.

100

80 y t i l

a 60 t r

o R1 M

49 R2

% 40 S1 S2 20

0

0 0.0084 0.084 0.84 8.4 84 Glyphosate Dose (kg glyphosate ae ha-1)

Glyphosate dose-response curves of glyphosate-resistant (R1 and R2) and glyphosate-sensitive (S1 and S2) common ragweed biotypes. Plants were visually scored live or dead 21 days after treatment (DAT) and data from two experimental runs were pooled. Each point represents the percent mortality of three to ten replicates. Curves were fit using a two-parameter log-logistic function (Equation [2.3]). Recommended field use rates of glyphosate range from 0.84 to 1.7 kg ae ha-1.

Figure 2.3. Fresh weight response curves of R and S2 common ragweed biotypes to varied doses of cloransulam in a greenhouse study. ) l 100 S o r t R n o c

80 f o

% (

t 60 h g i e 50 W

40 h s e r F

t 20 o o h S 0

0 0.0071 0.071 0.71 7.1 71 710 7100 Cloransulam Dose (g cloransulam ai ha-1)

Cloransulam-methyl dose-response curves of resistant (R) and sensitive (S) common ragweed biotypes. Fresh weights were measured 28 days after treatment (DAT). Each point represents the mean fresh weight of five replicates as a percentage of the respective control mean fresh weight. Curves were fit using a three-parameter log-logistic function (Equation [2.2]). The recommended field use rate of cloransulam- methyl is 7.1 g ai ha-1.

Figure 2.4. Mortality response curves of R and S2 common ragweed biotypes to varied doses of cloransulam in a greenhouse study.

100

80 y t

i S l

a 60 R t r o M 51

% 40

20

0

0 0.0071 0.071 0.71 7.1 71 710 7100 Cloransulam Dose (g cloransulam ai ha-1)

Cloransulam-methyl dose-response curves of resistant (R) and sensitive (S) common ragweed biotypes. Plants were scored live or dead 28 days after treatment (DAT). Each point represents the percent mortality of five replicates. Curves were fit using a two-parameter log-logistic function (Equation [2.3]). The recommended field use rate of cloransulam-methyl is 7.1 g ai ha-1.

Figure 2.5. Fresh weight response curves of R and S common ragweed biotypes to varied doses of fomesafen in greenhouse studies. ) l 100 R o r t S n o c

80 f o

% (

t 60 h g i e 52 W

40 h s e r F

t 20 o o h S 0

0 0.35 3.5 35 350 3500 35000 Fomesafen Dose (g fomesafen ai ha-1)

Fomesafen dose-response curves of resistant (R) and sensitive (S) common ragweed biotypes. Fresh weights were measured 28 days after treatment (DAT) and data from two experimental runs were pooled. Each point represents the mean fresh weight of five to sixteen replicates as a percentage of the respective control mean fresh weight. Curves were fit using a three-parameter log-logistic function (Equation [2.2]). Recommended field use rates of fomesafen range from 350 to 420 g ai ha-1.

Figure 2.6. Mortality response curves of R and S common ragweed biotypes to varied doses of fomesafen in greenhouse studies.

100

80 y t i l

a 60 t r o M

%

53 40

R 20 S

0

0 0.35 3.5 35 350 3500 35000 Fomesafen Dose (g fomesafen ai ha-1)

Fomesafen dose-response curves of resistant (R) and sensitive (S) common ragweed biotypes. Plants were scored live or dead 28 days after treatment (DAT) and data from two experimental runs were pooled. Each point represents the percent mortality of five to sixteen replicates as a percentage of the respective control mean fresh weight. Curves were fit using a two-parameter log-logistic function (Equation [2.3]). Recommended field use rates of fomesafen range from 350 to 420 g ai ha-1.

Chapter 3: Common Ragweed Target-Site Glyphosate-Resistance Mechanisms

3.1 Materials and Methods

3.1.1 Plant Materials

Common ragweed plants from the R1, R2, S1, and S2 progeny populations were grown in the greenhouse using Fafard 3B (Sun Gro) potting media (Canadian sphagnum peat moss, perlite, dolomitic limestone, composted bark, vermiculite, wetting agent, starter nutrient charge with gypsum), and supplemented with 700 g of Osmocote Plus

Lo-Start slow-release fertilizer (Scotts Professional) blended into each 79.3-L bag of media. Plants were grown at air temperatures of approximately 27˚C daytime and 20˚C nighttime, with a 16-hour photoperiod controlled by 1000-watt metal halide lamps that provided approximately 200 µmol m-2 s-1 of supplemental lighting. An automated overhead irrigation system (Dramm) was set to water three times daily, with supplemental hand watering as needed to maintain adequate soil moisture and prevent wilting.

3.1.2 Genomic DNA extraction

A 100-mg sample of young leaf tissue was collected in 1.5-mL tubes and immediately frozen in liquid nitrogen. This tissue was then ground to a fine powder using a pestle drill bit and an electric bench-top drill. The DNA was extracted from the powdered leaf tissue using the Qiagen DNeasy Plant Mini Kit and the manufacturer’s

54

protocol. Concentration of the extracted genomic DNA in buffer was determined spectrophotometrically using a Nanodrop 1000 (Thermo Fisher Scientific) and the manufacturer's protocol. An aliquot of each DNA sample was diluted to 4 ng µL-1 and the original and diluted samples were stored at −20 ̊C.

3.1.3 RNA extraction and complementary DNA synthesis

A 40-mg sample of young leaf tissue was collected in a 1.5-mL tube and immediately frozen in liquid nitrogen. This tissue was ground to a fine powder using a pestle drill bit and an electric bench-top drill. RNA was then extracted from the powder using the Qiagen RNeasy Mini Kit and the manufacturer’s plant RNA extraction protocol, except for the optional on-column DNase digestion. Concentration of the extracted RNA in buffer was determined spectrophotometrically using a Nanodrop 1000

(Thermo Fisher Scientific) and the manufacturer's protocol. An aliquot of each RNA sample containing 1 µg of RNA was transferred to 0.2-mL PCR tubes on ice and diluted to 8 µL in diethylpyrocarbonate (DEPC)-treated water. The remaining RNA was stored at −80 ̊C. To the aliquots of RNA, Invitrogen DNase I (Amplification Grade) and buffer were added, and DNase digestion and deactivation were performed according to the manufacturer's protocol. First-strand complementary DNA (cDNA) synthesis was conducted using these DNase-treated samples with the Promega Reverse Transcription

System. Each reaction contained 11 µL DNase-treated RNA, 4 µL 25-mM MgCl2, 2 μl

Reverse Transcription 10 Buffer, 2 μl 10-mM dNTP Mixture, 0.5 μl Recombinant

RNasin Ribonuclease Inhibitor, 15 units avian myeloblastosis virus (AMV) Reverse

55

Transcriptase (RT), and 0.5 µg Oligo(dT)15 Primer. No–RT controls were also included. cDNA was then stored at −20˚C.

3.1.4 PCR primer design

Most sequencing of the 5-enolpyruvylshikimate-3-phosphate synthase (EPSPS) gene was focused in the region typically containing a proline in the 106th amino acid position (Pro106), with numbering based on the maize EPSPS sequence. This region is near the phosphoenolpyruvate (PEP) binding site of EPSPS, and has been found in other weed species to contain a substitution for serine, threonine, or alanine, conferring a low level of glyphosate resistance (Preston et al., 2009). Polymerase chain reaction (PCR) primers for EPSPS in common ragweed had not been previously developed, and published genetic resources for Ambrosia species and closely related genera are limited.

Transcriptome data are available for two plants of common ragweed as short sequence read archives (SRA) of raw 454 pyrosequencing data on the National Center for

Biotechnology Information (NCBI) website, ncbi.nlm.nih.gov (accession numbers

SRX098769 and SRX096892). On an OSU server, the SRA files were extracted using

SRA Toolkit software (CentOS Linux 64-bit, version 2.1.2, available on the NCBI website), linked, and assembled with Mira software (version 3.0, http://chevreux.org/projects_mira.html). A “basic local alignment search tool” (BLAST) search of the assembled sequences was conducted using willowleaf sunflower

(Helianthus salicifolius; family Asteraceae; tribe Heliantheae) EPSPS protein sequence

(NCBI GenBank accession numbers AAT45238.1 and AAT45239.1). This returned eight contiguous sequences (contigs) of common ragweed EPSPS cDNA, ranging from

56

608-to-1808 base pairs (bp) in length. Four of these contained the region coding for

Proline 106. EPSPS primers for PCR were designed using each of these four contigs with Primer3, through the NCBI Primer-BLAST (Ye et al., 2012), and synthesized by

Eurofins MWG Operon (www.operon.com). Initial design of eight EPSPS primer pairs yielded no effective combinations, so a more rigorous approach of varied design parameters was conducted, again using Primer-BLAST and cross-referencing with

Integrated DNA Technologies (IDT) OligoAnalyzer Tool

(www.idtdna.com/analyzer/Applications/OligoAnalyzer/, 2012). With this approach,

16 forward primers and 22 reverse primers for EPSPS were designed and synthesized, with most calculated to be usable in various combinations. In a first screen, several pairs of EPSPS primers appeared to be usable.

PCR primers for acetolactate synthase (ALS) regions A and B sequencing were ordered from Operon, using published primer sequences (Patzoldt et al., 2001). Primers for fructan 1-exohydrolase IIa (FEH) were designed as described above for EPSPS, using sequence obtained by a BLAST search of the assembled common ragweed transcriptome sequences, with chicory (Cichorium intybus; family Asteraceae) FEH IIa sequence (NCBI GenBank accession number AY323935.1).

For real-time quantitative PCR (qPCR), new EPSPS primer pairs spanning approximately 100-to-200 bp were designed under stringent parameters using the software described previously, but using consensus EPSPS and FEH sequences obtained as described in the following section. Primers designed by the software were compared to sequence reads aligned using Sequencher 5.0 software (Gene Codes

57

Corp.), including two types of putative EPSPS alleles from multiple plants in the biotypes to be studied. Raw sequence chromatograms of the aligned reads were inspected for polymorphisms within the primer sequence. Primer sequences were then accepted, rejected, or modified manually and re-checked using the design software for predicted annealing temperatures and compatibility defects. Newly synthesized EPSPS primers were ordered as described previously, and FEH primers were chosen from previously purchased primer pairs.

3.1.5 Gene sequencing of EPSPS, acetolactate synthase (ALS), and fructan

1-exohydrolase IIa (FEH)

The primer pair chosen for initial sequencing reactions was EPSPS-H (Table

3.1). These primers span an intron of variable size among alleles, producing a PCR product ranging from approximately 400 bp to 1200 bp. 20-µL PCR reactions for sequencing contained 10 µL GoTaq Green Master Mix (Promega Corporation), 3 µL

PCR-pure water, 1 µL of each primer (5 µM in water), and 5 µL of genomic DNA template (4 µM in water), or alternatively, some reactions used 0.1 µL TaKaRa Ex Taq

DNA polymerase (TaKaRa Bio, Inc.), 9.9 µL PCR-pure water, 2 µL 10 Ex Taq Buffer,

2 µL (deoxyribonucleotide monomers (dNTPs) mixture (2.5 mM each of dATP, dTTP, dGTP, and dCTP), 1 µL of each primer (5 µM in water), and 5 µL of genomic DNA template (4 µM in water). Reactions with these primers were performed with a Veriti

(Applied Biosystems) or MJ Research PTC-200 (Bio-Rad) thermal cycler, equipped with a 96-well block. Amplification reactions began with an initial two-minute template denaturation step at 95ºC, followed by 35 cycles of 95ºC denaturation, 56ºC

58

annealing, and 72ºC extension steps of one minute each, and ending with a final extension period of seven minutes at 72ºC. PCRs required lower than calculated annealing temperatures to achieve acceptable EPSPS amplification. After the final extension step, samples were held at 4ºC to 10ºC until electrophoresis. Electrophoresis was performed with 1- to 2-percent agar mixed with 1 tris-acetate-EDTA (TAE) buffer, and run in 1 TAE buffer at 80-to-100-volts. 6 electrophoresis loading dye with glycerol loading-aid was added to samples if necessary, and a 1-kb Plus band size reference ladder (Invitrogen) was included in each row of samples. Ethidium bromide was blended with the gel and running buffer at 0.5 µg mL-1 for UV-detection of bands.

After electrophoresis, gels were photographed and desired bands were excised on a

UV-light box. All visible bands desired for sequencing were placed in 1.5-mL microcentrifuge tubes and weighed, then PCR products were extracted from the gel pieces following all recommended procedures with the QiaQuick Gel Extraction Kit

(Qiagen). When necessary, DNA concentrations of the purified samples were measured using a NanoDrop 1000 spectrophotometer (Thermo Fisher Scientific). Each sample was sequenced in both the forward and the reverse directions, with each of previously mentioned primers, respectively. Samples were sent for Sanger sequencing at Colorado

State University, or adjusted to specified concentrations based upon PCR product length and sent for Sanger sequencing at the Plant-Microbe Genomics Facility at Ohio State

University. Sequencher 5.0 software was used for multiple sequence alignment, raw chromatogram visualization, and polymorphism detection. Careful visual analysis of aligned raw sequence chromatogram alignments was necessary to determine whether indicated polymorphisms were consistently shown in the sample, or potentially

59

sequencing errors. Samples of cDNA were prepared for sequencing at the the Plant-

Microbe Genomics Facility at Ohio State University and analyzed as described previously, except three different EPSPS primer pairs (EPSPS-BBB, EPSPS-CCC, and

EPSPS-DDD), as well as two ALS primer pairs (ALS-A and ALS-B) and two FEH primer pairs (FEH54 and FEH34) ordered as described previously, were used, and the annealing temperature was lowered to 52ºC (Table 3.1).

3.1.6 EPSPS Enzyme Activity

For EPSPS activity assays, 30-to-40 g per plant of young, healthy leaf tissue was collected over time from greenhouse-grown glyphosate-sensitive (S1 and S2) and glyphosate-resistant (R1) common ragweed biotypes, weighed in 50-mL conical centrifuge tubes (BD Biosciences), frozen immediately in liquid nitrogen, and maintained at −80ºC until use. Protein extraction was performed similarly to Gaines et al. (2009). During tissue grinding, polyvinylpolypyrrolidone (PVPP) was added at a ratio of 1 g to 5 g leaf tissue (for adsorption of phenolics), and trypsin inhibitor was increased to 200 mg L-1 in the extraction buffer. All remaining extraction steps were carried out with chilled equipment, on ice or with refrigeration at 4ºC. Extraction buffer was added at a rate of 150 mL per 30 g of frozen, ground tissue powder, which was further ground in the buffer, then homogenized with an electric rotor-stator for five minutes. Homogenate was transferred to centrifuge tubes, and centrifuged for

30 minutes at 40,000 g. After centrifugation, the supernatant was poured through

Miracloth (EMD Millipore) into beakers to capture loose solids. Ammonium sulfate

((NH4)2SO4) precipitation was carried out following the procedures of Gaines et al.

60

(2009), with sequential additions to 45% and 70% of saturation, except stirred for

30 minutes and centrifuged at 40,000 g for 30 minutes. After dissolving the final centrifugation pellet in a minimal amount of extraction buffer, the extract was placed into 10,000-molecular weight (kD) cutoff dialysis tubing, and dialyzed for 16 hours at

4ºC in 4 L of dialysis buffer [10-mM 3-(N-morpholino) propanesulfonic acid (MOPS), adjusted to pH 7.0 with potassium hydroxide (KOH), 5-mM ethylenediaminetetraacetic acid (EDTA), 50-mM potassium chloride (KCl), 20% glycerin, 5-mM

β-mercaptoethanol (BME), 80 mg of benzamidine, 20 mg of pepstatin, 50 mg of trypsin inhibitor, and 2 mg of leupeptin]. Protein concentrations of the extracts were determined after dialysis by a Pierce Coomassie Plus Protein Assay (Thermo Fisher

Scientific), following the manufacturer’s protocol and using bovine serum albumin

(BSA) for the standard curve.

The EPSPS activity of the extract was measured using the EnzChek Phosphate

Assay Kit (Life Technologies) to spectrophotometrically monitor inorganic phosphate release (Webb, 1992) during the catalysis of shikimate-3-phosphate (S3P) and phosphoenolpyruvate (PEP) to 5-enolpyruvylshikimate-3-phosphate (Gaines et al.,

2009). A 2 reaction buffer was prepared, containing 100-mM MOPS-KOH pH 7.0,

1-mM magnesium chloride (MgCl2), 10% glycerin, 2-mM sodium molybdate

(Na2MoO4), and 200-mM sodium fluoride (NaF). HPLC-grade water was used throughout, to minimize phosphate contamination. From the EnzChek kit, stocks of

1-mM 2-amino-6-mercapto-7-methylpurine riboside (MESG) and 100-U mL-1 purine nucleoside phosphorylase (PNP) were prepared, along with 50-mM PEP and 10-mM

61

S3P. Final reactions contained 25 µL of EPSPS extract, 125 mL of assay buffer,

0.4-mM MESG, 2-U mL-1 PNP, 2.5-mM PEP, 1-mM S3P, and glyphosate (pH 7.0 with

KOH) concentrations ranging from 0.1 µM-to-200 mM with water to a final reaction volume of 250 µL in fused quartz (UV-grade) spectrophotometer cuvettes. Background absorbance and rates of phosphate release were checked adding only PEP or S3P as the last step, and this background rate was subtracted from all other measured reaction rates. S3P must bind EPSPS before PEP or glyphosate, so it was the last component added to all reactions measuring EPSPS activity. Absorbance at 360 nm was recorded continuously for at least two minutes, and linear regressions of absorbance on time were used to calculate slopes of relative reaction rates. Slopes were then regressed on glyphosate concentration using three-parameter log-logistic functions (lower limit zero).

Regression analyses were conducted using the drc package in “R” (R Core Team, 2014;

Ritz and Streibig, 2005), including estimation of glyphosate doses required to reduce

EPSPS activity by 50% (I50) given by Equation [3.1]:

d f (x) = [3.1] 1+ exp(b(log(x)- log(e))) where (x) is the concentration of glyphosate, (d) is the upper limit, (e) is the inflection point (equal to the I50), and (b) is the slope around (e). Euler’s constant is expressed as

(exp) here to differentiate from the parameter variable (e).

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3.1.7 EPSPS Enzyme Quantification

Young, fully expanded leaves of S1 and R1 common ragweed plants were sampled before flowering initiation for total soluble protein (TSP) extraction. These tissue samples were immediately frozen in liquid nitrogen and then ground to a fine powder by a mortar and pestle chilled with dry ice. A lysis buffer containing 4% sodium dodecyl sulfate (SDS), 20% glycerol, and 125-mM Tris-HCl buffer pH 6.8

[tris(hydroxymethyl)aminomethane adjusted to pH 6.8 with concentrated hydrochloric acid], was added in a ratio of 2 µL to 1 mg of frozen tissue, and samples were boiled for five minutes. The mixture was centrifuged to separate undissolved solids and allow removal of the TSP extract. Protein concentrations of the TSP extracts were determined by diluting 10-fold for a Pierce Coomassie Plus Protein Assay (Thermo Fisher

Scientific), following the manufacturer’s protocol using bovine serum albumin (BSA) for the standard curve.

Using the calculated protein concentrations, the extracts were diluted to

1 µg µL−1 with water and 2 Laemmli Sample Buffer (Bio-Rad) with added

β-mercaptoethanol (BME), according to the instructions of the manufacturer. The diluted samples were heated to 95ºC for five minutes then loaded into a Criterion 10%

Tris-HCl gel (Bio-Rad), with a duplicate gel for staining. A Precision Plus Protein

WesternC Standard (Bio-Rad) was used in each gel, along with previously examined glyphosate-sensitive and glyphosate-resistant Palmer amaranth (Amaranthus palmeri) samples as controls. Electrophoresis in Tris/Glycine/SDS buffer (Bio-Rad) was run for approximately 45 minutes at 200 volts. The duplicate gel was stained and destained

63

according to the instructions of the manufacturer with GelCode Blue Stain Reagent

(Thermo Fisher Scientific), and photographed on a ChemiDoc MP Imaging System

(Bio-Rad).

To prepare the immunoblot, an Invitrolon polyvinylidene difluoride (PVDF) membrane (Invitrogen) was wet with 100% methanol and rinsed with water. The membrane and gel for transfer were equilibrated separately for 30 minutes in a transfer buffer of 25-mM Trizma Base (Sigma-Aldrich) pH 8.0 (HCl), 192-mM glycine, and

20% methanol. A Criterion blotter (Bio-Rad) was used to transfer the gel-separated protein to the PVDF membrane for 30 minutes at a constant 1000 mA. Following transfer, the membrane was rinsed briefly in TBS buffer (20-mM Trizma base adjusted to pH 8.0 with HCl, and 150-mM NaCl), and then incubated overnight at room temperature in TBS buffer with the addition of 5% blotting grade blocker (nonfat dry milk, Bio-Rad) on a shaker. The membrane was rinsed twice for 15 seconds in TBSt

(TBS buffer with the addition of 0.05% Tween-20 nonionic surfactant), then incubated with anti-EPSPS primary antibodies (Monsanto) at 1:2000 dilution in TBSt with gentle shaking for one hour. Again, the membrane was rinsed twice for 15 seconds in TBSt, followed by two 10-minute rinses. After rinsing, the membrane was incubated for

45 minutes with horseradish peroxidase (HRP)-labeled goat anti-rabbit secondary antibodies (Invitrogen) diluted at a 1:20,000 ratio in TBSt. The membrane was again rinsed as described previously following primary antibody incubation. Following the instructions of the Pierce ECL Western Blotting Substrate kit, the reagents were mixed at a 1:1 ratio, and pipetted onto the surface of the membrane, incubated one minute for

6 4

activation of the chemiluminescence, and drained onto a paper towel. A ChemiDoc MP

Imaging System was used for fluorescence detection and imaging of the immunoblot.

Band intensity was quantified with Image Studio Lite 4.0.21 software (LI-COR, Inc.).

3.1.8 EPSPS relative genomic copy number determination

Real-time quantitative PCR (qPCR) was used to measure the relative genomic copy number of the EPSPS gene per somatic cell (2n) genome. Fructan 1-exohydrolase

IIa (FEH) was used as a low copy number reference gene, which would not be expected to undergo strong selective pressure for gene duplication (Maroufi et al., 2010).

Extensive testing was conducted to optimize qPCR primers and reaction efficiency using visualization of gels following standard PCR and electrophoresis, as above, and qPCR annealing temperature gradients. Primer pairs EPGq4 and FEH5 were chosen for the genomic copy number determination study (Table 3.1). Efficiency curves for each primer pair with each of four common ragweed plants sampled previously for EPSPS enzyme quantification were conducted using a dilution series of gDNA, with

2 replicates. Quantitative PCR reactions contained 12.5 µL of iQ SYBR Green

SuperMix (Bio-Rad), 0.5 µL of each primer (5 µM in water), and 5 µL of genomic

DNA template (4, 0.4, 0.04, or 0.004 µM in water) plus 6.5 µL of additional PCR-pure water. Reactions and analysis were carried out in Multiplate 96-well PCR plates covered with Microseal ‘B’ adhesive films (Bio-Rad), with an iQ5 Real-Time PCR

Detection System (Bio-Rad). The iQ SYBR Green SuperMix contains fluorescein as a passive reference dye, and this was used to collect dynamic well factors. Amplification reactions began with an initial 3-minute template denaturation and enzyme activation

65

step at 95ºC, followed by 45 cycles of 95ºC denaturation for 30 seconds, 58ºC annealing for 45 seconds, and 72ºC extension for 30 seconds. Fluorescence was measured at the end of the extension phase of each cycle. After the 45 cycles, a melt curve was generated by increasing the temperature in 0.5ºC-increments of 30 seconds each, from

51ºC to 95ºC, and continuously measuring fluorescence. Relative EPSPS gene copy

−��CT number (NEPSPS) was calculated using a deconstruction of the 2 method (Livak and

Schmittgen, 2001). Regression analysis of threshold cycle (CT) versus log10( dilution ) was conducted in R (R Core Team, 2014), including calculation of slope (b) and standard error of the slope (SEb) for each plant and primer pair. PCR efficiency (E) was calculated with Equation [3.2]:

-1 E = 10 b [3.2]

Standard error of the efficiency (SEE) was calculated using Equation [3.3]:

E[ln(10)SE ] SE = b [3.3] E b2

The qPCR was repeated as previously described, except only using the 4-µM dilution of

DNA. The mean CT of six replicates was used for calculation of initial template concentration (X0) of EPSPS and FEH in each plant using Equation [3.4]:

-CT X0 = E [3.4]

66

The ratio of the initial concentration of EPSPS (X0,EPSPS) to the initial concentration of

FEH (X0,FEH) was used to calculate relative copy number of EPSPS (NEPSPS) for each plant, which were all standardized relative to the control plant (S1).

Standard error of the mean CT values (SECT) was calculated using Equation [3.5]:

s SE = CT [3.5] CT n

where (�) is standard deviation of the CT values and (n) is the number of replicates.

Standard error of the initial template concentrations (SEX0) was calculated using

Equation [3.6]:

-CT 2 2 SEX = X0 SEE + ln(E)SEC [3.6] 0 E T

Standard error of the relative genomic copy number of EPSPS (��) was calculated using Equation [3.7]:

2 2 SE SE SE = N X0,EPSPS + X0,FEH [3.7] NEPSPS EPSPS X0,EPSPS X0,FEH

Calculated relative EPSPS copy numbers were compared using pairwise t tests (α =

0.05) in “R” (R Core Team, 2014).

67

3.2 Results and Discussion

3.2.1 EPSPS gene sequencing

For primer pair “EPSPS-H”, only the sequencing reactions utilizing the Primer

EPSPS-F7 produced usable data in the region of interest from genomic DNA, because intron regions were not conserved. Dissimilar lengths of introns led to varied lengths of

PCR products, occasionally allowing separation of alleles within a single plant sample into different gel bands during electrophoresis. Through a combination of variable- sized PCR-products and heterozygosity within sequencing results of single bands, at least four EPSPS alleles were determined to be present in some plants from both glyphosate-sensitive and glyphosate-resistant biotypes (results not shown).

Chromosome counts of meiotic cells in immature anthers have shown common ragweed to be diploid (2n) and the primitive chromosome number of the Ambrosieae is based on n = 18, but a common ancestor of this subtribe was likely a tetraploid (4n) of n = 9

(Payne et al., 1964). Therefore, it is logical that four or more copies of EPSPS exist within a typical common ragweed genome. All of the exon regions of these sequences were relatively similar, so these will collectively be referred to as the “Type-1” alleles.

Sequences of complementary DNA (cDNA) synthesized from mRNA contain no introns, so these data were generally usable with both forward and reverse primers.

In several cases, poor quality sequence data were obtained from reactions using one or both primers, likely due to polymorphisms within the PCR primer-binding region.

Unexpectedly, these EPSPS sequences appear to result from different loci than those that were previously examined, as evidenced by conserved amino acid substitutions and

68

silent polymorphisms across individuals and biotypes, relative to the first data, but are clearly representative of common ragweed EPSPS amino acid sequence (“Type-2”

EPSPS alleles). In addition, several plants from the Clinton County, Ohio glyphosate- resistant biotype contained nucleotide differences translating to substitutions of serine or threonine from proline in amino acid position-106 within these data. The proline-to- serine (P106S) and proline-to-threonine (P106T) substitutions have both been well documented as conferring glyphosate-resistance in biotypes of several species

(Sammons and Gaines, 2014). In addition, several of these glyphosate-resistant common ragweed plants contained a nucleotide difference that would translate as an alanine-to-leucine substitution at position-89 (A89L), which has not been reported previously in glyphosate-resistant biotypes of other species. Although position-89 of

EPSPS is less conserved among plant species (Figure 3.1), it is in a region close to the active site for glyphosate and PEP. It is possible for the greater size of the isopropyl (–

CH2–( CH3 )2 ) functional group of valine, relative to the simple methyl (–CH3 ) functional group of alanine, to subtly alter the binding of glyphosate or PEP to EPSPS, or work synergistically with other mutations to further decrease inhibition by glyphosate and/or lessen a likely decrease in the affinity for PEP (Sammons et al.,

2007a). Contrarily, this mutation may have no effect, and it is not known if all of the analyzed common ragweed EPSPS sequences form full-length, functional enzymes.

Aligned gDNA and cDNA sequences of the Type-1 alleles of a total of 14 plants from four Ohio biotypes of common ragweed contained 27 polymorphic nucleotides out of a 783-bp region of EPSPS (3.5% polymorphism rate), including 10 polymorphisms

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within ± 100 bp of Pro106 (5%), where most of the sequences were aligned. None of the latter 10 polymorphisms coded for amino acid substitutions within that group of sequences. In aligned cDNA sequences of Type-2 alleles, 44 polymorphisms in 783 bp

(5.6%) were detected among just six plants from two biotypes (three S1 and three R1), including 16 polymorphisms (8.0%) within ± 100 bp of amino acid position-106. When combined, 33 polymorphic nucleotide positions were found among all of the Type-1 and Type-2 alleles in the region ± 100 bp of position-106, or a 16.5% polymorphism rate. The amino acid sequence near Pro106 is generally considered a conserved region, and indeed, from positions-90 through -160 in these data, only the amino acid-106 was identified as polymorphic. And yet, extending the range from amino acids-64 through -

207, eleven additional amino acid positions were polymorphic, but with the only A89V substitution also exclusive to glyphosate-resistant plants (Figure 3.1). Similar results were reported in previous study of the ALS-gene sequence in common ragweed, where an unusually high polymorphism rate was detected (Tranel et al., 2004). In that study, eight common ragweed plants from two fields in each of three states (24 total plants from Ohio, Illinois, and Minnesota) were found to contain 48 polymorphic nucleotides in a 385-bp region of the ALS gene, but only two that translate to amino acid substitutions. In contrast, 24 common cocklebur (Xanthium strumarium) plants from the same fields, plus an additional common cocklebur plant from each of North

Carolina, New Mexico, Mississippi, and Washington, contained zero nucleotide polymorphisms in the same 385-bp region of ALS.

70

3.2.2 EPSPS Enzyme Activity

The glyphosate concentrations required to inhibit 50% of the activity of EPSPS

(I50) from all sampled plants were within the range of I50 concentrations observed for glyphosate-sensitive EPSPS in several other plant species (Sammons et al., 2007b). At

18-µM glyphosate, the I50 of the R1 biotype sample was approximately 2-fold higher than the I50 values of the S1 and S2 biotypes, which were 8 µM and 10 µM, respectively

(Table 3.2, Figure 3.2). These plants were not selected for testing with regard to EPSPS gene sequence, because the glyphosate-resistant biotype was not believed to contain individuals with Pro106 mutations until subsequent sequence analyses were conducted.

Sequences of this R1 sample analyzed after the enzyme activity assay still did not show any Pro106 mutations, but were heterozygous for the poorly understood Alanine-89-to-

Valine (A89V) substitution described previously. The true consequences of this mutation can only be speculated without comparisons of enzyme kinetics using cloned common ragweed EPSPS with and without this substitution. Subtle differences in the structures of EPSPS isoforms from different plant species lead to substantially different enzymatic efficiencies, in the presence and absence of glyphosate (Sammons and

Gaines, 2014; Sammons et al., 2007b). The potential effects of the A89V substitution may have avoided detection because it provided minimal benefit in EPSPS enzymes of plant species and microorganisms used in those studies. Additionally, the high diversity noted in common ragweed gene sequences makes it plausible that the differences observed among the common ragweed samples presented here may have been the result

71

of other polymorphisms outside of the sequenced region, and not related to the A89V substitution.

3.2.3 EPSPS Enzyme Quantification

As expected, the signal of the EPSPS band in the glyphosate-resistant Palmer amaranth control plant became saturated, measuring greater than 30-fold higher intensity than the band for glyphosate-sensitive Palmer amaranth biotype. EPSPS signal intensity for three out of four glyphosate-resistant common ragweed plants (R1-1,

R1-2, and R1-4) was similar to the glyphosate-sensitive control (S1). However, the fluorescent signal for plant “R1-3” was approximately 11-fold stronger than “S1”, indicating considerably higher expression of EPSPS in this plant (Figure 3.3).

3.2.4 Real-Time Quantitative PCR

PCR efficiency of the EPSPS2 primers and the FEH5 primers were 83% to 98% percent and 85% to 99% percent, respectively. This was deemed acceptable as these were calculated for each experimental plant individually, and analysis of melt curves produced minimal evidence of non-specific binding.

Supporting the EPSPS immunoblot evidence, the third glyphosate-resistant common ragweed plant (R1-3) contained approximately 3-fold more genomic copies of the EPSPS gene than the glyphosate-sensitive control plant (S1) and the other glyphosate-resistant plants (R1-1 and R1-2) (Figure 3.4). If normal somatic (2n) common ragweed cells contain four-to-six copies of the EPSPS gene, this translates to twelve-to-eighteen copies of EPSPS in plant R1-3. Together, this provides evidence

72

that EPSPS gene duplication resulting in overexpression of the EPSPS enzyme contributes to glyphosate resistance in some, but not all, common ragweed biotypes in the population from Clinton County, Ohio. These observations support the hypothesis that multiple mechanisms of glyphosate resistance, including gene mutations and duplications, have evolved in common ragweed.

73

Chapter 3 References

Gaines TA, Zhang W, Wang D, Bukun B, Chisholm ST, Shaner DL, Nissen SJ, Patzoldt WL, Tranel PJ, Culpepper AS, Grey TL, Webster TM, Vencill WK, Sammons RD, Jiang J, Preston C, Leach JE, Westra P (2009) Gene amplification confers glyphosate resistance in Amaranthus palmeri. Proceedings of the National Academy of Sciences 107:1029-1034

Livak KJ, Schmittgen TD (2001) Analysis of relative gene expression data using real- time quantitative PCR and the 2−ΔΔCT method. Methods 25:402-408

Maroufi A, Van Bockstaele E, De Loose M (2010) Validation of reference genes for gene expression analysis in chicory (Cichorium intybus) using quantitative real- time PCR. BMC molecular biology 11:15

Patzoldt WL, Tranel PJ, Alexander AL, Schmitzer PR (2001) A common ragweed population resistant to cloransulam-methyl. Weed Science 49:485-490

Payne WW, Raven PH, Kyhos DW (1964) Chromosome numbers in Compositae. IV. Ambrosieae. American Journal of Botany:419-424

Preston C, Wakelin AM, Dolman FC, Bostamam Y, Boutsalis P (2009) A decade of glyphosate-resistant Lolium around the world: Mechanisms, genes, fitness, and agronomic management. Weed Science 57:435-441

R Core Team (2014) R: A Language and Environment for Statistical Computing. R Foundation for Statistical Computing. Vienna, Austria. http://www.R- project.org/.

Ritz C, Streibig JC (2005) Bioassay analysis using R. Journal of Statistical Software 12:1-22

Sammons RD, Gaines TA (2014) Glyphosate resistance: state of knowledge. Pest Management Science

Sammons RD, Heering DC, Dinicola N, Glick H, Elmore GA (2007a) Sustainability and stewardship of glyphosate and glyphosate-resistant crops. Weed Technology 21:347-354

Sammons RD, Meyer J, Hall E, Ostrander E, Schrader S A simple continuous assay for EPSP synthase in plant tissue. Champaign, IL: North Central Weed Science Society

Tranel PJ, Jiang W, Patzoldt WL, Wright TR (2004) Intraspecific variability of the acetolactate synthase gene. Weed Science 52:236-241

74

Webb MR (1992) A continuous spectrophotometric assay for inorganic phosphate and for measuring phosphate release kinetics in biological systems. Proceedings of the National Academy of Sciences of the United States of America 89:4884- 4887

Ye J, Coulouris G, Zaretskaya I, Cutcutache I, Rozen S, Madden TL (2012) Primer- BLAST: a tool to design target-specific primers for polymerase chain reaction. BMC bioinformatics 13:134

75

Table 3.1. PCR primer pairs used for sequencing and real-time quantitative PCR of common ragweed.

Pair Primer Primer Sequence (5'–3') EPSPS-F7 AACTCTGGGCTTACGTGTTGAGGA EPSPS-H EPSPS-R3 AGGAAACAGTCGACATCAGCACCA EPSPS-F10 AAAGAGCAACTGTGGAAGGGTGGT EPSPS-BBB EPSPS-R15 TGTCCAGGTTACTTCTGCACCCAT EPSPS-F1 TTTCCGGTACTGTTAATTTGCCTG EPSPS-CCC EPSPS-R17 CAGCTACACGAACAGGTGGACAAT EPSPS-F3 AGGATCCTTCTTTTAGCTGCTCTTG EPSPS-DDD EPSPS-R15 TGTCCAGGTTACTTCTGCACCCAT ALS-A F AGCTCTGGAACGTGAAGGC ALS-A ALS-A R CGTGTTACCTCAACAATAGG ALS-B F ATGAACGTTCAAGAGTTAGC ALS-B ALS-B R CCTTCGGTGATCACATCCTTGAA

76 FEH-R4 ATACCCGCTATGCTATACACCG

FEH54 FEH-F5 TCACATGCATAACACTCCCACT FEH-R4 ATACCCGCTATGCTATACACCG FEH34 FEH-F3 CGTCCCGATCGTATACCAATCA EPGseq-F CTTGAGTTTCCACCAGCAGC EPGq4 EPCq4-R GTGTGGTGGTGCGTTTCC FEH-R5 CGAATTGTTGTAGGCGGTGAC FEH5 FEH-F5 TCACATGCATAACACTCCCACT

Table 3.2. EPSPS glyphosate-dose–response enzyme activity I50 values and R:S ratios.

Biotype I50 (µM) R1:S1 p R1:S2 p R1 17.8 ± 1.86 2.3 0.0023 1.7 0.0042 S1 7.8 ± 1.13 - - - - S2 10.2 ± 1.02 - - - -

Glyphosate concentrations causing 50% inhibition (I50) and R:S ratios obtained from curves shown in Figure 3.2 on partially purified 5-enolpyruvylshikimate-3-phosphate synthase (EPSPS) from three biotypes of common ragweed. R:S ratios are significant if p ≤ 0.05.

77

Figure 3.1. Alignment of predicted 5-enolpyruvylshikimate-3-phosphate synthase (EPSPS) amino acid sequences.

! ! ! 6!! 6!! ! ! 7!! 0!! ! ! 7!! 4!! ! ! 7!! 8!! ! ! 8!! 2!! ! ! 8!! 6!! ! ! ! ! |! ! ! |! ! ! |! ! ! |! ! ! |! ! ! |! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! R" V!! E! E! D" G! A" I! K! R! A! V" V! E! G! C! G! G! V! F! P! V! G! R" –!! Ragweed,!Type=1!alleles!a! N" =! =! =! N" =! E/A" =! =! =! =! T" =! =! =! =! =! =! =! =! =! =! =! R/K" –! Ragweed,!Type=2!alleles! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! 9!! 0!! ! ! 9!! 4!! ! ! 9!! 8!! ! ! 1!! 0!! 2!! ! 1!! 0!! 6!! ! 1!! 1!! 0!! ! ! ! |! ! ! |! ! ! |! ! ! |! ! |! ! |! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! E!! A!" K! D! E! I! Q! L! F! L! G! N! A! G! T! A! M! R! P" L! T! A! A! V!! –!! Ragweed,!Type=1!alleles! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! –! Ragweed,!Type=2!alleles!(S)! =! A/V" =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! P/T" =! =! =! =! =! –! Ragweed,!Type=2!alleles!(R1=A)! =! A/V" =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! –! Ragweed,!Type=2!alleles!(R1=B)! =! A/V" =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! P/S" =! =! =! =! =! –! Ragweed,!Type=2!alleles!(R1=C)! 78 =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! –! Willowleaf!sunflower!b!

=! S! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! –! Soybean!c! =! S! =! E! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! –! Petunia!d! D! G! =! E! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! –! Palmer!amaranth!e! D! S! =! S! D! =! E! =! Y! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! –! Arabidopsis!f! D! =! =! E! =! V! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! –! Maize!(B73)!g! D! =! =! E! =! V! =! =! =! =! =! =! =! =! I! =! =! =! S! =! =! =! =! =! –! 1st!generation!RR=Maize!h! D! =! =! E! =! V! K! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! –! Rigid!ryegrass!(S)!i! D! =! =! E! =! V! K! =! =! =! =! =! =! =! =! =! =! =! S! =! =! =! =! =! –! Rigid!ryegrass!(R)!j! D! =! =! E! =! V! K! =! =! =! =! =! =! =! =! =! =! =! L! =! =! =! =! =! –! Rigid!ryegrass!(R)!j! D! =! =! E! =! V! K! =! =! =! =! =! =! =! =! =! =! =! A! =! =! =! =! =! –! Rigid!ryegrass!(R)!k! D! =! =! E! =! V! K! =! =! =! =! =! =! =! =! =! =! =! T! =! =! =! =! =! –! Rigid!ryegrass!(R)!l!

Continued

1! 1! 4! 1! 1! 8! 1! 2! 2! 1! 2! 6! 1! 3! 0! 1! 3! 4! ! ! ! |! ! |! ! |! ! |! ! |! ! |! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! T!! A!! A! G! G! N! S! S! Y! I! L! D! G! V! P! R! M! R! E! R! P! I! G! D!! –!! Ragweed,!Type=1!alleles! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! –! Ragweed,!Type=2!alleles! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! 1! 3! 8! 1! 4! 2! 1! 4! 6! 1! 5! 0! 1! 5! 4! 1! 5!! 8!! ! ! ! |! ! |! ! |! ! |! ! |! ! |! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! L! V! T! G! L! K! Q! L! G! A! D! V! D! C! F! L! G! T! N! C! P! P!! V! R!! –!! Ragweed,!Type=1!alleles! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! –! Ragweed,!Type=2!alleles! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! 1! 6! 2! 1! 6! 6! 1! 7! 0! 1! 7! 4! 1! 7! 8! 1! 8!! 2!! ! ! ! |! ! |! ! |! ! |! ! |! ! |! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! V! V" G" G" G! G! L! P! G! G! K! V! K! L! S! G! S! I! S! S! Q! Y!! L! T!! –!! Ragweed,!Type=1!alleles! =! A" A" N" =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! –! Ragweed,!Type=2!alleles! 79

! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! 1! 8! 6! 1! 9! 0! 1! 9! 4! 1! 9! 8! 2! 0! 2! 2! 0!! 6!! ! ! ! |! ! |! ! |! ! |! ! |! ! |! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! A! L! L! M! A! S" P! L! A! L! G! D! V! E! I! E! I! I! D! K! L! I!! S! I!" –!! Ragweed,!Type=1!alleles! =! =! =! =! =! A" =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! =! V" –! Ragweed,!Type=2!alleles! Amino acid numbering is based upon the maize EPSPS numbering system. Positions 88–111 are aligned with other species for comparison. Dashes (-) represent amino acids that did not differ from “Ragweed, Type-1 alleles”. acommon ragweed (Ambrosia artemisiifolia), Type-1 and Type-2 alleles are assigned according to conserved differences (observed across S and R biotypes, unless noted); bHelianthus salicifolius, GenBank accession # AAT45239; cGlycine max, GenBank accession # XP_003521857; dPetunia xhybrida, GenBank accession # AAA33699; eAmaranthus palmeri, GenBank accession # AGE94064; fArabidopis thaliana (Columbia), GenBank accession # AEE32359; gZea mays (B73), GenBank accession # CAA44974; hGA21 Roundup-Ready glyphosate tolerance event (directed-mutagenesis “TIPS”, Monsanto), US Patent # 6,040,497; iLolium rigidum (glyphosate-sensitive allele), GenBank accession # ABC00782; Glyphosate-resistant L. rigidum alleles: j(Collavo and Sattin, 2012), kGenBank accession # ACB05442; lGenBank accession # ABC00790.

Figure 3.2. EPSPS glyphosate-dose–response enzyme activity assay.

100 S2 S1 R1 ) l

o 80 r t n o c

f o 60 % (

y t i v i t c 80 A

40

S P S P E 20

0

0 0.1 1 10 100 1000 Glyphosate Concentration (µM)

Three-parameter log-logistic dose-response curves (Equation [3.1]) for glyphosate on partially purified 5-enolpyruvylshikimate-3-phosphate synthase (EPSPS) extracted from three biotypes of common ragweed.

Figure 3.3. Relative EPSPS protein abundance determined by a western immunoblot.

A EPSPS Immunoblot GelCode Blue Stain

B 30

25

20

15

10 (relative to A.a.(S1) to (relative

5

EPSPS enzyme immunoblot fluorescence enzyme fluorescence immunoblot EPSPS 0

Specimen

A Western immunoblot was performed to visualize relative abundance of 5-enolpyruvylshikimate-3-phosphate synthase (EPSPS) protein extracted from young leaf tissue of glyphosate-sensitive (S) and glyphosate-resistant (R) biotypes of Palmer amaranth (A.p.) and common ragweed (A.a.). A) Precision Plus WesternC (Bio-Rad) was used as a size reference, run alongside the immunoblot and a duplicate gel stained with GelCode Blue Stain Reagent (Thermo Fisher Scientific) for visualization of total protein loading. Bands of EPSPS are visible near 50 kD. B) Fluorescence intensity of bands in the immunoblot, normalized to glyphosate- sensitive common ragweed [A.a.(S1)].

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Figure 3.4. EPSPS:FEH relative genomic copy number.

5

*

4

3

2

1 Scaled EPSPS:FEH relative genomic copy number copy genomic relative Scaled EPSPS:FEH 0 S1 R1-1 R1-2 R1-3 Common ragweed specimen

Copy number ± standard error of the 5-enolpyruvylshikimate-3-phosphate synthase (EPSPS) gene relative to fructan 1-exohydrolase IIa (FEH) of four common ragweed plants measured by real- time quantitative PCR of genomic DNA using primer pairs EPGq4 and FEH5 shown in Table 3.1, calculated from Equations [3.2] through [3.7], and scaled to S1 = 1.00. *Significantly different according to pairwise t-tests with a comparisonwise error rate of α = 0.05.

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Chapter 4: Common Ragweed Non–Target-Site Glyphosate-Resistance Mechanisms

4.1 Materials and Methods

4.1.1 Development of a sample population

In Autumn 2009, seed was collected from common ragweed plants that survived multiple glyphosate applications in a soybean field in Clinton County, Ohio, where a large population of glyphosate-resistant plants had developed over time. Seed from a presumably glyphosate-sensitive common ragweed biotype was purchased from Azlin

Seeds in Leland, Mississippi in 2010.

Prior to stratification, a sample of seeds from each common ragweed biotype were soaked for 16 hours in tap water with 0.1% v/v non-ionic surfactant in mesh packets submerged in a 1000-mL beaker, to improve penetration of water through the seed hulls.

The packets were removed from the water/surfactant solution and rinsed twice, and then covered in damp sand in a chest cooler (for moisture retention) at a temperature of 4˚C for 5 weeks. The stratified seeds were planted directly into 1.9-L greenhouse containers containing Fafard 3B potting media (Canadian sphagnum peat moss, perlite, dolomitic limestone, composted bark, vermiculite, wetting agent, starter nutrient charge with gypsum), and supplemented with 700 g of Osmocote Plus Lo-Start slow-release fertilizer blended into each 79.3-L bag of media. The containers were placed into a research greenhouse under 16-hour photoperiod controlled by 1000-watt metal halide lamps that 83

provided approximately 200 µmol m-2 s-1 of supplemental lighting. Air temperatures in the greenhouse were maintained at approximately 27˚C daytime and 20˚C nighttime.

Emerged plants were thinned to one seedling per container. Seedlings were approximately 16 cm tall with 3–5 pairs of true leaves four weeks after planting. At this time, 30 plants from the glyphosate-resistant Clinton County biotype and 10 plants from the presumed glyphosate-sensitive Leyland, Mississippi biotype were treated with glyphosate. The glyphosate (Roundup WeatherMAX®, Monsanto) was applied at a rate of 8.4 kg ae ha−1 (10 field use rate) with N-Pak ammonium sulfate solution at 5% v/v using a pneumatic track sprayer equipped with an even flat-nozzle (Teejet 8001EVS) calibrated to apply 140 L ha−1 of spray solution at a speed of 3.5 km hr−1. An additional

10 plants from the glyphosate-sensitive biotype were left untreated. Plant survival was assessed four weeks after treatment. Surviving plants from the glyphosate-resistant biotype were divided into two groups of 10 and 12 plants (R1 parents and R2 parents) based upon degree of stem dieback, and placed into two separate greenhouse compartments for pollination within groups. The ten untreated plants from the glyphosate-sensitive biotype were placed into a third greenhouse chamber for pollination

(S1 parents). Greenhouse chambers were maintained under conditions similar to those mentioned previously. At senescence, entire plants with seeds were harvested, placed into paper bags, and allowed to dry for one month in the greenhouse. Seeds were subsequently removed from the plants by hand, and cleaned with sieves and an air- powered seed separator. Dry seeds were stored in plastic bottles at 4˚C until needed.

Following a phenotypic observation experiment, some plants from the R1 and R2 progeny populations surviving 0.21, 0.84, and 3.36 kg ae ha−1 glyphosate applications 84

(Roundup WeatherMAX®, Monsanto) were saved for various self-pollination and cross- pollination attempts with each other and with untreated S plants. One individual from the

R2 group, surviving a glyphosate rate of 3.36 kg ae ha−1, and allowed to self-pollinate inside of a tent made from bamboo stakes and a clear plastic trash bag, produced enough viable seeds (labeled R 4 biotype) to be included in the 31phosphate nuclear-magnetic resonance (31P-NMR) studies described later.

4.1.2 14C-glyphosate uptake and translocation:

Common ragweed seeds from the R1 and S1 progeny populations were placed in mesh bags and stratified in damp sand for eight weeks, after which they were rinsed with water and a 500-ppm solution of X3 disinfectant, and sown in germination towels at

25˚C. Seeds were checked daily, and once radicals emerged, were placed into

5-cm 5-cm 5-cm cells of 36-cell propagation flats containing a 1:1:2 blend of all- purpose sand (Quikrete, The Home Depot), course calcined clay particles (Turface MVP,

Hummert International), and fine calcined clay particles (Turface Field & Fairway).

When the plants had two to three pairs of true leaves, the growth media was washed from the plant roots, and the plants were placed in a ⅛ solution of Murashige and Skoog

(MS) basal salt mixture (M524, Phyto Technology) to be maintained hydroponically.

Additionally, since the glyphosate treatment would be applied to the lowest pair of leaves, any damaged leaves were removed so that the lowest pair was healthy and undamaged. The plants were allowed to acclimate in the growth chamber for 48 hours before treatment. Growth chamber conditions were 14/10-hour day/night length, 28/26˚C day/night temperature cycle, with 50% relative humidity. A low dose of 14C-labeled

85

glyphosate was formulated using Code 11355 glyphosate (50 mCi mmol−1;

220,000 dpm µL−1) at a rate of 1.5 µg (984624 dpm) per 8 µL of solution, with glycerin and MON0818 (Monsanto Company) as surfactants at 10% and 0.1% v/v respectively. A high dose was formulated as above, with the addition of non-radiolabeled glyphosate

(MON76523, Monsanto Company) at 13.5 µg ae per 8-µL for a total of 15 µg ae plant−1.

Doses were applied as 2-µL droplet pairs on each side of the midribs of the lowest pair of leaves on each plant. Ten replicates of the R1 biotype and five replicates of the S1 biotype were harvested for each dose and harvest time. Plants were harvested at 6, 24, and 48 hours after treatment (HAT). The treated leaves were cut off, the surface of the leaves rinsed thoroughly with water followed by a quick methanol rinse, and the total volume of rinsate was adjusted to 30 or 40 mL (“wash”) and volume recorded. Fresh weight of the harvested leaves was measured. Plants were further dissected into the roots, the unexpanded leaves and apical meristem (“meristem”), the remaining tissue above the treated leaves (“above”), and the remaining tissue below the treated leaves

(“below”). Roots were rinsed into the hydroponic solution with methanol and blotted dry with paper towels, and fresh weights were measured for all tissues. The hydroponic solutions were weighed to determine their volume, and saved for scintillation counting.

To extract glyphosate from the tissues, 0.2-N hydrochloric acid (HCl) was added at 1 mL to treated leaves, 2 mL to meristems, 5 mL to “above”, 4 mL to “below”, and 13.5 mL to roots. Tissue with HCl solution was frozen and thawed three times, followed by vortexing after each thaw. Radioactivity in the samples was quantified with a Perkin

Elmer Tri-Carb 2910 TR scintillation counter using 15 mL of Ecolite (+) as a scintillation

86

cocktail with 200 µL of each treated leaf and meristem extract, 1 mL of the above, below, and root extracts, or 2 mL of the wash and hydroponic solutions.

4.1.3 Statistical analyses

Results were analyzed using the GLM procedure in SAS 9.3. Least squares means are reported for 14C-labeled glyphosate recovery, absorption, and total translocation. Raw data for translocation to individual plant sections failed Shapiro-Wilk normality tests, and were analyzed using a square-root transformation. Plant section means presented here are back-transformed from square root least squares means. The least squares means were separated according to Tukey’s HSD with p = 0.05.

4.1.4 In vivo 31P-NMR investigation

Plant tissue was treated with glyphosate by both foliar spray treatment and delivery by infusion for 31P-NMR studies. For analysis of common ragweed with the foliar spray treatment, plants from the R 4 (R4) and S1 biotypes were treated with glyphosate (Roundup WeatherMAX®, Monsanto) at 3.36 kg ae ha−1 (4 field use rate).

Immature leaves from the apical meristem (sink tissue) were protected from direct exposure to the herbicide spray with aluminum foil. Following treatment, plants were maintained in greenhouse conditions (14/10 hour day/night length, 30/20°C day/night temperature, 700–900 µE m−2 s−1 photosynthetically-active radiation). Source

(mature/treated) and sink (immature/covered) leaf tissues were harvested 24 HAT, washed repeatedly with deionized water, and vacuum infiltrated with the perfusion buffer

(50-mM sucrose, 12-mM MES adjusted to pH 5.0 with 2-M KOH). The tissue was then placed in a 10-mm NMR tube and fitted with a perfusion system for NMR studies at 87

11.74 T (Varian Inova-500) (Ge et al., 2012a; Ge et al., 2011; Ge et al., 2010). Under oxygenated perfusion conditions (flow rate 4 mL min−1), 31P-NMR spectra of leaf tissue were recorded in two-hour collection time blocks, up to 18 hours (42 HAT). Sink and source tissue for the same genotypes were harvested from different plants treated with foliar spray at 24-hour intervals. The relative glyphosate entry (as measured by comparing signal intensity to an external capillary reference of 20-mM methylene diphosphonate (MDP)) was established from signal magnitudes using Bayesian- probability-theory-based signal analysis algorithms and associated software (Bretthorst,

1990).

For glyphosate pulse-chase studies, where herbicide delivery is via perfusion infusion, untreated mature source leaf tissue was harvested from plants and vacuum infiltrated with 10-mM glyphosate buffer at pH 5.0. The tissue was placed in a 10-mm

NMR tube and perfused with 10-mM glyphosate buffer for 8 hours (the pulse period), and then replaced with a glyphosate-free perfusion buffer to observe glyphosate entry and possible partitioning. Data were collected in a similar manner for the next 16 hours (Ge et al., 2012a). Analyses performed by collaborators are presented in Appendix A.

4.2 Results and Discussion

4.2.1 14C-glyphosate uptake and translocation:

Total recovery of 14C-labeled glyphosate was variable, but within a biotype did not differ by dose of glyphosate or harvest timing. Recovery of 14C for the S1 and R1 biotypes was 74.8% and 72.4% of the applied dose, respectively. Absorption of

14C-glyphosate was highly correlated with the fresh weight of the treated leaves, so 88

weight of treated leaf was used as a covariate. A small difference in absorption was observed, with 44.1% of the recovered 14C-glyphosate absorbed by R1 plants versus

40.8% by the S1 plants. The glyphosate dose also affected absorption, as 46.7% of the recovered glyphosate was absorbed at the high dose of 15 µg, versus 38.2% at the low dose of 1.5 µg. Total absorption of glyphosate in both biotypes reached 39.4% by

8 HAT, which was significantly less than the absorption of 44.2% and 43.7% at 24 and

48 HAT, respectively (Figure 4.1). This was similar to the results observed by Brewer and Oliver (2009), where approximately 31% absorption of the recovered dose was observed at 6 HAT and 36% at 24 HAT, reaching a maximum of 38% at 48 HAT.

Translocation of the absorbed dose was similar between the R1 and S1 biotypes, and there were no biotype-by-harvest–time or biotype-by-dose-by-harvest interactions.

When combined over biotypes, some sections had small differences in translocation between the low dose of 1.5 µg and high dose of 15 µg of glyphosate (Table 4.1). For example, 53.2% of the absorbed 14C-glyphosate was localized within the treated leaves at the low dose, while only 45.0% remained within the treated leaves at the high dose.

However, within the remainder of the shoot tissue (“above”, “below”, and “meristem” sections combined), 15.9% of the absorbed 14C-glyphosate was found from the low dose, which was lower than the 23.1% found in this tissue at the high dose. There were no differences between the doses in the root tissue (Table 4.1).

As might be expected, there were differences among harvest times for all plant sections, pooling biotypes and doses (Table 4.2). At 8 HAT, 76.3% of the absorbed 14C- glyphosate was recovered from the washed treated leaves. By 24 and 48 HAT, this had

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dropped to 41.8% and 33.7%, respectively. The percentage of the absorbed dose in the meristem, above, and root sections increased with time, but only in the roots was there an increase between 24 and 48 HAT. The strongest sinks for glyphosate translocation in vegetative plants are generally the most active meristems, found at the shoot apexes and root tips. As expected in the current study, the largest percentages of the absorbed 14C- glyphosate (outside of the treated leaves) in common ragweed were found in the root and meristem regions at all harvest times.

4.2.2 Discussion

The results of the 14C-labeled glyphosate uptake and translocation study, combined with the 31P-NMR study, indicate that neither vacuole sequestration of glyphosate, nor reduced foliar absorption or translocation of glyphosate, are likely mechanisms of glyphosate resistance in the studied Ohio biotype of common ragweed. In biotypes of glyphosate-resistant horseweed (Conyza canadensis), rigid ryegrass (Lolium rigidum), and Italian ryegrass (Lolium multiflorum) where vacuole sequestration has previously been demonstrated to confer resistance, a large fraction of the cellular glyphosate is quickly loaded into the vacuole, preventing this glyphosate from entering the plastids or being translocated (Ge et al., 2012a; Ge et al., 2011; Ge et al., 2010). Even glyphosate-resistant horseweed shows a small fraction of the glyphosate entering the vacuole, so it is not likely that levels of glyphosate sequestration below detectable limits would be an important factor in glyphosate resistance (Ge et al., 2010; Ge et al., 2012c).

In all three of these species, 14C-glyphosate translocation studies had previously shown greatly differing glyphosate translocation patterns between S-biotypes and some

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R-biotypes (Feng et al., 2004; Koger and Reddy, 2005; Michitte et al., 2007; Nandula et al., 2008; Perez-Jones et al., 2007; Wakelin et al., 2004).

In both the 14C-glyphosate and pulse-chase 31P-NMR studies presented here, the R biotype of common ragweed absorbs glyphosate slightly more efficiently than the S biotype. However, the 31P-NMR signal for glyphosate was surprisingly small following spray treatment, and it is unknown how much glyphosate entering the cells is delivered to the chloroplasts. Currently, there is no way to detect the subtle chemical shift between cytoplasmic glyphosate and plastid-localized glyphosate (Ge et al., 2013), so chloroplast- exclusion has not been ruled out. In addition, visual observation and 31P-NMR data showing tissue degradation of R-plants sprayed with high rates of glyphosate indicate that some version of the source tissue sacrifice mechanism seen in some biotypes of giant ragweed (Ambrosia trifida) (Van Horn and Westra, 2014) may be involved in providing additional protection from high doses of glyphosate in some plants. However, this was not observed at the rates used in the 14C-glyphosate absorption and translocation study, and occurs less predictably than in giant ragweed biotypes. Other conceivable resistance mechanisms include EPSPS mutation (resulting in reduced inhibition by glyphosate) or increased production of the EPSPS enzyme (resulting in dilution of glyphosate relative to the target site and sequestration of glyphosate in a transition-state complex with EPSPS and S3P) (Sammons and Gaines, 2014; Sammons, personal communication).

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Chapter 4 References

Bretthorst GL (1990) Bayesian analysis. III. Applications to NMR signal detection, model selection, and parameter estimation. Journal of Magnetic Resonance 88:571-595

Brewer CE, Oliver LR (2009) Confirmation and resistance mechanisms in glyphosate- resistant common ragweed (Ambrosia artemisiifolia) in Arkansas. Weed Science 57:567-573

Feng PCC, Tran M, Chiu T, Douglas Sammons R, Heck GR, CaJacob CA (2004) Investigations into glyphosate-resistant horseweed (Conyza canadensis): retention, uptake, translocation, and metabolism. Weed Science 52:498-505

Ge X, d'Avignon DA, Ackerman JJ, Collavo A, Sattin M, Ostrander EL, Hall EL, Sammons RD, Preston C (2012a) Vacuolar glyphosate-sequestration correlates with glyphosate resistance in ryegrass (Lolium spp.) from Australia, South America, and Europe: a 31P NMR investigation. Journal of Agricultural and Food Chemistry 60:1243-50

Ge X, d'Avignon DA, Ackerman JJH, Duncan B, Spaur MB, Sammons RD (2011) Glyphosate-resistant horseweed made sensitive to glyphosate: low-temperature suppression of glyphosate vacuolar sequestration revealed by 31P NMR. Pest Management Science 67:1215-1221

Ge X, d'Avignon DA, Ackerman JJH, Ostrander E, Sammons RD (2013) Application of 31P-NMR spectroscopy to glyphosate studies in plants: Insights into cellular uptake and vacuole sequestration correlated to herbicide resistance. Pages 55-83 Handbook on Herbicides : Biological Activity, Classification and Health and Environmental Implications. Hauppauge, NY, USA: Nova Science Publishers, Inc.

Ge X, d'Avignon DA, Ackerman JJH, Sammons RD (2010) Rapid vacuolar sequestration: the horseweed glyphosate resistance mechanism. Pest Management Science:n/a-n/a

Ge X, d’Avignon DA, Ackerman JJH, Sammons RD (2012c) Observation and identification of 2-C-methyl-d-erythritol-2,4-cyclopyrophosphate in horseweed and ryegrass treated with glyphosate. Pesticide Biochemistry and Physiology 104:187-191

Koger CH, Reddy KN (2005) Role of absorption and translocation in the mechanism of glyphosate resistance in horseweed (Conyza canadensis). Weed science 53:84-89

Michitte P, De Prado R, Espinoza N, Pedro Ruiz-Santaella J, Gauvrit C (2007) Mechanisms of resistance to glyphosate in a ryegrass (Lolium multiflorum) biotype from Chile. Weed Science 55:435-440 92

Nandula VK, Reddy KN, Poston DH, Rimando AM, Duke SO (2008) Glyphosate tolerance mechanism in Italian ryegrass (Lolium multiflorum) from Mississippi. Weed Science 56:344-349

Perez-Jones A, Park K-W, Polge N, Colquhoun J, Mallory-Smith CA (2007) Investigating the mechanisms of glyphosate resistance in Lolium multiflorum. Planta 226:395-404

Sammons RD, Gaines TA (2014) Glyphosate resistance: state of knowledge. Pest Management Science

Van Horn CR, Westra P (2014) Updates on molecular response of glyphosate resistant giant ragweed (Ambrosia trifida). in Proceedings of the Weed Science Society of America and Canadian Weed Science Society. Vancouver, BC: Weed Science Society of America

Wakelin AM, Lorraine-Colwill DF, Preston C (2004) Glyphosate resistance in four different populations of Lolium rigidum is associated with reduced translocation of glyphosate to meristematic zones. Weed Research 44:453-459

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Table 4.1. Translocation of 14C-glyphosate in common ragweed plants receiving low or high doses of glyphosate. % of absorbed 14C-glyphosate Dose Treated Leaf Meristem Above Below Root Above+Below+Meristem Low 53.2 9.6 3.4 2.5 23.9 15.9 High 45.0 13.3 5.4 3.3 24.5 23.1 p= 0.0033 0.0070 <0.0001 0.0064 NS <0.0001 14C-labeled glyphosate translocation, expressed as a percentage of the absorbed dose, in treated leaf, meristem, above, below, and root sections, in pooled common ragweed R1 and S1 biotypes receiving low (1.5 µg glyphosate acid per plant) or high (15 µg glyphosate acid per plant) doses of glyphosate. Presented as back-transformed least squares means with p values for differences between doses in each section below each column. 94

Table 4.2. Translocation of 14C-glyphosate in common ragweed plants harvested 8-, 24-, or 48-HAT.

% of absorbed 14C-glyphosate

HAT Treated Leaf Meristem Above Below Root Hydroponic Meristem+Root+Hydroponic 8 76.3 a 6.6 b 3.0 b 3.5 a 13.1 c 0.3 c 18.2 c 24 41.8 b 15.7 a 5.4 a 3.0 ab 26.3 b 1.7 b 46.9 b 48 33.7 c 13.0 a 4.8 a 2.2 b 36.1 a 4.0 a 58.5 a 14C-labeled glyphosate translocation, expressed as a percentage of the absorbed dose, in treated leaf, meristem, above, below, and root sections, and hydroponic solutions, 8-, 24-, or 48-hours after treatment (HAT), in pooled common ragweed R1 and S1 biotypes receiving low and high doses of glyphosate. Back-transformed least squares means within each column followed by the same letter are not significantly different according to Tukey's HSD test at α = 0.05.

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Figure 4.1. Absorption of 14C-glyphosate in common ragweed plants over 48-hours.

50

* 40

glyphosate 30 - C 14

20 96

% of recoverd recoverd % of 10

0 0 12 24 36 48 Hours after treatment

14C-labeled glyphosate absorption, expressed as a percentage of the recovered dose, in pooled common ragweed R1 and S1 biotypes, presented as least squares means. * = p < 0.05 according to Tukey's HSD test.

Chapter 5: Conclusions

Unlike species that are highly adapted to narrow ecological niches, common ragweed appears to evolve using a strategy of outcrossing and broad inter- and intrapopulation genetic diversification to continually adapt to new environments and management strategies. The diverse genetics and self-incompatibility of common ragweed creates technical challenges for studying it in the laboratory, but its rapid evolution in response to herbicide selection pressure provides a rare opportunity to learn about evolutionary processes in a short time period.

Our primary study population of common ragweed from Clinton County, Ohio

(R) was compared to herbicide-susceptible populations from Leland, Mississippi (S1) and

Columbus, Ohio (S2). Whole-plant bioassays of herbicide dose-response in the greenhouse confirmed resistance, to glyphosate (EPSPS-inhibitor), cloransulam-methyl

(ALS-inhibitor), and fomesafen (PPO-inhibitor) herbicides, in the R population. These three herbicides are normally effective for common ragweed control, and this is the first reported common ragweed biotype with multiple resistance to three site-of-action (SOA) groups. Resistance to ALS-inhibiting herbicides is commonly encountered in common ragweed and has been documented in 144 other weed species worldwide (Heap, 2014), aided by the fact that various single-nucleotide polymorphisms (SNPs) in the ALS gene provide near-immunity to ALS-inhibitors (Zheng et al., 2005). Interestingly, Rousonelos

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et al. (2012) reported that resistance to PPO-inhibitors in a common ragweed population is conferred by a SNP in the PPO gene, yet common ragweed is one of only 6 species with known PPO-resistant biotypes (Heap, 2014).

In the study of herbicide-resistance, glyphosate-resistance mechanisms have proven to be particularly interesting because more mechanisms have been reported for glyphosate resistance than for any other herbicide mode of action, but none of these confer high levels of resistance (Sammons and Gaines, 2014). Potential mechanisms resulting in changes to the target-site of glyphosate (EPSPS) were investigated here using molecular genetic and enzymology approaches. Studies of the EPSPS gene using PCR and sequencing revealed a high rate of nucleotide polymorphisms in all common ragweed populations sampled in the present study. A high polymorphism rate in common ragweed genes has also been reported by others studying ALS (Tranel et al., 2004), PPO

(Rousonelos et al., 2012), and pollen coat proteins (Léonard et al., 2010), so our finding of polymorphisms is not exclusive to common ragweed populations or EPSPS.

Polymorphisms can reduce the efficiency of PCRs if they occur in a PCR-primer binding- site, and we experienced difficulty working with DNA and mRNA transcripts, as did the aforementioned authors. However, this high level of genetic diversity could have roles in the adaptability of common ragweed and the rapid evolution of herbicide resistance.

Although no mutations coding for amino acid substitutions were detected initially,

PCRs conducted using a different pair of primers amplified a second type of EPSPS alleles. We detected alleles of this second type coding for substitutions at two amino acid positions (89 and 106), exclusively in plants from the R population of common ragweed.

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Previous studies by other researchers have reported the P106S and P106T amino acid substitutions to confer glyphosate resistance in numerous other species, but the A89V substitution we reported here has not been previously documented.

A crude EPSPS extract from a single R common ragweed plant surprisingly required 2-fold higher glyphosate concentrations to inhibit enzymatic activity than extracts from S1 and S2 plants. In a subsequent analysis of EPSPS gene sequences from the R plant sample, the A89V mutation was detected, but the P106 mutations were not observed. This supports the hypothesis that the A89V mutation provides a small benefit in glyphosate resistance, but other differences in the gene sequence outside of the amplified fragments or post-translational modifications to the enzyme could also contribute to the resistance. Future studies involving cloning, sequencing, and expressing the entirety of various isoforms of common ragweed EPSPS for enzyme kinetics experiments could be employed to confidently describe the effects of these mutations.

An EPSPS immunoblot assay of electrophoresis-separated total soluble protein from one S1 and four R common ragweed plants, as well as Palmer amaranth S and

EPSPS-overexpressing R controls, showed a single plant from the R common ragweed population with increased EPSPS expression. Quantitative real-time PCR (qPCR) of genomic DNA was used to compare EPSPS gene copy number relative to FEH of this S1 and first three R common ragweed plants. Results showed an increased relative EPSPS gene copy number in the same R plant. The quantitative PCR requires an assumption that all copies of each gene were amplified at a similar efficiency within each plant sample.

The DNA sequence chromatograms obtained from our previous experiments with these

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common ragweed populations were used to carefully select qPCR primers. Some alleles of either gene could have been poorly detected in the qPCR experiments because of the problems with primer binding described previously. Attempts to measure the relative abundance of EPSPS mRNA transcripts by qPCR of complementary DNA were abandoned because PCR problems were compounded by the challenges of working with

RNA. However, the agreement of the genomic DNA qPCR and EPSPS immunoblot data increases our confidence that increased gene copy number is a likely factor contributing to glyphosate resistance in some plants of the common ragweed R population.

Variably expressed necrosis of glyphosate-treated foliage, followed by regrowth from undamaged meristems, has been observed within the R population of common ragweed in whole-plant dose-response bioassays and other experiments, but not in glyphosate-sensitive populations. This necrosis appears similar, but not identical, to a more rapid and consistent response being studied by other labs in some biotypes of giant ragweed. Therefore, we hypothesized that this rapid necrosis was a non–target-site mechanism of glyphosate-resistance in common ragweed, and we examined this with

14C-labeled-glyphosate absorption and translocation studies and 31P-NMR spectroscopy.

In the 14C-glyphosate studies, the only detectable differences between R and S common ragweed was slightly lower recovery of applied 14C and slightly higher absorption of recovered 14C in the R biotype than the S biotype. We do not believe that these small differences represent a non–target-site resistance mechanism. However, no necrosis was observed in any of the plants, and we have not elucidated the plant, environment, and glyphosate-application conditions required to trigger this response in common ragweed.

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31P-NMR studies of mature common ragweed leaf tissue with glyphosate delivered by perfusion showed similar uptake of glyphosate into the leaf cells and no vacuolar sequestration in all common ragweed biotypes, with higher accumulation of shikimate-3-phosphate in glyphosate-susceptible common ragweed plants. An atypical biotype (R4) from the R population, which was identified from progeny of a single plant that was successfully self-pollinated, had somewhat consistently displayed necrosis of mature leaf tissue when sprayed with high rates of glyphosate. Mature source leaves collected from glyphosate-sensitive plants 24 hours after spray treatment behaved similarly to glyphosate-perfused leaves, but R4 mature leaves absorbed very little glyphosate and began to show signs of cell death. Initially, immature sink leaf tissue that was shielded from the glyphosate spray applications and collected 24 HAT appeared similar to glyphosate-perfused mature leaves, except that both biotypes had received very little glyphosate. However, unlike the spray-treated mature leaves, the S biotype began to show signs of cell death, whereas the R4 tissue remained relatively healthy during the observation period.

The overall findings presented in this dissertation indicate that common ragweed evolved multiple mechanisms of resistance within one population, and likely within individual plants. Common ragweed appears to exhibit the type of genetic diversity employed by other successful invasive species in the process of rapid evolutionary adaptation (Prentis et al., 2008). Several hypothetical scenarios exist for the sequence of trait selection (Olson-Manning et al., 2012), including:

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a) All mechanisms existed as standing genetic variation within the field, and

survived the selective sweep of a single glyphosate application.

b) An initial mechanism was selected, followed by introgression of other traits from

the extended common ragweed population of untreated plants surrounding the

field.

c) An initial mechanism was selected, and other mechanisms arose from new

mutations and/or recombination in subsequent years.

The most important step was the selection of the initial mechanism. This mechanism could have been sufficient to survive an ideal application of a recommended rate of glyphosate, but this is not an absolute requirement. For a variety of reasons, a lower effective dose of glyphosate could have been delivered to one or more individual plants, and this would have increased the likelihood of selection for any weak mechanism(s) that existed in among the genetic variation of the population. Target-site glyphosate- resistance mechanisms would be expected to display incomplete dominance, meaning that they would have a weaker effect in a heterozygous state than a homozygous state. If a plant was only capable of surviving a reduced effective dose of glyphosate, recombination and/or epigenetic variation would have been required for its progeny to reach the level of glyphosate resistance that we observed. This is similar to the principle of quantitative disease resistance (Poland et al., 2009), and consistent with the integrative mechanistic model of glyphosate resistance presented by Sammons and Gaines (2014).

Our studies suggest that the apparent strategy of broad genetic diversification that has made common ragweed a successful invasive species, is also central to its development of herbicide resistance. Whether this occurred through standing genetic variation or a

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high rate of new mutations is an intriguing question, and the most productive strategy for future research may be through utilization of modern techiques in the field of population genetics.

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Chapter 5 References

Heap I (2014) The international survey of herbicide resistant weeds. http://www.weedscience.com. Accessed November 5, 2014

Léonard R, Wopfner N, Pabst M, Stadlmann J, Petersen BO, Duus J, Himly M, Radauer C, Gadermaier G, Razzazi-Fazeli E, Ferreira F, Altmann F (2010) A New Allergen from Ragweed (Ambrosia artemisiifolia) with Homology to Art v 1 from Mugwort. Journal of Biological Chemistry 285:27192-27200

Olson-Manning CF, Wagner MR, Mitchell-Olds T (2012) Adaptive evolution: evaluating empirical support for theoretical predictions. Nat Rev Genet 13:867-877

Poland JA, Balint-Kurti PJ, Wisser RJ, Pratt RC, Nelson RJ (2009) Shades of gray: the world of quantitative disease resistance. Trends in Plant Science 14:21-29

Prentis PJ, Wilson JRU, Dormontt EE, Richardson DM, Lowe AJ (2008) Adaptive evolution in invasive species. Trends in Plant Science 13:288-294

Rousonelos SL, Lee RM, Moreira MS, VanGessel MJ, Tranel PJ (2012) Characterization of a common ragweed (Ambrosia artemisiifolia) population resistant to ALS- and PPO-inhibiting herbicides. Weed Science 60:335-344

Sammons RD, Gaines TA (2014) Glyphosate resistance: state of knowledge. Pest Management Science

Tranel PJ, Jiang W, Patzoldt WL, Wright TR (2004) Intraspecific variability of the acetolactate synthase gene. Weed Science 52:236-241

Zheng D, Patzoldt WL, Tranel PJ (2005) Association of the W574L ALS substitution with resistance to cloransulam and imazamox in common ragweed (Ambrosia artemisiifolia). Weed Science 53:424-430

104

References

Alarcón-Reverte R, García A, Urzúa J, Fischer AJ (2012) Resistance to glyphosate in junglerice (Echinochloa colona) from California. Weed Science 61:48-54

Baerson SR, Rodriguez DJ, Biest NA, Tran M, You J, Kreuger RW, Dill GM, Pratley JE, Gruys KJ (2002a) Investigating the mechanism of glyphosate resistance in rigid ryegrass (Lolium ridigum). Weed Science 50:721-730

Baerson SR, Rodriguez DJ, Tran M, Feng Y, Biest NA, Dill GM (2002b) Glyphosate- resistant goosegrass. Identification of a mutation in the target enzyme 5- enolpyruvylshikimate-3-phosphate synthase. Plant Physiol 129:1265-75

Baskin JM, Baskin CC (1980) Ecophysiology of secondary dormancy in seeds of Ambrosia artemisiifolia. Ecology 61:475-480

Bass DJ, Delpech V, Beard J, Bass P, Walls RS (2000a) Late summer and fall (March- May) pollen allergy and respiratory disease in Northern New South Wales, Australia. Annals of allergy, asthma & immunology : official publication of the American College of Allergy, Asthma, & Immunology 85:374-381

Bass DJ, Delpech V, Beard J, Bass P, Walls RS (2000b) Ragweed in Australia. Aerobiologia 16:107-111

Bassett IJ, Crompton CW (1975) The biology of Canadian weeds.: 11. Ambrosia artemisiifolia L. and A. psilostachya DC. Canadian Journal of Plant Science 55:463-476

Bassett IJ, Terasmae J (1962) Ragweeds, Ambrosia species, in Canada and their history in postglacial time. Canadian Journal of Botany 40:141-150

Belz RG, Duke SO (2014) Herbicides and plant hormesis. Pest Management Science 70:698-707

Bethel JD (2013) Evaluation of glyphosate resistant giant ragweed (Ambrosia trifida) in Ohio soybean (Glycine max) fields. MS Thesis. Columbus, OH: The Ohio State University

Binkholder KM, Fresenburg BS, Teuton TC, Xiong X, Smeda RJ (2011) Selection of glyphosate-resistant annual bluegrass (Poa annua) on a golf course. Weed Science 59:286-289

105

Bostamam Y, Malone JM, Dolman FC, Boutsalis P, Preston C (2012) Rigid ryegrass (Lolium rigidum) populations containing a target site mutation in EPSPS and reduced glyphosate translocation are more resistant to glyphosate. Weed Science 60:474-479

Boulet LP, Turcotte H, Laprise C, Lavertu C, Bedard PM, Lavoie A, Hébert J (1997) Comparative degree and type of sensitization to common indoor and outdoor allergens in subjects with allergic rhinitis and/or asthma. Clinical & Experimental Allergy 27:52-59

Brammall RA, Higgins VJ (1988) The effect of glyphosate on resistance of tomato to Fusarium crown and root rot disease and on the formation of host structural defensive barriers. Canadian Journal of Botany 66:1547-1555

Bretthorst GL (1990) Bayesian analysis. III. Applications to NMR signal detection, model selection, and parameter estimation. Journal of Magnetic Resonance 88:571-595

Brewer CE, Oliver LR (2009) Confirmation and resistance mechanisms in glyphosate- resistant common ragweed (Ambrosia artemisiifolia) in Arkansas. Weed Science 57:567-573

Burbach GJ, Heinzerling LM, Röhnelt C, Bergmann KC, Behrendt H, Zuberbier T (2009) Ragweed sensitization in Europe – GA2LEN study suggests increasing prevalence. Allergy 64:664-665

Burton WN, Conti DJ, Chen C-Y, Schultz AB, Edington DW (2001) The Impact of Allergies and Allergy Treatment on Worker Productivity. Journal of Occupational and Environmental Medicine 43:64-71

Carvalho LB, Alves PL, Duke SO (2013) Hormesis with glyphosate depends on coffee growth stage. Anais da Academia Brasileira de Ciências 85:813-822

Chauvel B, Dessaint F, Cardinal-Legrand C, Bretagnolle F (2006) The historical spread of Ambrosia artemisiifolia L. in France from herbarium records. Journal of Biogeography 33:665-673

Chikoye D, Weise SF, Swanton CJ (1995) Influence of common ragweed (Ambrosia artemisiifolia) time of emergence and density on white bean (Phaseolus vulgaris). Weed Science 43:375-380

Clewis SB, Askew SD, Wilcut JW (2001) Common ragweed interference in peanut. Weed Science 49:768-772

Coble HD, Williams FM, Ritter RL (1981) Common ragweed (Ambrosia artemisiifolia) interference in soybeans (Glycine max). Weed Science 29:339-342

106

Collavo A, Sattin M (2012) Resistance to glyphosate in Lolium rigidum selected in Italian perennial crops: bioevaluation, management and molecular bases of target-site resistance. Weed Research 52:16-24

Cowbrough MJ, Brown RB, Tardif FJ (2003) Impact of common ragweed (Ambrosia artemisiifolia) aggregation on economic thresholds in soybean. Weed Science 51:947-954

Cseh A, Cernák I, Taller J (2009) Molecular characterization of atrazine resistance in common ragweed (Ambrosia artemisiifolia L.). Journal of Applied Genetics 50:321-327

Csontos P, Vitalos M, Barina Z, Kiss L (2010) Early distribution and spread of Ambrosia artemisiifolia in Central and Eastern Europe. Bot. Helv. 120:75-78

Culpepper AS, Grey TL, Vencill WK, Kichler JM, Webster TM, Brown SM, York AC, Davis JW, Hanna WW (2006) Glyphosate-resistant Palmer amaranth (Amaranthus palmeri ) confirmed in Georgia. Weed Science 54:620-626

Culpepper AS, Whitaker JR, MacRae AW, York AC (2008) Distribution of glyphosate- resistant Palmer amaranth (Amaranthus palmeri) in Georgia and North Carolina during 2005 and 2006. J. Cotton Sci 12:306-310

Deen W, Hunt T, Swanton CJ (1998) Influence of temperature, photoperiod, and irradiance on the phenological development of common ragweed (Ambrosia artemisiifolia). Weed Science 46:555-560

Dickerson CT, Jr, Sweet RD (1971) Common ragweed ecotypes. Weed Science 19:64-66

Dinelli G, Marotti I, Bonetti A, Catizone P, Urbano JM, Barnes J (2008) Physiological and molecular bases of glyphosate resistance in Conyza bonariensis biotypes from Spain. Weed Research 48:257-265

Dinelli G, Marotti I, Bonetti A, Minelli M, Catizone P, Barnes J (2006) Physiological and molecular insight on the mechanisms of resistance to glyphosate in Conyza canadensis (L.) Cronq. biotypes. Pesticide Biochemistry and Physiology 86:30-41

DiTommaso A (2004) Germination behavior of common ragweed (Ambrosia artemisiifolia) populations across a range of salinities. Weed Science 52:1002- 1009

Duke SO, Powles SB (2008a) Glyphosate-Resistant Weeds and Crops. Pest Management Science 64:317-318

Duke SO, Powles SB (2008b) Glyphosate: a once-in-a-century herbicide. Pest Management Science 64:319-325

107

Eom SH, DiTommaso A, Weston LA (2013) Effects of soil salinity in the growth of Ambrosia artemisiifolia biotypes collected from roadside and agricultural field. Journal of Plant Nutrition 36:2191-2204

Falco SC, McDevitt RE, Chui CF, Hartnett ME, Knowlton S, Mauvais CJ, Smith JK, Mazur BJ (1989) Engineering herbicide-resistant acetolactate synthase. Dev Ind Microbiol 30:187-194

Feng PCC, Pratley JE, Bohn JA (1999) Resistance to glyphosate in Lolium rigidum. II. Uptake, translocation, and metabolism. Weed Science 47:412-415

Feng PCC, Tran M, Chiu T, Douglas Sammons R, Heck GR, CaJacob CA (2004) Investigations into glyphosate-resistant horseweed (Conyza canadensis): retention, uptake, translocation, and metabolism. Weed Science 52:498-505

Friedman J, Barrett SCH (2008) High outcrossing in the annual colonizing species Ambrosia artemisiifolia (Asteraceae). Annals of Botany 101:1303-1309

Fuchs MA, Geiger DR, Reynolds TL, Bourque JE (2002) Mechanisms of glyphosate toxicity in velvetleaf (Abutilon theophrasti medikus). Pesticide Biochemistry and Physiology 74:27-39

Gaines TA, Shaner DL, Ward SM, Leach JE, Preston C, Westra P (2011) Mechanism of resistance of evolved glyphosate-resistant Palmer amaranth (Amaranthus palmeri). Journal of Agricultural and Food Chemistry 59:5886-5889

Gaines TA, Ward SM, Bukun B, Preston C, Leach JE, Westra P (2012) Interspecific hybridization transfers a previously unknown glyphosate resistance mechanism in Amaranthus species. Evolutionary Applications 5:29-38

Gaines TA, Zhang W, Wang D, Bukun B, Chisholm ST, Shaner DL, Nissen SJ, Patzoldt WL, Tranel PJ, Culpepper AS, Grey TL, Webster TM, Vencill WK, Sammons RD, Jiang J, Preston C, Leach JE, Westra P (2009) Gene amplification confers glyphosate resistance in Amaranthus palmeri. Proceedings of the National Academy of Sciences 107:1029-1034

Gaudeul M, Giraud T, Kiss L, Shykoff JA (2011) Nuclear and chloroplast microsatellites show multiple introductions in the worldwide invasion history of common ragweed, Ambrosia artemisiifolia. PloS one 6:e17658

Ge X, d'Avignon DA, Ackerman JJ, Collavo A, Sattin M, Ostrander EL, Hall EL, Sammons RD, Preston C (2012a) Vacuolar glyphosate-sequestration correlates with glyphosate resistance in ryegrass (Lolium spp.) from Australia, South America, and Europe: a 31P NMR investigation. Journal of Agricultural and Food Chemistry 60:1243-50

108

Ge X, d'Avignon DA, Ackerman JJH, Duncan B, Spaur MB, Sammons RD (2011) Glyphosate-resistant horseweed made sensitive to glyphosate: low-temperature suppression of glyphosate vacuolar sequestration revealed by 31P NMR. Pest Management Science 67:1215-1221

Ge X, d'Avignon DA, Ackerman JJH, Ostrander E, Sammons RD (2013) Application of 31P-NMR spectroscopy to glyphosate studies in plants: Insights into cellular uptake and vacuole sequestration correlated to herbicide resistance. Pages 55-83 Handbook on Herbicides : Biological Activity, Classification and Health and Environmental Implications. Hauppauge, NY, USA: Nova Science Publishers, Inc.

Ge X, d'Avignon DA, Ackerman JJH, Sammons RD (2010) Rapid vacuolar sequestration: the horseweed glyphosate resistance mechanism. Pest Management Science:n/a-n/a

Ge X, d’Avignon DA, Ackerman JJH, Collavo A, Sattin M, Ostrander EL, Hall EL, Sammons RD, Preston C (2012b) Vacuolar glyphosate-sequestration correlates with glyphosate resistance in ryegrass (Lolium spp.) from Australia, South America, and Europe: A 31P NMR investigation. Journal of Agricultural and Food Chemistry 60:1243-1250

Ge X, d’Avignon DA, Ackerman JJH, Sammons RD (2012c) Observation and identification of 2-C-methyl-d-erythritol-2,4-cyclopyrophosphate in horseweed and ryegrass treated with glyphosate. Pesticide Biochemistry and Physiology 104:187-191

Genton BJ, Shykoff JA, Giraud T (2005) High genetic diversity in French invasive populations of common ragweed, Ambrosia artemisiifolia, as a result of multiple sources of introduction. Molecular Ecology 14:4275-4285

González-Torralva F, Gil-Humanes J, Barro F, Domínguez-Valenzuela J, De Prado R (2014) First evidence for a target site mutation in the EPSPS2 gene in glyphosate- resistant Sumatran fleabane from citrus orchards. Agron. Sustain. Dev. 34:553- 560

Harrison SK, Regnier EE, Schmoll JT, Webb JE (2001) Competition and fecundity of giant ragweed in corn. Weed Science 49:224-229

Heap I (2014) The international survey of herbicide resistant weeds. http://www.weedscience.com. Accessed November 5, 2014

Hodgins KA, Rieseberg L (2011) Genetic differentiation in life-history traits of introduced and native common ragweed (Ambrosia artemisiifolia) populations. Journal of Evolutionary Biology 24:2731-2749

Jalaludin A, Han H, Powles S (2013) Evolution in action: a double amino acid substitution in the EPSPS gene endows high-level glyphosate resistance. Pages 109

35-35 in Proceedings of the Global Herbicide Resistance Challenge. Fremantle, Australia: Australian Herbicide Resistance Initiative

Joly M, Bertrand P, Gbangou RY, White M-C, Dubé J, Lavoie C (2011) Paving the way for invasive species: road type and the spread of common ragweed (Ambrosia artemisiifolia). Environmental Management 48:514-522

Jones KL (1936) Studies on Ambrosia, I. The inheritance of floral types in the ragweed, Ambrosia elatior L. American Midland Naturalist 17:673-699

Karis PO (1995) Cladistics of the subtribe Ambrosiinae (Asteraceae: Heliantheae). Systematic Botany 20:40-54

Kaundun SS, Dale RP, Zelaya IA, Dinelli G, Marotti I, McIndoe E, Cairns A (2011) A novel P106L mutation in EPSPS and an unknown mechanism(s) act additively to confer resistance to glyphosate in a South African Lolium rigidum population. Journal of Agricultural and Food Chemistry 59:3227-3233

Kaundun SS, Zelaya IA, Dale RP, Lycett AJ, Carter P, Sharples KR, McIndoe E (2008) Importance of the P106S target-site mutation in conferring resistance to glyphosate in a goosegrass (Eleusine indica) population from the Philippines. Weed Science 56:637-646

Kil JH, Shim KC, Park SH, Koh KS, Suh MH, Ku YB, Suh SU, Oh HK, Kong HY (2004) Distributions of naturalized alien plants in South Korea. Weed Technology 18:1493-1495

Kiss L, Béres I (2006) Anthropogenic factors behind the recent population expansion of common ragweed (Ambrosia artemisiifolia L.) in Eastern Europe: is there a correlation with political transitions? Journal of Biogeography 33:2156-2157

Koger CH, Reddy KN (2005) Role of absorption and translocation in the mechanism of glyphosate resistance in horseweed (Conyza canadensis). Weed science 53:84-89

Lavoie C, Jodoin Y, De Merlis AG (2007) How did common ragweed (Ambrosia artemisiifolia L.) spread in Québec? A historical analysis using herbarium records. Journal of Biogeography 34:1751-1761

Legleiter TR, Bradley KW (2008) Glyphosate and multiple herbicide resistance in common waterhemp (Amaranthus rudis) populations from Missouri. Weed Science 56:582-587

Léonard R, Wopfner N, Pabst M, Stadlmann J, Petersen BO, Duus J, Himly M, Radauer C, Gadermaier G, Razzazi-Fazeli E, Ferreira F, Altmann F (2010) A New Allergen from Ragweed (Ambrosia artemisiifolia) with Homology to Art v 1 from Mugwort. Journal of Biological Chemistry 285:27192-27200

110

Leskovšek R, Datta A, Knezevic SZ, Simončič A (2012) Common ragweed (Ambrosia artemisiifolia) dry matter allocation and partitioning under different nitrogen and density levels. Weed Biology and Management 12:98-108

Liu L, Punja ZK, Rahe JE (1997) Altered root exudation and suppression of induced lignification as mechanisms of predisposition by glyphosate of bean roots (Phaseolus vulgaris L.) to colonization by Pythium spp. Physiological and Molecular Plant Pathology 51:111-127

Livak KJ, Schmittgen TD (2001) Analysis of relative gene expression data using real- time quantitative PCR and the 2−ΔΔCT method. Methods 25:402-408

Lorraine-Colwill DF, Powles SB, Hawkes TR, Hollinshead PH, Warner SAJ, Preston C (2002) Investigations into the mechanism of glyphosate resistance in Lolium rigidum. Pesticide Biochemistry and Physiology 74:62-72

Makra L, Juhász M, Béczi R, Borsos Ek (2005) The history and impacts of airborne Ambrosia (Asteraceae) pollen in Hungary. Grana 44:57-64

Maroufi A, Van Bockstaele E, De Loose M (2010) Validation of reference genes for gene expression analysis in chicory (Cichorium intybus) using quantitative real-time PCR. BMC molecular biology 11:15

Maryushkina VY (1991) Peculiarities of common ragweed (Ambrosia artemisiifolia L.) strategy. Agriculture, Ecosystems & Environment 36:207-216

Mátyás K, Taller J, Cseh A, Poczai P, Cernák I (2011) Development of a simple PCR- based assay for the identification of triazine resistance in the noxious plant common ragweed (Ambrosia artemisiifolia) and its applicability in higher plants. Biotechnology Letters 33:2509-2515

McAndrews JH (1988) Human disturbance of North American forests and grasslands: The fossil pollen record. Pages 673-697 in Huntley B, Webb T, III, eds. Vegetation history: Springer Netherlands

Michitte P, De Prado R, Espinoza N, Pedro Ruiz-Santaella J, Gauvrit C (2007) Mechanisms of resistance to glyphosate in a ryegrass (Lolium multiflorum) biotype from Chile. Weed Science 55:435-440

Mulligan GA (1957) Chromosome numbers of Canadian weeds. I. Canadian Journal of Botany 35:779-789

Munoz SE, Gajewski K (2010) Distinguishing prehistoric human influence on late- Holocene forests in southern Ontario, Canada. The Holocene 20:967-981

111

Myers MW, Curran WS, VanGessel MJ, Calvin DD, Mortensen DA, Majek BA, Karsten HD, Roth GW (2004) Predicting weed emergence for eight annual species in the northeastern United States. Weed Science 52:913-919

Mylonas PN, Giannopolitis CN, Efthimiadis PG, Menexes GC, Madesis PB, Eleftherohorinos IG (2014) Glyphosate resistance of molecularly identified Conyza albida and Conyza bonariensis populations. Crop Protection 65:207-215

Nandula VK, Ray JD, Ribeiro DN, Pan Z, Reddy KN (2013) Glyphosate resistance in tall waterhemp (Amaranthus tuberculatus) from Mississippi is due to both altered target-site and nontarget-site mechanisms. Weed Science 61:374-383

Nandula VK, Reddy KN, Poston DH, Rimando AM, Duke SO (2008) Glyphosate tolerance mechanism in Italian ryegrass (Lolium multiflorum) from Mississippi. Weed Science 56:344-349

Ng CH, Wickneswari R, Salmijah S, Teng YT, Ismail BS (2003) Gene polymorphisms in glyphosate-resistant and -susceptible biotypes of Eleusine indica from Malaysia. Weed Research 43:108-115

Nol N, Tsikou D, Eid M, Livieratos IC, Giannopolitis CN (2012) Shikimate leaf disc assay for early detection of glyphosate resistance in Conyza canadensis and relative transcript levels of EPSPS and ABC transporter genes. Weed Research 52:233-241

Norsworthy JK, Griffith GM, Scott RC, Smith KL, Oliver LR (2008) Confirmation and control of glyphosate-resistant Palmer amaranth (Amaranthus palmeri) in Arkansas. Weed Technology 22:108-113

Okada M, Jasieniuk M (2014) Inheritance of glyphosate resistance in hairy fleabane (Conyza bonariensis) from California. Weed Science 62:258-266

Olson-Manning CF, Wagner MR, Mitchell-Olds T (2012) Adaptive evolution: evaluating empirical support for theoretical predictions. Nat Rev Genet 13:867-877

Patzoldt WL, Tranel PJ (2002) Molecular analysis of cloransulam resistance in a population of giant ragweed. Weed Science 50:299-305

Patzoldt WL, Tranel PJ, Alexander AL, Schmitzer PR (2001) A common ragweed population resistant to cloransulam-methyl. Weed Science 49:485-490

Payne WW (1963) The morphology of the inflorescence of ragweeds (Ambrosia- Franseria: Compositae). American Journal of Botany 50:872-880

Payne WW, Raven PH, Kyhos DW (1964) Chromosome numbers in Compositae. IV. Ambrosieae. American Journal of Botany:419-424

112

Perez-Jones A, Park K-W, Polge N, Colquhoun J, Mallory-Smith CA (2007) Investigating the mechanisms of glyphosate resistance in Lolium multiflorum. Planta 226:395-404

Poland JA, Balint-Kurti PJ, Wisser RJ, Pratt RC, Nelson RJ (2009) Shades of gray: the world of quantitative disease resistance. Trends in Plant Science 14:21-29

Powles SB, Lorraine-Colwill DF, Dellow JJ, Preston C (1998) Evolved resistance to glyphosate in rigid ryegrass (Lolium rigidum) in Australia. Weed Science 46:604- 607

Prentis PJ, Wilson JRU, Dormontt EE, Richardson DM, Lowe AJ (2008) Adaptive evolution in invasive species. Trends in Plant Science 13:288-294

Preston C, Wakelin AM, Dolman FC, Bostamam Y, Boutsalis P (2009) A decade of glyphosate-resistant Lolium around the world: Mechanisms, genes, fitness, and agronomic management. Weed Science 57:435-441

R Core Team (2014) R: A Language and Environment for Statistical Computing. R Foundation for Statistical Computing. Vienna, Austria. http://www.R-project.org/.

Riar DS, Norsworthy JK, Johnson DB, Scott RC, Bagavathiannan M (2011) Glyphosate resistance in a johnsongrass (Sorghum halepense) biotype from Arkansas. Weed Science 59:299-304

Ritz C, Strebig J (2013) Package ‘drc’.

Ritz C, Streibig JC (2005) Bioassay analysis using R. Journal of Statistical Software 12:1-22

Rosenbaum KK, Miller GL, Kremer RJ, Bradley KW (2014) Interactions between glyphosate, Fusarium infection of common waterhemp (Amaranthus rudis), and soil microbial abundance and diversity in soil collections from Missouri. Weed Science 62:71-82

Rousonelos SL, Lee RM, Moreira MS, VanGessel MJ, Tranel PJ (2012) Characterization of a common ragweed (Ambrosia artemisiifolia) population resistant to ALS- and PPO-inhibiting herbicides. Weed Science 60:335-344

Saint-Louis S, DiTommaso A, Watson AK (2005) A common ragweed (Ambrosia artemisiifolia) biotype in southwestern Québec resistant to linuron. Weed Technology 19:737-743

Salas RA, Dayan FE, Pan Z, Watson SB, Dickson JW, Scott RC, Burgos NR (2012) EPSPS gene amplification in glyphosate-resistant Italian ryegrass (Lolium perenne ssp. multiflorum) from Arkansas. Pest Management Science 68:1223- 1230 113

Sammons RD, Gaines TA (2014) Glyphosate resistance: state of knowledge. Pest Management Science

Sammons RD, Heering DC, Dinicola N, Glick H, Elmore GA (2007a) Sustainability and stewardship of glyphosate and glyphosate-resistant crops. Weed Technology 21:347-354

Sammons RD, Meyer J, Hall E, Ostrander E, Schrader S A simple continuous assay for EPSP synthase in plant tissue. Champaign, IL: North Central Weed Science Society

Santos G, Oliveira RS, Constantin J, Constantin Francischini A, Machado MFPS, Mangolin CA, Nakajima JN (2014) Conyza sumatrensis: A new weed species resistant to glyphosate in the Americas. Weed Biology and Management 14:106- 114

Schafer JR, Hallett SG, Johnson WG (2012) Response of giant ragweed (Ambrosia trifida), horseweed (Conyza canadensis), and common lambsquarters (Chenopodium album) biotypes to glyphosate in the presence and absence of soil microorganisms. Weed Science 60:641-649

Schafer JR, Hallett SG, Johnson WG (2013) Soil microbial root colonization of glyphosate-treated giant ragweed (Ambrosia trifida), horseweed (Conyza canadensis), and common lambsquarters (Chenopodium album) biotypes. Weed Science 61:289-295

Sharon A, Amsellem Z, Gressel J (1992) Glyphosate suppression of an elicited defense response: Increased susceptibility of Cassia obtusifolia to a mycoherbicide. Plant Physiology 98:654-659

Simard M-J, Benoit DL (2012) Potential pollen and seed production from early- and late- emerging common ragweed in corn and soybean. Weed Technology 26:510-516

Simard M-J, Panneton B, Longchamps L, Lemieux C, Légère A, Leroux GD (2009) Validation of a management program based on a weed cover threshold model: Effects on herbicide use and weed populations. Weed Science 57:187-193

Simard MJ, Benoit DL (2010) Distribution and abundance of an allergenic weed, common ragweed (Ambrosia artemisiifolia L.), in rural settings of southern Quebec, Canada. Canadian Journal of Plant Science 90:549-557

Simarmata M, Penner D (2008) The basis for glyphosate resistance in rigid ryegrass (Lolium Rigidum) from California. Weed Science 56:181-188

Smith DA, Hallett SG (2006) Interactions between chemical herbicides and the candidate bioherbicide Microsphaeropsis amaranthi. Weed Science 54:532-537

114

Stachler JM (2008) Characterization and management of glyphosate-resistant giant ragweed (Ambrosia trifida L.) and horseweed [Conyza canadensis (L.) Cronq.]. Ph.D Dissertation. Columbus, OH: The Ohio State University

Steinrücken HC, Amrhein N (1980) The herbicide glyphosate is a potent inhibitor of 5- enolpyruvylshikimic acid-3-phosphate synthase. Biochemical and Biophysical Research Communications 94:1207-1212

Stephenson GR, Dykstra MD, McLaren RD, Hamill AS (1990) Agronomic practices influencing triazine-resistant weed distribution in Ontario. Weed Technology 4:199-207

Stoller EW, Wax LM (1973) Periodicity of germination and emergence of some annual weeds. Weed Science 21:574-580

Tokarska-Guzik B, Bzdęga K, Koszela K, Żabińska I, Krzuś B, Sajan M, Sendek A (2011) Allergenic invasive plant Ambrosia artemisiifolia L. in Poland: threat and selected aspects of biology. Biodiversity: Research and Conservation 21:39-48

Tranel PJ, Jiang W, Patzoldt WL, Wright TR (2004) Intraspecific variability of the acetolactate synthase gene. Weed Science 52:236-241

Tranel PJ, Wright TR (2002) Resistance of weeds to ALS-inhibiting herbicides: what have we learned? Weed Science 50:700-712

Traveset A (1992) Sex expression in a natural population of the monoecious annual, Ambrosia artemisiifolia (Asteraceae). American Midland Naturalist 127:309-315

Týr Š, Vereš T, Lacko-Bartošová M (2009) Occurrence of common ragweed (Ambrosia artemisiifolia L.) in field crops in the Slovak Republic. Herbologia 10:1-9

Urbano JM, Borrego A, Torres V, Leon JM, Jimenez C, Dinelli G, Barnes J (2007) Glyphosate-resistant hairy fleabane (Conyza bonariensis) in Spain. Weed Technology 21:396-401

Van Horn CR, Westra P (2014) Updates on molecular response of glyphosate resistant giant ragweed (Ambrosia trifida). in Proceedings of the Weed Science Society of America and Canadian Weed Science Society. Vancouver, BC: Weed Science Society of America

VanGessel MJ (2001) Glyphosate-resistant horseweed from Delaware. Weed Science 49:703-705

Velini ED, Alves E, Godoy MC, Meschede DK, Souza RT, Duke SO (2008) Glyphosate applied at low doses can stimulate plant growth. Pest Management Science 64:489-496

115

Waite J, Thompson CR, Peterson DE, Currie RS, Olson BLS, Stahlman PW, Al-Khatib K (2012) Differential kochia (Kochia scoparia) populations response to glyphosate. Weed Science 61:193-200

Wakelin AM, Lorraine-Colwill DF, Preston C (2004) Glyphosate resistance in four different populations of Lolium rigidum is associated with reduced translocation of glyphosate to meristematic zones. Weed Research 44:453-459

Wakelin AM, Preston C (2006) A target-site mutation is present in a glyphosate-resistant Lolium rigidum population. Weed Research 46:432-440

Walker S, Bell K, Robinson G, Widderick M (2011) Flaxleaf fleabane (Conyza bonariensis) populations have developed glyphosate resistance in north-east Australian cropping fields. Crop Protection 30:311-317

Weaver SE (2001) Impact of lamb’s-quarters, common ragweed and green foxtail on yield of corn and soybean in Ontario. Canadian Journal of Plant Science 81:821- 828

Webb MR (1992) A continuous spectrophotometric assay for inorganic phosphate and for measuring phosphate release kinetics in biological systems. Proceedings of the National Academy of Sciences of the United States of America 89:4884-4887

Werle R, Sandell LD, Buhler DD, Hartzler RG, Lindquist JL (2014) Predicting emergence of 23 summer annual weed species. Weed Science 62:267-279

Wiersma AT (2012) Regional whole plant and molecular response of Kochia scoparia to glyphosate. MS Thesis. Fort Collins, CO: Colorado State University

Willemsen RW (1975) Effect of stratification temperature and germination temperature on germination and the induction of secondary dormancy in common ragweed seeds. American Journal of Botany 62:1-5

Wopfner N, Gadermaier G, Egger M, Asero R, Ebner C, Jahn-Schmid B, Ferreira F (2005) The spectrum of allergens in ragweed and mugwort pollen. International archives of allergy and immunology 138:337-346

Wright TR, Bascomb NF, Sturner SF, Penner D (1998) Biochemical mechanism and molecular basis for ALS-inhibiting herbicide resistance in sugarbeet (Beta vulgaris) somatic cell selections. Weed Science:13-23

Ye J, Coulouris G, Zaretskaya I, Cutcutache I, Rozen S, Madden TL (2012) Primer- BLAST: a tool to design target-specific primers for polymerase chain reaction. BMC bioinformatics 13:134

116

Yu Q, Abdallah I, Han H, Owen M, Powles S (2009) Distinct non-target site mechanisms endow resistance to glyphosate, ACCase and ALS-inhibiting herbicides in multiple herbicide-resistant Lolium rigidum. Planta 230:713-723

Yu Q, Cairns A, Powles S (2007) Glyphosate, paraquat and ACCase multiple herbicide resistance evolved in a Lolium rigidum biotype. Planta 225:499-513

Yuan JS, Abercrombie LLG, Cao Y, Halfhill MD, Zhou X, Peng Y, Hu J, Rao MR, Heck GR, Larosa TJ, Sammons RD, Wang X, Ranjan P, Johnson DH, Wadl PA, Scheffler BE, Rinehart TA, Trigiano RN, Stewart CN (2010) Functional genomics analysis of horseweed (Conyza canadensis) with special reference to the evolution of non-target-site glyphosate resistance. Weed Science 58:109-117

Zar JH (1996) Biostatistical Analysis. 3rd edn. Upper Saddle River, NJ: Prentice-Hall

Zheng D, Patzoldt WL, Tranel PJ (2005) Association of the W574L ALS substitution with resistance to cloransulam and imazamox in common ragweed (Ambrosia artemisiifolia). Weed Science 53:424-430

Ziska LH, Schlesinger WH, Epstein PR (2009) Rising CO2, climate change, and public health: Exploring the links to plant biology. Environmental Health Perspectives 117:155-158

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Appendix A: In vivo 31P-NMR Results

X. Ge1, D.A. D'Avignon1, J.T. Parrish2, M.M. Loux2, R.D. Sammons3 1Washington University, St. Louis, MO; 2The Ohio State University, Columbus, OH; 3Monsanto Company, St. Louis, MO.

A.1 31P-NMR observations for source tissue following glyphosate infusion

Mature expanded leaves, from both R4 and S1 plant biotypes, were perfused with buffer solution containing glyphosate and studied via 31P-NMR. The 31P-NMR spectra, collected in two-hour blocks, are presented in Figure A.1. The spectra show the anticipated NMR signals for ATP, UDPG, sugar phosphates, inorganic phosphate, and glyphosate. The tissue in all cases was viable and healthy as evidenced by the presence of ATP and a pH gradient demonstrated by the chemical shift difference between phosphate in the plant cellular cytosol (2.85 ppm) and vacuole (0.92 ppm). Substantial inorganic phosphate occurred in the vacuole for both biotypes. Both the R4 and S1 biotypes exhibited similar and moderate glyphosate uptake under perfusion infusion conditions; the ATP appeared to be somewhat higher in the S1 than the R4 biotypes. The glyphosate NMR signal was consistent with that previously observed for glyphosate occurring in the cell cytosol. We observed no evidence for vacuolar sequestration in either biotype under these conditions and for the time-course investigated. There was an increase over time of the signal attributed to shikimate-3-phosphate (S3P) at 4.56 ppm in the S1 biotype, with much less S3P accumulation observed in the R biotype. For the

24 hours of investigation, the S1 sample S3P accumulates sufficiently to decrease the 118

large signal corresponding to the vacuolar phosphate pool, as phosphorylation of shikimate to S3P indirectly consumes inorganic phosphate.

A.2 31P-NMR observations of source tissue following glyphosate spray treatment

31P-NMR data collected in two-hour time blocks is shown in Figure A.2 for an

18-hour period starting at 26 HAT. The NMR observations for leaves from the sprayed plants indicated tissue viability (ATP and pH gradient between cytosol and vacuole) at the early collection times for both biotypes. This changed with time, in that the

R4 cytosol became more acidic (pH change of 0.5 unit), whereas the pH of the S1 biotype was unchanged for the full 18 hours. The R4 biotype showed behavior consistent with cell death whereas the S1 biotype maintained apparent health, even at 18 hours. Similar to the glyphosate infusion experiments, there was greater buildup of S3P in the S1 biotype than the R4 biotype after spraying, though not as dramatic of a difference as in the infused glyphosate trials.

The primary difference between the sprayed and infused tissue was that there was little glyphosate observable by NMR in the R4 biotype 24 HAT, but a moderate amount in the S1 biotype. A secondary difference was the occurrence of 2-C-methyl-

D-erythritol-2,4--cyclopyrophosphate (MEcPP), as well as an apparent variant thereof, in both the S1 and R4 tissue of the sprayed plants. With tissue death occurring in the later collection times for R4, this substrate disappears along with ATP and any evidence of

S3P, presumably because it moved from the cell into the circulation perfusate bath

(representing a vastly large reservoir compared to the tissue cell volume).

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A.3 31P-NMR observations for sink tissue following glyphosate spray treatment

Sink tissue that had been protected during the herbicide spraying process, and thus able to gain access to glyphosate only through translocation, was harvested 24 HAT.

These plants were maintained in greenhouse conditions prior to tissue harvest. Harvested tissue was confined to immature that were light green in color and easily distinguished from the mature dark green fully differentiated leaf tissue. The 31P-NMR spectra obtained from this tissue in two-hour time block collection is shown in Figure A.3.

Comparisons between the NMR of R4 and S1 sink tissue shared some commonality with source tissue in that it was alive (indicated by presence of ATP and a cellular pH gradient), and the S3P buildup is substantial in the sensitive biotype. Again, so much

S3P accumulated in the sensitive tissue that the large inorganic phosphate signal corresponding to the vacuole pool (occurring at 0.65 ppm) is greatly diminished. The R4 tissue showed a decrease in ATP towards the end of the 18-hour trial, but pH did not change. The S1 tissue loses its pH gradient after 12 hours of perfusion and S3P exited the intact cell as the tissue became progressively more compromised. The cytosolic glyphosate measured by 31P-NMR from the data contained in Figure A.1 through Figure

A.3 is displayed graphically in Figure A.4.

The R4 biotype demonstrates a survival of uncovered sink tissue 12 days after treatment (DAT) as shown in Figure A.5. The sink tissue, in this case, is identified as the lighter green immature shoots that appear in both S and R plants. The S1 plant biotype is completely dead at 12 DAT. The S1 and R4 biotypes maintained in greenhouse conditions appeared visually unaffected 24 HAT with a 4 glyphosate spray, but they

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exhibited substantial changes in appearance 48 HAT. At 48 HAT, the R4 biotype leaves begin to lose their rigidity and appear water-soaked (except for sink tissue that maintains its lighter green color); S1 tissue looks normal and healthy at the same time point.

Twelve days later, leaf tissue of the S1 biotype was dead and only the sink tissue (not protected during spraying) in the R4 biotype was alive (Figure A.5). This is a somewhat remarkable biotype (R4), in that the young growing sink tissue does not share the apparently heritable degradation of the source tissue, in response to the herbicide treatment. The rapid degeneration of source leaf tissue in R4 due to glyphosate treatment does not coincide with the expected slow timescale of observed tissue degradation anticipated from the known mode of action of glyphosate blocking the shikimate pathway. The normal time course of plant death from glyphosate treatment is shown by the apparent healthy appearance of S1 plants 48 HAT, with subsequent slow and steady degradation. The rapid response of R4 is more like a hypersensitive response seen in plant-pathogen interactions, or the action of a cell-membrane disrupting herbicide, which may render the R4 source tissue incapable of effective translocation over many hours.

Conversely, the S1 biotype is anticipated to engage in effective translocation based on its healthy appearance in the early days after treatment, when 14C-glyphosate studies have shown the majority of translocation occurs. The 31P-NMR data obtained from source leaf tissue post-spraying supports this trend. But regardless of translocation, the immature sink tissue in the R4 biotype has a developmental difference that does not elicit the apparent hypersensitive response following glyphosate treatment observed in the R4 mature leaf tissue. This is apparently linked to the ability of the R4 biotype to survive substantial glyphosate doses.

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The 31P-NMR spectra from infusion studies on S1 and R4 source leaf tissue show moderate uptake of glyphosate by both tissue types, but substantial S3P accumulation in the S1 biotype. This observation may suggest the R4 biotype demonstrates some degree of chloroplast exclusion, or a target-site modification involving EPSPS (such as a mutation resulting in reduced inhibition by glyphosate or increased expression of the

EPSPS enzyme). This same observation occurred for the sink tissue studies where glyphosate had been delivered only through translocation (selected sink tissue protected during spray treatment). The S3P buildup was considerable, resulting in the reduction of the cell vacuole inorganic phosphate level by more than 50%. In previous NMR research with many other plant species, S3P accumulation varied considerably in source leaf tissue

(Ge et al., 2013). Between the biotypes studied here, trends in S3P accumulation were similar. However, S3P accumulation differs between the S1 and R4 biotypes in sink tissue where shikimate pathway flux is anticipated to be highest, suggesting the substantial S3P buildup in S1 and minimal S3P buildup in R4 tissue is in fact an important difference between the S1 and R4 biotypes.

The glyphosate observed by 31P-NMR 24 hours after spray treatment in R4 source tissue was very low compared with the S1 biotype. This was unexpected and appears reproducible. For reasons that are not clear (based on the absence of this observation from the infusion studies) the uptake of glyphosate in R4 in the presence of light appeared to be far less than in the S1 biotype. We assume, from the visual appearance of the plant 48 HAT, that enough glyphosate entered the plant cell to promote the overall unhealthy appearance of R4, but at 24 HAT, only a small NMR signal was observed

(Figure A.2). At 24 HAT, the S1 tissue maintained consistent cytosolic pH, suggesting 122

an overall healthier plant compared with the R4, for which pH changed. Although visual degeneration in R4 source leaf tissue was not apparent 24 HAT, the occurrence of this degeneration 48 HAT is consistent with the pH change observed by NMR in R4 mature leaf tissue 24 HAT. The opposite trend was observed in sink tissue that was protected during treatment, and therefore received glyphosate only via translocation. In this case, the R4 tissue maintained consistent pH over the 18 hours of collection, whereas the S1 tissue showed clear signs of cell death (loss of pH gradient, loss of cellular S3P) after

12 hours. The difference in sink tissue of the biotypes is striking, based on both the

NMR results and visual inspection following treatment. Taken together, it cannot be ruled out that the R4 biotype exhibits a toxic, possibility hypersensitive, response to glyphosate that adversely affects translocation, but also an altered EPSPS enzyme or restricted or delayed access of glyphosate to EPSPS, leading to its effective high resistance.

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Figure A.1. 31P-NMR spectra of perfused S and R common ragweed mature leaves following an 8-hour treatment with 10-mM glyphosate.

A) 31P-NMR spectra of the R4 biotype collected in 2-hour blocks. B) 31P-NMR spectra of the S1 biotype collected in 2-hour time blocks. Abbreviations: MDP, Methylenediphosphonate; *, residual buffer glyphosate; CG, cytoplasmic glyphosate; Sugar-P, sugar phosphates; S3P, shikimate-3-phosphate; CPi, cytoplasmic inorganic phosphate; VPi, vacuolar inorganic phosphate; α-, β-, γ-ATP refer to corresponding ATP phosphate groups; UDPG, uridine 5-diphosphoglucose.

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Figure A.2. 31P-NMR spectra of common ragweed mature (source) leaves 24-hours after spray treatment with 3.36 kg ae ha−1 glyphosate.

Tissue was studied for 18 hours. A) 31P-NMR spectra collected in 2-hour time blocks for treated R4 leaves perfused with glyphosate free buffer. B) 31P-NMR spectra collected in 2-hour time blocks for S1 leaves perfused with glyphosate free buffer. Abbreviation: MEcPP (Ge et al., 2012c), 2-C-methyl-D-erythritol-2,4-cyclopyrophosphate, (only aP is shown, bP is overlapped with NAD(P) and UDPG region at −11 ppm); other abbreviations see Figure A.1.

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Figure A.3. 31P-NMR spectra of common ragweed shielded immature (sink) leaves 24 hours after spray treatment of mature leaves with 3.36 kg ae ha−1 glyphosate.

A) 31P-NMR spectra from 2-hour time blocks of R4 common ragweed sink leaves. B) 31P-NMR spectra from 2-hour time block collections of S1 common ragweed sink leaves. Abbreviations see Figure A.1.

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Figure A.4. 31P-NMR measured cytosolic glyphosate relative to reference (MDP) 24 hours after treatment in common ragweed leaf tissues.

A) The uptake of glyphosate in common ragweed mature leaf after 10-mM glyphosate 8 h infiltration; B) The uptake of glyphosate in common ragweed mature leaf 24 HAT; C) The uptake of glyphosate in common ragweed sink leaf 24 HAT through glyphosate translocation from source leaf.

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Figure A.5. Photo of mature R4 and S1 common ragweed plants 12-days after spray treatment with 3.36 kg ae ha-1 glyphosate.

R4 (left) and S1 (right) mature common ragweed plants 12 days after spray treatment with 3.36 kg ae ha-1 glyphosate. R4 shows the survival of young growing tissue but S1 was killed.

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