The Induction and Photoregulation of Flavonoid Synthesis in Poa Trivialis L

The Induction and Photoregulation of Flavonoid Synthesis in Poa trivialis L. and its Impact on Salt Stress Sensitivity

DISSERTATION

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

By Dominic P. Petrella, BSc. Graduate Program in Horticulture and Crop Science

The Ohio State University 2017

Dissertation Committee: Dr. David S. Gardner, Advisor Dr. Joshua J. Blakeslee, Advisor Dr. James D. Metzger Dr. Anne E. Dorrance

Abstract

Flavonoids are a group of plant secondary metabolites that are in great demand for their health promoting properties, and specific flavonoids, like anthocyanins, are in demand because of their potential as natural dyes. However, sources of these metabolites are limiting, and new alternative plant species are needed to meet industry needs.

Turfgrasses are a group of monocots that could serve as biological production systems due to their unique anatomy. These plants, however, do not naturally produce large quantities of flavonoids, but some turfgrass species like rough bluegrass (Poa trivialis L.) exhibit greater flavonoid quantities relative to other turfgrasses. Flavonoids, in general, are highly regulated by light, and the application of wavelength-specific light treatments to rough bluegrass could increase flavonoid content to acceptable levels. While flavonoids are used in consumer products for their color or anti-oxidant properties, the function of flavonoids in plants is still not completely understood. However, flavonoids have been shown to interact with the transport and metabolism of the phytohormone auxin, and flavonoids may therefore act as modulators of plant tropic growth.

Halotropism, a recently defined tropic growth response, occurs following exposure to a directional salt stress, and roots will bend/grow away from the area of high salt concentration due to changes in auxin transport and metabolism. For turfgrasses, the use of re-claimed water as an alternate irrigation strategy is on the rise due to restrictions on fresh water irrigation, but re-claimed water can contain large concentrations of salt. ii

Salt exclusion and sequestration are well-understood mechanisms of salt tolerance.

However, turfgrass seedlings may exhibit halotropic growth as an additional mechanism to evade saline soil, and flavonoids may modulate this growth response. Our objectives were to first determine what wavelengths of light regulate flavonoid metabolism in the turfgrass species rough bluegrass, second to determine if the trait for flavonoid upregulation exists in natural populations of this species, and last to evaluate the impact of light and flavonoid up-regulation on halotropic growth in rough bluegrass.

My results have shown that blue wavelengths of light are responsible for regulating flavonoid synthesis in rough bluegrass, and red light can modulate flavonoid synthesis in the presence of blue light. This ability to increase flavonoid concentrations under blue light treatment, however, has only been observed in production cultivars of rough bluegrass and in accessions originating from Germany. When exposed to a salt gradient, rough bluegrass exhibits halotropic growth at NaCl concentrations greater than

350 mM. My results also show that blue light modulates halotropism; blue light increases the degree to which roots bend away from NaCl. However, exposure to red light results in the loss of halotropism. High pressure liquid chromatography (HPLC) profiles show that blue light significantly increases both flavonoid and anthocyanin content in root tissue, while white light treated roots only exhibit trace amounts of flavonoids. In red light treated roots, flavonoids are below the limit of detection, and display a secondary metabolite profile similar to pre-treatment samples.

iii

Dedication To Marie Tuvell and Mary Petrella

Acknowledgements

I would like to thank the James B. and Harriet Beard Fellowship for supporting my graduate research. Dr. Gardner, thank you for every opportunity that you have given me since day one. Thank you for supporting research that many saw as not important, thank you for always having an open door, and thanks for always having a sense of humor.

Thank you Dr. Blakeslee for helping me to think about plant science in a different way, and for helping me to take my skills to the next level. Dr. Metzger, you have helped me so much I truly don’t even know where to start to say thank you. Thank you for shifting my perspective on plant biology, and thank you for all of the advice you have given to me over the years.

To Dr. Ed Nangle, you brought me into the lab on a whim to help with some work, and now we are brothers. I hope we can get the band back together one day. To

Eun-Hyang Han and Yun Lin, thank you for allowing me to come into your lab, teaching me more than many others ever have, and for being my friends. To Jim Vent, thank you for all of the help and most importantly, all of the laughs. Arly, Matt, Pam, and of course

Dr. John Street thank you for all of your help. To Jon Szallai thank you for being my friend for the past 30 years, and for helping me stay grounded. Thank you to my sister,

Andrea, for all of her support. Finally, I would like to thank my parents Andrew and

Millie Petrella, you have backed every decision I have made, good and bad, you taught me how to be my own person, and I owe you more than I could ever pay back.

Vita

October 13, 1985...... Born, Youngstown Ohio, USA 2004...... Girard High School

2008...... BSc. Biology Pre-medical, Youngstown

State University

2011...... BSc. in Agriculture, Turfgrass Science,

The Ohio State University

2011 to present...... Graduate Research Associate, The Ohio

State University

Publications

Petrella, Dominic P., Metzger, James D., Blakeslee, Joshua J., Nangle, Edward, J., and Gardner, David S. 2016. Effects of Blue Light and Phenotype on Anthocyanin Accumulation in Accessions and Cultivars of Rough Bluegrass. Crop Science.

Petrella, Dominic P., Metzger, James D., Blakeslee, Joshua J., Nangle, Edward, J., and Gardner, David S. 2016. Anthocyanin Production Using Rough Bluegrass Treated with High Intensity Light. HortScience. 51(9) 1111-1120. 2016.

Nangle, Edward J., Gardner, David S., Metzger, James D., Rodrigues-Sanoa, L., Petrella, Dominic P., Danneberger, Tom K., and Cisar, J. 2016. Cool-Season Turfgrass Color and Growth Habit Response to Elevated Levels of Ultraviolet-B Radiation. HortScience. 51: 439-443.

Nangle, Edward J., Gardner, David S., Metzger, James D., Rodrigues-Sanoa, Luis, Giusti, Maria M., Danneberger, Tom K., and Petrella, Dominic P. 2015. Pigment Changes in Cool-Season Turfgrass in Response to Ultraviolet-B Light Irradiance. Agronomy Journal.

Fields of Study

Major: Horticulture and Crop Science

vii

Table of Contents

Abstract...... ii

Dedication...... iv

Acknowledgements...... v

Vita...... vi

List of Tables...... xiii

List of Figures...... xv

Chapter 1: Literature Review...... 1

Medicinal and Industrial Use of Flavonoids...... 1

Flavonoid/Anthocyanin Biological Production Systems...... 5

Biosynthesis and Regulation of Flavonoid Synthesis...... 8

Photoreceptors and Their Biological Roles...... 11

Salt Stress in the Turfgrass System...... 23

Salt Stress Responses...... 27

Auxin Biology and Its Modulation by Flavonoids...... 33

Summary...... 45

Reference...... 47

Chapter 2: Anthocyanin Production Using Rough Bluegrass Treated With High Intensity

Light...... 72

viii

Abstract...... 72

Introduction...... 73

Materials and Methods...... 76

Results...... 85

Discussion...... 94

Reference...... 105

Chapter 3: Effects of Blue Light and Phenotype on Anthocyanin Accumulation in

Accessions and Cultivars of Rough Bluegrass...... 111

Abstract...... 111

Introduction...... 112

Materials and Methods...... 115

Results...... 121

Discussion...... 130

Reference...... 138

Chapter 4: Modulation of Halotropic Growth in Rough Bluegrass by Blue and Red

Light...... 142

Abstract...... 142

Introduction...... 143

Materials and Methods...... 150

Results...... 162

Discussion...... 185

Reference...... 195

Chapter 5: Overall Conclusions...... 205

Abstract...... 205

Discussion...... 206

Complete Reference...... 211

Appendix A: Gravitropic Root Growth (Poa trivialis L.)...... 245

Appendix B: LED Light Strip Wiring Diagram...... 247

Appendix C: Blue LED Spectral Distribution Figure...... 249

Appendix D: Blue LED Spectral Distribution Table...... 251

Appendix E: Red LED Spectral Distribution Figure...... 253

Appendix F: Red LED Spectral Distribution Table...... 255

Appendix G: Far-Red LED Spectral Distribution Figure...... 257

Appendix H: Far-Red LED Spectral Distribution Table...... 259

Appendix I: Far-Red LED Wiring Diagram...... 261

Appendix J: Cool White/Warm White (50:50) LED Spectral Distribution Figure...... 263

Appendix K: Cool White/Warm White (50:50) LED Spectral Distribution Table...... 265

Appendix L: Combination LED Strip Light Wiring Diagram...... 267

Appendix M: Anthocyanin Accumulation in Rough Bluegrass Treated With High

Intensity Light...... 269

Appendix N: Lumigrow Blue LED Light Spectral Distribution Figure...... 271

Appendix O: Lumigrow Blue LED Light Spectral Distribution Table...... 273

Appendix P: Lumigrow Red LED Light Spectral Distribution Figure...... 275

Appendix Q: Lumigrow Red LED Light Spectral Distribution Table...... 277

Appendix R: Lumigrow White LED Light Spectral Distribution Figure...... 279

Appendix S: Lumigrow White LED Light Spectral Distribution Table...... 281

Appendix T: Natural Light Spectral Distribution Figure and Table (11/17/2016, 13:00,

Wooster OHIO, 44691, - 40.8051° N, 81.9351° W)...... 283

Appendix U: LED Light Box Design: LED Enclosure...... 285

Appendix V: LED Light Box Design: Treatment Enclosure...... 287

Appendix W: LED Light Box Design: Fan and Baffling Arrangement...... 289

Appendix X: LED Light Box Design: Completed Image...... 291

Appendix Y: Halotropism Media Preparation...... 293

Appendix Z: Halotropic Root Growth (Poa trivialis L.)...... 295

Appendix AA: 12 Hour Flavonoid and Hydroxycinammic Acid Quantification in Rough

Bluegrass...... 297

Appendix BB: 24 Hour Flavonoid and Hydroxycinammic Acid Quantification in Rough

Bluegrass...... 299

Appendix CC: Chromatograms of Anthocyanins Present in Rough Bluegrass Roots....301

Appendix DD: Example Chromatograms of Non-Anthocyanin Phenolic Metabolites

Accumulated in Rough Bluegrass Treated with 350 mM NaCl and Exposed to Blue or

Blue/Red Light for 24 Hours...... 303

Appendix EE: Example Chromatograms of Non-Anthocyanin Phenolic Metabolites

Accumulated in Rough Bluegrass Treated with 350 mM NaCl and Exposed to White or

Red Light for 24 Hours...... 305

Appendix FF: Example Chromatograms of Non-Anthocyanin Phenolic Metabolites

Present in Rough Bluegrass Prior to treatment and following Treatment with 350 mM

NaCl Under Dark Conditions...... 307

Appendix GG: UV-vis Absorption Spectra of Anthocyanin, Non-Anthocyanin Phenolic

Standards, and Phenolic Metabolites Present in Rough Bluegrass Roots...... 309

Appendix HH: Zone Specific Anthocyanin Accumulation in Roots of Rough Bluegrass

Treated with Blue light and 350 mM NaCl for 24 hours...... 311

xii

List of Tables

Table 2.1: Anthocyanin concentration (cyanidin-3-glucoside eq.) of rough bluegrass following light treatment...... 85

Table 2.2: Liquid chromatography – mass spectrometry (LC-MS) identification of anthocyanins accumulated in rough bluegrass following light treatment...... 90

Table 3.1: List of rough bluegrass accessions (Plant Introduction – PI) and cultivars used in these experiments with their country of origin...... 117

Table 3.2: Summary statistics for rough bluegrass accession and cultivar phenotypic data. Total flavonoid content, total phenolic content, specific leaf area (SLA), leaf cuticular wax (Wax), and the total chlorophyll:total carotenoid ratio

(Chl:Carot)...... 125

Table 3.4: Principle component analysis (PCA) of phenotypic trait data of accessions and cultivars of rough bluegrass that were not treated with blue light...... 129

Table 4.1: Categories for ranking data for degree of bending statistics...... 161

Table 4.2: Categories for ranking data for percent of roots bending statistics...... 161

Table 4.3: Salt concentration test. 12 hours post treatment...... 163

xiii

Table 4.4: Salt concentration test. 24 hours post treatment...... 164

Table 4.5: Salt concentration test. 48 hours post treatment...... 165

Table 4.6: 12 hour light box experiment; percent of root distribution...... 169

Table 4.7: 24 hour light box experiment; percent of root distribution...... 172

Table 4.8: Anthocyanin concentration of roots treated with 24 hours of light and 0 or

350 mM NaCl...... 174

Table 4.9: Mass spectrometry data for anthocyanins induced in rough bluegrass roots exposed to 350 mM NaCl under blue light...... 179

Table 4.10: Mass spectrometry data for non-anthocyanin phenolics induced in rough bluegrass roots exposed to 350 mM NaCl under blue light...... 183

xiv

List of Figures

Figure 1.1: Flavonoid and anthocyanin agylcones and their conjugates...... 4

Figure 1.2: The reaction of phenylalanine ammonia-lyase – PAL...... 10

Figure 1.3: Hydroxycinammic acids, alcohols, aldehydes and their conjugates...... 12

Figure 1.4: The flavonoid and anthocyanin biosynthetic pathways...... 14

Figure 1.5: Phytochrome light dependent photoconversion...... 16

Figure 1.6: The cryptochrome photocycle...... 20

Figure 1.7: The United States Golf Association (USGA) root zone and its associated perched water table...... 26

Figure 2.1: High performance liquid chromatography (HPLC) chromatograms of rough bluegrass...... 88

Figure 2.2: Chlorophyll a and b content (A), and chlorophyll a:b ratios (B) of dark and light grown rough bluegrass seedlings. Relative anthocyanin concentration of light grown (C) and dark grown (D) rough bluegrass seedlings treated with a white light

(control), blue, far-red, or red LED light...... 91

Figure 2.3: Relative anthocyanin concentration of light grown rough bluegrass seedlings treated with blue LED light of increasing intensity...... 93

Figure 2.4: Relative anthocyanin concentration of light grown seedlings (A) and mature

(3 month old) rough bluegrass (B) plants...... 95

Figure 2.5: Relative anthocyanin concentration of light grown (A) and dark grown (B) rough bluegrass seedlings treated with or without 2.8 mg L-1 amicarbazone (AMI), and exposed to increasing red light...... 96

Figure 2.6: Relative anthocyanin concentration of light grown (A) and dark grown (B) rough bluegrass seedlings treated with or without 2.8 mg L-1 amicarbazone (AMI), and exposed to increasing blue light...... 97

Figure 3.1: A 95% confidence interval separating accessions and cultivars based on total anthocyanin content (cyanidin-3-glucoside eq. - mg 100 g-1 DW)...... 122

Figure 3.2: Cluster dendrogram using Ward’s Hierarchical method to group 25 accessions and cultivars of rough bluegrass based on phenotypic traits associated with anthocyanin content...... 127

Figure 3.3: Linear regression of principle component 2 and total anthocyanin concentration following irradiance with five days of constant blue light...... 130

Figure 4.1: The degree of root bending following 12, 24. And 48 hours of treatment with various concentrations of NaCl (mM)...... 166

Figure 4.2: The concentration of salt in close proximity to the roots of rough bluegrass following placement (0 hours) onto the salt gradient plates and up to 48 hours following placement...... 168

Figure 4.3: The degree of root bending following 12 hours of treatment with 0 or 350 mM NaCl in combination with white light, blue light, red light, or the combination of blue and red light...... 171

xvi

Figure 4.4: The degree of root bending following 24 hours of treatment with 0 or 350 mM

NaCl in combination with white light, blue light, red light, or the combination of blue and red light...... 173

Figure 4.5: Quantification of flavonoids accumulated in rough bluegrass roots following

A) 12 hours of treatment and B) 24 hours of treatment...... 176

Figure 4.6: Quantification of hydroxycinammic acids accumulated in rough bluegrass roots following A) 12 hours of treatment and B) 24 hours of treatment...... 177

Figure 4.7: Chromatogram (520 nm) of anthocyanins induced in rough bluegrass roots exposed to 350 mM NaCl and blue light...... 179

Figure 4.8: Chromatograms (340 nm) of A) Standard phenolic mix, B) Luteolin 7- glucoside standard, C) Rough bluegrass non-anthocyanin phenolic leaf extract, and C)

Rough bluegrass non-anthocyanin phenolic...... 181

Figure 4.9: UV-vis absorption spectra of A) Rutin, B) Luteolin 7-glucoside, C) Peak 3 from rough bluegrass leaf tissue extract, and D) Peak 3 from rough bluegrass root tissue extract, E) Caffeic acid, and E) Peak 4 from rough bluegrass root extract...... 182

Figure 4.10: Biomass percent of control (0 mM NaCl) for rough bluegrass treated with 25 mM NaCl...... 184

xvii

Chapter 1: Literature Review

Portions of the salt stress response and auxin biology sections have been included in a

publication co-authored by Eun-Hyang and Dominic Petrella; Han et al., 2017.

“Bending” Models of Halotropism: Incorporating Protein Phosphatase 2A, ABCB

Transporters, and Auxin Metabolism. Journal of Experimental Botany.

Medicinal and Industrial Use of Flavonoids

There is increasing demand for “natural” or “organic” foods as the public views them as being healthier, more safe, and more environmentally sustainable (Siro et al.,

2008; Kraus, 2015). Along with greater demand for healthier foods, there has also been an increase in the consumption of nutraceuticals and/or functional foods, which were estimated to have a global market value of $47.6 billion dollars in the early 2000s (Siro et al., 2008). Nutraceuticals are broadly classified and include dietary supplements, functional foods themselves, and medicinal foods (Liang et al., 2015). More confusing is the concept of “dietary supplements”. The national institute of health (NIH) defines a dietary supplement as any product that is meant to supplement the diet, and the dietary supplement must contain at least vitamins, minerals, herbs botanicals compounds, or amino acids (NIH). This broad definition, in itself, allows for a wide range of food type products to be labeled as nutraceuticals; however, the use of specific plants to supplement

1 the diet, and the use of botanical extracts is known to aid in decreasing the prevalence of some diseases (He and Giusti, 2008).

One group of naturally occurring compounds used for its health promoting properties are flavonoids. Flavonoids are a diverse group of plant secondary metabolites, characterized based on whether or not they contain a ketone group, as well as other structural modifications including hydroxyl distribution (Fig. 1.1) (Winkel-Shirley, 2001;

Saito et al., 2013). Anthocyanins are a type of flavonoid classified as plant pigments due to producing red, purple, and blue coloration in plant tissues (Dooner and Robbins, 1991;

Delgado-Vargas et al., 2000). According to Castañeda-Ovando et al. (2009), 23 anthocyanin agylcones have been identified (6 being most common: cyanidin, delphinidin, malvidin pelargonidin, peonidin, petunidin), and further structural variation lies within conjugation to sugars, acyl groups, and their combinations (Fig. 1.1).

Glycosylation and acylation provide stability to both anthocyanins and flavonoids, and these modifications are also thought to increase their ability to prevent human diseases

(Castañeda-Ovando et al., 2009).

Anthocyanins and flavonoids are potent anti-oxidants, and have been shown to have anti-oxidant activity similar to or greater than vitamin E, and cyanidin 3-glucoside in particular has been shown to decrease lipid peroxidation in both blood serum and hepatic cells of rats (Tsuda et al., 2000, He and Giusti; 2008; Mauray et al., 2012). This anti-oxidant activity has been demonstrated to decrease the oxidation of low-density lipoproteins (LDL), decreasing the risk of heart disease, and consumption of both anthocyanins and flavonoids have been show to lower the risk of heart disease through

2 decreasing blood pressure (He and Giusti, 2008; Cassidy et al., 2016; Liobikas et al.,

2016). Furthermore, anthocyanin extract from purple corn (rich in both cyanidin-3- glucoside and cyanidin 3-malonylglucoside) has been shown to decrease the formation of new adipose (fat tissue) as well as hyperglycemia in mice fed a high fat diet (Tsuda et al.,

2003; Tsuda, 2008).

Anthocyanin extracts, including from purple corn, have been shown to decrease the growth of HT-29 human colon cancer cells, especially anthocyanin agylcones and monoglycosylated anthocyanins (Jing et al., 2008). Similarly, extracts from red berries, including strawberry and raspberry, have been shown to decrease the growth and proliferation of tumor lines, and results have demonstrated that this could be through modulation of the cell cycle (Seeram et al., 2006; Wang et al., 2011). Studies clearly show that anthocyanins and flavonoids positively impact human health through their antioxidant capacity as well as unknown mechanisms. However, besides their use as dietary supplements, anthocyanins and flavonoids alike have significant potential in the development of other new and emerging technologies.

Figure 1.1: Flavonoid and anthocyanin agylcones and their conjugates (modified from

Ferrer et al., 2008 4

One of the largest markets for anthocyanin extract(s) is food coloring (worth an estimated $200 million worldwide in 2002), and demand for natural colorants has been estimated to increase by 5-15% every year (Deroles, 2009). The replacement of synthetic colorants (i.e. FD&C red 40) with natural pigments like anthocyanins has been increasing, and consumers have been shown to rate food safety (food without “toxic” ingredients) as the highest priority for food quality (Kraus, 2015; Rodriguez-Amaya,

2016). Anthocyanins are also photosensitize, and are becoming increasingly used as natural dyes for solar cells (Abdou et al., 2013; Narayan, 2012). In the textile industry, approximately $23 billion dollars is spent yearly on hazardous fabric dyes.

Consequently, cost effective and environmentally friendly anthocyanin dyes have become increasingly used (Bechtold et al., 2007; Mansour et al., 2013; Hwa-Hong, 2015).

Flavonoid/Anthocyanin Bio-production Systems

The potential wide range of uses for anthocyanins is however limited by the small number of plant species they can be commercially obtained from (Deroles, 2009).

Currently, plants used for anthocyanin production include fruit and vegetable crops including: grape (Vitis spp.), blackberry (Rubus spp.), raspberry (Rubus spp.), chokeberry

(Aronia spp.), red cabbage (Brassica oleracea), and red lettuce (Latuca sativa) (Wu et al.,

2006; Rodriguez-Amaya, 2016). However, these plants lack the ability to meet current and future anthocyanin demand due to being directly consumed, exhibiting slow growth and limited yield, and many require precise growing conditions. The use of plant tissue culture has been proposed as a means for large scale anthocyanin production. However,

5 tissue culture techniques have not been able to produce anthocyanins in high concentration, production may be difficult to mechanize and scale up, and the process is costly (Callebaut et al., 1997; Delgado-Vargas 2009; Ramachandra Rao and Ravishankar,

2002; Vogelien et al., 1990; Yamamoto et al., 1982). In cell culture, it is necessary to maintain cells in an undifferentiated state to maximize metabolite production, but at the same time cell growth needs to be maintained (Deroles, 2009). Enhancing growth, to increase cell biomass, however, may lead to differentiation, resulting in the cell culture based bioprodution system losing its benefit (Deroles, 2009). Cell culture based anthocyanin production of carrot, grape, strawberry, Ajuga reptans, sweet potato, and many other plants has been evaluated but in all cases the system is limited by simple factors including nutrient and hormone supplementation (Deroles, 2009; Delgado-Vargas,

2009)

On the other hand, atypical turfgrasses uniquely possess anatomical attributes that could aid in creating a higher yielding and more sustainable anthocyanin/flavonoid biological production system. Turfgrasses are a group of perennial monocots that exhibit a crown type growth habit, and are known for a high degree of recuperative potential even when compared to other grasses (Beard, 1973). The shoot apical meristem of the turfgrass plant lies at or just beneath the soil surface compared to grasses with a culm type growth habit (a growth habit where the apical meristem grows well above the soil surface), preventing damage from aggressive defoliation, and allowing a large proportion of tissue to be removed at a given time (Younger, 1969; Stiff and Powell, 1974).

However, if the upper portion of the shoot apical meristem is damaged axillary buds will

6 become uninhibited, and increased tillering will occur (Jameson, 1964). Grasses in general also possess intercalary meristems at the base of leaf blades, at the leaf collar, or where the leaf blade becomes distinct from the leaf sheath (Younger, 1969). This meristematic region allows for rapid elongation of the leaf even following clipping.

A biological production system using turfgrasses can be described as follows: A stand (a large, dense area of turfgrass) of turfgrass is maintained under low to moderate intensity, leaf tissue is harvested (mowed) and extracted for the desired metabolite, plants back in the field are allowed to re-grow leaf tissue, and the process can be re-started once leaves are grown back to a desirable height. Many turfgrasses are known to accumulate anthocyanin content under stress, and during certain seasons. Bermudagrass (Cynodon dactylon), a C4 photosynthetic turfgrass, is known to increase anthocyanin synthesis in leaf tissue during winter or when exposed to sub-optimal temperatures, and

Bermudagrass can also be found with highly pigmented stolons (Dudeck and Murdoch,

1998). Creeping bentgrass (Agrostis stolonifera L.) notably produces anthocyanin in the late fall and early spring when day temperatures begin to decrease, but instantaneous light intensity is still relatively high (Dernoden, 1995). Anthocyanin production in creeping bentgrass is, however, highly dependent on the cultivar as well as overall light conditions

(Nangle et al., 2015). Rough bluegrass (Poa trivialis L.) is known to constitutively produce anthocyanin in the leaf sheath, and purple/red coloration can be seen in this species throughout the year (Fossen et al., 2002; Hurley, 2010). Rough bluegrass is highly intolerant to drought and high temperature, and vigorous stolon production by this species allows for survival under harsh conditions (Rutledge et al., 2010). When exposed

7 to abiotic stress, rough bluegrass will exhibit summer decline which results in leaf tissue dieback, accompanied by increased anthocyanin accumulation.

While other grasses are known for producing anthocyanin, some in high quantity

(ornamental grasses in particular), grass species other than “crown type” grasses and turfgrasses do not possess the unique to rapidly recuperate following aggressive defoliation ability, and if used in production systems many of these grasses would limit harvesting to once per season (Fossen et al., 2002; Boldt, 2014). Turfgrasses possess unique anatomy and growth patterns that would allow for a potentially high yielding biological production system. To serve as a source of anthocyanins/flavonoids, however, production of these metabolites would need to be increased to a greater degree as turfgrasses do not naturally accumulate large quantities of anthocyanin during all times of the year (Fossen et al., 2002).

Biosynthesis and Regulation of Flavonoid Synthesis

The induction of anthocyanin synthesis is regulated by a number of abiotic and biotic factors at multiple committed steps throughout the biosynthetic pathway. The pathway of both flavonoid and anthocyanin synthesis can be divided into two separate steps: 1) the phenylpropanoid pathway, and 2) the flavonoid pathway (Ferrer et al.,

2008). The first committed step in the production of phenylpropanoid compounds like flavonoids and anthocyanins is the deamination of phenylalanine (or tyrosine) producing cinnamic acid (or para-coumaric acid if tyrosine is the starting product) through the

8 enzyme phenylalanine-ammonia lyase (PAL) (or tyrosine-ammonia lyase - TAL)

(Croteau et al., 2000; Ferrer et al., 2008) (Fig 1.2).

PAL is transcriptionally regulated, it exhibits feedback inhibition by cinnamic acid, and PAL is also post-translationally regulated through ubiquination (Zhang and Liu,

2015). Transcriptional regulation of PAL has been shown to be controlled by numerous environmental factors (Rabino and Mancinelli, 1986; Chalker-Scott, 1999; Zhang and

Liu, 2015; Zhang et al., 2011). Compared to most environmental factors PAL regulation by light has been studied the most intensively, and in many cases light is required for

PAL transcription (Lamb, 1979; Mancinelli, 1985; Tobin and Silverthorne, 1985; Ma et al., 2001). The products of the phenylpropanoid pathway are not only precursors for flavonoids and anthocyanins, but also include hydroxycinammic acids and their conjugates which lead to the formation lignin (Boerjan et al., 2003) (Fig. 1.3).

The second step in anthocyanin and flavonoid production takes place through the flavonoid biosynthetic pathway. The enzyme chalcone synthase (CHS) catalyzes the first committed step in this pathway using the products from the phenylpropanoid pathway

(para-coumaric acid), and malonyl-CoA, a key intermediate in fatty acid synthesis, to produce the flavanone naringenin (Winkel-Shirley, 2001; Ferrer et al., 2008) (Fig. 1.4).

Just like PAL, CHS is also transcriptionally regulated by the environment. Both ultraviolet (UV) and blue light are considered predominate wavelengths of light that regulate CHS, and this enzyme has also been shown to be regulated by environmentally linked factors such as sucrose concentration (Tsukaya et al., 1991; Mol, 1996).

Figure 1.2: The reaction of phenylalanine ammonia-lyase – PAL (modified from Ferrer et al., 2008).

Besides CHS, another key enzyme in anthocyanin synthesis is dihydroflavonol-4- reductase (DFR). This enzyme is responsible for the reduction in the ketone group of the

C ring of dihydroflavonols: dihydrokaempferol, dihydroquercetin, and dihydromyricetin

(Winkel-Shirley, 2001) (Fig. 1.4). This enzyme performs a reversible reaction and is not considered rate limiting like CHS and PAL; however, it is responsible for the production of the “leuco” (colorless) anthocyanins (Winkel-Shirley, 2001). PAL and CHS serve as control points for limiting the downstream production of metabolites that are only needed during specific developmental stages or under stressful environmental conditions.

Most of the genes within these pathways are also transcriptionally regulated through regulatory genes or transcription factors, and flavonoid biosynthesis may also be spatially regulated through metabolon formation (Holton and Cornish, 1995; Winkel,

2004; Stommel et al., 2009; Petroni and Tonelli, 2011). The same transcription factors

(bHLH, MYB, MYC, and WD40) have been shown to control transcription of CHS,

DFR, flavanone-3-hydroxylase (FHT), and anthocyanin synthase (ANS) in certain species

(Holton and Cornish, 1995; Stommel et al., 2009; Cominelli et al., 2007; Petroni and

Tonelli, 2011) (Fig. 1.4). In general, light is a common factor that induces the production of regulatory genes that will induce the transcription of flavonoid and anthocyanin biosynthesis related enzymes, and light is required for the production of flavonoids and anthocyanins (Downs and Siegelman, 1962; Lange et al., 1971; Rabino et al., 1977;

Drumm and Mohr, 1978; Mancinelli, 1985; Mol et al., 1996; Chalker-Scott, 1999; Vyas

2014).

Photoreceptors and Their Biological Roles

In plants and animals, the perception and signal transduction of light takes place through photoreceptors that in turn can regulate transcript of downstream genes like PAL and CHS. Photoreceptors sense the amount of light (quantity) as well as specific wavelengths of light (quality) through a chromophore (the absorbing moiety of a pigment possessing a conjugated double system), and transduction of the perceived light signal takes place through a peptide associated with the chromophore as well as through alternate pathways (Galvão and Frankhauser, 2015; Li and Mathews, 2016). In plants, photoreceptors for red, far-red, and blue light are present.

Figure 1.3: Hydroxycinammic acids, alcohols, aldehydes and their conjugates (modified from Boerjan et al., 2003.

Phytochromes are photoreceptors of both red and far-red light (Galvão and

Frankhauser, 2015). The perception of blue light occurs through multiple flavoprotein photoreceptors including cryptochromes and phototropins (Christie et al., 2015; Galvão and Frankhauser, 2015). In Arabidopsis, there are five phytochrome genes (PHYA –

PHYE), whereas in grasses like rice only three phytochromes have been identified, PHYA

- PHYC (Takano et al., 2000). Phytochrome isoforms can be further classified as type 1 or II based on their abundance in dark or light grown tissue (Hughes, 2007; Pratt et al.,

1991). Type I phytochromes are abundant in dark grown tissues, are light labile (rapidly degrade under light), and are only encoded by PHYA (Kevei et al., 2007). Type II phytochromes are abundant in light grown tissues, are light stable, and are encoded by

PHYB-PHYE (Kevei et al., 2007). The phytochrome chromophore is a linear tetrapyrrole, phytochromobilin, synthesized in the chloroplast, and functional phytochrome is constructed from a dimerization of two holoproteins (Davis et al., 2001;

Galvão and Frankhauser, 2015).

Figure 1.4: The flavonoid and anthocyanin biosynthetic pathways (modified from Ferrer et al., 2008)

When first synthesized, phytochromobilin exists as a cis-isomer with a maximum absorption at 660 nm, and is termed phytochrome-red absorbing or Pr (Burgie et al.,

2014; Hughes 2007; Takano et al., 2009). Upon perception of red light, isomerization occurs at carbons 15 & 16 to produce phytochrome-far-red absorbing (Pfr) with a maximum absorption at 730 nm (Fig. 1.5) (Burgie et al., 2014; Hughes, 2007; Rockwell,

2006). In most phytochrome isoforms (i.e. PHYB), Pfr will photoconvert back to Pr when far red light is absorbed, and biological responses are correlated to the equilibrium of Pfr/P total (Chen et al., 2004; Yanovsky et al., 1997). Change in chromophore conformation leads to a subsequent change in the conformation of the apoprotein, leading to the exposure of a nuclear localization signal (NLS) on the C-terminal domain that results in Pfr transport into the nucleus (Hughes, 2007; Quail, 2002; Rockwell et al.,

2006). Phytochrome is also a serine/threonine kinase, and when in the Pfr confirmation phytochrome can phosphorylate other proteins (Rockwell et al., 2006).

In oat (Avena sativa L.), the N-terminal extension of PHYA has been shown to be autophosphorylated in vivo (Frankhasuer, 2000; Casal et al., 2002). While the function of phytochrome autophosphorylation is still unclear, multiple hypotheses have been presented (Rockwell, 2006). Autophosphorylation may lead to the exposure of the NLS leading to nuclear re-localization; and autophosphorylation is thought to release phytochrome to a cytoplasmic anchor allowing re-localization (Frankhasuer, 2000;

Rockwell, 2006). For PHYA, phosphorylated Pfr is also rapidly ubiquinated in the nucleus leading to degradation; therefore, in this case phosphorylation may serve to attenuate the signal (Schepens et al., 2004; Han et al., 2010). Phytochrome has also been

15 shown to phosphorylate other proteins (phytochrome kinase substrates (PKSs), nucleoside phosphate dikinase 2 (NPDK2), phytochrome interacting factors (PIFs), and blue light photoreceptors (cryptochromes), and this is thought to function in further downstream signaling or protein stabilization as in the case of cryptochrome

(Frankhauser, 2000).

Figure 1.5: Phytochrome light dependent photoconversion (modified from Rockwell et al., 2006)

When in the nucleus, PHYA and PHYB directly interact with nuclear proteins.

Both PHYA and PHYB bind to Phytochrome Interacting Factors (PIFs), transcription factor repressors of irradiance responses, leading to PIF degradation or decreased PIF activity (Leivar and Monte, 2014; Lee and Choi, 2017). PIFs actively promote skotomorphogenesis (etiolation; opposite of photomorphogenesis) along with shade avoidance, plants overexpressing various PIFs exhibit constitutive etiolation and shade avoidance phenotypes through promotion of auxin related responses (Casal, 2013).

Phytochromes also interact with the nuclear localized Constitutive

Photomorphogenic/De-etiolated/Fusca protein complex (COP/DET/FUS), an E3 ubiquitin ligase that promotes ubiquination of photomorphogenesis related transcription factors (Chen et al., 2004; Lau and Deng, 2012). As PHYA and PHYB accumulate in the nucleus, COP1 (Constitutive Photomorphogenic 1), specifically, is excluded from the nucleus, preventing the repression of transcription of factors that promote photomorphogenesis including: long Hypocotyl 5 (HY5), Long After Far-red 1 (LAF1), and long Hypocotyl in Far-Red 1 (HFR1) (Osterlund et al., 2000; Seo et al., 2003; Chen et al., 2004; Jang et al., 2005; Lau and Deng, 2012; Pacin et al., 2013)

The blue light photoreceptor cryptochrome perceives blue as well as ultraviolet A

(UVA) light, and is part of a conserved gene family that includes DNA photolyases that are found in animals, bacteria, and fungi (Chaves et al., 2011; Galvão and Frankhauser,

2015). Rather than repair DNA following light absorption, plant cryptochromes function in environmental sensing and signal transduction (Chaves et al., 2011; Galvão and

Frankhauser, 2015). In Arabidopsis, three cryptochrome genes have been identified,

CRY1-3 (Ahmad et al., 1998; Chaves et al., 2011; Galvão and Frankhauser, 2015). In rice, three cryptochrome genes have been identified, OsCRY1a, OsCRY1b, and OsCRY2

(Hirose et al., 2006). CRY1 and CRY2 have many overlapping physiological functions; however, CRY2 rapidly degrades under light while CRY1 is stable (Chaves et al., 2011;

Xu et al., 2009).

Cryptochromes consists of two chromophores, a methenyltetrahydrofolate

(MTHF) molecule, and a flavin adenine dinucleotide (FAD) that binds non-covalently to the apoprotein (Lin et al., 2003; Liu et al., 2011). Reduction of MTHF through photon absorption does not result in conformational change in the cryptochrome protein, indicating that MTHF is not the primary chromophore of cryptochrome (Chaves et al.,

2011). The active cryptochrome chromophore, FAD, is fully oxidized under dark conditions, absorbs maximally at 450 nm, and cryptochrome signaling is dependent on the redox state of FAD (Fig. 1.6) (Bouly et al., 2007; Conrad et al., 2014; Liu et al.,

2011). When FAD absorbs a single photon, one nitrogen is reduced, producing semi- reduced FADH˙, resulting in auto-phosphorylation (through a bound ATP) and cryptochrome dependent signaling (Fig. 1.6) (Chaves et al., 2011; Conrad et al., 2014;

Liu et al., 2011). Semi-reduced FADH˙ is short lived, and is either oxidized to attenuate signaling, or fully reduced (FADH2) by green light resulting in inhibition of blue light signal transduction (Liu et al., 2011). In contrast to phytochromes, CRY1 and CRY2 are nuclear localized; however, CRY1 is transported to the cytosol when exposed to blue light; whereas CRY2 is nuclear localized irrespective of light conditions (Lin et al., 2003;

Xu et al., 2009). On the other hand, OsCRY1 is localized to both the nucleus and the

18 cytosol regardless of ambient light conditions (Xu et al., 2009). Cryptochrome function is also dependent on dimerization, similar to phytochrome, but cryptochromes lack kinase activity (Lin et al., 2003; Liu et al., 2011).

Similar to phytochrome nuclear signaling, cryptochromes also interact with the

COP/DET/FUS E3 ubiquitin ligase complex, COP1 specifically, promoting photomorphogenesis through transcription factors like HY5 and HYH (HY5 Homolog), which is thought to be more specific for blue light (Holm et al., 2002; Lau and Deng,

2012). Cytoplasmic signaling may also occur through the cryptochrome photocycle (Fig.

1.6). Reactive oxygen species (ROS) are produced in the presence of oxygen as FADH- is re-oxidized to FAD in dark, producing H2O2 (Muller and Ahamd, 2011). This process is in competition with FAD reduction reactions in the light, and therefore the ROS produced by cryptochrome may serve as signaling molecules (Miao et al., 2006;

Maathuis, 2014; Ahmad, 2016).

The second most studied blue light photoreceptors are the phototropins (Galvão and Frankhauser, 2015). The primary function of phototropins is to optimize photosynthesis under the given light environment through tropic bending, chloroplast re- distribution, and stomatal control (Christie, 2007; Liscum et al., 2014). Two phototropin genes have been identified in Arabidopsis and rice, PHOT1 and PHOT2 (Christie, 2007;

Jain et al., 2006; Liscum et al., 2014). Both PHOT1 and PHOT 2 function under low light conditions, while only PHOT2 is functional under high light intensity (Christie,

2007). Compared to phytochromes and cryptochromes, phototropins are localized to the plasma membrane, and lack nuclear localization (Chen et al., 2004; Christie, 2007). The

19 apoprotein consists of two light-oxygen-voltage (LOV) domains that each non-covalently bind one flavin mononucleotide (FMN), and a serine/threonine protein kinase domain

(PKD) necessary for signaling (Christie, 2007; Liscum et al., 2014).

Figure 1.6: The cryptochrome photocycle (modified from Conrad et al., 2014)

FMN shows a similar signaling photocycle compared to FAD, and absorbs maximally at 447 nm; however, following blue light absorption, FMN covalently binds a conserved cysteine residue on the LOV domain (Christie, 2007). The LOV2 domain has been shown to be required for signaling, whereas the importance for LOV1 is still unknown (Christie, 2007). Following FMN photoactivation of phototropin, interaction between LOV2 and the PKD leads to autophosphorylation, and transport of phototropin into the cytosol where it localized to the Golgi apparatus (Christie, 2007; Liscum et al.,

2014). Phototropins primarily signal through interaction and phosphorylation of membrane proteins, transporters in particular, resulting in ion flux and changes in phytohormone transport. PHOT1 has also been shown to phosphorylate plasma membrane H+-ATPase, ATP binding cassette B-19 (ABCB19) proteins that mediate auxin efflux, and the auxin efflux protein PIN-Formed 3 (PIN3) in roots (Christie et al.,

2015; Spalding, 2013; Zhang et al., 2013). These photoreceptors not only respond to specific wavelength(s), but they also show distinct responses dependent on photon flux and length of exposure.

Physiological responses associated with these photoreceptors are separated into three distinct categories: very low fluence responses (VLFR), low fluence responses

(LFR), and high irradiance responses (HIR). Very low fluence responses saturate at very low photon flux, .001 - 1 µmol m-2 s-1, are mediated by Pr, and are not photoreversible

(Chen et al., 2004; Smith and Whitelam, 1990). These responses are associated with

PHYA (Type I phytochrome), and are mediated by Pfr/Ptotal ratios, below 0.1%, caused by limited red light in vegetative shade situations (Casal et al., 1997). VLFRs are

21 exhibited in seedlings that may germinate below the soil, or in plants that germinate within a pre-existing dense canopy (Casal et al., 1997).

LFRs are regulated by PHYB, and signaling occurs through Pfr, and a high

Pfr/Ptotal (Casal, 2013). The absorption spectra of phytochrome shows that Pfr also absorbs red light; therefore, Pfr/Ptotal can never be 100%, and maximum Pfr is approximately 88% (Butler et al., 1964; Roux et al., 2010). PHYB associated responses include de-etiolation, and, most importantly, shade avoidance (Casal et al., 2013; Casal et al., 1997; Chen et al., 2004). Cryptochromes and phototropins are also actively involved in low fluence responses; however, blue light responses are associated to a greater degree with HIRs (Mancinelli and Rabino, 1978).

HIRs require high light intensity, are dependent on the fluence rate, and are not characterized by red far-red reversibility (Smith and Whitelam, 1990). PHYA is known to act in HIRs with far-red light given over a prolonged period of time or in intermittent pulses over a 24 hour period, and the FR-HIR response is notably associated with de- etiolation of seedlings enriched in PHYA as well as during flowering of long day plants

(Shinomura et al., 2000). However, one of the most studied HIRs is associated with blue light and the production of flavonoids and anthocyanins.

Blue and red light HIRs are known to regulate flavonoid and anthocyanin synthesis; for example, cry1 knockout mutants lack anthocyanin accumulation and overexpression of cry1 has been shown to increase anthocyanin accumulation (Lin, 2002;

Chatterjee et al., 2006). Phenylalanine ammonia-lyase (PAL), the first committed enzymatic reaction in phenylpropanoid biosynthesis is transcriptionally regulated by both

22 red and blue light (Croteau et al., 2000; Delgado-Vargas et al., 2000; Mol et al., 1996;

Rabino and Mancinelli, 1986; Zhang and Liu, 2015). Chalcone synthase, the first committed enzymatic reaction in flavonoid synthesis, is however known to be regulated to a greater degree by both UV and blue light rather than red light (Mol et al., 1996;

Croteau et al. 2000; Wade et al., 2001). Some plants may increase anthocyanin content under red light, while others exhibit greater anthocyanin content under blue light irradiance, and the application of red light in combination with blue light has also been shown to increase anthocyanin content relative to blue light by itself, even when red light does not exhibit a response (Kerchoffs and Kendrick, 1997; Mancinelli, 1985; Neff and

Chory, 1998; Wade et al., 2001; Oh et al., 2014;).

Flavonoids and anthocyanins are regulated by environmental conditions other than light, and greater concentrations of these phenolic metabolites can be observed in turfgrasses under low temperature and drought stress. This suggests that flavonoids may have specific functions in stressed plant tissues (Chalker-Scott, 1999). Interestingly, flavonoids and anthocyanins have been shown to increase during salt stress, and flavonoids are known to modulate hormone metabolism and transport (Murphy et al.,

2000; Eryilmaz, 2006).

Salt Stress in the Turfgrass System

Turfgrasses have been estimated to cover approximately 50 million acres in the

United States, and are presumably the largest irrigated crop in the country (Breuninger et al., 2013; Milesi et al., 2005). Turfgrasses, however, are also economically important

23 crops, and for many states they produce more income than other agricultural commodities

(Breuninger et al., 2013). Turfgrasses have been shown to decrease stress and anxiety through the promotion of increased physical activity, and through greater person-to- person interaction during sporting events (Tyrvainen et al., 2014). Playing sports, like soccer, has also shown to be decrease child hood obesity and improve cardio-respiratory function (Seabra et al., 2016).

Greater demands for fresh water (water containing ≤ 1000 PPM total dissolved solids or ≤ 0.05% dissolved salts), however, are limiting the use of supplemental irrigation for turfgrasses (American Meteorological Society, 2012). Because of this, there is increased use of re-claimed wastewater for irrigation, or simply the physical decrease in irrigation. The application of re-claimed water, which often contains NaCl concentrations of 30 mM or higher, to irrigate turfgrasses will increase soluble salts in the turfgrass root zone, and reductions in irrigation will also lead to the aggregation of salts currently in the soil (Hayes et al., 1990; Harivandi 2004; Mancino and Pepper, 1992).

Climate change has currently, and is predicted, to further increase drought and salt stress.

For example, climate change is projected to cause prolonged increases in extreme temperatures, especially increases in surface heating, and greater occurrences of sporadic, yet intense, rainfall events leading to increased risk of drought and/or salinity stress

(Meehl and Stocker, 2007).

While water availability and precipitation may be difficult to model, it is well- accepted that increases in evapotranspiration (ET), due to rising temperatures, as well as alterations in precipitation patterns will increase soil drying and salt aggregation (Meehl

24 and Stocker, 2007; Trenberth et al., 2013). Additionally, changes in global temperatures have increased and are predicted to further increase the use of fresh water, and result in increasingly limited irrigation (Meehl and Stocker, 2007; MacDonald, 2010; Sabo et al.,

2010; Trenberth et al., 2013). Exposure to high concentrations of NaCl can cause considerable amounts stress for turfrgasses, resulting in decreased recovery, reductions in turfgrass quality, and potential economic losses. Unfortunately, the turfgrass species used for sports, golf, or home lawns are already not very tolerant to NaCl stress

(Harivandi et al., 1992; Munns and Tester, 2008).

For golf courses in particular, where turfrgasses are maintained with a minimal root system due to constant mowing, greater concentrations of salt in the soil can result in turfgrass decline. Many times, the root zone of a golf course green is constructed to meet specific requirements for playability. The most common construction method follows the

United States Golf Association (USGA) method (USGA Staff, 2004). With this method, the rootzone consists of a 400 mm subgrade, 100 mm of gravel, and is topped off with

300 mm of a sand based root zone mix (Fig. 1.7). This design results in a firm surface that allows for adequate drainage, and benefits a well stuck golf shot. The standard

USGA golf green confers a perched water table due to the difference in pore size between the gravel layer and the root zone (Hummel, 1993; Schlossberg and Karnok, 2002) (Fig.

1.7).

The perched water table, which has been shown to exist up to six days following watering, allows for water to move back up towards the root system by capillary action, thereby providing water when supplemental irrigation is withheld (Schlossberg and

Karnok, 2002; McCoy and McCoy, 2009). In the summer months, golf greens are irrigated “deep-and-infrequent,” meaning greens are heavily irrigated approximately once per week. This helps to decrease disease occurrence and prevent over-watering soils that exhibit increased temperature. Under these circumstances, the perched water table helps to provide water to the roots when irrigation is not regularly applied. However, salt stress can be an issue when irrigating in this fashion by A) increasing salt aggregation in the drying soil and B) by creating a gradient of soluble salt originating from the perched water table (Fig. 1.7).

Figure 1.7: The United States Golf Association (USGA) root zone and its associated perched water table (modified from USGA, 2004).

NaCl and other minerals (iron in particular) have been shown to accumulate in perched water tables (Ben-Hur et al., 2001; Obear et al., 2013). Soluble salts, like NaCl, in the perched water have also been shown to move closer to roots by capillary action, and as water rises closer to the surface, roots will be exposed to an increasing concentrations of NaCl (Ben-Hur et al., 2001). This can be overcome through the application of large volumes of irrigation to flush the salt out of the soil; however, as fresh irrigation water is not available in some parts of U.S. for turfgrass irrigation, and because fresh water is becoming more restricted overall this practice may be limited. The

NaCl gradient that can be found in the turfgrass root zone may cause fluctuating levels of salt along with regions of various salt concentrations. Responses to these conditions include cellular trafficking and transport of NaCl, and alterations in root architecture that decrease salt uptake.

Salt Stress Responses

Salinity stress from sodium chloride (NaCl) results in both osmotic and ionic stresses in plants which are sensed through different pathways (Blumwald, 2000; Munns and Tester, 2008). Perception of NaCl takes place at the plasma membrane as well as in the cytosol, however true sensors of Na+ have yet to be identified (Deinlein et al., 2014;

Henderson and Gilliham, 2015; Maathuis, 2014). Loss in turgor from cellular desiccation due to salt stress results in plasma membrane retraction from the cell wall (plasmolysis), and results in mechanical sensation of osmotic stress specifically (Christmann et al.,

2013; Hamant et al., 2008; Maathuis, 2014). Shrink-stretch forces on the plasma

27 membrane caused by fluctuation in turgor may be perceived through constraint or relaxation of microtubules, and plasmolysis may lead to the activation of MAPK phosphorylation cascades (Chefdor et al., 2006; Hamant et al., 2008; Laurie et al., 2002;

Maathuis, 2014). Plasma membrane plasmolysis may also cause altered protein-protein interaction, or the production of lipid signaling molecules through the action of phospholipase enzymes (Christmann et al., 2013; Haswell et al., 2011; Kung, 2005).

Sensation of ion accumulation is less clear; however, increases in Na+ may be perceived at the plasma membrane by transport proteins, perception could take place through rapid shifts in membrane potential, or cytosolic perception may occur through rapid changes in the Na+: K+ ratio (Munns and Tester, 2008; Yeo, 1998).

The osmotic stress generated by increased levels of NaCl in the soil results from changes in the osmotic potential in both the soil and apoplast, and is considered the first phase of salt stress. Exposure to this osmotic stress decreases the ability of roots to absorb water, and is therefore similar in several aspects to drought stress (Munns and

Termaat, 1986; Munns, 1993; 2002; Tester and Davenport, 2003). As salinity stress progresses, NaCl begins to accumulate in root and shoot tissues, causing additional osmotic stress and resulting in loss of turgor pressure and rapid accumulation of Na+ in the cytosol (Munns and Termaat, 1986; Munns, 1993; 2002; Tester and Davenport,

2003). Accumulations of Na+ and Cl- in the cytosol are considered the second stage of salt stress (Munns and Termaat, 1986; Munns, 1993; 2002; Tester and Davenport, 2003).

The cellular uptake of Na+ occurs passively through non-selective cation channels, as well as via uptake by members of the Na+/K+-specific high affinity K+ transporter (HKT)

28 family of plasma membrane carriers (Gassmann et al., 1996; Uozumi et al., 2000; Laurie et al., 2002; Haro et al., 2005; Munns and Tester, 2008).

Uptake of Na+ ions results in a rapid depolarization of the plasma membrane, followed by a sharp increase in the Na+: K+ ratio driven by the passive efflux of K+ to restore membrane potential, and decreases in the rate of K+ influx (Blumwald, 2000;

Shabala and Cuin, 2007). The increased Na+: K+ ratio generated by the symplastic accumulation of Na+ initiates the production of reactive oxygen species (ROS), resulting in enzyme inhibition, reductions in stomatal conductance, and decreases in photosynthesis (Maathuis and Antmann, 1999; James et al., 2002; Tester and Davenport,

2003). Decreasing ROS has been shown to increase NaCl in turfgrasses. The expression of antioxidant enzymes [Superoxide Dismutase (SOD), Catalase (CAT), Ascorbate

Peroxidase (APX), etc.] has been shown to increase in both C3 and C4 turfgrasses when exposed to 200 and 400 mM NaCl, and higher antioxidant enzyme activity is known to increase NaCl tolerance (Hu et al., 2011; Hu et al., 2012a).

Physiological responses to NaCl stress fall into three categories: 1) removal of

NaCl from the cytosol through vacuolar sequestration or transport into the apoplast; 2) exclusion of NaCl from shoots and roots; and 3) NaCl avoidance through root growth away from regions of high salt concentrations. Of these mechanisms, the sequestration of

NaCl in the vacuole and the transport of Na+ ions into the apoplast have been the most studied in recent years. Na+ ions can be transported directly into the vacuole for sequestration by Na+/H+ Exchanger 1 (NHX1), a passive tonoplast localized Na+/H+ antiporter which exhibits increased expression in the presence of either salt or abscisic

29 acid (ABA) (Apse et al., 1999; Shi and Zhu, 2002; Apse et al., 2003; Zhu, 2003).

Interestingly, under non-saline conditions both NHX1 and the related transporter NHX2

(also localized to the vacuole) have been shown to play a role in maintaining both K+ homeostasis and cytosolic pH, suggesting that both of these transporters serve a dual purpose, and that their Na+ transport activity is only activated during times of ionic imbalance (Leidi et al., 2010; Bassil et al., 2011; Barragan et al., 2012). NHX1 homologs have also been identified in turfgrasses. Zoysiagrass (Zoysia japonica L.) ZjNHX1 was shown to exhibit greater expression during salt stress, (Du et al, 2009). Treatment of the

C3 turfgrass red fescue (Festuca rubra ssp. litoralis L.) with 500 mM NaCl was shown to increase expression of FrNHX1 in endodermal and vascular tissues, while a vacuolar

ATPase (FrVHA-B) was also shown to exhibit greater levels of expression (Diedhiou et al., 2009).

Like in red fescue, proton gradients are important for Na+ sequestration, and it has been demonstrated that the overexpression of the vacuolar H+ pyrophosphatase AVP1, which pumps protons into the vacuole, increases the salt tolerance of multiple species

(Gaxiola et al., 2001; Bao et al., 2009; Schilling et al., 2014). This phenomenon is thought to be the result of increased NHX1-facilitated NaCl sequestration into the vacuole resulting from the increased proton gradient generated across the vacuolar membrane by AVP1 overexpression (Gaxiola et al., 2001; Bao et al., 2009; Schilling et al., 2014). In addition to direct transport into the vacuole by NHX-type transporters, Na+ ions can be sequestered in the vacuole through a vesicular trafficking pathway. The

NHX5 and NHX6 Na+/H+ (also K+/H+) antiporters located on vesicles found in the Golgi

30 and trans-Golgi network can mediate the influx of Na+ ions into vesicles moving through the Golgi network. As the NHX5- and NHX6-containing vesicles are transported to, and fuse with the vacuole, accumulated Na+ is released into the vacuole (Bassil et al., 2011).

In addition to sequestration in the vacuole, Na+ can also be removed from the cytosol by transport across the plasma membrane and into the apoplast, a process mediated by the SOS (Salt-Overly-Sensitive) family of proteins (Liu and Zhu, 1998; Zhu,

2002; Qiu et al., 2003; Julkowska and Testerink, 2015). Salt stress induces the rapid release of calcium stores into the cytosol, where they interact with SOS3 (Liu and Zhu,

1998). Once bound to calcium, SOS3 then interacts with and activates the SOS2 serine/threonine protein kinase, resulting in increased transcription and activation of the

Na+/H+ antiporter SOS1 at the plasma membrane and, ultimately, increased removal of

Na+ into the apoplast (Liu and Zhu, 1998; Zhu, 2002; Qiu et al., 2003). Arabidopsis genes in the SOS pathway have been expressed in tall fescue (Festuca arundinacea L.), and results showed that overexpressing lines reduced Na+ accumulation compared to wild-type plants (Ma et. al., 2014).

Both the NHX and SOS1 transporters function to increase salt tolerance by reducing the cytosolic concentration of Na+. Plants can decrease salt concentrations on a larger scale by altering root architecture to minimize exposure to saline conditions (i.e., adaptive growth). Most of these salt stress responses rely, at least in part, on salt stress- induced alterations to phytohormone metabolism and transport. For example, during salt stress, abscisic acid (ABA) functions to alter root architecture, upregulate expression of stress-related genes, promote stomatal closure, and decrease shoot growth (Grill and

Himmelbach, 1998; Xiong et al., 2002; Duan et al., 2013; Julkowska and Testerink,

2015). In Arabidopsis, ABA synthesis was increased in root cortical and endodermal cells during salt stress, and ABA signaling has been shown to result in the suppression of lateral root initiation, but not primary root elongation, during salt stress (Duan et al.,

2013; Ruiz-Sola et al., 2014). The role of ABA in altering root architecture is, however, highly dependent upon the species, the amount of salt present, and the developmental stage of the root tissue exposed to saline conditions (i.e. age and region of the root exposed to salt) (Duan et al., 2013; Geng et al., 2013). For example, in Arabidopsis,

ABA may function to maintain lateral root development under low salt concentrations, but inhibit lateral root development at higher salt concentrations (Zolla et al., 2010;

McLoughlin et al., 2012). Consistent with this, ABA-dependent decreases in lateral root length, which decrease the overall surface area of the root exposed to saline conditions, have been shown to positively correlate with lower rates of NaCl uptake in Arabidopsis roots, even under conditions where the total number of lateral roots was unaffected

(Julkowska et al., 2014). In rice, however, reduced lateral root number has been shown to be positively correlated to increased salt tolerance (Faiyue et al., 2010).

The phytohormone auxin is also a key regulator of root architecture and development during salt stress. Modulation of auxin metabolism and transport are essential components in root adaptive growth responses, particularly the initiation of lateral roots and tropic growth. Halotropism is a recently defined tropic response in which, upon exposure to a salt gradient, roots exhibit directional growth away from areas of increased salt concentration (Galvan-Ampudia et al., 2013; Rosquete and Kleine-

Vehn, 2013). This tropic growth is dependent on the asymmetrical redistribution of auxin from the side of the root nearest to the salt to the side of the root opposite the salt, resulting in the growth of the root tip away from the area of high salt concentration.

(Galvan-Ampudia et al., 2013; Rosquete and Kleine-Vehn, 2013).

Auxin Biology and Its Modulation by Flavonoids

Long term changes in architecture that lead to plant adaption are dependent on changes in auxin metabolism (biosynthesis, storage, and degradation) and transport.

While in the early stages of plant development auxin can be produced in many cells and tissues, throughout most of the plant life-cycle, however, the majority of auxin synthesis has been shown to occur in root and shoot meristems, expanding young leaves, and root tissues (Ljung et al., 2001; Ljung et al., 2002; Ljung, 2013). Biosynthesis of the primary auxin present in most plants, indole-3-acetic acid (IAA), has been proposed to take place through two major routes; the tryptophan (Trp)-independent pathway(s) and the Trp- dependent pathways (Soeno et al., 2010; Ljung, 2013; Spiess et al., 2014). Both inhibitor studies and reverse genetic investigations using auxin synthesis mutants have supported the hypothesis that the Trp-dependent pathways are the primary IAA biosynthesis pathways in most plants; and many of the genes encoding the biosynthetic enzymes within these pathways have been identified (Zhao et al., 2001; Zhao et al., 2002;

Stepanova et al., 2008; Tao et al., 2008; Nemoto et al., 2009; Mano and Nemoto, 2012;

Ljung, 2013).

The biosynthesis of indole-3-acetic acid is environmentally regulated at the transcriptional level, as well as through regulation of the subcellular localization of the biosynthetic enzymes (Kriechbaumer et al., 2012; Kriechbaumer et al., 2016;

Kriechbaumer et al., 2015). Additionally, carbon flux and high temperature have been shown to regulate IAA synthesis through the action of phytochrome interacting factors

(PIFs), which also function in red light-dependent, phytochrome-mediated regulation of

IAA synthesis (Franklin et al., 2011; Hornitschek et al., 2012; Lilley et al., 2012;

Sairanen et al., 2012). Auxin responses are not only controlled through biosynthesis, but are also finely tuned through IAA degradation and conjugation, which function to diffuse the auxin signal.

The most abundant IAA degradation products are 2-oxindole-3-acetic acid

(oxIAA) and its associated glucose or hexose conjugates (Normanly, 2010; Peer et al.,

2013). The exact mechanisms responsible for IAA oxidation are still not entirely defined, but evidence indicates that it occurs both enzymatically and/or through oxidative decarboxylation (Beffa et al., 1990; Normanly, 2010; Zhang et al., 2016). Oxidative decarboxylation is thought to occur following localized, transient, accumulations of auxin, which generally result in the rapid production of reactive oxygen species (ROS) at the site of auxin aggregation. ROS are thought to cause the oxidative decarboxylation of indole-3-acetic acid into oxIAA; and accumulations of oxIAA in plant tissues are commonly observed following auxin-induced oxidative bursts, a process that is inhibited by the presence of anti-oxidants, such as flavonoids, in these tissues (Ostin et al., 1998;

Peer and Murphy, 2007; Peer et al., 2013).

Flavonoids modulate auxin metabolism. For example, in the Arabidopsis flavonoid over-accumulating (tt3) and flavonoid-deficient (tt4) mutants, it has been shown that anti-oxidant flavonoids (aglycone flavonols in particular) can mitigate IAA oxidation by ROS, and decrease oxIAA production (Peer et al., 2013). The synthesis of flavonoids has also been linked to greater tolerance to salt stress in both Arabidopsis, soybean, and some halophytic plants (Van Oosten et al., 2013; Yan et al., 2014; Ismail et al., 2016). oxIAA can also be generated through enzymatic activity. An enzyme responsible for the conversion of IAA to oxIAA, dioxygenase for auxin oxidation (DAO; a 2-oxoglutarate-dependent-Fe (II) dioxygenase), has been recently identified in rice, and the ability of this enzyme to convert IAA to oxIAA was confirmed both in vitro and in a heterologous E. coli expression system (Zhao et al., 2013).

Additionally, the Arabidopsis DAO1 ortholog (dioxygenase for auxin oxidation 1;

DAO1) has also been demonstrated to catalyze the synthesis of oxIAA from IAA (Zhang et al., 2016). The Arabidopsis dao1 mutant exhibited a 95% reduction in IAA levels and had larger cotyledons, greater lateral root density, and elongated pistils compared to wild- type plants (Zhang et al., 2016). The dao1 mutant also showed alternations in developmental patterns, including delayed sepal opening, and reduced fertility of the primary inflorescence (Zhang et al., 2016). In addition to conversion into oxIAA, auxin signals can be attenuated by conjugating IAA into forms that allow for inactive storage, or lead to its degradation (Ludwig-Muller, 2011; Korasick et al., 2013).

Ester-linked sugar conjugates to IAA in the form of glucose conjugation (IAA-

Glc) lead to degradation, but in maize this conjugate has been shown to be hydrolyzed

35 back into IAA, suggesting that it may serve as a storage form in certain situations

(Jakubowska and Kowalczyk, 2005; Korasick et al., 2013; Ludwig-Muller, 2011).

Members of the Gretchen Hagen 3 (GH3) group of acyl amido synthetase enzymes, first identified in soybean, conjugate amino acids (Ala, Leu, Asp, Glu, and Trp) to free IAA

(Hagen et al., 1984; Korasick et al., 2013; Normanly, 2010). Both IAA-Ala and IAA-Leu are thought to be storage forms off IAA that can readily hydrolyzed back into the active form via members of the IAA-Leucine resistant 1 (ILR1) and members of the IAA-

Alanine resistant 3 (IAR3) family of amidohydrolases (Bartel and Fink, 1995; Davies et al., 1999; Rampey et al., 2004). Conjugation to Asp (IAA-Asp) or Glu (IAA-Glu) results in permanent inactivation, and these conjugates are destined for degradation (Ostin et al.,

1998; Rampey et al., 2004; Staswick et al., 2005). IAA-Trp conjugates have been shown to antagonize auxin responses, and suppress IAA inhibition of root growth as well as inhibit the lateral root growth stimulated by IAA (Staswick, 2009).

IAA conjugation is transcriptionally regulated, and is also regulated through enzyme compartmentalization, by auxin quantities, and by abiotic stress (Ludwig-Muller,

2011). GH3-5, in particular, exhibits rapidly increased expression following treatment with exogenous IAA as well as ABA, but not by any other phytohormones, including GA or Me-JA (Park et al., 2007). Similarly, GH3-5 increases expression under abiotic stress including: cold, heat, drought, and salt stress (Park et al., 2007). The GH3-5 overexpressing Arabidopsis line, wes1-D, also shows greater tolerance to high salt conditions as well as high temperature (Park et al., 2007).

In the salt tolerant desert poplar, Populus euphratica, IAA conjugates have been shown to increase by 29% under salt stress relative to the salt intolerant grey poplar, P. x canescens, which exhibited a 55% decrease in IAA conjugates (Junghans et al., 2006). P. x canescens was also shown to have an 80% loss in free IAA under salt stress, while P. euphratica only exhibited a 37% decrease; this difference in IAA content showing the importance of auxin homeostasis under salt stress by maintaining auxin pools through conjugation (Junghans et al., 2006). Previous gene expression analysis of P. euphratica under high salt concentrations revealed greater expression of an auxin amidohydrolase

(ILL3), again suggesting the importance of having available free IAA under salt stress through maintaining auxin storage pools (Brosche et al., 2005; Junghans et al., 2006).

However, when comparing expression of ILL3 from both P. euphratica and P. x canescens under salt stress, results showed that there was no significant difference in expression between the salt tolerant and in tolerant poplar species; however, when overexpressed in Arabidopsis, ILL3 increased salt tolerance through maintained root elongation (Junghans et al., 2006).

Although many tissues and cell types are capable of producing indole-3-acetic acid, under most growing conditions, however, the majority of IAA production is limited to the root and shoot apical meristems, as well as young tissues at early stages of development (Ljung et al., 2001; Peer et al., 2011). The distribution of IAA regulates organ initiation and development, and non-uniform re-distribution of IAA in response to stressful conditions leads to changes in growth that allow for escape and/or adaptations to new conditions (Blakeslee et al., 2005; Zazimalova et al., 2010; Peer et al., 2011). Long

37 distance auxin transport can take place from source to sink by means of bulk flow through phloem sieve tubes, which is dependent on sucrose production as well as sink strength (Baker, 2000; De Schepper et al., 2013). Auxin can also be transported both short and long distances by cell-to-cell movement through chemiosmotic H+ gradients and through membrane transporters that may be either localized in polar or apolar fashions (Blakeslee et al., 2005; Peer et al., 2011).

Indole-3-acetic acid is a weak acid (pKa = 4.75), and is found in a protonated, non-polar state, within the apoplast due to the acidic pH (pH 5.0 – 6.0) generated by plasma membrane ATPase activity (Lomax et al., 1985). The non-polar nature and multi- ringed structure of protonated IAA allow it to freely diffuse through the plant cell plasma membrane into the cytosol, where in the neutral pH of the cytoplasm, IAA is de- protonated and effectively “trapped” in the cell, as the ionic form of IAA can no longer diffuse through the cell membrane and can therefore only exit the cytosol through efflux proteins (Luschnig et al., 1998; Blakeslee et al., 2005). In tissues where IAA concentrations are high, or where IAA may exist in the apoplast in both protonated and de-protonated states, IAA influx can be supplemented through protein driven uptake via the AUXIN RESISTANT1/LIKE AUX1 (AUX1/LAX) family of H+ symporters (Lomax et al., 1985; Swarup et al., 2004). AUX1/LAX transporters are actively involved in regulating root growth and development, and are required for root gravitropism, with

Arabidopsis aux1 and lax3 mutants exhibiting loss of gravitropism and reduced lateral root initiation (Appendix A) (Swarup et al., 2004; Swarup et al., 2008). Interestingly, changes in AUX1 patterning may also impact (or increase) stress tolerance; and in rice

AUX1 helps to maintain root growth under heavy metal stress, particularly cadmium

(Cd) stress (Yu et al., 2015). Further, in Arabidopsis, both expression of AUX1 transcripts and the levels of AUX1 protein present in roots have recently been shown to increase following NaCl stress (Galvan-Ampudia et al., 2013; van den Berg et al., 2016).

Modeling of root tissue during salt stress responses has further indicated the importance of AUX1-mediated re-distribution of auxin at the root tip in regulating changes in root architecture (van den Berg et al., 2016). In order to maintain root growth and development under both normal and stressed conditions, however, it is essential to balance influx of auxin into plant cells with transporter-mediated auxin efflux.

Auxin efflux is mediated primarily by two classes of proteins, ATP-binding cassette type B/P-glycoprotein (ABCB/PGP) transporters and PINFORMED (PIN) transporters (Blakeslee et al., 2005). PIN efflux proteins were the first documented IAA efflux carrier proteins, and have been shown to exist in all plant species, with the

Arabidopsis genome containing eight PIN, and the rice genome encoding at least twelve, with three (PIN9, PIN10a, PIN10b) being monocot specific (Wang et al., 2009a;

Zazimalova et al., 2010; Peer et al., 2011). PIN proteins are generally polarly localized and mediate directional auxin transport in the cells and tissues where they are expressed.

For example, in Arabidopsis seedlings, PIN1 proteins are polarly localized on the rootward side of the plasma membrane in both cells of the xylem parenchyma and adjacent cortical cells, where they are responsible for long distance IAA transport from shoots to roots, as well as vascular tissue development (Bennett et al., 1995; Galweiler et al., 1998; Peer et al., 2004; Blilou et al., 2005). Salt stress has been demonstrated to

39 decrease auxin levels in root tissues (Sun et al., 2008), and recent evidence has suggested that PIN1 may play a role in this process, and that loss of PIN1 may increase salt tolerance by altering root meristem cell size and number (Liu et al., 2015).

PIN proteins play an essential role in maintaining root growth and root tip architecture, and the co-ordinated activity of multiple PIN proteins (PIN1, PIN2, PIN3,

PIN4, and PIN7) has been hypothesized to generate a “reverse fountain” reflux loop in which auxin delivered from the shoot apical meristem moves rootward into the root tip/apical meristem (Blilou et al., 2005). Shoot-derived auxin, along with auxin produced in the apical meristem is then moved in a shootward direction through root cortical and epidermal cells, until it reaches the distal elongation zone, where it is routed back into the central rootward bound auxin transport stream (Blakeslee et al., 2005; Krecek et al.,

2009; Peer et al., 2011). PIN2 plays a particularly important role in this process, mediating the shootward transport of auxin in root tips (Krecek et al., 2009). PIN2 is localized on the shootward (upper, “apical”) side of epidermal and root cap cells, and functions to re-distribute auxin in a shootward direction tissue, promoting root gravitropism and allowing for the formation of an IAA reflux loop described above

(Chen et al., 1998; Luschnig et al., 1998; Blilou et al., 2005; Rahman et al., 2010; Peer et al., 2011). Similar to PIN1, PIN2 expression is impacted by root stress; and decreases in

PIN2 transcript levels have been observed in Arabidopsis during iron stress (i.e., exposure to excessive levels of iron), a phenomenon postulated to aid in decreasing lateral root formation in the presence of high levels of iron (Sun et al., 2008; Wang et al.,

2009b; Li et al., 2015).

While both PIN1 and PIN2 have been demonstrated to function in salt stress responses, the role of other PIN proteins during salt stress is currently somewhat unclear.

Under normal growing conditions in Arabidopsis, PIN3 is increased in root endodermal, pericycle, and columellar cells and functions to re-distribute auxin laterally into the root cortex for shootward transport, or back into vascular tissues allowing for rootward reflux

(Friml et al., 2002b; Krecek et al., 2009). PIN3 also exhibits an apolar/non-symmetrical lateral localization in columellar cells, where it contributes to the non-symmetrical re- distribution of IAA involved in directional root growth (i.e. gravitropism) (Friml et al.,

2002b; Peer et al., 2011). Additionally, in rice, increased expression of PIN3 has also been shown to confer greater tolerance to drought (Zhang et al., 2012).

In roots, PIN4 has been shown to exhibit polar localization in the cells of the quiescent center, and PIN4 is thought to regulate auxin levels in the root by helping to generate an auxin maximum at the root tip (Friml et al., 2002a; Blakeslee et al., 2005;

Blilou et al., 2005). PIN7 is thought to maintain rootward auxin transport, helping to establish and maintain the polar auxin stream needed for normal development (Friml et al., 2003; Blakeslee et al., 2005; Blilou et al., 2005). Based on these data, it is possible, indeed likely, that PIN3 and PIN4 activity is also altered during root salt stress and halotropism responses, but to date this has not been investigated. PINs 5, 6, and 8, are truncated short PINs, lacking a central hydrophilic region, that localize to the endomembrane system, and are thought to maintain cytosolic auxin homeostasis by transporting auxins during the biosynthetic process (Mravec et al., 2009; Ljung, 2013).

The second large family of auxin transporters are the ATP binding cassette-B

(ABCB) family of transporters, at least three members of which, ABCB1, ABCB19,

ABCB4 (and possibly ABCB21) have been demonstrated to actively transport auxin, sometimes in a synergistic co-ordination with PIN proteins (in areas where ABCB transporters and PINs co-localize) (Blakeslee et al., 2005; Geisler and Murphy, 2006;

Zazimalova et al., 2010; Peer et al., 2011; Kamimoto et al., 2012). Where PIN proteins are polarly localized and function in the directional movement of auxins, ABCB transporters tend to be more apolarly localized and function in the energy-dependent active transport of auxin against its concentration gradient (Blakeslee et al., 2007; Peer et al., 2011).

For example, in light-grown Arabidopsis seedlings, ABCB1 and 19 are found primarily in meristematic tissues in both roots and shoots, where they mediate the transport of auxin out of the meristems. Loss of these ABCB transporters results reduced auxin transport and concomitant reductions in growth in Arabidopsis, sorghum, and maize (Geisler et al., 2003; Multani et al., 2003; Noh et al., 2003; Geisler et al., 2005;

Bandyopadhyay et al., 2007; Titapiwatanakun et al., 2009; Yang and Murphy, 2009; Peer et al., 2011). Unlike ABCB1 and ABCB19, in Arabidopsis, ABCB4 is found in primarily in flowers and roots (Terasaka et al., 2005). Additionally, unlike ABCB1 and ABCB19, or indeed almost all other ABCB transporters, ABCB4 appears to function as a bi- directional, concentration dependent transporter, mediating auxin uptake when concentrations of auxin are low, but IAA efflux as concentrations of auxin increase

(Kubes et al., 2012). In roots, ABCB4 functions in shootward IAA transport within

42 cortical and epidermal cells; and where ABCB4 co-localizes with PIN2, these transporters are thought to work synergistically to mediate auxin efflux (Terasaka et al.,

2005; Blakeslee et al., 2007; Titapiwatanakun et al., 2009; Yang and Murphy, 2009).

Loss of ABCB4 results in decreased lateral root formation, increased root hair length, reduced root elongation, and delayed gravitropism that is thought to be a result of slowed root growth (Terasaka et al., 2005; Cho et al., 2007; Yang and Murphy, 2009; Kubes et al., 2012).

There is increasing evidence that ABCB transporters function in stress responses in multiple plant species. In rice, ABCB4 expression has been shown to decrease under drought stress, while expression of both ABCB1 and 19 was increased under these conditions (Chai and Subudhi, 2016). Salt stress has also been demonstrated to alter the expression patterns of ABCBs in rice, particularly ABCB1, which showed decreased expression levels 5-hours post-salt stress (levels of ABCB4 and ABCB19 transcripts were relatively unchanged under these conditions) (Chai and Subudhi, 2016). The alterations in ABCB gene expression observed in rice plants during stress responses appear to be conserved in other monocots, such as maize. In a recent study, in response to drought, salt, and cold stresses the majority of maize ABCB transporters monitored showed increased expression in leaf tissues, but dramatically decreased expression in roots (Yue et al., 2015).

ABCB and PIN transporters can interact both directly (in membranes) and indirectly (through ABCB-based modulation of lipid environments). Loss of ABCB19 leads to enhanced gravi- and phototropism, in part due to mis-localization of PIN1

43 resulting in greater lateral distribution of IAA (Noh et al., 2003; Bandyopadhyay et al.,

2007; Titapiwatanakun et al., 2009). Further, ABCB1 has been shown to co-localize with

PIN1 and PIN2, and ABCB19 has been isolated in detergent resistant microdomains

(DRMs, also known as “lipid rafts”) with both PIN1 and PIN2 (Noh et al., 2003;

Bandyopadhyay et al., 2000; Blakeslee et al., 2007; Titapiwatanakun et al., 2009).

Membrane composition and trafficking serve as one layer of regulation of both ABCB- and PIN-mediated auxin transport. An additional level of control of auxin transport is provided by controlling the activity of the transporters.

Flavonoids also have been shown to modulate auxin transport (Jacobs and

Rubery, 1988; Murphy et al., 2000; Buer et al., 2014). The Arabidopsis tt4 mutant as previously mentioned does not synthesize flavonoids (Peer et al., 2001). This mutation results in an increase in auxin transport from shoot to root as well as slowed gravitropic bending (Peer et al., 2004). Here, flavonoids were shown to function though regulation of PIN1 and PIN4 localization by modulating PIN trafficking to the plasma membrane

(Brown et al., 2001; Buer et al., 2004; Peer et al., 2004). The Arabidopsis tt7 mutant, which hyper accumulates flavonoids, exhibits decreased auxin transport and de- localization of PIN4 from the plasma membrane (Peer et al., 2004). A similar mutant in tomato, are (anthocyanin reduced), is deficient in flavonoid 3 hydroxylase (F3H), the enzyme that produces flavonols, and this mutant also exhibits increased shoot-to-root auxin transport (Maloney et al., 2014). PINs, however, are less sensitive to flavonoids compared to ABCB transporters (Peer and Murphy, 2007). Flavonoids are known to directly bind the ATP binding sites of ABCB proteins, and ABC transporters exhibit

44 greater flavonoid sensitivity because flavonoids themselves are transported into the vacuole by ABC type C (ABCC) transporters and Multidrug and Toxin Extrusion

(MATE) transporters (Williams et al., 2004).

The auxin transporters ABCB1 and ABCB4 both bind flavonols, and this interaction has been shown to regulate ABCB activity (Geisler et al., 2005; Geisler and

Murphy, 2006). The Arabidopsis ABCB4 mutant exhibits enhanced gravitropism, and when stacked with the tt4 mutation (which exhibits delayed gravitropism) there is no change in phenotype compared to abcb4 alone; therefore, abcb4 is epistatic to tt4 and flavonols exhibit direct regulation of this ABCB transporter (Lewis et al., 2007).

Flavonoids also interact with other proteins, including kinases and phosphatases, and small quantities of flavonoids inhibit their function (Estrada et al., 2005; Holder et al.,

2007; Peer and Murphy, 2007; Hou and Kumamoto, 2010). As both kinases (PINOID) and phosphatases (PP2A) regulate polarity and trafficking of auxin transporters, it is also very likely that flavonoids modulate auxin transporters through regulating their phosphorylation state (Peer and Murphy, 2007).

Summary

Flavonoids and anthocyanins alike are increasing in demand for use as natural substitutes for synthetic industrial dye and food colorants, and these phenolic metabolites are being used more and more for their medicinal properties. However, in order to meet the demand for flavonoids across multiple industries it is necessary to find alternate plant sources. Turfgrasses are a group of plants that could serve as flavonoid bio-factories in

45 that their leaves can be easily harvested, and turfgrass tissues grow back relatively fast allowing for continual harvest. To increase flavonoid production in turfrgasses to greater levels, plants can be treated with specific light regimes that promote flavonoid and anthocyanin synthesis. Other than light, flavonoid production is, in general, upregulated by a number of biotic and abiotic stresses.

One such stress where flavonoids are found to accumulate to a greater degree is salt stress. During salt stress, flavonoids increase in root tissues where they serve as potent anti-oxidants, and halophytic plants have been shown to contain greater quantities of flavonoids during salt stress responses. Under conditions of excessive NaCl, plants will also alter root architecture. Changes in root architecture including, development and growth of lateral roots, and root tropic growth are all dependent on alterations in phytohormone transport and metabolism. Interestingly, flavonoids are known to directly modulate both the metabolism and the transport of the phytohormone auxin. Not only may turfgrasses serve as flavonoid bio-factories, but turfgrasses may also be model plants for studying the function of flavonoids during stress responses.

Reference

Abdou, E.M. H.S. Hafez, E. Bakir, M.SA. Abdel-Mottaleb. 2013. Photostability of low cost dye-sensitized solar cells based on natural and synthetic dyes. Spectrochimica Acta Part A: Molecular and Biomolecular Spectroscopy. 115: 202-207.

Ahmad, M., J.A. Jarillo, O. Smirmova, and A.R. Cashmore. 1998. Cryptochrome blue- light photoreceptors of Arabidopsis implicated in phototropism. Nature. 392: 720- 723.

Ahmad, M. 2016. Photocycle and signaling mechanism of plant cryptochrome. Curr. op. in plant biol. 33: 108-115.

American Meteorological Society. 2012. Meteorology glossary. http://glossary.ametsoc.org/wiki/Freshwater.

Apse, M.P., G.S. Aharon., W.A. Snedden, and E. Blumwald. 1999. Salt tolerance conferred by overexpression of a vacuolar Na+/H+ antiport in Arabidopsis. Science. 285.

Apse, M.P., J.B. Scottosanto, and E. Blumwald. 2003. Vacoular cation/H+ exchange, ion homeostasis, and leaf development are altered in a T-DNA insertional mutant of AtNHX1, the Arabidopsis vacoular Na+/H+ antiporter. The Plant Journal. 36: 229-239.

Baker, D.A. 2000. Vascular transport of auxins and cytokinins in Ricinus. Plant Growth Regulation. 32: 157-160.

Bandyopadhyay, A., J.J. Blakeslee, O.R. Lee, J. Mravec, M. Sauer, B. Titapiwatanakun, S.N. Makam, R. Bouchard, M. Geisler, E. Martinoia, J. Friml, W.A. Peer, and A.S. Murphy. 2007. Interactions of PIN and PGP auxin transport mechanisms. Biochem. Soc. Trans. 35: 137-141.

Bao, A.K., S.M. Wang, G.Q. Wu, J.J. Xi, J.L. Zhang, and C.M. Wang. 2009. Overexpression of the Arabidopsis H+-PPase enhanced resistance to salt and drought stress in transgenic alfalfa (Medicago sativa L.). Plant Science. 176: 232- 240.

Barragan, V., E.O. Leidi E, Z. Andres, L. Rubio, A. De Luca, J.A. Fernandez, B. Cubero, and J.M Pardo. 2012. Ion Exchangers NHX1 and NHX2 Mediate Active Potassium Uptake into Vacuoles to Regulate Cell Turgor and Stomatal Function in Arabidopsis. Plant Cell. 24: 1127-1142.

Bartel, B., and G.R. Fink.. 1995. Ilr1, an Amidohydrolase That Releases Active Indole-3- Acetic-Acid from Conjugates. Science. 268: 1745-1748.

Bassil, E., M.A. Ohto, T. Esumi, H. Tajima, Z. Zhu, O. Cagnac, M. Belmonte, Z. Peleg, T. Yamaguchi, and E. Blumwald. 2011. The Arabidopsis intracellular Na+/H+ antiporters NHX5 and NHX6 are endosome associated and necessary for plant growth and development. Plant Cell. 23: 224-239.

Beard, J.B. 1973. Chapter 2: Growth and development pgs. 25-53. In. Turfgrass: Science and culture. J.B. Beard. Prentice-Hall. Edgewood Cliffs, NJ. USA.

Bechtold, T., A. Mahmud-Ali, and R. Mussak. 2007. Anthocyanin dyes extracted from grape pomace for the purpose of textile dyeing. J. Sci. Food Agr. 87: 2589-2595.

Beffa, R., H.V. Martin, and P.E. Pilet. 1990. In vitro oxidation of indoleacetic acid by soluble auxin-oxidases and peroxidases from maize roots. Plant Physiol. 94: 485- 491.

Ben-Hur, M., F.H. Li, R. Keren, and G. Shalit. 2001. Water and salt distribution in a field irrigation with marginal under high water table conditions. Soil Sci. Soc. of Am. J. 65: 191-198.

Bennett, S.R.M, J. Alvarez, G. Bossinger, and D.R. Smyth. 1995. Morphogenesis in pinoid mutants of Arabidopsis thaliana. The Plant Journal. 8: 505-520.

Blilou, I., J. Xu, M. Wildwater, V. Willemsen, I. Paponov, J. Friml, R. Heidstra, M. Aida, K. Palme, and B. Scheres. 2005. The PIN auxin efflux facilitator network controls growth and patterning in Arabidopsis roots. Nature. 433: 39-44.

Blakeslee, J,J., W.A. Peer, and A.S. Murphy. 2005. Auxin transport. Curr. Op. Plant Biol. 8: 494-500.

Blakeslee, J.J., A. Bandyopadhyay, O.R. Lee, J. Mravec, B. Titapiwatanakun, M. Sauer, S.N. Makam, Y. Cheng, R. Bouchard, J. Adamec, M. Geisler, A. Nagashima, T. Sakai, E. Martinoia, J. Friml, W.A. Peer, and A.S. Murphy. 2007. Interactions among PIN-FORMED and P-Glycoprotein Auxin Transporters in Arabidopsis. Plant Cell. 19: 131-147.

Blumwald E. 2000. Sodium transport and salt tolerance in plants. Curr. Op. in Cell Biol. 12: 431-434.

Boerjan, W., J. Ralph, and M. Baucher. 2003. Lignin biosynthesis. Ann. Rev. Plant Biol. 54: 519-546.

Boldt, J.K., M.H. Meyer, and J.E. Erwin. 2014. Foliar anthocyanins: A horticultural review, p. 209-251. In: J. Janick (ed.). Hort. Rev. Wiley, New York, NY.

Bouly, J.E. Scheicher, M. Dionisio-Sese, F. Vandenbussche, D. Van Der Straeten, N. Bakrim, S. Meier, A. Batschaeur, P. Galland, R. Bittl, and M. Ahmad. 2007. Cryptochrome blue light photoreceptors are activated through interconversion of flavin redox states. J. of Biol. Chem. 282(13): 9389-9391.

Breuninger, J.M., M.S. Welterlen, B.J. Augustin, V. Cline, and K. Morris. 2013. The turfgrass industry, p. 37-103. In: Turfgrass: Biology, use, and management. J.C. Stier, B.P. Horgan, and S.A. Bonos (eds.). American Society of Agronomy, Soil Science Society of America, Crop Science Society of America monographs. Madison WI., USA.

Brosche, M., B. Vinocur, E.R. Alatalo, A. Lamminmaki, T. Teichmann, E.A. Ottow, D. Djilianov, D. Afif, M.B. Bogeat-Triboulot, A. Altman, A. Polle, E. Dreyer, S. Rudd, L. Paulin, P. Auvinen, and J. Kangasjarvi. 2005. Gene expression and metabolite profiling of Populus euphratica growing in the Negev desert. Genome Biol. 6: R101.

Brown, D.E., AM. Rashotte, A.S. Murphy, J. Normanly, B.W. Tague, W.A. Peer, L. Taiz, and G.K. Muday. 2001. Flavonoids act as negative regulators of auxin transport in vivo in Arabidopsis. Plant Physiol. 126: 524-535.

Buer, C.S. and G.K. Muday. 2004. The transparent testa4 mutation prevents flavonoid synthesis and alters auxin transport and the response of Arabidopsis roots to gravity and light. The Plant Cell. 16: 1191-1205.Butler, W. L., S.B. Hendricks, and H.W. Siegelman. 1964. Action Spectra of Phytochrome in vitro. Photochem. and Photobiol. 3: 521-528.

Buer, C.S. and G.K. Muday. 2014. The transparent testa4 mutation prevents flavonoid synthesis and alters auxin transport and the response of Arabidopsis roots to gravity and light. The Plant Cell. 16: 1191-1205.Butler, W. L., S.B. Hendricks, and H.W. Siegelman. 1964. Action Spectra of Phytochrome in vitro. Photochemistry and Photobiology. 3: 521-528.

Burgie, E.S., A.N. Bussell, J.M. Walker, J. Dubiel, and R. D. Vierstra. 2014. PNAS. 111(28): 10179-10184.

Butler, W.L., S. Hendricks, and H.W. Siegelman. 1964. Action spectra of phytochrome in vitro. Phytochem. and Photobiol. 3: 521-528.

Callebaut, A., N. Terahara, M. de Haan, and M. Decleire. 1997. Stability of anthocyanin composition in Ajuga reptans callus and cell suspension cultures. Plant, Cell, Tissue, and Organ Cult. 50: 195-201.

Casal, J.J., R.A. Sanchez, and M.J. Yanovsky. 1997. The function of phytochrome A. Plant Cell and Environ. 20: 813-819.

Casal, J.J. S.J. Davis, D. Kirchenbauer, A. Viczian, M.J. Yanovsky, R.C. Clough, S. Kircher, E.T. Jordan-Beebe, E. Schafer, F. Nagy, and R.D. Viestra. 2002. The serine-rich N-terminal domain of oat phytochrome A helps regulate light responses and subnuclear localization of the photoreceptor. Plant Physiol. 129: 1127-1137.

Casal, J.J. 2013. Photoreceptor signaling networks in plant responses to shade. Ann. Rev. Plant Biol. 64: 403-427.

Cassidy, A., M. Bertoia, S. Chiuve, A. Flint, J. Forman, and E.B. Rimm. 2016. Habitual intake of anthocyanins and flavanones and risk of cardiovascular disease in men. Am. J. Clinical Nutrition. 104: 587-594.

Castandea-Ovando, A., L. Pacheco-Hernandez, E. Paez-Hernandez, J.A. Rodriguez, and C.A. Galan-Vidal. 2009. Chemical studies of anthocyanin: A review. Food Chem. 113: 859-871.

Chai. C., and P.K. Subudhi. 2016. Comprehensive Analysis and Expression Profiling of the OsLAX and OsABCB Auxin Transporter Gene Families in Rice (Oryza sativa) under Phytohormone Stimuli and Abiotic Stresses. Front Plant Sci. 7: 593.

Chalker-Scott, L. 1999. Environmental significance of anthocyanins in plant stress responses. Photochem. and Photobiol. 70(1): 1-9.

Chaves, I., R. Pokorny, M. Byrdin, N. Hoang, T. Ritz, K. Brettel, L. Essen, G.T.J. van der Horst, A. Batschauer, and M. Ahmad. 2011. The cryptochromes: Blue light photoreceptors in plants and animals. Ann. Rev. Plant Biol. 62: 335-364.

Chefdor, F., H. Benedetti, C. Depierreux, F. Delmotte, D. Morabito, and S. Carpin. 2006. Osmotic stress sensing in Populus: components identification of a phosphorelay system. FEBS Lett. 580: 77-81.

Chen, R., P. Hilson, J. Sedbrook, E. Rosen, T. Caspar, and P. Masson. 1998. The Arabidopsis thaliana AGRAVITROPIC 1 gene encodes a component of the polar- auxin-transport efflux carrier. PNAS. 95: 15112-15117.

Chen, M., J. Chory, and C. Frankhauser. 2004. Light signal transduction in higher plants. Ann. Rev. Genet. 38: 84-117.

Cho, M., S.H. Lee, and H.T. Cho. 2007. P-glycoprotein4 displays auxin efflux transporter-like action in Arabidopsis root hair cells and tobacco cells. Plant Cell. 19: 3930-3943.

Christie, J.M. 2007. Phototropin blue-light receptors. Ann. Rev. Plant Biol. 58: 21-45.

Christie, J.M., H.B. Yang, G.L. Richter, S. Sullivan, C.E. Thomson, J.S. Lin, B. Titapiwatanakun, M. Ennis, E. Kaiserli, O.R. Lee, J. Adamec, W.A. Peer, and A.S. Murphy. 2011. phot1 Inhibition of ABCB19 Primes Lateral Auxin Fluxes in the Shoot Apex Required For Phototropism. Plos Biology. 9.

Christie, J.M., L. Blackwood, J. Peterson, and S. Sullivan. 2015. Plant flavoprotein photoreceptors. Plant and Cell Physiol. 1-13.

Christmann, A., E. Grill, and J. Huang. 2013. Hydraulic signals in long-distance signaling. Curr. Op. in Plant Biol. 16: 293-300.

Conrad, K.S., C.C. Manahan, and B.R. Crane. 2014. Photochemistry of flavoprotein light sensors. Nature Chem. Biol. 10: 801-809.

Croteau, R., T.M. Kutchan, and N.G. Lewis. 2000. Natural Products (Secondary Metabolites). In. Biochemistry and Molecular Biology of Plants. Buchanan, Gruissem, and Jones, John Wiley & Sons, Inc.

Davies, R.T.,D.H. Goetz, J. Lasswell, M.N. Anderson, and B. Bartel . 1999. IAR3 encodes an auxin conjuate hydrolase from Arabidopsis. Plant Cell. 11: 365-376.

Davis, S.J., S.H. Bhoo, A.M. Durski, J.M. Walker, and R.D. Vierstra. 2001. The Heme- Oxygenase family required for phytochrome chromophore biosynthesis is necessary for proper photomorphogenesis in higher plants. Plant Physiol. 126: 656-669.

Delgado-Vargas, F., A.R. Jiménez, and O. Paredes-López. 2000. Natural pigments: Carotenoids, anthocyanins, and betalains - characteristics, biosynthesis, processing, and stability. Crit. Rev. Food Sci. Nutr. 40(3): 173–289.

Dernonden, P.H. 1995. Why my putting greens appear purplish winter. Turfax. 4-6.

De Schepper, V., T. De Swaef, I. Bauweraerts, and K. Steppe. 2013. Phloem transport: a review of mechanisms and controls. J. Exp. Bot. 64: 4839-4850.

Deroles, S. 2009. Anthocyanin biosynthesis in plant cell cultures: A potential source of natural colourants. In. Gould, K., K. Davies, C. Winfield (eds.). Anthocyanins: Biosynthesis, functions, and applications. Springer, New York, NY. 107-158.

Diedhiou, C., O.V. Popova, and D. Golldack. Transcript profiling of the salt-tolerant Festuca rubra ssp. litoralis reveals a regulatory network controlling salt acclimation. J. of Plant Physiol. 166: 679-711.

Dooner, H.K., T.P. Robbins, and R.A. Jorgensen. 1991. Genetic and developmental control of anthocyanin biosynthesis. Ann. Rev. Genetics. 25: 173-199.

Downs, R.J. and H.W. Siegelman. 1962. Photocontrol of anthocyanin synthesis in Milo seedlings. Plant Physiol. 25-30.

Drumm, H., and H. Mohr. 1978. The mode of interaction between blue (UV) light photoreceptor and phytochrome in anthocyanin formation of the Sorghum seedling. Phytochem. and Photobiol. 27: 241-248

Duan, L.N., D. Dietrich, C.H. Ng, P.M.Y. Chan, R. Bhalerao, M.J. Bennett, and J.R. Dinneny. 2013. Endodermal ABA Signaling Promotes Lateral Root Quiescence during Salt Stress in Arabidopsis Seedlings. Plant Cell. 25: 324-341. doi:10.1105/tpc.112.107227.

Du, Y., Q. Hei, Y. Liu, H. Zhang, K. Xu, and T. Xia. 2010. Isolation and characterization of a putative vacuolar Na+/H+ antiporter gene from Zoysia japonica L. J. Plant Biol. 53(4): 251-258.

Dudeck, A.E. and C.L. Murdoch. 1988. Registration of ‘FloralDwarf’ Bermudagrass. Crop Sci. 38: 538.

Estrada, O., M. Hasegawa, D. Gonzalez-Mujica, N. Motta, E. Perdomo, A. Solorzano, J. Mendez, and E.G. Zea. 205. Evaluation of flavonoids from Bauhinia megalandra leaves as inhibitors of glucose-6-phosphotase system. Phytotherapy Res. 19: 859- 863.

Eryilmaz, F. 2006. The relationships between salt stress and anthocyanin accumulation in higher plants. Biotech. And Biotech. Equip. 20(1): 47-52.

Faiyue, B., M.J. Al-Azzawi, and T.J. Flowers. 2010. The role of lateral roots in bypass flow in rice (Oryza sativa L.). Plant Cell and Environ. 33: 702-716.

Ferrer, J.L., M.B. Austin, C. Stewart, and J.P. Noel. 2008. Structure and function of enzymes involved in the biosynthesis of phenylpropanoids. Plant Physiol. and Biochem. 46: 356-370.

Fossen, T., R. Slimestad, D.O. Øvstedal D, and Ø.M. Anderson. 2002. Anthocyanins of grasses. Biochem. Systematics and Ecol. 30: 855-864.

Frankhauser, C. 2000. Phytochromes as light-modulated protein kinases. Cell and Dev. Biol. 11: 467-473.

Franklin, K.A., S.H. Lee, D. Patel, S.V. Kumar, A.K. Spartz, C. Gu, S. Ye, P. Yu, G. Breen, J.D. Cohen, P.A. Wigge, and W.M. Gray. 2011. Phytochrome-interacting factor 4 (PIF4) regulates auxin biosynthesis at high temperature. PNAS. 108: 20231-20235.

Friml, J., E. Benkova, I. Blilou, J. Wisniewksa, T. Hamann, K. Ljung, S. Woody, G. Sandberg, B. Scheres, G. Jurgens, and K. Palme. 2002a. AtPIN4 mediates sink- driven auxin gradients and root patterning in Arabidopsis. Cell. 108: 661-673.

Friml, J., A. Vieten, M. Sauer, D. Weijers, H. Schwarz, T. Hamann, R. Offringa, and G. Jurgens. 2003. Efflux-dependent auxin gradients establish the apical–basal axis of Arabidopsis. Nature. 426: 147-153.

Friml, J., J. Wisniewska, E. Benkova, K. Mendgen, and K. Palme. 2002b. Lateral relocation of auxin efflux regulator PIN3 mediates tropism in Arabidopsis. Nature. 415: 806-809.

Friml, J., X. Yang, M. Michniewicz, D. Weijers, A. Quint, O. Tietz, R. Benjamins, P.B. Ouwerkerk, K. Ljung, G. Sandberg, P.J. Hooykaas, K. Palme, and R. Offringa. 2004. A PINOID-dependent binary switch in apical-basal PIN polar targeting directs auxin efflux. Science. 306: 862-865.

Galvan-Ampudia, C.S., M.M. Julkowska, E. Darwish, J. Gandullo, R.A. Korver, G. Brunoud, M.A. Haring, T. Munnik, T. Vernoux, and C. Testerink. 2013. Halotropism is a response of plant roots to avoid a saline environment. Curr. Biol. 23: 2044-2050.

Galvão, V.C. and C. Frankhauser. 2015. Sensing the light environment in plants: Photoreceptors and early signaling steps. Curr. Op. in Neurobiology. 34: 46-53.

Galweiler, L., C. Guan, A. Muller, E. Wisman, K. Mendgen, A. Yephremov, and K. Palme. 1998. Regulation of polar auxin transport by AtPIN1 in Arabidopsis vascular tissue. Science. 282: 2226-2230.

Gassmann, W., F. Rubio, and J.I. Schroeder. 1996. Alkali cation selectivity of the wheat root high-affinity potassium transporter HKT1. The Plant Journal. 10: 869-882.

Gaxiola, R.A., J. Li, S. Undurraga, L.M Dang, G.J. Allen, S.L. Alper, and G.R. Fink. 2001. Drought- and salt-tolerant plants result from overexpression of the AVP1 H+-pump. PNAS. 98: 11444-11449.

Geisler, M., J.J. Blakeslee, R. Bouchard, O.R. Lee, V. Vincenzetti, A. Bandyopadhyay, B. Titapiwatanakun, W.A. Peer, A. Bailly, E.L. Richards, K.F.K. Ejenda,A.P. Smith, C. Baroux, U. Grossniklaus, A. Muller, C.A. Hrycyna, R. Dudler, A.S. Murphy, and E. Martinoia. 2005. Cellular efflux of auxin catalyzed by the Arabidopsis MDR/PGP transporter AtPGP1. The Plant Journal. 44: 179-194.

Geisler, M., H.U. Kolukisaoglu, R. Bouchard, K. Billion, J. Berger, B. Saal, N. Frangne, Z. Koncz-Kalman, C. Koncz, R. Dudler, J.J. Blakeslee, A.S. Murphy, E. Martinoia, and B. Schulz. 2003. TWISTED DWARF1, a unique plasma membrane-anchored immunophilin-like protein, interacts with Arabidopsis multidrug resistance-like transporters AtPGP1 and AtPGP19. Mol. Biol. Cell. 14: 4238-4249.

Geisler, M., and A.S. Murphy. 2006. The ABC of auxin transport: The role of p- glycoproteins in plant development. FEBS Letters. 580: 1094-1102.

Geng, Y., R. Wu, C.W. C.W. Wee, F. Xie, X. Wei, P.M. Chan, C. Tham, L. Duan, and J.R. Dinneny. 2013. A spatio-temporal understanding of growth regulation during the salt stress response in Arabidopsis. Plant Cell. 25: 2132-2154.

Grill, E., and A. Himmelbach. 1998. ABA signal transduction. Curr. Op. in Plant Biol. 1: 412-418.

Hagen, G., A. Kleinschmidt, and T. Guilfoyle. 1984. Auxin-regulated gene expression in intact soybean hypocotyl and excised hypocotyl sections. Planta. 162: 147-153.

Hamant, O., M.G. Heisler, H. Jonsson, P. Krupinski, M. Uyttewaal, E.M. Meyerowitz, Y. Couder, and J. Traas. 2008. Developmental patterning by mechanical signals in Arabidopsis. Science. 322: 1650-1655.

Han, Y., H. Kim, P. Song, and J. Kim. 2010. Autophosphorylation desensitizes phytochrome signal transduction. Plant Signaling and Behavior. 5(7): 868-871.

Harivandi, M.A., J.D. Butler, and L. Wu. 1992. Salinity and turfgrass culture, pgs. 207- 229. In: turfgrass agronomy monograph no. 32. G.A. Peterrson, P.S. Baenziger, and R.J. Luxmore (eds.). American Society of Agronomy, Inc., Crop Science Society of America, Inc., Soil Science Society of America, Inc. Madison WI., USA.

Harivandi, M.A. 2004. Evaluating recycled waters for golf course irrigation. USGA green section record. Nov.-Dec.

Haro, R., M.A. Banuelos, M.E. Senn, J. Barrero-Gil, and A. Rodriguez-Navarro. 2005. HKT1 mediates sodium uniport in roots. Pitfalls in the expression of HKT1 in yeast. Plant Physiol. 139: 1495-1506.

Haswell, E.S., R. Phillips, and D.C. Rees. 2011. Mechanosensitive channels: what can they do and how do they do it? Structure. 19: 1356-1369.

Hayes, A.R., C.F. Mancino, and I.L. Pepper. 1990. Irrigation of turfgrass with secondary sewage effluent: I. Soil and leachate water quality. Agron. J. 82: 939-943.

Henderson, S.W., and M. Gilliham. 2015. The "gatekeeper" concept: cell-type specific molecular mechanisms of plant adaptation to abitotic stress. Hoboken, N.J.: John Wiley and Sons.

He, J. and M. Giusti. 2010. Anthocyanin: Natural colorants with health-promoting properties. Annu. Rev. Food Sci. Technol. 1: 163-187.

Hirose, F., T. Shinomura, T. Tanabata, H. Shimada, and M. Takano. 2006. Involvement of rice cryptochromes in de-etiolation responses and flowering. Plant Cell Physiol. 47(7): 915-925.

Holm, M., L.G. Ma., L.J. Qu., and X.W. Deng. 2002. Two interacting bZIP proteins are direct targets of COP1-mediated control of light-dependent gene expression in Arabidopsis. Genes and Dev. 16: 1247-1259.

Holder, S., M. Zemskova, C. Zhang, M. Tabrizizad, R. Bremer, J.W. Neidigh, and M.B. Lilly. 2007. Characterization of a potent and selective small-molecule inhibitor of the PIM1 kinase. Mol. Cancer. Ther. 6(1): 163-172.

Holton, T.A. and E.C. Cornish. 1995. Genetics and Biochemistry of Anthocyanin Biosynthesis. Plant Cell. 7: 1071-1083.

Hornitschek, P., M.W. Kohnen, S. Lorrain, J. Rougemont, K. Ljung, I. Lopez-Vidriero, J.M. Franco-Zorrilla, R. Solano, M. Trevisan, S. Pradervand, I. Xenarios, and C. Fankhauser. 2012. Phytochrome interacting factors 4 and 5 control seedling growth in changing light conditions by directly controlling auxin signaling. Plant J. 71: 699-711.

Hou, D. and T. Kumamoto. 2010. Flavonoids as protein kinase inhibitors for cancer chemoprevention: Direct binding and molecular modeling. Antiox. and Redox Signaling. 13(5): 691-719.

Hu, T., H. Li, X. Zhang, H. Luo, and J. Fu. 2011. Toxic effect of NaCl on ion metabolism, antioxidative enzymes and gene expression of perennial ryegrass. Ecotoxicology and Environ. Safety. 74: 2050-2056.

Hu, L., T. Hu, Z. Zhang, H. Pang, and J. Fu. 2012a. Exogenous glycine betaine ameliorates the adverse effect of salt stress on perennial ryegrass. J. Amer. Soc. Hort. Sci. 137(1): 38-46.

Hu, L., Z. Huang, S. Liu, and J. Fu. 2012b. Growth response and gene expression in antioxidant-related enzymes in two Bermudagrass genotypes differing in salt tolerance. J. Amer. Soc. Hort. Sci. 137(3): 134-143.

Hughes, N.M., K.L. Carpenter, and J.G. Cannon. 2013. Estimating contribution of anthocyanin pigments to osmotic adjustment during leaf reddening. J. of Plant Physiol. 170: 230-233.

Hummel, N.W. 1993. Rationale for the revisions of the USGA green construction specifications. USGA green section record. March-April.

Hurley, R. 2010. Rough Bluegrass (Poa trivialis L.), p. 67-73. In: Casler, M.D. and R.R. Duncan (eds.). Turfgrass Biology, Genetics, and Breeding. Wiley, New York, NY

Hwa-Hong, K. 2015. Preparation and properties of cotton and wool fabrics dyed by black rice extract. Textile Res. J. 1-9.

Ismail, H., J.D. Maksimovic, V. Maksimovic, L. Shabala, B.D. Zivanovic, Y. Tian, S. Jacobsen, and S. Shabala. 2016. Rutin, a flavonoid with antioxidant activity, improves plant salinity tolerance by regulating K+ retention and Na+ exclusion from leaf mesophyll in quinoa and broad beans. Func. plant biol. 43: 75-86.

Jacobs, M. and P.H. Rubery. 1988. Naturally occurring auxin transport regulators. Science. 241: 346-349.

Jakubowska, A., and S. Kowalczyk. 2005. A specific enzyme hydrolyzing 6-O(4-O)- indole-3-ylacetyl-beta-D-glucose in immature kernels of Zea mays. Journal of Plant Physiol. 162: 207-213.

James, R.A., A.R. Rivelli, R. Munns, and S.V. Caemmerer. 2002. Factors affecting CO2 assimilation, leaf injury and growth in salt-stressed durum wheat. Functional Plant Biology. 29: 1393.

Jameson, D.A. 1964. Effect of defoliation on forage plant physiology. Crop Sci. 67-80.

Jang, I., J. Yang, H.S. Seo, and N. Chua. 2005. HFR1 is targeted by COP1 E3 ligase for post-translational proteolysis during phytochrome A signaling. Genes and Dev. 19: 593-602.

Jing, P. and M.M. Giusti. 2007. Effects of extraction conditions on improving yield and quality of an anthocyanin-rich purple corn (Zea mays L.) color extract. J. Food Sci. 72(7): 363-368.

Julkowska, M.M., H.C. Hoefsloot, S. Mol, R. Feron, G.J. de Boer, M.A. Haring, and C. Testerink. 2014. Capturing Arabidopsis root architecture dynamics with ROOT- FIT reveals diversity in responses to salinity. Plant Physiol. 166: 1387-1402.

Julkowska, M.M., and C. Testerink. 2015. Tuning plant signaling and growth to survive salt. Trends Plant Sci. 20: 586-594.

Junghans, U., A. Polle, P. Duchting, E. Weiler, B. Kuhlman, F. Gruber, and T. Teichmann. 2006. Adaptation to high salinity in poplar involves changes in xylem anatomy and auxin physiology. Plant Cell and Environment. 29: 1519-1531.

Kamimoto, Y., K. Terasaka, M. Hamamoto, K. Takanashi, S. Fukuda, N. Shitan, A. Sugiyama, H. Suzuki, D. Shibata, B. Wang, S. Pollmann, M. Geisler, and K. Yazaki. 2012. Arabidopsis ABCB21 is a facultative auxin importer/exporter regulated by cytoplasmic auxin concentration. Plant Cell Physiol. 53: 2090-2100.

Kerckhoffs, L.H.J. and R.E. Kendrick. 1997. Photocontrol of anthocyanin biosynthesis in tomato. J. Plant Res. 110: 141-149.

Kevei, E., E. Schafer, and F. Nagy. 2007. Light-regulated nucleo-cytoplasmic partitioning of phytochromes. J. Exp. Bot. 58(2): 3113-3124.

Korasick, D.A., T.A. Enders, and L.C. Strader. 2013. Auxin biosynthesis and storage forms. J. Exp. Bot. 64: 2541-2555.

Kraus, A. 2015. Development of functional food with the participation of the consumer. Motivators for consumption of functional products. Intl. J. Consumer Studies. 39: 2-11.

Krecek, P., P. Skupa, J. Libus, S. Naramoto, R. Tejos, J. Friml, and E. Zazimalova. 2009. The PIN-FORMED (PIN) protein family of auxin transporters. Genome Biol. 10: 249.

Kriechbaumer, V., P. Wang, C. Hawes, and B.M. Abell. 2012. Alternative splicing of the auxin biosynthesis gene YUCCA4 determines its subcellular compartmentation. Plant J. 70: 292-302.

Kriechbaumer, V., S.W. Botchway, and C. and C. Hawes. 2016. Localization and interactions between Arabidopsis auxin biosynthetic enzymes in the TAA/YUC- dependent pathway. J. Exp. Bot. 67: 4195-4207.

Kriechbaumer, V., H. Seo, W.J. Park, and C. Hawes. 2015. Endoplasmic reticulum localization and activity of maize auxin biosynthetic enzymes. J. Exp. Bot. 66: 6009-6020.

Kubes, M., H. Yang, G.L. Richter, Y. Cheng, E. Mlodzinska, X. Wang, J.J. Blakeslee, N. Carraro, J. Petrasek, E. Zazimalova, K. Hoyerova, W.A. Peer, and A.S. Murphy. 2012. The Arabidopsis concentration-dependent influx/efflux transporter ABCB4 regulates cellular auxin levels in the root epidermis. The Plant Journal. 69: 640- 654.

Kung, C. 2005. A possible unifying principle for mechanosensation. Nature. 436: 647- 654.

Lamb, C.J. 1979. Regulation of enzyme levels in phenylpropanoid biosynthesis: Characterization of the modulation by light and pathway intermediates. Archives of Biochem. and Biophysics. 192(1): 311-317.

Lange, H., W. Shiropshire, and H. Mohr. 1971. An analysis of phytochrome-mediated anthocyanin synthesis. Plant Physiol. 47: 649-655.

Lau, O.S. and X.W. Deng. 2012. The photomorphogenic repressors COP1 and DET1: 20 years later. Trends in Plant Sci. 17(10): 584-593.

Laurie, S., K.A. Feeney, F.J. Maathuis, P.J. Heard, S.J. Brown, and R.A. Leigh. 2002. A role for HKT1 in sodium uptake by wheat roots. The Plant Journal. 32: 139-149.

Lee, N. and G. Choi. 2017. Phytochrome-interacting factor from Arabidopsis to liverwort. Curr. Op. in Plant Biol. 35: 54-60.

Leidi, E.O., V. Barragan, L. Rubio, A. El-Hamdaoui, M.T. Ruiz, B. Cubero, J.A. Fernandez, R.A. Bressan, P.M. Hasegawa, F.J. Quintero, and J.M. Pardo. 2010. The AtNHX1 exchanger mediates potassium compartmentation in vacuoles of transgenic tomato. The Plant Journal. 61: 495-506.

Leivar, P. and E. Monte. 2014. PIFs: Systems integrators in plant devlopement. Plant Cell. 26: 56-78.

Lewis, D.R., N.D. Miller, B.L. Splitt, G. Wu, and E.P. Spalding. 2007. Separating the roles of acropetal and basipetal auxin transport on gravitropism with mutations in two Arabidopsis multidrug resistance-like ABC transporter genes. Plant Cell. 19: 1838-1850.

Liang, V., W. Go, D.M. Harris, and P. Srihari. 2015. Global overview of the role of nutraceuticals in cancer, p. 1-10. In: Sarkar, F.H (ed.). Nutraceuticals and Cancer. Springer-Verlag, New York, NY. 1-10.

Li, G., H. Song, B. Li, H.J. Kronzucker, and W. Shi. 2015. Auxin Resistant1 and PIN- FORMED2 Protect Lateral Root Formation in Arabidopsis under Iron Stress. Plant Physiol. 169: 2608-2623.

Li, F. and S. Matthews. 2016. Evolutionary aspects of plant photoreceptors. J. Plant Res.

Lilley, J.L., C.W. Gee, I. Sairanen, K. Ljung, J.L. Nemhauser. 2012. An endogenous carbon-sensing pathway triggers increased auxin flux and hypocotyl elongation. Plant Physiol. 160: 2261-2270.

Lin, C. and D. Shalitin. 2003. Cryptochrome structure and signal transduction. Annu. Rev. Plant Biol. 54: 469-496.

Liu J, Zhu JK. 1998. A calcium sensor homolog required for plant salt tolerance. Science. 280: 1943-1945.

Liu, H., B. Liu, C. Zhao, M. Pepper, and C. Lin. 2011. The action of plant cryptochromes. Trends in Plant Sci. 16(12): 684-691.

Liu, W., R.J. Li, T.T Han, W. Cai, Z.W. Fu, and Y.T. Lu. 2015. Salt stress reduces root meristem size by nitric oxide-mediated modulation of auxin accumulation and signaling in Arabidopsis. Plant Physiol. 168: 343-356.

Liobikas, J., K. Skemiene, S. Trumbeckaite, and V. Borutaite. 2016. Anthocyanins in cardioprotection: A path through mitochondria. Pharma. Res. 113: 808-815.

Liscum, E., S.K. Askinosie, D.L. Leuchtman, J. Morrow, K.T. Willenburg, and D.R. Coats. 2014. Phototropism: Growing towards an understanding of plant movement. Plant Cell. 26: 38-55.

Ljung, K, R.P. Bhalerao, and G. Sandberg. 2001. Sites and homeostatic control of auxin biosynthesis in Arabidopsis during vegetative growth. The Plant Journal. 28: 465- 474.

Ljung, K, A.K. Hull, M. Kowalczyk, A. Marchant, J. Celenza, J.D. Cohen, and G. Sandberg. 2002. Biosynthesis, conjugation, catabolism and homeostasis of indole- 3-acetic acid in Arabidopsis thaliana. Plant Mol. Biol. 49: 249-272.

Ljung, K. 2013. Auxin metabolism and homeostasis during plant development. Development 140, 943-950.

Lomax, T.L., R.J. Mehlhorn, W.R. Briggs. 1985. Active auxin uptake by zucchini membrane vesicles: Quantitation using ESR volume and ApH determinations. PNAS. 82: 6541-6545.

Ludwig-Muller, J. 2011. Auxin conjugates: their role for plant development and in the evolution of land plants. J. Exp. Bot. 62: 1757-1773.

Luschnig, C., R.A. Gaxiola, P. Grisafi, and G.R. Fink. 1998. EIR1, a root-specific protein involved in auxin transport, is required for gravitropism in Arabidopsis thaliana. Genes and Development. 12: 2175-2187.

Ma, D., W. Zu, H. Li, F. Jin, L. Guo, J. Wang, H. Dai, and X. Xu. 2014. Co-expression of the Arabidopsis SOS genes enhances salt tolerance in transgenic tall fescue (Festuca arundinacea Schreb.). Protoplasma. 251: 219-231.

Maathuis, F.J., and A. Antmann. 1999. K+ nutrition and Na+ toxicity: the basis of cellular K+/Na+ ratios. Annals of Botany. 84: 123-133.

Maathuis, F.J. 2014. Sodium in plants: perception, signalling, and regulation of sodium fluxes. J. Exp. Bot. 65: 849-858.

MacDonald, G.M. 2010. Climate Change and water in Southwestern North America special feature: water, climate change, and sustainability in the southwest. PNAS. 107: 21256-21262.

Maloney, G.S., K.T. DiNapoli, and G.K. Muday. 2014. The anthocyanin reduced tomato mutant demonstrates the role of flavonols in tomato lateral root and root hair development. Plant Physiol. 166: 614-631.

Mancino, C.F., and I.L. Pepper. 1992. Irrigation of turfgrass with secondary sewage effluent: Soil quality. Agron. J. 84: 650-654.

Mancinelli, A.L. and I. Rabino. 1978. The “High Irradiance Responses” of plant photomorphogenesis. The Botanical Rev. 44 (2): 129-180.

Mancinelli, A.L. 1985. Light-dependent anthocyanin synthesis: A model system for the study of plant photomorphogenesis. The Botanical Rev. 51(1): 107-157.

Mansour, R., B. Ezzili, and M. Farouk. 2013. Dyeing properties of wool fabrics dyes with Vitis vinifera L. (Black Grenache) leaves extract. Fibers and Polymers. 14(5): 786-792.

Mano, Y., and K. Nemoto. 2012. The pathway of auxin biosynthesis in plants. J. Exp. Bot. 63: 2853-2872.

Mauray, A., C. Felgines, C. Morand, A. Mazur, A. Scalbert, and D. Milenkovic. 2012. Bilberry anthocyanin-rich extract alters expression of genes related to atherosclerosis development in aorta of apo E-deficient mice. Nutrition, Metabolism, and Cardiovascular Diseases. 22: 72-80.

McCoy, E.L., and K.R. McCoy. 2009. Simulation of putting-green soil water dynamics: Implications for turfgrass water use. Ag. Water Management. 96: 405-414.

McLoughlin, F., C.S. Galvan-Ampudia, M.M. Julkowska, L. Caarls, D. van der Does, C. Lauriere, T. Munnik, M.A. Haring, and C. Testerink. 2012. The Snf1-related protein kinases SnRK2.4 and SnRK2.10 are involved in maintenance of root system architecture during salt stress. The Plant Journal. 72: 436-449.

Meehl, G.A., and T.F. Stocker. 2007. Global Climate Projections. Climate Change 2007: The Physical Science Basis. 747-845.

Miao, Y., D. Lv, P. Wang, X.C. Wang, J. Chen, C. Miao, and C. Song. 2006. An Arabidopsis glutathione peroxidase functions as both a redox transducer and a scavenger in abscisic acid and drought stress responses. Plant Cell. 18: 2749- 2766. doi:10.1105/tpc.106.044230.

Mol., J., G. Jenkins, E. Schäfer, and D. Weiss. 1996. Signal perception, transduction, and gene expression involved in anthocyanin biosynthesis. Critical Rev. in Plant Sci. 15(5 and 6): 525-557.

Milesi, C., S.W. Running, C.D. Elvidge, J.B. Dietz, B.J. Tuttle, and R.R. Nemani. 2005. Environmental Management. 32 (3): 426-438.

Mravec, J., P. Skupa, A. Bailly, K. Hoyerova, P. Krecek, A. Bielach, J. Petrasek, J. Zhang, V. Gaykova, Y.D.Stierhof, P.I. Dobrev, K. Schwarzerova, J. Rolcik, D. Seifertova, C. Luschnig, E. Benkova, E. Zazimalova, M. Geisler, and J. Friml. 2009. Subcellular homeostasis of phytohormone auxin is mediated by the ER- localized PIN5 transporter. Nature. 459: 1136-U1127.

Muller, P. and M. Ahmad. 2011. Light activated cryptochrome reacts with molecular oxygen to form a flavin-superoxide radical pair consistent with magnetoperception. J. biol. Chem. 286(34): 21033-21040.

Multani, D.S., S.P. Briggs, M.A. Chamberlin, J.J. Blakeslee, A.S. Murphy, and G.S. Johal. 2003. Loss of an MDR transporter in compact stalks of maize br2 and sorghum dw3 mutants. Science. 302: 81-84.

Munns, R., and A. Termaat. 1986. Whole-plant responses to salinity. Australian J. Plant Physiol. 13: 143-160.

Munns, R. 1993. Physiological processes limiting plant growth in saline soils: some dogmas and hypotheses. Plant Cell and Environ. 16: 15-24.

Munns, R. 2002. Comparative physiology of salt and water stress. Plant Cell and Environ. 25: 239-250.

Munns, R, Tester M. 2008. Mechanisms of salinity tolerance. Ann. Rev. Plant Biol. 59: 651-681.

Murphy, A.S., W.A. Peer, and Taiz, L. 2000. Regulation of auxin transport by aminopeptidases and endogenous flavonoids. Planta. 211: 315-324.

Nangle, E.J., D.S. Gardner, J.D. Metzger, L. Rodriquez-Saona, M.M. Giusti, T.K. Danneberger, and D.P. Petrella. 2015. Pigment changes in cool-season turfgrasses in response to ultraviolet-B light. Agron. J. 107(1): 41-50.

Narayan, M.R. 2012. Review: Dye sensitized solar cells based on natural photosensitizers. Renewable and Sustainable Energy Reviews. 16: 208– 215.

National Institutes of Health (NIH). Dictionary of dietary supplement terms. https://ods.od.nih.gov/HealthInformation/dictionary.aspx.

Neff, M.M. and J. Chory. 1998. Genetic interactions between phytochrome A, phytochrome B, and cryptochrome 1 during Arabidopsis development. Plant Physiol. 118: 27-36.

Nemoto, K., M. Hara, M. Suzuki, H. Seki, T. Muranaka, and Y. Mano. 2009. The NtAMI1 gene functions in cell division of tobacco BY-2 cells in the presence of indole-3-acetamide. FEBS Lett. 583: 487-492.

Noh, B., A. Bandyopadhyay, W.A. Peer, E.P. Spalding, and A.S. Murphy. 2003. Enhanced gravi- and phototropism in plant mdr mutants mislocalizing the auxin efflux protein PIN1. Nature. 423: 999-1002.

Normanly, J. 2010. Approaching Cellular and Molecular Resolution of Auxin Biosynthesis and Metabolism. Cold Spring Harbor Perspectives in Biology 2.

Obear, G.R., A.E. Hartemink, and D.J. Soldat. Soils with iron-cemented layers on golf courses in the USA. Geoderma. 232-234: 198-207.

Oh, S., S.N. Warnasooriya, and B.L. Montgomery. 2014. Mesophyll-Localized Phytochromes gate stress- and light-inducible anthocyanin accumulation in Arabidopsis thaliana. Plant Signaling and Behavior. 9: 1-9.

Ostin, A., M. Kowalczyk, R.P. Bhalerao, and G. Sandberg. 1998. Metabolism of indole- 3-acetic acid in Arabidopsis. Plant Physiol. 118, 285-296.

Osterlund, M.T., C. Hardtke, N. Wei, and X.W. Dang. 2000. Targeted destabilization of HY5 during light-regulated development of Arabidopsis. Nature. 405: 462-466.

Pacin, M., M. Legris, and J.J. Casal. 2013. COP1 re-accumulates in the nucleus under shade. The Plant Journal. 75: 631-641.

Park, JE, Park JY, Kim YS, Staswick PE, Jeon J, Yun J, Kim SY, Kim J, Lee YH, Park CM. 2007. GH3-mediated auxin homeostasis links growth regulation with stress adaptation response in Arabidopsis. J. Biol. Chem. 282: 10036-10046.

Peer, W.A., A. Bandyopadhyay, J.J. Blakeslee, S.N. Makam, R.J. Chen, P.H. Masson, and A.S. Murphy. 2004. Variation in expression and protein localization of the PIN family of auxin efflux facilitator proteins in flavonoid mutants with altered auxin transport in Arabidopsis thaliana. Plant Cell. 16: 1898-1911.

Peer, W.A., and A.S. Murphy. 2007. Flavonoids and auxin transport: modulators or regulators? Trends in Plant Science. 12: 556-563.

Peer, W.A., J.J. Blakeslee, H.B. Yang, and A.S. Murphy. 2011. Seven Things We Think We Know about Auxin Transport. Molecular Plant. 4: 487-504.

Peer, W.A., Y. Cheng, and A.S. Murphy. 2013. Evidence of oxidative attenuation of auxin signalling. J. Exp. Bot. 64: 2629-2639.

Petroni, K. and G. Tonelli. 2011. Recent advances on the regulation of anthocyanin synthesis in reproductive organs. Plant Sci. 181: 219-229.

Pratt, L.H., S.J. Stewart, Y. Shimazaki, Y. Wang, and M. Cordonnier. 1991. Monoclonal antibodies directed to phytochrome from green leaves of Avena sativa L. cross- react weakly or not at all with the phytochrome that is most abundant in etiolated seedlings of the same species. Planta. 184: 87-95.

Qiu, Q.S., B.J. Barkla, R. Vera-Estrella, J.K. Zhu, K.S. Schumaker. 2003. Na+/H+ exchange activity in the plasma membrane of Arabidopsis. Plant Physiol. 132: 1041-1052.

Quail, P.H. 2002. Phytochrome signaling networks. Nature Rev: Mol. Cell Biol. 3: 85-93.

Rabino, I., A.L. Mancinelli, and K.M. Kuzmanoff. 1977. Photocontrol of anthocyanin synthesis: VI. Spectral sensitivity, irradiance dependence, and reciprocity relationship. Plant Physiol. 59: 569-573.

Rabino, I. and A.L. Mancinelli. 1986. Light, temperature, and anthocyanin production. Plant Physiol. 81: 922-924

Rahman, A., M. Takahashi, K. Shibasaki, S. Wu, T. Inaba, S. Tsurumi, and T.I. Baskin. 2010. Gravitropism of Arabidopsis thaliana roots requires the polarization of PIN2 toward the root tip in meristematic cortical cells. Plant Cell. 22: 1762-1776.

Ramachandra Rao, S. and G.A. Ravishankar. 2002. Plant cell cultures: Chemical factories of secondary metabolites. Biotech. Adv. 20: 101-153.

Rampey, R.A., S. LeClere, M. Kowalczyk, K. Ljung, G. Sandberg, and B. Bartel. 2004. A family of auxin-conjugate hydrolases that contributes to free indole-3-acetic acid levels during Arabidopsis germination. Plant Physiol. 135: 978-988.

Rockwell, N.C., Y. Su, and J.C. Lagarias. 2006. Phytochrome structure and signaling mechanisms. Ann. Rev. Plant Biol. 57: 837-858.

Rodriguez-Amaya, D.B. 2016. Natural pigments and colorants. Curr. Op. Food Sci. 7: 20-26.

Rosquete, M.R., and J. Kleine-Vehn. 2013. Halotropism: turning down the salty date. Curr. Biol. 23: R927-929.

Roux, S., J. Silverthorne, and T. Brutnell. 2010. Phytochrome and light control of plant development. In. Plant Physiology 5th edition. Eds. Taiz L. and E. Zeiger. Sinauer. 493-516.

Ruiz-Sola, M.A., V. Arbona, A. Gomez-Cadenas, M. Rodriguez-Concepcion, and A. Rodriguez-Villalon. 2014. A Root Specific Induction of Carotenoid Biosynthesis Contributes to ABA Production upon Salt Stress in Arabidopsis. PLoS One 9.

Rutledge, J.M. D.V. Weisenberger, and Z.J. Reicher. 2010. Response of seedling roughstalk bluegrass and creeping bentgrass to bispyribac-sodium or sulfosulfuron. HortSci. 45(2): 288-292.

Sabo, J.L., T. Sinha, L.C. Bowling, G.H.W. Schoups, W.W. Wallender, M.E. Campana, K.A. Cherkauer, P.L. Fuller, W.L. Graf, J.W. Hopmans, J.S. Kominoski, C. Taylor, S.W. Trimble, R.H. Webb, and E.E. Wohl. 2010. Reclaiming freshwater sustainability in the Cadillac Desert. PNAS. 107: 21263-21269.

Saito, K., K. Yonekura-Sakakibara, R. Nakabayshi, Y. Higashi, M. Yamazaki, T. Toghe, and A.R. Fernie. 2013. The flavonoid biosynthetic pathway in Arabidopsis: Structural and genetic diversity. Plant Physiol. and Biochem. 72: 21-34.

Sairanen, I., O. Novak, A. Pencik, Y. Ikeda, B. Jones, G. Sandberg, and K. Ljung. 2012. Soluble carbohydrates regulate auxin biosynthesis via PIF proteins in Arabidopsis. Plant Cell. 24: 4907-4916.

Schepens, I., P. Duek, and C. Frankhauser. Phytochrome-mediated light signaling in Arabidopsis. Curr. Op. in Plant Biol. 7: 564-569.

Seo, H.S., J. Wang, M. Ishikawa, C. Bolle, M.L. Ballesteros, and N. Chua. 2003. Nature. 423(26): 995-999.

Schilling, R.K., P. Marschner, Y. Shavrukov, B. Berger, M. Tester, S.J. Roy, and D.C. Plett. 2014. Expression of the Arabidopsis vacuolar H(+)-pyrophosphatase gene (AVP1) improves the shoot biomass of transgenic barley and increases grain yield in a saline field. Plant Biotechnology J. 12: 378-386.

Seabra, A., P. Katzmarzyk, MK. Cavalho, A. Seabra, M. Coelho-E-Silva, and S. Abreu. Effects of 6-month soccer and traditional physical activity programmes on body composition, cardiometric risk factors, inflammatory, oxidative stress markers, and cardiorespiratory fitness in obese boys. 2016. J. of Sports Sci. 34(19).

Seeram, N.P., L.S. Adams, Y. Zhang, R. Lee, D. Sand, H.S. Scheuller, and D. Heber. 2006. Blackberry, black raspberry, blueberry, cranberry, red raspberry, and strawberry extract inhibit growth and stimulate apoptosis of human cancer cells in vitro. J. Agr. Food Chem. 54: 9329-9339.

Shabala, S. and T.A. Cuin. 2007. Potassium transport and plant salt tolerance. Physiologia Plantarum. 133: 651-669.

Shinomura, T., K. Uchida, M. Furuya. 2000. Elementary processes of photoperception by phytochrome A for high-irradiance response of hypocotyl elongation in Arabidopsis. Plant Physiol. 122: 147-156.

Siro, I., E. Kapolna, B. Kapolna, and A. Lugasi. 2008. Functional food. Product development, marketing and consumer acceptance - A review. Appetite. 51: 456- 467.

Smith, H. and G.C. Whitelam. 1990. Phytochrome, a family of photoreceptors with multiple physiological roles. Plant Cell and Environ. 13: 695-707.

Spalding, E.P. 2013. Diverting the downhill flow of auxin to steer growth during tropisms. Am. J. of Botany. 100(1): 203-214.

Spiess, G.M., A. Hausman, P. Yu, J.D. Cohen, R.A. Rampey, and B.K. Zolman. 2014. Auxin Input Pathway Disruptions Are Mitigated by Changes in Auxin Biosynthetic Gene Expression in Arabidopsis. Plant Physiol. 165: 1092-1104.

Soeno, K., Goda H, Ishii T, Ogura T, Tachikawa T, Sasaki E, Yoshida S, Fujioka S, Asami T, Shimada Y. 2010. Auxin biosynthesis inhibitors, identified by a genomics-based approach, provide insights into auxin biosynthesis. Plant Cell Physiol. 51: 524-536.

Staswick, P.E., B. Serban, M. Rowe, I. Tiryaki, M.T. Maldonado, M.C. Maldonado, and W. Suza. 2005. Characterization of an Arabidopsis enzyme family that conjugates amino acids to indole-3-acetic acid. Plant Cell. 17: 616-627.

Staswick, P.E.. 2009. The Tryptophan Conjugates of Jasmonic and Indole-3-Acetic Acids Are Endogenous Auxin Inhibitors. Plant Physiol. 150: 1310-1321.

Stepanova, A.N., J. Robertson-Hoyt, J. Yun, L.M. Benavente, D.Y. Xie, K. Dolezal, A. Schlereth, G. Jurgens, J.M. Alonso. 2008. TAA1-mediated auxin biosynthesis is essential for hormone crosstalk and plant development. Cell. 133: 177-191.

Stiff, M.L. and J.B. Powell. 1974. Stem anatomy of turfgrass. Crop Sci. 14: 181-186.

Stommel, J.R., G.J. Lightbourn, B.S. Winkel. R.J. and Griesback. 2009. Transcription Factor Families Regulate the Anthocyanin Biosynthetic Pathway in Capsicum annuum. J. Amer Soc. Hort. Sci. 134(2):244–251.

Sun, F., W. Zhang, H. Hu, B. Li, Y. Wang, Y. Zhao, K. Li, M. Liu, and X. Li. 2008. Salt modulates gravity signaling pathway to regulate growth direction of primary roots in Arabidopsis. Plant Physiol. 146: 178-188.

Swarup, K., E. Benkova, R. Swarup, I. Casimiro, B. Peret, Y. Yang, G. Parry, E. Nielsen, I. De Smet, S. Vanneste, M.P. Levesque, D. Carrier, N. James, V. Calvo, K. Ljung, E. Kramer, R. Roberts, N. Graham, S. Marillonnet, K. Patel, J.D. Jones, C.G. Taylor, D.P. Schachtman, S. May, G. Sandberg, P. Benfey, J. Friml, I. Kerr , T. Beeckman, L. Laplaze, M.J. Bennett. 2008. The auxin influx carrier LAX3 promotes lateral root emergence. Nat Cell Biol. 10: 946-954.

Swarup, R., J. Kargul, A. Marchant, D. Zadik, A. Rahman, R. Mills, A. Yemm, S. May, L. Williams, P. Millner, S. Tsurumi, I. Moore, R. Napier, I.D. Kerr, and M.J. Bennett. 2004. Structure-function analysis of the presumptive Arabidopsis auxin permease AUX1. Plant Cell. 16: 3069-3083.

Takano, M., N. Inagaki, X. Xie, S. Klyota, A. Baba-Kasai, T. Tanabata, and T. Shinomura. 2009. Phytochromes are the sole photoreceptors for perceiving red/far-red light in rice. PNAS. 106(34): 14705-14710.

Tao, Y., J.L. Ferrer, K. Ljung, F. Pojer, F. Hong, J.A. Long, L. Li, J.E. Moreno, M.E. Bowman, L.J. Ivans, Y. Cheng, J. Lim, Y. Zhao, C.L. Ballare, G. Sandberg, J.P. Noel, J. Chory. 2008. Rapid synthesis of auxin via a new tryptophan-dependent pathway is required for shade avoidance in plants. Cell. 133: 164-176.

Terasaka, K., J.J. Blakeslee, B. Titapiwatanakun, W.A. Peer, A. Bandyopadhyay, S.N. Makam, O.R. Lee, E.L. Richards, A.S. Murphy, F. Sato, and K. Yazaki. 2005. PGP4, an ATP binding cassette P-glycoprotein, catalyzes auxin transport in Arabidopsis thaliana roots. Plant Cell. 17: 2922-2939.

Tester, M., and R.J. Davenport. 2003. Na+ Tolerance and Na+ Transport in Higher Plants. Annals of Botany. 91: 503-527.

Tobin, E.M. and J. Silverthrorne. 1985. Light regulation of gene expression in higher plants. Ann. Rev. Plant Physiol. 36: 569-593.

Titapiwatanakun, B., J.J. Blakeslee, A. Bandyopadhyay, H. Yang, J. Mravec, M. Sauer, Y. Cheng, J. Adamec, A. Nagashima, M. Geisler, T. Sakai, J. Friml, W.A. Peer, and A.S. Murphy. 2009. ABCB19/PGP19 stabilises PIN1 in membrane microdomains in Arabidopsis. The Plant Journal. 57: 27-44.

Trenberth, K.E., A. Dai, G. van der Schrier, P.D. Jones, J. Barichivich, K.R. Briffa, and J. Sheffield. 2013. Global warming and changes in drought. Nature Climate Change 4: 17-22.

Tsuda, T, F. Horio, and T. Osawa. 2000. The role of anthocyanins as an antioxidant under oxidative stress in rats. BioFactors. 13: 133-139.

Tsuda, T., F. Horio, K. Uchida, H. Aoki, and T. Osawa. 2003. Dietary cyanidin 3-O-beta- D-glucoside-rich purple corn color prevents obesity and ameliorates hyperglycemia in mice. J. Nutr. 133: 2125–2130.

Tsuda, T. 2008. Regulation of adipocyte function by anthocyanins; possibility of preventing the metabolic syndrome. J. Agric. Food Chem. 56: 642–646.

Tsukaya, H., T. Oshima, S. Naito, M. Chino, and Y. Komeda. 1991. Sugar-dependent expression of CHS-A gene for chalcone synthase from petunia in transgenic Arabidopsis. Plant Physiol. 97: 1414-1421.

Tyrvainen, L., A. Ojala, K. Korpela, T. Lanki, Y. Tsunetsugu, and T. Kagawa. 2014. The influence of urban green environments on stress relief measures: A field experiment. J. Environ. Psychology. 38: 1-9.

United States Golf Association staff. 2004. USGA recommendation for a method of putting green construction. USGA green section record.

Van Oosten, M.J., A. Sharkhuu, G. Batelli, R.A. Bressan, and A. Maggio. 2013. The Arabidopsis thaliana mutant air1 implicates SOS3 in the regulation of anthocyanins under salt stress. Plant Mol. Biol. 83, 405-415.

Vogelien, D.L., G. Hrazdina, S. Reeves, and D.K. Dougall. 1990. Phenotypic differences in anthocyanin accumulation among clonally related cultured cells of carrot. Plant Cell, Tissue, and Organ Cult. 22: 213-222.

Vyas, P., I. Haque, and M. Kumar. 2014. Photocontrol of differential gene expression and alterations in foliar anthocyanin accumulation: A comparative study using red and green forma Ocimum tenuiflorum. Acta Physiol. Plant. 36: 2091-2102.

Wade, H.K., T.N. Bibikova, W.J. Valentine, and G.I. Jenkins. 2001. Interactions within a network of phytochrome, cryptochrome and UV-B phototransduction pathways regulate chalcone synthase gene expression in Arabidopsis leaf tissue. The Plant Journal. 25(6): 675-685.

Wang, J.R., H. Hu, G.H. Wang, J. Li, J.Y. Chen, and P. Wu. 2009a. Expression of PIN genes in rice (Oryza sativa L.): tissue specificity and regulation by hormones. Mol. Plant. 2: 823-831.

Wang, Y., K. Li, and X. Li. 2009b. Auxin redistribution modulates plastic development of root system architecture under salt stress in Arabidopsis thaliana. J Plant Physiol. 166: 1637-1645.

Wang, L., M. Arnold, Y. Huang, C. Sardo, C. Seguin, E. Martin, T.H.M. Huang, K. Riedl, S. Schwartz, W. Frankel, D. Pearl, Y. Xu, J. Winston, G. Yang, and G. Stoner. 2011. Modulation of genetic and epigenetic biomarkers of colorectal cancer in humans by black raspberry: A phase 1 pilot study. Clin. Cancer Res. 17(3): 598-610.

Williams, R.J., J.P.E. Spencer, and C. Rice-Evans. 2004. Flavonoids: Antioxidants or signaling molecules? Free Radical Biology and Medicine. 36(7): 838-849.

Winkel-Shirley, B. 2001. Flavonoid biosynthesis. A colorful model of genetics, biochemistry, cell biology, and biotechnology. Plant Physiol. 126: 485-493.

Winkel, B.S.J. 2004. Metabolic channeling in plants. Ann. Rev. Plant Biol. 55: 85-107.

Wu, X., G. R. Beecher, J.M. Holden, D.B Haytowitz, S.E. Gebhardt, and R.L. Prior. 2006. Concentrations of anthocyanins in common foods in the United States and estimation of normal consumption. J. Agr. Food Chem. 54: 4069-4075.

Xiong, L., K.S. Schumaker, and J.K. Zhu. 2002. Cell signaling during cold, drought, and salt stress. Plant Cell. 165-183.

Xu, P., Y. Xiang, H. Zhu, H. Zhu, Z. Zhang, C. Zhang, Lixia, Zhang, and Z. Ma. 2009. Wheat cryptochromes: Subcellular localization and involvement in photomorphogenesis and osmtic stress responses. Plant Physiol. 149: 760-774.

Yamamoto, Y. and R. Mizuguchi. 1982. Selection of a high and stable pigment producing strain of Euphorbia millii cells. Theoretical Appl. Genet. 61: 113-116.

Yang, H., and A.S. Murphy. 2009. Functional expression and characterization of Arabidopsis ABCB, AUX 1 and PIN auxin transporters in Schizosaccharomyces pombe. The Plant Journal. 59: 179-191.

Yan, J.H., B.A. Wang, Y.N. Jiang, L.J. Cheng, and T.L. Wu. 2014. GmFNSII-Controlled Soybean Flavone Metabolism Responds to Abiotic Stresses and Regulates Plant Salt Tolerance. Plant and Cell Physiol. 55: 74-86.

Yanovsky, M.J., J.J. Casal, and J.P. Luppi. 1997. The VLF loci, polymorphic between ecotypes of Landsberg erecta and Columbia, dissect two branches of phytochrome A signal transduction that correspond to very-low-fluence and high-irradiance responses. The Plant Journal. 12(3): 659-667.

Yeo, A. 1998. Molecular biology of salt tolerance in the context of whole-plant physiology. J. Exp. Bot. 49: 915-929.

Younger, V.B. 1969. Physiology of growth and development, p. 187-216. In: Hanson A.A. and F.V. Juska (ed). Turfgrass Science. Amer. Soc. of Agron. Madison, WI.

Yu, C., C. Sun, C. Shen, S. Wang, F. Liu, Y. Liu, Y. Chen, C. Li, Q. Qian, B. Aryal, M. Geisler, A. Jiang de, and Y. Qi. 2015a. The auxin transporter, OsAUX1, is involved in primary root and root hair elongation and in Cd stress responses in rice (Oryza sativa L.). The Plant Journal. 83: 818-830.

Yue, R., S. Tie, T. Sun, L. Zhang, Y. Yang, J. Qi, S. Yan, X. Han, H. Wang, and C. Shen. 2015. Genome-wide identification and expression profiling analysis of ZmPIN, ZmPILS, ZmLAX and ZmABCB auxin transporter gene families in maize (Zea mays L.) under various abiotic stresses. PLoS One. 10: e0118751.

Zazimalova, E., A.S. Murphy, H.B. Yang, K. Hoyerova, and P. Hosek P. 2010. Auxin Transporters - Why So Many? Cold Spring Harbor Perspectives in Biology 2.

Zhang, J., J.E. Lin, C. Harris, F. Campos Mastrotti Pereira, F. Wu, J.J. Blakeslee, and W.A. Peer. 2016. DAO1 catalyzes temporal and tissue-specific oxidative inactivation of auxin in Arabidopsis thaliana. PNAS. 113: 11010-11015.

Zhang, K., H. Xu, T. Yuan, L. Zhang, and Y. Lu. 2013. Blue-light-induced PIN3 polarization for root negative phototropic response in Arabidopsis. The Plant Journal. 76: 308-321.

Zhang, Q., J. Li, W. Zhang, S. Yan, R. Wang, J. Zhao, Y. Li, Z. Qi, Z. Sun, and Z. Zhu. 2012. The putative auxin efflux carrier OsPIN3t is involved in the drought stress response and drought tolerance. The Plant Journal. 72: 805-816.

Zhang, X. and C. Liu. 2015. Multifaceted regulations of gateway enzyme phenylalanine ammonia-lyase in the biosynthesis of phenylpropanoids. Mol. Plant. 8: 17-27.

Zhang, Y., S. Zheng, Z. Liu, L. Wang, and Y. Bi. 2011. Both HY5 and HYH are necessary regulators for low temperature-induced anthocyanin accumulation in Arabidopsis seedlings. J. Plant Physiol. 168: 367-374.

Zhao, Y., S.K. Christensen, C. Frankhauser, J.R. Cashman, J.D. Cohen, D. Weigel, and J. Chory. 2001. A role for flavin monooxygenase-like enzymes in auxin biosynthesis. Science. 291.

Zhao, Z., Y. Zhang, X. Liu, X. Zhang, S. Liu, X. Yu, Y. Ren, X. Zheng, K. Zhou, L. Jiang, X. Guo, Y. Gai, C. Wu, H. Zhai, H. Wang, and J. Wan. 2013. A role for a dioxygenase in auxin metabolism and reproductive development in rice. Dev. Cell. 27: 113-122.

Zhao, YD, Hull AK, Gupta NR, Goss KA, Alonso J, Ecker JR, Normanly J, Chory J, Celenza JL. 2002. Trp-dependent auxin biosynthesis in Arabidopsis: involvement of cytochrome P450s CYP79B2 and CYP79B3. Genes & Development. 16, 3100- 3112.

Zhu, J.K. 2002. Salt and drought stress signal transduction in plants. Ann. Rev. Plant Biol. 53: 247-273.

Zhu, J.K. 2003. Regulation of ion homeostasis under salt stress. Curr. Op. in Plant Biol. 6: 441-445.

Zolla, G., Y.M. Heimer, and S. Barak. 2010. Mild salinity stimulates a stress-induced morphogenic response in Arabidopsis thaliana roots. J. Exp. Bot. 61: 211-224.

Chapter 2: Anthocyanin Production Using Rough Bluegrass Treated With High Intensity Light

Published September 2016: HortScience 51(9):1111–1120. 2016. doi: 10.21273/HORTSCI10878-16. Permission to re-publish granted 11/17/2016.

Abstract

Anthocyanins are plant pigments that are in demand for medicinal and industrial uses.

However, anthocyanin production is limited due to the harvest potential of the species currently used as anthocyanin sources. Rough bluegrass (Poa trivialis L.) is a perennial turfgrass known for accumulating anthocyanins, and may have the potential to serve as a source of anthocyanins through artificial light treatments. The objectives of this research were to determine optimal light conditions that favor anthocyanin synthesis in rough bluegrass, and to determine the suitability of rough bluegrass as a source of anthocyanins.

When exposed to high intensity white light, rough bluegrass increased anthocyanin content by 100-fold on average, and anthocyanin contents greater than 0.2% of dry tissue weight were observed in some samples. Blue light, at intensities between 150 – 250

µmol m-2 s-1, was the only wavelength that increased anthocyanin content. However, when red light was applied with blue light at 30 or 50% of the total light intensity, anthocyanin content was increased compared to blue light alone. Further experiments demonstrated that these results may be potentially due to a combination of

72 photosynthetic- and photoreceptor-mediated regulation. Rough bluegrass is an attractive anthocyanin production system since leaf tissue can be harvested while preserving meristematic tissues that allow new leaves to rapidly grow; thereby allowing multiple harvests in a single growing season and greater anthocyanin yields.

Introduction

Anthocyanins have become sought-after natural products due to potential for medicinal and industrial uses. These metabolites have a number of health promoting properties; increasing demand for nutraceuticals, fruits, and vegetables containing anthocyanins (Deroles, 2009; He and Giusti, 2010; Wrolstad and Culver 2012).

Production of textiles, cosmetics, and solar panels are examples of industrial applications where anthocyanins are also being increasingly used to replace synthetic dyes (Hao et al.,

2006; Wongcharee et al., 2007; Mansour et al., 2013).

The use of anthocyanin extracts for the above applications is limited by the small number plant sources of anthocyanins as well as the cultural limitations of these species: annual life cycle, slow growth, limited harvest, high input, etc. (Deroles, 2009). The use of plant tissue culture has been proposed as a means of large scale anthocyanin production; however, to date, these techniques have not been able to produce anthocyanins at levels sufficient to meet current industrial needs (Yamamoto et al., 1982;

Vogelien et al., 1990; Delgado-Vargas, 2000). One way to meet the increasing demand for anthocyanins is to employ non-conventional plant species, such as Poaceous grasses.

The anatomy and perennial nature of turfgrasses make them attractive anthocyanin production systems.

Turfgrasses accumulate anthocyanins in leaves that originate from meristematic tissues that sit at or below the soil surface (Christians, 2011). Leaf tissue can therefore be harvested while keeping meristematic tissues intact, allowing for year-round production from the same stand of plants. Cool-season turfgrasses (C3 photosynthetic) devote greater than 60% of photosynthate towards leaf and sheath growth, and the potential yield of anthocyanin containing tissue could be upwards of 3 Mg ha -1 following a single harvest (Younger, 1969; Krans and Beard, 1980; Landschoot and Waddington, 1987).

Relative to fruit crops, currently the most used anthocyanin source, turfgrasses provide numerous advantages. For example, turfgrasses could be harvested for anthocyanins within weeks of seeding, and leaf tissue could be harvested at least once per month.

Further, since turfgrasses do not undergo secondary growth, a greater proportion of photosynthate could be devoted towards anthocyanin synthesis.

Rough bluegrass (Poa trivialis L.) is known to constitutively produce the anthocyanins cyanidin-3-glucoside and cyanidin-3-malonylglucoside in the leaf sheath

(Fossen et al., 2002; Hurley, 2010). This turfgrass is a fast growing perennial under field conditions, and high tissue yield could therefore be expected (Atkin et al., 1996). Still, to employ this species as an industrial crop, it would be necessary to increase anthocyanin production in rough bluegrass to levels greater than those currently observed in the field.

Environmental stress, light in particular, is one factor that is known to increase anthocyanin synthesis (Boldt et al., 2014).

Transient anthocyanin accumulation occurs with changes in light quantity and/or quality, and has been documented in a variety of plants including: tomato, Arabidopsis, maize, sorghum, rye, red cabbage, and mustard (Mancinelli, 1985; Mol et al., 1996;

Chalker-Scott, 1999). Light is a requirement for anthocyanin synthesis, and anthocyanin production is photoregulated (Downs and Siegelman, 1962; Lange 1971; Rabino et al.,

1977; Mancinelli, 1985; Vyas, 2014). Photomanipulation could therefore be used to increase anthocyanin content in rough bluegrass.

Phytochrome has been shown to regulate anthocyanin synthesis through the absorption of red or far-red light (Mancinelli, 1985; Kerchoffs and Kendrick, 1997; Neff and Chory, 1998; Wade et al., 2001; Oh et al., 2014). However, blue light also regulates anthocyanin synthesis, although this is accomplished through the activity of multiple photoreceptors, including both cryptochromes and phototropins (Galvão and

Frankhauser, 2015). Cryptochromes are well known to regulate anthocyanin synthesis, whereas phototropins have only been recently implicated in anthocyanin regulation

(Poppe et al., 1998; Vandenbussche et al., 2007; Hong et al., 2009; Fox et al., 2012;

Kadomura-Ishikawa et al., 2013; Folta and Carvalho 2015).

Treatment with red and blue light may also increase anthocyanin synthesis through photoreceptor co-action. In other words, red light may be non-inductive on its own, but when applied with blue light, anthocyanin synthesis may be increased compared to blue light alone (Drumm and Mohr, 1978; Mohr and Drumm-Herrel, 1983; Mancinelli,

1985; Wade et al., 2001). Additionally, anthocyanin synthesis has also been shown to be regulated through photosynthesis and increased under high intensity light (Schneider and

Stimson, 1971; Mancinelli et al., 1976; Mancinelli and Rabino, 1978; Mancinelli, 1985;

Weiss and Halevy, 1991 Kumar Das et al., 2011). Therefore, anthocyanin content may increase through a combination of photosynthetic- and photoreceptor- regulation when blue, red, and/or combinations of blue and red light are applied.

Given the previously established regulation of anthocyanin synthesis in several monocot crops (i.e., sorghum and maize), we hypothesized that anthocyanin production in rough bluegrass could be manipulated through exposure to specific light regimes. The objectives of this research were to determine conditions that favor anthocyanin synthesis in rough bluegrass by first evaluating whether or not treatment with high intensity light could increase anthocyanin content. Second, the wavelength(s) of light capable of upregulating anthocyanin synthesis was determined to optimize light conditions. Finally, the role of photosynthesis on anthocyanin production in rough bluegrass was evaluated.

Materials and Methods

Metal halide lighting

Phillips advance high intensity discharge (HID) 400 W ballasts were used for full spectrum lighting experiments, along with metal halide (400 W) lamps (Phillips

Advance® Rosemont IL, USA). For growth chamber experiments, ballasts were kept outside of the growth chamber, and wiring was passed through front-mounted instrument ports to allow for lamp mounting inside of the growth chamber. Light intensity was

76 measured using a LI-COR quantum sensor and LI-2189 light meter (LI-COR®

Biosciences, Lincoln NE USA).

Light Emitting Diode (LED) array construction

Blue and red 5050 LEDs (5.0 x 5.0 mm) were purchased pre-mounted on flexible metal-core printed circuit strips - 60 LEDs m-1 (Torchstar Inc. ® La Puente, CA, USA).

LED strips were cut into lengths of 40 cm and were attached side by side to sheets of cardboard (150 x 60 x 0.64 cm) using 3M® 8805 thermally conductive tape (3M®

Medina, OH, USA). LED strips were wired in parallel, and were powered using a 12 V

AC-DC switching power supply (LED Wholesalers® Hayward, CA, USA). For experiments combining red and blue light, LED strips were wired to separate dimming units (LED Wholesalers® Hayward, CA, USA) to allow for adjustment in spectral quality

(Appendix B and L).

High power, 10 W far-red (FR) LEDs (LED Engin® Inc., San Jose, CA, USA) were mounted on aluminum sheet metal (150 x 60 x .16 cm) using thermally conductive tape (Appendix I). Four FR LEDs were wired in parallel with a 15 W 4.7 Ω wire wound cement filled resistor (Xicon® Passive Components, Arlington TX, USA) soldered in between each negative terminal. Lenses were attached to induvial LEDs (LED Engin®

High Uniformity Lighting Lens) using thermal adhesive (Arctic Silver® Inc. Visalia, CA,

USA). Four groups of parallel wired FR LEDs (16 total LEDs) were used for each array, and each group was powered using an AC-DC switching power supply. LED arrays for all experiments were constructed in three replicates. LED arrays were mounted within

Conviron E15 growth chambers set to 15°C (Controlled Environment Ltd., Winnipeg,

Canada) by hanging them underneath a pre-existing adjustable light rack. Ventilation fans, 425 m3 h-1 or greater, were placed on top of the light rack and provided a constant flow of air over the LED arrays.

Light measurement

LED array spectral distribution and quantum flux were measured using a

Stellarnet® BLACK- Comet spectroradiometer, a CR2 cosine receptor, and spectra were analyzed using SpectraWiz® software (Stellarnet® Inc., Tampa, FL, USA). Blue LEDs produced a bandwidth of 400-525 nm with a peak wavelength of 453 nm (3.38% of the spectrum) (Appendix C and D). Red LEDs produced a bandwidth of 550-700 nm with a peak wavelength of 635 nm (4.61% of the spectrum) (Appendix E and F). Far-red LEDs produced a bandwidth of 670-800 nm with a peak wavelength of 732 nm (3.13% of the spectrum) (Appendix G and H).

Light intensity was controlled through LED dimming, and by adjusting the distance between the plant canopy and the LED array. When light intensity and distance were set, canopy temperature was measured using a Kestrel 3000 weather meter (Kestrel

Meters, Birmingham, MI, USA). On average, the canopy temperatures under an LED array consisting of 100% red light were 20.5 ± 2.7°C, under 100% blue light they were

20.6°C ± 2.5°C, and under dichromatic arrays (i.e. 50% red and blue light) they were

20.4°C ± 2.5°C (not statistically different when compared using Student’s T-test; P =

.05).

Plant materials and growth conditions

Rough bluegrass, cv. ‘Havana’ (J.R. Simplot Company, Boise, ID, USA), plants were grown in 2.5 x 12 cm Cone-tainers™ (Stuewe and Sons, Tangent, OR, USA) using soilless media (Farfard® 3B Mix, Sungro® Horticulture, Agawam, MA, USA). However for experiment one, plants were grown to maturity (approximately 3 months old) in standard 15 x 11 cm plastic pots using a seeding rate of 1.8 kg (seed) 93 m-2 according to the methods of Nangle et al. (2015). Mature plants used in experiments other than experiment one were grown under greenhouse conditions, and were maintained as single plants (tillers removed) per Cone-tainer™ rather than a stand of multiple plants.

Seedlings seven days past germination (DPG) were also used in experiments two through five. Light grown seedlings were grown within a Conviron E15 growth chamber: 12 hour photoperiod, 200 µmol m-2 s-1, and 20°C day/night temperature. Plants were watered daily, and were not fertilized. Dark grown seedlings were also grown under the same growth chamber conditions by placing Cone-tainers™ under four layers of black polyester cloth. Dark conditions were verified using a LI-COR quantum sensor and LI-

2189 light meter. Two to three seedlings were grown per Cone-tainer™.

Pigment extractions and analysis

Chlorophylls were extracted using N, N- Dimethylformamide (DMF), analyzed, and spectrophotometrically quantified (Shimadzu UV1800 spectrophotometer, Shimadzu

Corp., Colombia, MD) according to the methods of Wellburn (1994). Fresh leaf tissue was weighed into a glass test tube, 10 mL of DMF was added, and samples were allowed

79 to incubate for 24 hours in the dark 4°C. Following extraction an aliquot of extract was analyzed for chlorophyll content spectrophotometrically. Total anthocyanins were extracted using 1% acidified methanol (HCl). Fresh shoot tissue was weighed and extracted in 10 mL of extraction solvent for 24 hours in the dark at 4°C (Rabino and

Mancinelli 1986). Aliquots were analyzed spectrophotometrically, and relative anthocyanin content was estimated using the following equation: (Abs.530 – .25 Abs.657) g-1 FW (Rabino and Mancinelli, 1986).

Cyanidin-3-glucoside equivalents were determined using the pH differential method (Giusti and Wrolstad, 2001). Tissue extracts for the pH differential method were prepared as previously described (Nangle et al., 2015). Freeze dried tissue was ground in liquid nitrogen, and was suspended in 10 mL of 100% acetone. Samples were vortexed for 30 seconds, and were centrifuged at 10,000 G for 5 minutes. The supernatant was collected, and the pellet was re-extracted two more times in 70% (v/v) aqueous acetone containing 1% (v/v) HCl. All extracts were pooled, and vacuum filtered. The filtered extract was phase separated in a sepratory funnel using a 2:1 ratio of chloroform to extract, and was allowed to incubate for 24 hours in the dark at 4°C. After being incubated the aqueous layer was collected, and any remaining acetone was removed under reduced pressure at 40°C for 5-10 minutes. The remaining extract was brought to a known volume using 0.1% (v/v) acidified water, and was used to determine total monomeric anthocyanin content. The pH differential method was performed by combing a known volume of the resultant extract in 1) KCl buffer (pH 1.0), and by combining a known volume of extract in 2) sodium acetate buffer (pH 4.5). The solutions were

80 allowed to stand for 10 minutes and absorption at 510 and 700 nm was monitored using a spectrophotometer (Shimadzu UV1800 spectrophotometer, Shimadzu Corp., Colombia,

MD). Cyanidin-3-glucoside equivalents were calculated using a previously defined formula: 1) Absorbance = (A510 – A700) pH 1.0 – (A510 – A700) pH 4.5; 2) Anthocyanin content

(mg L-1) = (Absorbance x 449.2 x Dilution Factor x 1000) / (26,900) (Giusti and

Wrolstad, 2001).

To identify the anthocyanins present, the extract from above was purified for

HPLC and LC-MS analysis. Briefly, C-18 solid phase extraction (SPE) cartridges

(Biotage, Charlotte, NC) were conditioned on a vacuum manifold with two volumes of methanol and three volumes of 0.01% acidified (HCl) water. The extract was passed through the column, was washed using two volumes of 0.01% (v/v) acidified (HCl) water, and was washed with a half volume of ethyl acetate. Anthocyanins were then eluted using 0.01% (v/v) acidified (HCl) methanol. The elutant was then concentrated under reduced pressure at 40°C, was re-suspended in 0.01% (v/v) acidified (HCl) water, and was filtered using a 0.22 µm syringe filter. Aliquots of rough bluegrass extract were saponified to determine if anthocyanins present were acylated using previously described methods (de-Pascual-Teresa et al., 2002). A portion of sample was combined with 10%

KOH until the color of solution was blue, the tube was filled with nitrogen gas, and was allowed to stand for 10 minutes. Following incubation, concentrated HCl was added dropwise until the extract was red. The saponified sample was then re-purified using the

SPE methods described above.

The following method was used for anthocyanin identification. A Shimadzu

2010EV LC-ESI MS coupled with a Shimadzu SPD-M20A photodiode array (PDA) was used to identify anthocyanins. A Varian C18-A (150 mm x 4.6 mm I.D.) column was used (Varian Inc., Lake Forest, CA). A binary flow mobile phase consisting of (A) 4.5% aqueous formic acid and (B) 100% acetonitrile with a flow rate of 0.8 mL min-1 was used, and 20 µL of sample was injected onto the column. Anthocyanins were separated using the following gradient: 12% B 1-25 minutes, 35% B 25-30 minutes, a 35%-12% decrease in B from 30-38 minutes, and 12% B until minute 45. UV-vis spectral profiles, retention times, and mass fragmentation patterns were compared to those previously published for anthocyanin identification (Giusti and Wrolstad 2001; de-Pascual-Teresa et al., 2002;

Nangle et al., 2015). A purple corn extract was also run under the same conditions to compare retention times and absorption spectra to the rough bluegrass sample (de-

Pascual-Teresa et al., 2002).

Experimental design, setup, and statistics

Five separate experiments were performed to determine the role of light in upregulating anthocyanin content in rough bluegrass. Experiment one was designed to evaluate whether or not high intensity white light could increase anthocyanin content in a rough bluegrass. Six pots of mature rough bluegrass (15 x 11 cm - 1.8 kg seed 93 m-2) were constantly exposed to 1000 µmol m-2 s-1 of light from two 400 W metal halide lamps placed 61 cm above the plant canopy for five days. Following treatment, leaf tissue was harvested down to the crown of the plant, and analyzed for cyanidin-3-

82 glucoside equivalents. All pots of rough bluegrass were then transferred back to the greenhouse, and leaves were re-grown to 3 cm. Rough bluegrass plants were then treated under high light conditions again for replication over time, and to simulate a potential production system. Using these methods, experiment one was replicated three times in total. For a given replicate, the six rough bluegrass samples treated were extracted and analyzed separately, and resulting cyanidin-3-glucoside equivalents were averaged for statistical analysis. One growth chamber was used for high light treatment in this experiment, and the experiment was rotated to a new growth chamber for each replicate.

Plants grown under high light were compared to untreated plants that were grown under standard greenhouse conditions.

Experiment two was designed to determine what wavelengths of light were able to upregulate anthocyanin production. Light and dark grown seedlings (7 DPG) were exposed to either 200 µmol m-2 s-1 blue, red, or far-red LED light constantly for five days.

Dark grown seedlings were used as an added treatment alongside light grown seedlings because tissues exhibiting lower quantities of chlorophyll a and b may exhibit different patterns of anthocyanin accumulation (Hughes et al., 2007). In experiment three, an anthocyanin dose response curve was generated using a range of blue light intensities.

Light grown seedlings (7 DPG) were constantly exposed to five days of the following blue LED light treatments: 25, 50, 100, 150, 200, 250, 300, and 350 µmol m-2 s-1 of blue light. Experiment four was designed to evaluate co-action between blue and red light by altering spectral quality. Mature (3 month old) and light grown seedlings (7 DPG) were treated for five days with 200 µmol m-2 s-1 of constant LED light consisting of the

83 following blue/red spectral distributions: 0% blue (100% red), 30% blue (70% red), 50% blue (50% red), 70% blue (30% red), or 100% blue (0% red).

Experiment five was designed to evaluate the effects of photosynthesis on anthocyanin synthesis by altering spectral distribution, light intensity, and photosynthetic depression using the photosystem II inhibitor amicarbazone (4-Amino-N-(tert-butyl)-3- isopropyl-5-oxo-4,5-dihydro-1H-1,2,4-triazole-1-carboxamide). Amicarbazone was applied at a concentration of 2.8 mg L-1 using the commercially available product

Xonerate (70% amicarbazone by wt., Arysta LifeScience©, Cary, NC, USA) 4 hours prior to light treatment using an atomizing spray applicator (Dayan et al., 2009). Light and dark grown seedlings (7 DPG) were treated for five days with 170 µmol m-2 s-1 blue light plus 0, 10, 25, 50, or 100 µmol m-2 s-1 red light using dimmable LED arrays. The experiment was also performed in reverse using 170 µmol m2 s-1 red light plus 0, 10, 25,

50, or 100 µmol m-2 s-1 blue light. Experiments two though five had a total of six plant samples per treatment that were pooled for analysis.

Plants were dark adapted for 24 hours prior to light treatments, and were watered daily. Growth chamber doors were open on average for five minutes per day, and light intensity outside of the growth chamber was approximately 3 µmol m-2 s-1. For experiments using more than one LED array treatment, LED arrays were separated in each growth chamber using cardboard sheets covered in aluminum foil.

Spectroradiometeric measurements were taken before and after the separation sheet was installed to verify that light from neighboring treatments was not being transmitted or interfering. All experiments were analyzed as randomized complete block designs with

84 three replicates. Separate Conviron E15 growth chambers served as blocking units, and three separate chambers were used simultaneously for experiments other than experiment one (replicated over time). All statistics were performed using Minitab software v. 17.

Data were analyzed using General Linear Model ANOVA (GLM ANOVA), and Tukey’s honest significant difference (HSD) test was used for mean separation (P =.05).

Results

Anthocyanin upregulation under high intensity white light

When exposed to constant high intensity white light (1000 µmol m-2 s-1 - metal halide lighting) rough bluegrass plants significantly increased anthocyanin concentration compared to untreated plants (Table 2.1) (Appendix M). In untreated plants, the low levels of anthocyanins present were localized primarily to sheath tissue, a trend similar to what has been previously observed in field grown rough bluegrass (Hurley, 2010).

Light treatment Cyanidin-3-glucoside eq. (mg 100 g -1 DW ) Avg. fold difference

z Untreated 1.32 ± 0.64 N/A

y HID metal halide 155.14 ± 88.45*** 117.64 ± 67.07 z Mean ± Standard deviation (n = 3) y Statistical significance (P = .05) determined using a 2-sample t-test (* = P ≤ 0.05; ** = P ≤ 0.01; *** = P ≤ 0.001). DW = dry weight

Table 2.1: Anthocyanin concentration (cyanidin-3-glucoside eq.) of rough bluegrass following light treatment. 85

Light treated plants, however, exhibited an average 117.64 fold increase in anthocyanin content, and accumulated anthocyanins in both leaf blades and sheath tissue

(Table 2.1). These increases were independent of the growth stage of the tissue, and were observed in plants of all ages and developmental stages within the stand. Anthocyanins were identified via LC-MS, and the presence of cyanidin-3-glucoside and cyanidin-3- malonylglucoside were confirmed based on comparison of mass fragmentation data

(Table 2.1), retention times, and UV-vis spectral profiles (Fig. 2.1) to a purple corn standard containing both cyanidin-3-glucoside and cyanidin-3-malonylglucoside, as well as comparison to previously published retention times and spectral profiles for turfgrasses

(de Pascual-Teresa et al., 2002; Nangle et al., 2015).

In rough bluegrass tissue, cyanidin-3-glucoside accounted for 12.7% of the total peak area (UV-vis 520 nm) and cyanidin-3-malonylglucoside accounted for 89.3% of the total peak area (Table 2.2). To further confirm the presence of cyanidin-3- malonylglucoside, the acyl group was removed through saponification, and the corresponding release of cyanidin-3-glucoside was measured using LC-MS (Fig. 2.1).

Comparing chromatograms of rough bluegrass, saponified rough bluegrass, and purple corn confirmed mass spectrometry results through compound retention time (Fig. 2.1A-

C). UV-vis spectral profiles of both cyanidin-3-glucoside and cyanidin-3- malonylglucoside from rough bluegrass and purple corn also exhibited the same spectral profiles and maxima (Fig. 1D-G).

Wavelength specificity

To determine the primary wavelength(s) of light responsible for upregulating anthocyanin synthesis in rough bluegrass, dark grown and light grown rough bluegrass seedlings were exposed to blue, red, and far-red LED light. Analysis of chlorophyll content showed that seedlings grown under dark conditions had significantly lower concentrations of chlorophyll a, while chlorophyll b content was not different between light and dark grown seedlings (Fig. 2.2A). These results are reflected in the chlorophyll a:b ratios calculated for light and dark grown seedlings. Dark grown seedlings had a significantly lower chlorophyll a:b ratio compared to light grown seedlings (Fig. 2.2B).

When exposed to 200 µmol m-2 s-1 of blue, far-red, and red LED light, only blue light increased anthocyanin content in light grown seedlings (Fig. 2.2C). Likewise, dark grown seedlings only showed a significant increase in anthocyanin content when exposed to blue light (Fig. 2.2D). For both dark grown and light grown seedlings, anthocyanins visually accumulated in mesocotyl, sheath, and leaf tissues. Interestingly, in dark grown seedlings a majority of pigmentation was concentrated in the mesocotyl region, a trend that was not observed in light grown seedlings.

Figure 2.1: High performance liquid chromatography (HPLC) chromatograms of rough bluegrass (RB); cyanidin-3-glucoside; Cy3g (1), cyanidin-3-malyonlyglucoside; Cy3mg

(2). Rough bluegrass anthocyanins; B) saponified rough bluegrass anthocyanins; and C) purple corn (PCorn) anthocyanins. Spectral profiles of cyanidin-3-glucoside from D) rough bluegrass and E) purple corn exhibited the same profile and maxima (279 and 517 nm). Spectral profiles of cyanidin-3-malonylglucoside from F) rough bluegrass and G) purple corn exhibited the same profile and maxima (279 and 519 nm). Anthocyanins were identified using mass spectrometry data (Table 2.2) along with retention times and

UV-vis absorption spectral profiles. 88

Blue light dose response

Light grown seedlings were treated with blue LED light ranging in intensity from

25 – 350 µmol m-2 s-1, and results showed that anthocyanins did not begin to accumulate in high quantities until plants were exposed to 150 µmol m-2 s-1 blue light (Fig. 2.3). We determined that the threshold blue light intensity needed to stimulate anthocyanin synthesis in rough bluegrass lies somewhere between 150 and 250 µmol m-2 s-1. Levels of anthocyanin observed following exposure to 150 µmol m-2 s-1 of blue light were not statistically different than those observed at 100 or 200 µmol m-2 s-1.

Further, anthocyanin levels at 200 µmol m-2 s-1 were not statistically different from those seen at 250 µmol m-2 s-1. Taken together, these data indicate that anthocyanin production is stimulated at blue light intensities between 150 and 250 µmol m-2 s-1. Our data also indicate that the response begins to saturate above 250 µmol m-2 s-1 of blue light

(Fig. 2.3).

z Peak + y x w Peak area (520nm) v Identity Tr (min) [M] Mass Fragment UV-vis (nm)

1 6.41 449 287 279,517 12.7% Cy3g

2 9.86 535 287 279,519 89.3% Cy3mg z Retention time y Molecular ion x Aglycone fragment w Peak absorption of compounds in UV and PAR regions v Percentage of anthocyanin identified based on peak area of UV-vis (520nm) chromatogram Cy3g = Cyanidin-3-glucoside Cy3mg = Cyanidin-3-malyonlyglucoside

Table 2.2: Liquid chromatography – mass spectrometry (LC-MS) identification of anthocyanins accumulated in rough bluegrass following light treatment.

Co-action between blue and red light

To determine whether or not red light could modulate the blue light response, light grown seedlings were exposed to 200 µmol m-2 s-1 LED light treatments consisting of 0% blue (100% red), 30% blue (70% red), 50% blue (50% red), 70% blue (30% red), or 100% blue (0% red). Applications of blue and red light together increased anthocyanin synthesis compared to blue light alone (Fig. 2.4).

(control), blue, far-red, or red LED light (200 µmol m2 s-1). Relative anthocyanin

-1 concentration = (Abs.530 - .25 Abs.657) g FW. Error bars represent treatment standard error (SE), and letters represent mean separation using Tukey’s HSD (n = 3). Different letters indicate a significant statistical difference (P = .05).

Specifically the addition of 30% or 50% red light significantly increased anthocyanin content compared to 100% blue light in light grown seedlings (Fig. 2.4A).

Experiments were repeated using mature rough bluegrass plants (3 month old), and the results showed a similar trend. In the case of mature plants however, only the addition of

30% red light increased anthocyanin content compared to 100% blue light (Fig. 2.4B).

Photosynthetic contribution to anthocyanin synthesis

Dark grown and light grown seedlings were exposed to LED light treatments with a fixed quantity of blue light (170 µmol m-2 s-1), and increasing quantities of red light (0,

10, 25, 50, or 100 µmol m-2 s-1). A subset of rough bluegrass seedlings were also treated with the photosystem II inhibitor amicarbazone. For light grown seedlings, anthocyanin content increased as the quantity of red light increased (Fig. 2.5A). The addition of 100

µmol m-2 s-1 of red light led to a significant increase in anthocyanin concentration compared to all other treatments, suggesting that the response may be due to an increase in photosynthesis rather than the involvement of phytochrome.

When light grown plants were treated with amicarbazone, anthocyanin synthesis saturated at 25 µmol m-2 s-1, supporting a potential role for red light in increasing anthocyanin content through photosynthesis (Fig. 2.5B). Results for dark grown seedlings showed a similar trend to those observed in light grown seedlings.

Anthocyanin content increased in a linear fashion with increasing red light intensity, and the addition of 100 µmol m-2 s-1 led to a significant increase in anthocyanin concentration compared to 0, 10 and 25 µmol m-2 s-1 red light treatments (Fig. 2.5C). Following

92 treatment with amicarbazone, anthocyanin content saturated at 25 µmol m-2 s-1 and exhibited a sharp decline with increasing quantities of red light, again suggesting photosynthetic regulation (Fig. 2.5D). In reverse experiments, when seedlings were exposed to a fixed amount of red light (170 µmol m-2 s-1) and increasing blue light (0,

10, 25, 50, 100 µmol m-2 s-1), light grown seedlings only increased anthocyanin synthesis when exposed to an additional 100 µmol m-2 s-1 of blue light (Fig. 6A).

Figure 2.3: Relative anthocyanin concentration of light grown rough bluegrass seedlings treated with blue LED light of increasing intensity. Plants were treated using the following light intensities: 25, 50, 100, 150, 200, 250, 300, or 350 µmol m2 s-1. Relative

-1 anthocyanin concentration = (Abs.530 - .25 Abs.657) g FW. Error bars represent treatment standard error (SE), and letters represent mean separation using Tukey’s HSD

(n = 3). Different letters indicate a significant statistical difference (P = .05).

The application of amicarbazone did not change this trend when applied, 100

µmol m-2 s-1 blue light increased anthocyanin production, although overall concentration was reduced (Fig. 2.6B). This indicates that while photosynthesis may impact the degree to which blue light stimulates anthocyanin production, blue light alone is sufficient to induce anthocyanin synthesis. For dark grown seedlings, the addition of 100 µmol m-2 s-1 blue light led to an increase in anthocyanin content compared to all other LED treatments

(Fig. 2.6C). Similar to other treatments, when amicarbazone was applied to dark grown seedlings, only 100 µmol m-2 s-1 treatments accumulated anthocyanins in larger quantities

(Fig. 2.6D). These results indicate that 100 µmol m-2 s-1 of blue light is necessary for the response to occur, and the addition of amicarbazone does not eliminate or reduce the response, as was observed in experiments with increasing red light.

Discussion

The objectives of this research were to determine artificial light conditions capable of inducing anthocyanin synthesis in rough bluegrass in order to evaluate the suitability of this turfgrass as a source of anthocyanin. Using high intensity white light, rough bluegrass was able to increase anthocyanin content upwards of 0.06% fresh weight, or 64 mg 100 g -1 FW (0.24% dry weight, or 243 mg 100 g -1 DW). Previous research has shown that cyanidin-3-malonylglucoside is the primary anthocyanin produced by rough bluegrass under field conditions, and results here show that this is also true under high intensity white light treatment (Fossen et al., 2002). For fruits and vegetables, anthocyanin content has been shown to range from 0.1 – 1% of dry weight (Delgado-

Vargas et al., 2000). Our data therefore show that the anthocyanin content of rough bluegrass following high light treatment is comparable to many common fruits and vegetables; particularly red leaf lettuce, where cyanidin-3-malonylglucoside is also the primary anthocyanin produced (Delgado-Vargas et al., 2000; Giusti and Wrolstad, 2001;

Wu et al., 2006).

Figure 2.4: Relative anthocyanin concentration of light grown seedlings (A) and mature

(3 month old) rough bluegrass (B) plants. Relative anthocyanin concentration = (Abs.530

-1 - .25 Abs.657) g FW. Error bars represent treatment standard error (SE), and letters represent mean separation using Tukey’s HSD (n = 3). Different letters indicate a significant statistical difference (P = .05).

-1 Abs.657) g FW. Error bars represent treatment standard error (SE), and letters represent mean separation using Tukey’s HSD (n = 3). Different letters indicate a significant statistical difference (P = .05).

A major advantage of using turfgrass, e.g. rough bluegrass, for anthocyanin production is the ability to harvest leaves containing anthocyanin numerous times throughout a single growing season. Results from experiment one show that rough bluegrass plants maintain the ability to synthesize large quantities of anthocyanin over time, even following aggressive harvesting. A conservative estimate of yield from one harvest, based on the amount of tissue harvested from experiment one in this study, would be 6 Mg (FW) ha -1. Because of the aggressive growth of this species, it is reasonable to assume anthocyanin containing leaves could be harvested once per month under a moderate input management system if the methods described were used in an indoor vertical farm. Using these figures, we calculate that approximately 72 Mg (FW) ha -1 extractable tissue could be produced annually, with an estimated anthocyanin yield of 45 kg. Furthermore, since anthocyanin concentrations of 90 mg 100 g FW -1 were observed following high light treatment of rough bluegrass (Table 2.1), potential anthocyanin yields of up to 65 kg per year may be possible.

Blackberry, blueberry, and grape generally produce greater concentrations of anthocyanin compared to rough bluegrass on average. However, relatively low fruit yield

(compared to the mass of turfgrass tissue that could be harvested) ultimately results in less anthocyanin per hectare in the same time frame (Giusti and Wrolstad, 2001; de

Pascual-Teresa 2008; USDA 2015). For example, on average blackberry may yield approximately 20 kg anthocyanin per year, blueberry 18 kg, and grape 66 kg relative to the potential 45-65 kg from rough bluegrass (Giusti and Wrolstad, 2001; de Pascual-

Teresa 2008; USDA, 2015). Some tropical monocots, particularly palm trees such as

Euterpe species, may also produce greater quantities and more diverse anthocyanins compared to rough bluegrass (Del Pozo-Insfran et al., 2004). However, these palm trees only grow in very specific environments, they produce limited quantities of anthocyanin containing fruit, and only produce anthocyanins in a small portion of the plant (i.e. the fruit).

While blueberry and grape produce high levels of anthocyanin, the specific anthocyanin compounds in these species are different from those produced by rough bluegrass. As specific anthocyanins are often used in particular industries, rough bluegrass-derived anthocyanins could be used to meet industry specific needs. Red leaf lettuce has been demonstrated to produce a complement of anthocyanins similar to that of bluegrass (i.e. cyanidin-3-malonylglucoside), but yield data for this species is lacking; although the few published reports indicate that anthocyanin levels are substantially lower that what was found in rough bluegrass (2.2 mg 100 g-1 FW from lettuce vs. 64 mg

100 g -1 FW from rough bluegrass) from our study (Wu et al., 2006).

Additionally, other perennial grasses may also have the potential to serve as anthocyanin production systems; including varieties of pigmented ornamental grasses

(Pennisetum setaceum, Panicum virgatum, Miscanthus sinensis, etc.). Currently however, data regarding anthocyanin yield in grass species is limited. Interestingly, red varieties of switchgrass (Panicum virgatum) have been shown to produce anthocyanins in quantities ranging from 60 – 200 mg 100 g-1 FW indicating that this grass may also serve as a potential source of anthocyanin (Boldt, 2013).

Blue light is necessary and sufficient to increase anthocyanin synthesis in rough bluegrass. While red light is not required for anthocyanin synthesis in this species, treatment with blue and red light increased anthocyanin accumulation compared to blue light alone and exhibited a co-action response. However, the results of experiment four

(Fig. 2.4) showed no statistical difference in anthocyanin production between plants treated with 100% blue or 100% red light, and this may be primarily the result of variability in “resting levels” of anthocyanins present in plants prior to LED treatments.

Interestingly, these results indicate that growth conditions of rough bluegrass prior to light treatment (e.g., light conditions, temperature, water status, nitrogen availability, etc.) can impact light-induced anthocyanin production.

Combined treatment with blue and red light also decreased the threshold of blue light needed for anthocyanin synthesis to occur in experiments four and five. When 50%

(100 µmol m-2 s-1 blue light) and 70% (140 µmol m-2 s-1 blue light) blue light treatments were applied (i.e., a total of 200 µmol m-2 s-1 combined blue and red light), there was an evident increase in anthocyanin content compared to 100% blue light (Fig. 2.4). Both of these treatments induced anthocyanin synthesis at quantities of blue light that were lower than that what was shown to be required from our dose response experiment (Fig. 2.3 - between 150-250 µmol m-2 s-1). These data show that red light modulates the blue light threshold required to induce anthocyanin synthesis. Previous research suggests that this co-action effect could be mediated through phytochrome; however, our data suggest that the red light induced modulation is primarily through photosynthesis (Schneider and

Stimson, 1971; Kumar Das et al., 2011; Oh et al., 2014).

100

As this increase in anthocyanin content was saturated by the addition of amicarbazone in experiment five it is likely that the increase in accumulation observed when red light and blue light are applied together (Fig. 2.5 and 2.6) is due to photosynthetic regulation. Loss of anthocyanin synthesis following application of diuron

(DCMU – a PSII inhibitor related to amicarbazone) has been previously shown in

Arabidopsis, petunia, and maize (Schneider and Stimson, 1971; Weiss and Halevy, 1991;

Jeong et al., 2010; Kumar Das et al., 2011). For Arabidopsis in particular, the redox state of the plastaquinone (PQ) pool has been shown to be a potential signal for anthocyanin synthesis; diuron oxidizes the PQ pool and decreases anthocyanin content, while the inhibitor dibromothymoquinone (DPMIB) reduces the PQ pool and does not change anthocyanin content (Jeong et al., 2010). For rough bluegrass, if treatment with red light were impacting anthocyanin synthesis through modulation of phytochrome signaling, increases in anthocyanin synthesis would be expected to saturate at lower light intensities, which was not seen here. Our results show that anthocyanin content increases with increasing intensities of red light applied with constant blue light, indicating that a co-action response is acting primarily through photosynthetic and cryptochrome signaling.

Our data also suggest that dark grown plants may also exhibit increased anthocyanin content compared to light grown plants, although the statistical significance of this trend was not clear. Dark grown seedlings, however, showed a relatively large decline in anthocyanin content when treated under 170 µmol m-2 s-1 blue light in conjunction with simultaneous treatment with amicarbazone and either 50 or 100 µmol

101 m-2 s-1 of added red light (Fig. 2.5). Dark grown plants are more susceptible to both light stress and inhibition of photosynthesis by amicarbazone due to immature photosystems.

Generally, photosynthetically impaired tissues accumulate greater quantities of anthocyanin, and we hypothesize that the addition of amicarbazone to these light sensitive seedlings during excessive light treatment led to irreversible photodamage resulting in impairment of signaling (Chalker-Scott, 1999; Boldt, 2013).

In experiment two, seedlings accumulated greater quantities of anthocyanin compared to mature plants, which may be due to differences in quantities of chlorophyll a and b present in immature plants that compete for similar wavelengths of light (Merzylak et al., 2009). The photosynthetic machinery of seedlings is not fully established, and may require increased photoprotection through the accumulation of anthocyanins (Chalker-

Scott, 1999; Boldt, 2013). Additionally, when the levels of anthocyanin in light grown seedlings are compared across experiments, anthocyanins from plants treated under LED light never reached or exceeded levels of anthocyanins quantified in plants treated under high intensity metal halide lighting.

The decreased anthocyanin content of LED-treated rough bluegrass may result from the fact that plants grown under metal halide lamps were not only exposed to increased light intensity compared to LED treatments, but were also exposed to increased canopy temperatures and mild drought stress due to the heat generated by the lamps. It is well known that low temperature and drought stresses both have the ability to individually increase anthocyanin content in many plant species, including turfgrasses

(Mol et al., 1996; Chalker-Scott, 1999; Hughes et al., 2013; Boldt et al., 2014). While

102 high temperatures are generally associated with degradation and loss of anthocyanin over time, rather than long term increases in quantity; high temperatures are also known to be causative agents of photoinhibtion and may therefore be potentially increasing short term anthocyanin synthesis through photosynthesis (Rabino and Mancinelli, 1986; Yamane et al., 2006; Takahashi and Murata, 2008; Lin-Wang et al., 2011).

Our results showed that high intensity metal halide lighting significantly increased anthocyanin content, and that wavelength specific light treatments using LEDs may also increase the efficiency of anthocyanin synthesis. Currently, the use of LED systems to modulate anthocyanin production in turfgrasses would require greater upfront investment compared to traditional lighting. However, the upfront cost of the LED arrays would be largely offset over time due to decreased operating and maintenance costs, increasing the economic sustainability of anthocyanin production (Morrow, 2008). Similarly, the cost of LED lighting is estimated to decrease by 10-fold every decade, making this a much more affordable alternative in the future (Haitz and Tsao, 2011; Stutte, 2015).

The research presented here has defined the optimal light conditions necessary for inducing large quantities of anthocyanins in rough bluegrass, and this data may also potentially be used in other perennial grass systems (e.g. other turfgrasses, ornamental grasses, or prairie grasses) to increase anthocyanin content. Our data show that white light at 1000 μmol m-2 s-1 is sufficient for increasing anthocyanin content in rough bluegrass, and that treatment with 70% blue and 30% red light increases the efficiency of anthocyanin production compared to other wavelength applications. Our LED experiments showed that the application of 200 μmol m-2 s-1 of combined blue and red

103 light significantly increased anthocyanin concentration relative to blue light alone; therefore, increasing the light intensity of this LED treatment may increase anthocyanin content beyond what was seen with white light treatment. While, anthocyanin concentrations produced by rough bluegrass under these conditions may not be as high as currently used plant sources, the advantage of using a turfgrass is the ability to maximize anthocyanin yield over an entire growing season (through multiple harvests). Because of this, we calculate that the use of turfgrasses over an entire growing season could potentially increase anthocyanin yield by two fold over currently used plant sources.

These methods may help increase both the environmental and economic sustainability of anthocyanin extract production.

While this research has focused on this use of supplemental light to induce and optimize anthocyanin production in rough bluegrass, efforts to increase production could be made through germplasm evaluation. These experiments only focused on anthocyanin production using one cultivar of rough bluegrass. Natural accessions of rough bluegrass and/or other cultivars may produce greater concentrations of anthocyanin following light treatment, and they may increase production with decreased light fluence or decreased exposure duration. The evaluation of rough bluegrass germplasm may also provide further insight into the regulation of anthocyanin synthesis in this turfgrass. Together this could help develop a breeding program focused on the generation of rough bluegrass varieties that produce greater concentrations of anthocyanin using decreased input.

104

Reference

Atkin, O.K., B. Botman, and H. Lambers. 1996. The causes of inherently slow growth in alpine plants: an analysis based on the underlying carbon economies of alpine and lowland Poa species. Functional Ecol. 10(6): 698-707.

Boldt, J.K. 2013. Foliar anthocyanins in coleus and ornamental grasses: Accumulation, localization, and function. Univ. of Minnesota, Minneapolis, MN. Ph.D. Diss.

Boldt, J.K., M.H. Meyer, and J.E. Erwin. 2014. Foliar anthocyanins: A horticultural review, p. 209-251. In: J. Janick (ed.). Hort. Rev. Wiley, New York, NY.

Chalker-Scott, L. 1999. Environmental significance of anthocyanins in plant stress responses. Photochemistry and Photobiology. 70(1): 1-9.

Christians, N. E. 2011. Chapter 2: Introduction to the grasses, p. 9-31. In. N. E. Christians (ed.). Fundamentals of Turfgrass Management – 4th edition. Wiley, New York, NY.

Dayan, F.E., M.L.B, Trindade, and E.D. Velini. 2009. Amicarbazone, a new photosystem II inhibitor. Weed Sci. 57(6): 579-583.

Del Pozo-Insfran, D., C.H. Brenes, and S.T. Taloctt. 2004. Phytochemical composition and pigment stability of acai (Euterpe oleracea Mart.). J. of Ag. And Food. Chem. 52: 1539-1545. de Pascual-Teresa, S., C. Santos-Buelga, and J.C. Rivas-Gonzalo. 2002. LC-MS analysis of anthocyanins from purple corn cob. J. of the Sci. of Food and Ag. 82: 1003- 1006. de Pascual-Teresa, S., M.T. Sanchez-Ballesta. 2008. Anthocyanins: From plant to health. Phytochem. Rev. 7: 281-299.

105

Deroles, S. 2009. Anthocyanin biosynthesis in plant cell cultures: A potential source of natural colourants, p. 107-168. In. Gould, K., K. Davies, C. Winfield (eds.). Anthocyanins: Biosynthesis, functions, and applications. Springer-Verlag, New York, NY.

Downs, R.J. and H.W. Siegelman. 1962. Photocontrol of anthocyanin synthesis in Milo seedlings. Plant Physiol. 25-30.

Drumm, H., and H. Mohr. 1978. The mode of interaction between blue (UV) light photoreceptor and phytochrome in anthocyanin formation of the Sorghum seedling. Phytochem. and Photobiol. 27: 241-248

Folta, K.M. and S.D. Carvalho. 2015. Photoreceptors and control of horticultural plant traits. HortScience. 50(9): 1274-1280.

Fossen, T., R. Slimestad, D.O. Øvstedal D, and Ø.M. Anderson. 2002. Anthocyanins of grasses. Biochem. Systematics and Ecol. 30: 855-864.

Fox, A.R., G.C. Soto, A.M. Jones, J.J. Casal, J.P. Muschietti, and M.A. Mazzella. 2012. Cry1 and GPA1 signaling genetically interact in hook opening and anthocyanin synthesis in Arabidopsis. Plant Mol. Biol. 20: 315-324.

Galvão, V.C. and C. Frankhauser. 2015. Sensing the light environment in plants: Photoreceptors and early signaling steps. Curr. Opinion in Neurobiology. 34: 46- 53.

Giusti, M., and R.E. Wrolstad. 2001. Characterization and Measurement of Anthocyanins by UV-Visible Spectroscopy. Current Protocols in Food Anal. Chem. 2: 1-13.

Haitz, R. and J.Y. Tsao. 2011. Solid-state lighting: ‘The case’ 10 years after and future prospects. Phys. Status Solidi. 1: 17-29.

Hao, S., J. Wu, Y. Huang, and J. Lin. 2006. Natural dyes as photosensitizers for dye- sensitized solar cell. Solar Energy. 80: 209-214.

He, J. and M. Giusti. 2010. Anthocyanin: Natural colorants with health-promoting properties. Annu. Rev. Food Sci. Technol. 1: 163-187.

Hong, G., W. Hu, J. Li, Z. Chen, and L. Wang. 2009. Increased accumulation of artemisinin and anthocyanins in Artemisia annua expressing Arabidopsis blue light receptor CRY1. Plant Mol. Biol. Rpt. 27: 334-341.

106

Hughes, N.M., C.B. Morley, and W.K. Smith. 2007. Coordination of anthocyanin decline and photosynthetic maturation in juvenile leaves of three deciduous tree species. New Phytol. 175: 675-685.

Hughes, N.M., K.L. Carpenter, and J.G. Cannon. 2013. Estimating contribution of anthocyanin pigments to osmotic adjustment during leaf reddening. J. of Plant Physiol. 170: 230-233.

Hurley, R. 2010. Rough Bluegrass (Poa trivialis L.), p. 67-73. In: Casler, M.D. and R.R. Duncan (eds.). Turfgrass Biology, Genetics, and Breeding. Wiley, New York, NY.

Jeong, S., P. Kumar Das, S.C. Jeoung, J. Song, H.K. Lee, Y. Kim, W.J. Kim, Y.I. Park, S. Yoo., S. Choi, G. Choi, and Y. Park. 2010. Ethylene suppression of sugar induced anthocyanin pigmentation in arabidopsis. Plant Physiol. 154: 1514-1531.

Kadomura-Isikawa, Y., K. Miyawaki, S. Noji, and A. Takahashi. 2013. Phototropin 2 is involved in blue light-induced anthocyanin accumulation in Fragaria x ananassa fruits. J. Plant Res. 126: 847-857.

Kerckhoffs, L.H.J. and R.E. Kendrick. 1997. Photocontrol of anthocyanin biosynthesis in tomato. J. Plant Res. 110: 141-149.

Krans, J.V. and J.B. Beard. 1980. The effects of stage of seedling development on selected physiological and morphology parameters in Kentucky bluegrass and red fescue, p. 89–95. Proc. of the Third Intl. Turfgrass Res. Conf. Amer. Soc. of Agron., Crop Sci. Soc. of Amer., Soil Sci. Soc. of Amer., and the Intl. Turfgrass Soc.

Kumar Das, P., B. Geul, S. Choi, S. Yoo, and Y. Park. 2011. Photosynthesis-dependent anthocyanin pigmentation in Arabidopsis. Plant Signaling and Behavior. 6(1): 23- 25.

Landschoot, P.J. and D.V. Waddington. 1987. Response of turfgrass to various nitrogen sources. Soil Sci. Soc. of Amer. J. 51: 225-230.

Lange, H., W. Shiropshire, and H. Mohr. 1971. An analysis of phytochrome-mediated anthocyanin synthesis. Plant Physiol. 47: 649-655.

Lin-Wang, K., D. Micheletti, J. Palmer, R. Voltz, L. Lozano, R. Espley, R.P. Hellens, D. Chagne, D.D. Rowan, M.A. Troggio, I. Iglesias, and A.C. Allan. 2011. High temperature reduces apple fruit color via modulation of the anthocyanin regulatory network. Plant Cell and Envrion. 34: 1176-1190.

107

Mancinelli, A.L., C. H. Yang, I. Rabino, and K.M. Kuzmanoff. 1976. Photocontrol of anthocyanin synthesis: V. Further evidence against the involvement of photosynthesis in high irradiance reaction anthocyanin synthesis of young seedlings. Plant Physiol. 58: 241-217.

Mancinelli, A.L. and I. Rabino. 1978. The “High Irradiance Responses” of plant photomorphogenesis. The Botanical Rev. 44 (2): 129-180.

Mancinelli, A.L. 1985. Light-dependent anthocyanin synthesis: A model system for the study of plant photomorphogenesis. The Botanical Rev. 51(1): 107-157.

Mansour, R., B. Ezzili, and M. Farouk. 2013. Dyeing properties of wool fabrics dyes with Vitis vinifera L. (Black Grenache) leaves extract. Fibers and Polymers. 14(5): 786-792.

Merzylak, M.N., O.B. Chivkunova, A.E. Solovchenko, and K.R. Naqvi. 2008. Light absorption by anthocyanins in juvenile, stressed, and senescing leaves. J. of Exp. Bot. 59(14): 3903-3911.

Mohr, H., and H. Drumm-Herrel. 1983. Coaction between phytochrome and blue/UV light in anthocyanin synthesis in seedlings. Physiol. Plant. 58: 408-414.

Mol., J., G. Jenkins, E. Schäfer, and D. Weiss. 1996. Signal perception, transduction, and gene expression involved in anthocyanin biosynthesis. Critical Rev. in Plant Sci. 15(5 and 6): 525-557.

Morrow, R.C. 2008. LED lighting in horticulture. HortScience. 43(7): 1947-1950.

Neff, M.M. and J. Chory. 1998. Genetic interactions between phytochrome A, phytochrome B, and cryptochrome 1 during Arabidopsis development. Plant Physiol. 118: 27-36.

Poppe, C., U. Sweere, H. Drumm-Herrel, and E. Schäfer. 1998. The blue light receptor cryptochrome 1 can interact independently of phytochrome A and B in Arabidopsis thaliana. The Plant Journal. 16(4): 466-471.

108

Rabino, I. and A.L. Mancinelli. 1986. Light, temperature, and anthocyanin production. Plant Physiol. 81: 922-924.

Schneider, M.J. and W.R. Stimson. 1971. Contribution of photosynthesis and phytochrome to the formation of anthocyanin in turnip seedlings. Plant Physiol. 48: 312-315.

Stutte, G.W. Commercial transition to LEDs: A pathway to high-value products. HortSci. 50(9): 1297-1300.

Takahashi, S. and N. Murata. 2008. How do environmental stresses accelerate photoinhibtion? Trends in Plant Sci. 13(4): 178-182.

U.S. Department of Agriculture. 2015. Noncitrus fruits and nuts 2014 summary. US. Dept. of Agr. Washington D.C.

Vandenbussche, F., Habricot, Y., A.S. Condiff, R. Maldiney, D. Van Der Straeten, and M. Ahmad. 2007. HY5 is a point of convergence between cryptochrome and cytokinin signaling pathways in Arabidopsis thaliana. The Plant J. 49: 428-441.

Weiss, D. and A.H. Halevy. 1991. The role of light reactions in the regulation of anthocyanin synthesis in Petunia corollas. Physiologia Plantarum. 81: 127-133.

Wellburn, A.R. 1994. The spectral determination of chlorophylls a and b, as well as total carotenoids, using various solvents with spectrophotometers of different resolution. J. Plant Physiol. 144: 307-313.

109

Wongcharee, K., V. Meeyoo, and S. Chavadej. 2007. Dye-sensitized solar cell using natural dyes extracted from rosella and blue pea flowers. Solar Energy and Materials and Solar Cells. 91: 566-571.

Wrolstad, R.E. and C.A. Culver. 2012. Alternatives to those artificial FD&C food colorants. Annu. Rev. Food Sci. 3: 59-77.

Wu, X., G.R. Beecher, J.M. Holden, D.B. Hautowitz, S.E. Gebhardt, and R.L. Prior. 2006. Concentrations of anthocyanins in common foods consumed in the United States and estimation of normal consumption. J. of Agr. and Food Chem. 54: 4069-4075.

Yamamoto, Y. and R. Mizuguchi. 1982. Selection of a high and stable pigment producing strain of Euphorbia millii cells. Theoretical Appl. Genet. 61: 113-116.

Yamane, T., S. Tae Jeong, N. Goto-Yamamoto, Y. Koshita, and S. Kobayashi. 2006. Effects of temperature on anthocyanin biosynthesis in grape berry skins. Am. J. of Enol. and Viticult. 57(1): 54-59.

Younger, V.B. 1969. Physiology of growth and development, p. 187-216. In: Hanson A.A. and F.V. Juska (ed). Turfgrass Science. Amer. Soc. of Agron. Madison, WI.

110

Chapter 3: Effects OF Blue Light and Phenotype on Anthocyanin Accumulation in Accessions and Cultivars of Rough Bluegrass

Published December 2016: Crop Science doi:10.2135/cropsci2016.05.0438. Permission to re-publish granted 11/21/2016.

Abstract

Anthocyanins are increasingly being used as natural alternatives in medicinal, food, and industrial products. However, production of anthocyanin extract is often inefficient due to agronomic limitations. The use of turfgrasses for anthocyanin production, on the other hand, has been suggested to increase yield by 2-fold. Rough bluegrass (Poa trivialis L.), cv. Havana, has been shown to increase anthocyanin content by 117-fold under high-light treatment, exhibiting concentrations similar to current anthocyanin sources, and could be an alternative source of anthocyanin. The objectives of this research were to evaluate variation in anthocyanin content in twenty accessions and five cultivars of rough bluegrass treated with blue light, and to evaluate phenotypes associated with anthocyanin content in this species to help develop germplasm screening tools. Following blue light treatment, rough bluegrass U.S. cultivars Laser, ProAm, Sabre, Colt, Havana, and accessions originating from Germany produced statistically greater quantities of anthocyanin compared to other accessions. Phenotypes associated with anthocyanins, including: total phenolic content, total flavonoid content, specific leaf area (SLA), leaf

111 cuticular wax, and the chlorophyll:carotenoid ratio were variable for all accessions and cultivars. However, accessions producing greater quantities of anthocyanin exhibited less cuticular wax. Results of a principle component analysis (PCA) showed that there was a significant negative correlation between leaf cuticular wax and anthocyanin content.

These data demonstrate that phenotypes associated with increased anthocyanin content, especially leaf cuticular waxes, could serve as screening tools for evaluating other turfgrasses as anthocyanin sources, and may therefore help to maximize industrial anthocyanin production.

Introduction

Anthocyanins are plant pigments that produce red, purple, and blue coloration in reproductive and vegetative tissues, and are increasingly being used in industrial, medicinal, and food products (Dooner and Robbins, 1991; Delgado-Vargas et al., 2000;

Narayan, 2012; Mansour et al., 2013; He and Giusti 2010). Alternative uses of anthocyanins are rising globally, and use is predicted to increase by 5-15% every year

(Deroles, 2009; Liang et al., 2015). The use of anthocyanins across the range of technologies described is predicted to increase by 5-15% every year, and these alternative uses have traditionally been limited by the small number of plant species used to obtain anthocyanin extracts (Deroles, 2009).

To meet industrial demand for anthocyanins, a plant species capable of producing high quantities of anthocyanin over a short period of time is needed. Turfgrasses are a group of perennial monocots that can withstand aggressive harvesting while still allowing

112 the growth of new leaves due to the presence of intercalary meristems, and compounds of interest (i.e. anthocyanin) could be harvested multiple times using the same stand of plants; therefore, potentially producing greater amounts of anthocyanin, relative to current anthocyanin sources, over an entire growing season (Petrella et al., 2016). Rough bluegrass (Poa trivialis L.) is known to produce cyanidin-3-glucoside and cyanidin-3- malonylglucoside (Fossen et al., 2002; Hurley, 2010; Petrella et al., 2016). However, anthocyanins do not accumulate to suitable levels under natural growing conditions

(Petrella et al., 2016). Previous data has shown that rough bluegrass has the ability to increase anthocyanin content by over 100-fold when exposed to high intensity white light; up to 0.24% of dry wt. (Petrella et al., 2016). Constant exposure to blue light for a five day period was also shown to increase anthocyanin content compared to white and red light (Petrella et al., 2016). It is possible that further increases in cyanidin-3- glucoside and cyanidin-3-malonylglucoside could be achieved through genetic selection or engineering for phenotypes associated with increased anthocyanin content.

In addition to increasing anthocyanin production, blue light also induces morphological changes that may result in changes in spectral absorption within leaves.

These changes function to decrease photoinhibitory related stresses (Mullineaux and

Karpinski, 2002). For example, leaf thickness increases with intensity of blue light, and as a result a light gradient is created within the leaf due to greater spectral absorption by the cells first irradiated (Sinclair et al., 1973; Vogelmann and Olof Björn, 1986; Sæbø et al., 1995; Vogelmann, 1996; Hogewoning et al., 2010). Plants with naturally thinner leaves have altered light distribution within the leaf compared to thicker leaves, and

113 therefore a larger number of cells may be exposed to greater quantities of blue light leading to increased anthocyanin content. Leaf thickness can be difficult to measure; however, the specific leaf area (SLA) can serve an estimate of leaf thickness (Evans and

Poorter, 2001; Vile et al., 2005).

Quantities of chlorophyll and carotenoids are known to increase with blue light exposure, and may impact the induction of anthocyanin synthesis through competitive light absorption, and/or alterations in light reflection (Sæbø et al., 1995; Steyn et al.,

2002; Hogewoning et al., 2010). Similarly, leaf cuticular waxes have been demonstrated to increase with greater photosynthetic photon flux (PPF) (Hull et al., 1975; Barnes et al.,

1996). Leaf cuticular waxes scatter and reflect light (especially ultra-violet (UV) light) at the leaf surface; therefore, leaves that naturally exhibit greater quantities of cuticular wax may exhibit decreased anthocyanin synthesis as a result of reduced PPF (Reicosky and

Hanover, 1978; Vogelmann, 1993; Pilon et al., 1999; Long et al., 2003; Holmes and

Keiller, 2002; Close and Beadle, 2003). Phenolics and flavonoids, anthocyanin precursors, are known to increase anthocyanin synthesis when applied supplementally, specifically in plant cell and tissue culture media; therefore, plants with naturally greater quantities of precursors may accumulate more anthocyanins under blue light (Forkmann,

1977; McCormick, 1978; Vogelien et al., 1990; Dédaldéchamp and Uhel, 1999;

Luczkiewcz and Cisowski, 2001).

Variants within a species can exhibit differences in anthocyanin content, and patterns of anthocyanin accumulation may differ due to differences in phenotypes described above (Kalt et al., 1999; Lachman et al., 2009). Analysis of accessions and

114 cultivars of rough bluegrass has previously shown that there are high degrees of genetic and phenotypic variation within this species (Rajasekar et al., 2006). We therefore hypothesized that rough bluegrass accessions/cultivars would exhibit variations in anthocyanin accumulation as a result of their differences in phenotype. The objectives of this research were to evaluate variation in anthocyanin content in accessions and cultivars of rough bluegrass to help discover easily detectable phenotypes that would allow for initial screening of turfgrass germplasm with potential for accumulating increased anthocyanins following light treatment.

Materials and Methods

Light emitting diode (LED) array construction

Blue (472 nm peak λ) LED’s were purchased pre-mounted on flexible metal-core printed circuit strips - 5.0 x 5.0 mm LEDs, 60 LEDs m-1 (Torchstar Inc.® La Puente, CA,

USA). LED strips were cut to lengths of 117 cm, and were mounted on aluminum sheet metal (122 x 61 x .64 cm) according to the methods of Petrella et al. (2016) (Appendix

B). The final LED array consisted of 54 LED strips, placed side-by-side with no space between individual strips. Four LED strips were then soldered together in parallel, and each group of four strips was independently powered using a 12V AC-DC switching power supply (LED Wholesalers® Hayward, CA, USA). The LED array was mounted within a growth chamber (Conviron E15, Controlled Environment Ltd., Winnipeg,

Canada) according to the methods of Petrella et al. (2016).

115

Plant materials

Experiments were performed using seeded samples of nineteen accessions and five cultivars of rough bluegrass obtained from the USDA National Plant Germplasm

System (NPGS) collections at the Western Regional Plant Introduction Station (WRIPS), and an additional production cultivar, Havana, was obtained from J.R. Simplot (J.R.

Simplot Company, Boise, ID, USA) for a total of 25 accessions and cultivars used in this study (Table 3.1). Four replicates of each sample were grown from seed (7.5 g m-²) in standard 15 x 11 cm plastic pots using soilless media (Farfard® 3B Mix, Sungro®

Horticulture, Agawam, MA, USA). Plants were mowed, fertilized, and treated with pesticide according to the methods of Nangle et al. (2015). Stands of rough bluegrass were allowed to mature for five months prior to initiating light experiments.

LED treatment and experimental design

Five month-old rough bluegrass accessions/cultivars were exposed to 450 ± 50

µmol m-2 s-1 of blue light (a light gradient existed across the LED array, with light intensity varying between 400 and 450 µmol m-2 s-1) constantly for five days to induce and increase anthocyanins according to previous methods (Petrella et al., 2016)

(Appendix C and D). The LED array described above was suspended 5 cm above the turfgrass canopy within the growth chamber, and light intensity was measured at the turfgrass canopy using a quantum sensor and light meter (LI-COR LI-2189, LI-COR®

Biosciences, Lincoln NE USA) once per day during irradiance treatments. The growth chamber was set to a constant 15°C, and canopy temperature was 20.5° ± 2.5 C. Canopy

116 temperature was monitored with a handheld weather meter (Kestrel 3000, Kestrel Meters,

Birmingham, MI, USA) once per day during irradiance treatments. Plants were also watered daily within the growth chamber.

I.D. z PI number Country of origin NPGS collection datey 1 221908 Afghanistan Sept. 9th 1954 2 221915 Afghanistan Sept. 9th 1954 3 221951 Afghanistan Sept. 9th 1954 4 659652 Czech Republic Aug. 15th 2006 5 225826 Denmark May 18th 1955 6 W62820 Germany Aug. 15th 2006 7 W628122 Germany Aug. 15th 2006 8 W628268 Germany Aug. 15th 2006 9 227672 Iran Aug. 26th 1955 10 227858 Iran Sept. 30th 1955 11 229719 Iran Nov. 1st 1955 12 251407 Iran Sept. 25th 1958 13 380993 Iran Sept. 9th 1972 14 254908 Iraq Jan. 9th 1959 15 659892 Kyrgyzstan March 7th 2007 16 422592 Morocco Feb. 1st 1978 17 578852 - ‘Polis’ Netherlands May 1st 1977 18 250982 Macedonia Sept. 8th 1958 19 251167 Bosnia and Herzegovina Sept. 16th 1958 20 204484 Turkey Jan. 6th 1953 21 537439 - ‘Laser’ USA May 17th 1994 22 592521 - ‘ProAm’ USA Dec. 12th 1995 23 594396 - ‘Sabre’ USA May 20th 1994 24 601315 - ‘Colt’ USA May 13th 1994 25 ‘Havana’ USA N/A zNumbers used for identifying accessions and or cultivars yDate accessions were collected by the national plant germplasm system (NPGS)

Table 3.1: List of rough bluegrass accessions (Plant Introduction – PI) and cultivars used in these experiments with their country of origin.

117

The blue LED irradiance experiment consisted of 25 accessions/cultivars, with four replicates (pots) of each accession/cultivar (100 pots total). Due to the light gradient under the LED array, blocking was necessary within the growth chamber, and a maximum of 28 pots, arranged in four rows (blocks) of seven pots each, (i.e. four replicates of seven accessions/cultivars) could be placed under the LED array at a given time; the experiment was, therefore, set up as an incomplete randomized block design to accommodate for the number of plants being analyzed. As a result, experimental runs were divided into four groups to allow analysis of all accessions and cultivars. Groups 1-

3 consisted of six accessions/cultivars, and the fourth group contained the remaining seven. Accessions and cultivars were randomly assigned to a group for blue light treatment, and all replicates of accessions/cultivars were arranged within the four blocks under the LED array. An experimental run was defined as a single treatment of all four groups described above. Two experimental runs were completed using the same stand of plants; plant material was approximately seven months old at the initiation of the second experimental run. Experimental run one was initiated on May 13th 2013, and treatment of groups two and three began on May 19th and May 25th respectively. Experimental run two was initiated on Aug. 19th 2013, and treatment of groups two and three began on 25th and Sept. 1st respectively. To account for growth chamber variability, the LED array was moved to a second growth chamber for run two, and plants were re-randomized into new groups.

118

Tissue extraction and analysis

Phenolic and flavonoid compounds were extracted and analyzed as described previously (Nangle et al., 2015). Approximately 25 mg of freeze dried leaf tissue was suspended in 5 mL of methanol, homogenized, and vortexed for 30 seconds. The extract was centrifuged at 10,000 G for 10 minutes, and placed in the dark prior to use. For total phenolic content, 0.5 mL of extract was combined with 4 mL of Folin’s reagent, and 4 mL of 1M sodium carbonate. The extract was incubated in the dark for approximately 30 minutes, and the absorption was read at 765 nm. Quantification of total phenolics was made with a gallic acid standard curve. For total flavonoids, 0.5 mL of the methanol extract was first diluted with 1.5 mL methanol, and was then combined with 0.1 mL 10% aluminum chloride, 0.1 mL 1M potassium acetate, and 2.8 mL ddH2O. The extract was incubated in the dark for approximately 45 minutes, and the absorption was read at 415 nm. Quantification of total flavonoids was made with a quercetin standard curve.

Total chlorophylls and total carotenoids were extracted as described previously in chapter 2 (Wellburn, 1994; Petrella et al., 2016). Total epicuticular waxes were extracted and analyzed according to the methods of Ebercon et al. (1976). Approximately 0.5 g of fresh leaf tissue was placed in a 50 mL beaker, and 20 mL of chloroform was added.

Leaves were extracted in chloroform for 15 seconds, and the chloroform extract was filtered through two layers of cheese cloth. Standards of carnauba wax were run simultaneously with the turfgrass leaf extracts. Chloroform extracts were placed on a boiling water bath until the chloroform had evaporated. Potassium dichromate reagent (5 mL) (20g potassium dichromate dissolved in 40 mL ddH2O, combined with 1L

119 concentrated sulfuric acid, and heated until a clear amber solution was obtained) was then added. The solution was heated on a boiling water bath for 30 minutes, allowed to cool to room temperature, and 12 mL of ddH2O was added. Once cooled the sample was analyzed at 590 nm using a spectrophotometer, and quantification was based on a carnauba wax standard curve.

Specific leaf area (SLA) was measured an alternative to leaf thickness by obtaining five random leaf samples from across the entire turfgrass canopy for each accession/cultivar. Leaves were clipped, length and width measured using digital calipers, and tissue was oven dried at 55°C for 24 hours. Following drying, samples were weighed and SLA was calculated by dividing leaf area by dry mass (Garnier et al., 2001).

For analysis of total anthocyanins, all plant material 2.5 cm above the soil surface was harvested, flash frozen in liquid nitrogen, freeze dried, and ground to a powder.

Cyanidin-3-glucoside equivalents (eq.) were determined spectrophotometrically using the pH differential method (Giusti and Wrolstad, 2001). Tissue extracts were prepared as described previously in Chapter 2 (Nangle et al., 2015).

Data Analysis

Data obtained for total monomeric anthocyanin concentrations were subjected to analysis of variance (ANOVA) as an incomplete randomized block design using PROC

MIXED and SAS 9.2 (SAS Institute, Cary NC, USA). The statistical model included replicate, group, experimental run, and their interactions as random effects. Means were separated using 95% confidence intervals. Because data for anthocyanin content were

120 not statistically different between experimental runs, data for experimental runs one and two were combined. Data taken for phenotypic traits of accessions and cultivars that were not treated with blue light were subjected to ANOVA as a randomized complete block design using PROC GLM; means were separated using Tukey’s honest significant difference - HSD (P =0.05). Principle component analysis was performed using PROC

PRINCOMP, Pearson’s correlation coefficients were generated using PROC CORR, and linear regression was performed using PROC REG. Cluster analysis was performed using Ward’s Hierarchical method in R version 3.0.2 (R Foundation for Statistical

Computing©, Vienna, Austria).

Results

Blue light irradiance experiment

Results showed that the pattern of blue light induced anthocyanin accumulation differed across rough bluegrass accessions and/or cultivars (Fig. 3.1). Many accessions showed increased anthocyanin content following treatment with blue light relative to untreated controls (plants maintained under greenhouse conditions not treated with blue light). Accessions 1-3, 11, 14, 15, and 20, however, did not exhibit this increase. The data show that rough bluegrass accessions and cultivars fall into three groups based on their capacity to increase anthocyanin content following blue light treatment (Fig. 3.1).

Group one produced the lowest quantities of anthocyanin following light treatment, and consisted of accessions 1, 2, 11, 14, 18, and 20. Group two produced the highest quantities of anthocyanin (statistically greater quantities compared to group one), and

121 consisted of accessions 7, 8, and 21-25. Finally, group three produced quantities of anthocyanin that were not different from either group one or two, and consisted of accessions 3-6, 9-13, 15-17, and 19.

Figure 3.1: A 95% confidence interval separating accessions and cultivars based on total anthocyanin content (cyanidin-3-glucoside eq. - mg 100 g-1 DW). See Table 3.1 for accession labels. Bars represent sample averages, and error bars represent 95% confidence intervals (n=8). Non-overlapping error bars represent significant difference

(P = 0.05). Black shaded bars represent total anthocyanin content of plants that were not treated with blue light (n=8). There was no significant difference found in total anthocyanin content among these untreated controls (P = 0.05).

122

Blue light induced anthocyanin accumulation was associated with the geographic origin for several accessions and cultivars (Fig. 3.1, Table 3.1). U.S. bred rough bluegrass cultivars Laser, ProAm, Sabre, Colt, and Havana (cultivars 21-25) accumulated anthocyanins in large quantities when exposed to blue light. Additionally, German accessions (7 and 8) accumulated quantities of anthocyanin greater than most other accessions. In particular, accession 8 produced the largest quantity of anthocyanin on average; however, quantities for these two German accessions were still not statistically different from U.S. cultivars 21-25. The Danish cultivar, Polis (cultivar 17), accumulated quantities of anthocyanin that were not different from U.S. bred cultivars, but anthocyanin production in this cultivar was significantly lower compared to German accession 8.

Accessions from Afghanistan (accessions 1-3) exhibited anthocyanin accumulation following blue light treatment, but anthocyanin content from these accessions was significantly lower compared to U.S. cultivars 21-25 and German accessions 7 and 8. Similar to Afghani cultivars, Iranian accessions (accessions 9-13) accumulated levels of anthocyanin that were statistically lower than U.S. cultivars 21-25 and German accession 7 and 8 following blue light treatment. Interestingly however,

Iranian accessions exhibited increased variability in light induced anthocyanin synthesis compared to accessions from Afghanistan.

123

Phenotypic trait analysis

Accessions and cultivars not treated with blue light were analyzed for phenotypic traits potentially associated with anthocyanin accumulation, specifically changes in the levels of total phenolic content, total flavonoid content, specific leaf area (SLA), leaf cuticular wax, and the total chlorophyll:total carotenoid ratio. For total flavonoids, data show that accessions 2 and 25 contain statistically greater quantities of flavonoids compared to accession 8 only (Table 3.2). There were no statistical differences in accessions and cultivars for total phenolic content. Data for SLA ranged from 2.60 - 3.74 cm2 mg-1 for all accessions and cultivars (Table 3.2). Accessions 4, 5, 7, 8, 17, 24 exhibited the thinnest leaf tissue on average compared to accessions 9-15, which exhibited the thickest leaves.

Analysis of leaf cuticular waxes indicated that only accession 1, which did not increase anthocyanin production following blue light treatment, produced greater quantities of wax compared to accessions that did increase anthocyanin content following blue light treatment (German accessions 7 and 8 as well as U.S. cultivars 21-25) (Table

3.2). There were no significant differences in cuticular wax content observed across most other accessions and cultivars. Accession 6 exhibited a lower total chlorophyll:total carotenoid ratio compared to accession 12, while other accessions and cultivars were not different from one another (Table 3.2).

124

I.D.z Flavonoids Phenolics SLA Wax Chl:Carot -1 2 -1 -2 mg g dry wt. cm mg dry wt. µg cm 1 20.24ab‡ 12.79 2.92cdef 6.41a 7.67ab 2 20.93a 12.54 3.04abcd 4.14abc 6.22ab 3 19.46ab 12.8 3.06abcdef 5.08ab 7.52ab 4 18.11ab 12.99 3.72ab 4.29abc 7.20ab 5 19.16ab 13.79 3.44abcd 4.37abc 7.80ab 6 19.37ab 13.71 2.89cdef 5.02ab 5.20b 7 17.75ab 11.91 3.74a 2.55bc 6.87ab 8 15.23b 11.46 3.32abcdef 2.34bc 7.98ab 9 18.39ab 13.13 2.56f 4.55abc 7.67ab 10 20.05ab 14.49 2.60f 4.85abc 6.68ab 11 18.9ab 11.6 2.65f 3.49abc 6.61ab 12 19.36ab 13.56 2.77def 3.45abc 9.46a 13 19.53ab 15.4 2.89cdef 4.30abc 7.13ab 14 16.42ab 12.58 2.76def 3.44abc 7.73ab 15 17.04ab 14.39 2.60f 2.99bc 7.59ab 16 19.11ab 13.37 3.16abcdef 3.31bc 7.12ab 17 18.1ab 14.5 3.38abcd 2.88bc 7.42ab 18 20.75ab 15.36 2.97cdef 1.97c 6.60ab 19 17.87ab 15.08 3.11abcdef 3.04bc 6.84ab 20 19.55ab 11.99 2.82def 4.18abc 8.00ab 21 19.54ab 14.47 2.68ef 2.72bc 6.11ab 22 18.13ab 12.44 3.09abcdef 3.18bc 7.67ab 23 17.8ab 12.26 3.31abcde 3.92abc 8.31ab 24 17.15ab 10.77 3.57abc 2.72bc 8.74ab 25 20.84a 12.37 2.95cdef 2.46bc 6.73ab Nsy zAccession/cultivar I.D. (see table 1) yns = not significant ‡Means followed by the same letter are not significantly different (P = 0.05) as determined by Tukey’s HSD (n = 4)

125

A cluster analysis was performed using the previously analyzed phenotypic traits to examine potential relationships among the 25 rough bluegrass accessions and cultivars, and the resultant cluster dendrogram showed five clusters (Fig. 3.2). These clusters were aligned with the geographic sites of origin of the accessions and cultivars, illustrating the potential impact of genotype on phenotype (Table 3.1). For example, accessions 1-3, all from Afghanistan, formed a single cluster. Those from Iran (accessions 9-13) formed a single cluster, while the U.S. cultivars (21-25) used in these experiments formed a third cluster. Accession 20 (originating from Turkey), clustered with U.S. cultivars 21-25, yet this accession was not morphologically similar to the rough bluegrass cultivars. Danish cultivar Polis (accession 17) did not cluster with U.S. cultivars, but instead clustered with accessions from Iraq (14), Kyrgyzstan (15), Morocco (16), and Serbia/Montenegro (18 and 19). Accessions within this cluster were morphologically disparate yet shared phenotypic traits associated with anthocyanin production following blue light treatment.

Using principal component analysis (PCA), phenotypic trait data were simplified to examine associations between these data and the levels of anthocyanins accumulated under blue light. Results within the correlation matrix showed that total phenolic content, total flavonoid content, leaf cuticular waxes, and SLA exhibited negative correlations with blue light induced anthocyanin production; with only total flavonoid content and leaf cuticular waxes exhibiting statistically significant correlations with total anthocyanin concentration (Table 3.3).

126

Figure 3.2: Cluster dendrogram using Ward’s Hierarchical method to group 25 accessions and cultivars of rough bluegrass based on phenotypic traits associated with anthocyanin content (data taken on plants that were not treated with blue light): total phenolic content, total flavonoid content, cuticular waxes, specific leaf area (SLA), and the total chlorophyll:total carotenoid ratio. Accessions and cultivars are labeled based on sample I.D. number (See table 3.1). Clusters are labeled based on geographic origin of accessions.

127

Eigenvalues from the PCA showed five principal components. The first three components accounted for 75% of the variation, and components with eigenvalues below

0.9 were not considered for further analysis; only principle components 1-3 were therefore used for further analyzed (Table 3.4).

Correlation matrix Phe Flav Wax SLA Chl:Carot Antho Pearson’s correlation coefficient Phe 1.000 0.327 0.117 -0.186 -0.074 -0.262 Flav 0.327 1.000 -0.020 -0.268 -0.093 -0.407* Wax 0.117 -0.020 1.000 0.279 -0.099 -0.377* SLA -0.186 -0.268 0.279 1.000 -0.047 -0.147 Chl:Carot -0.074 -0.093 -0.099 -0.047 1.000 0.255 Antho -0.262 -0.407 -0.377 -0.147 0.255 1.000 *, **, *** Significant at P ≤ 0.05, 0.01, or 0.001, respectively

Table 3.3: Correlation matrix examining relationships between phenotypic trait data of rough bluegrass accessions and cultivars that were not treated with blue light, and anthocyanin content following blue light treatment. Total phenolic content (Phe), total flavonoid content (Flav), leaf cuticular wax (Wax), specific leaf area (SLA), and the total chlorophyll:total carotenoid ratio (Chl:Carot), and anthocyanin content of rough bluegrass plants following blue light treatment (Antho).

128

Principle component 1 was highly loaded for total phenolic content, total flavonoid content, and SLA (Table 3.4). Principle component 2 was highly loaded with leaf cuticular waxes, and principle component 3 was highly loaded with the total chlorophyll:total carotenoid ratio (Table 3.4). Principal component scores for principle components 1-3 were used for linear regression analysis where anthocyanin concentration data was set as the dependent variable. Only principle component 2 showed a significant

(P = 0.05) negative correlation with anthocyanin concentration (Fig. 3.3). Even though the correlation coefficient was relatively low (r2 = 0.28), the correlation was still significant (p = 0.0064).

PCA results Principle Component 1 2 3 Eigenvalue 1.55 1.27 .93 Cumulative 0.31 0.56 0.75 Phe 0.656 0.406 0.308 Flav 0.758 0.186 0.03 Wax -0.218 0.784 0.354 SLA -0.693 0.429 0.072 Chl:Carot -0.118 -0.526 0.835

Table 3.4: Principle component analysis (PCA) of phenotypic trait data of accessions and cultivars of rough bluegrass that were not treated with blue light. Total phenolic content (Phe), total flavonoid content (Flav), leaf cuticular wax (Wax), specific leaf area

(SLA), and the total chlorophyll:total carotenoid ratio (Chl:Carot).

129

Figure 3.3: Linear regression of principle component 2 (loaded with leaf cuticular wax –

See table 3.3) and total anthocyanin concentration following irradiance with five days of constant blue light (450 µmol m-2 s-1). The predictably of the relationship is relatively weak (r2 = 0.28); however the correlation is significant (p = 0.0064).

Discussion

The objective of this research was to evaluate variation in anthocyanin content in accessions/cultivars of rough bluegrass to help discover easily detectable phenotypes that would allow for initial screening of turfgrasses that exhibit potential for increasing anthocyanin content following blue light treatment. Rough bluegrass cv. Havana has been shown to accumulate anthocyanins in large quantities when treated with high- intensity white, blue, and with combined application of blue and red light; therefore, rough bluegrass may produce as much or more anthocyanin compared to current plant 130 sources due to the ability to repeatedly harvest anthocyanin containing tissue using the turfgrass system (Petrella et al., 2016). It was not known, however, if all accessions or cultivars of rough bluegrass have an equivalent ability to increase anthocyanin synthesis following light treatment. The data presented here show that all rough bluegrass U.S. cultivars and German accessions 7 and 8 exhibited the ability to increase anthocyanin content when exposed to constant blue light. Rough bluegrass U.S. cultivars and German accessions may provide germplasm capable of increasing anthocyanin production or enabling breeding efforts to increase anthocyanin levels of rough bluegrass following light treatment. Further, our results indicate that accessions with other geographic sites of origin (Afghanistan and Iran) would be less suitable as anthocyanin sources.

An anthocyanin production system using a turfgrass species as the model plant relies on the generation of large tissue yields to produce greater seasonal amounts of anthocyanin compared to plants currently used (Petrella et al., 2016). Because of this, experiments here were designed to evaluate rough bluegrass grown at densities that would potentially be used in a physical production system. Densely packed plants, on the other hand, this may influence blue light induced anthocyanin production due to differences in maturity within the stand along with competition for light. It has been previously shown, however, that plants varying in age as well as developmental stage still produced high quantities of anthocyanin when grown at a high density and exposed to high-intensity white light (Petrella et al., 2016). The use of a lower plant density may, in fact, overestimate the quantity of anthocyanin that would be produced in a potential production system; our data, therefore, reflect anthocyanin concentrations that could be

131 expected when using a rough bluegrass based production system. Still, this system may be further refined through the development of optimized seeding rates for rough bluegrass U.S. cultivars and German accessions, potentially allowing for greater anthocyanin concentration per leaf without decreasing the quantity of harvestable tissue.

Potential phenotypic traits associated with anthocyanin content were analyzed in this study for the potential development of rapid screening tools for selecting turfgrasses that may increase anthocyanin content following light treatment. For total flavonoid content, German accession 8 (produced high concentrations of anthocyanin under blue light) exhibited the lowest flavonoid concentration, while Afghani accession 2 (which produced low concentrations of anthocyanin under blue light) produced large quantities of flavonoids, and U.S. cultivar 25 (which produced high concentrations of anthocyanin under blue light) produced large quantities of flavonoids. At first, these data suggest that accession 8 may exhibit low quantities of flavonoids because it is consistently producing larger quantities of the end product, anthocyanin. However, the fact that U.S. cultivar 25 exhibited both high flavonoid and anthocyanin levels does not support the hypothesis.

For SLA, German accessions 7 and 8, which produced high concentrations of anthocyanin under blue light, exhibited leaf tissue that was thinner compared to accessions that accumulated little anthocyanin following light treatment (i.e. accessions

11-13). However, accession 8 was not statistically different from these accessions; other accessions that did not produce large amounts of anthocyanin (i.e. Afghani accessions) also exhibited SLA that were statistically the same as accession 7. Light attenuation by adaxial mesophyll may be decreased in thinner leaves when compared to thicker leaves,

132 and blue light exposure to individual mesophyll cells may increase in thinner leaves due to changes in spectral distribution (Vogelmann, 1993). As a result, in plants with a larger

SLA, blue light treatments would therefore potentially irradiate a greater proportion of cells, leading to increased anthocyanin content across the entire leaf surface (Vogelmann and Olof Björn, 1986). Our data did not suggest that greater SLA increased anthocyanin content in rough bluegrass under blue light irradiance; however, SLA is only an approximation of leaf thickness and further investigation is required to use leaf thickness as an anthocyanin selection tool.

Cluster analysis based on phenotypic data showed a relationship between accession origin and phenotypic traits associated with anthocyanin content. Previous genetic analysis of rough bluegrass accessions has shown that accession 5 (Denmark) and

U.S. cultivar 22 (ProAm) were the most similar based on random amplified polymorphic

DNA (RAPD) marker analysis, whereas the U.S. cultivars ProAm and Sabre were much less genetically similar (Rajasekar et al., 2006). Rough bluegrass U.S. cultivar ProAm was also more genetically similar to accessions 18 (Macedonia) and 19 (Bosnia and

Herzegovina) compared to the U.S. cultivar Sabre (Rajasekar et al., 2006). Interestingly, phenotypic data presented here do not indicate a relationship between the U.S. cultivars

ProAm and accessions 5 (Demark), 18, and 19 (Macedonia and Bosnia and Herzegovina respectively) based on blue light induced anthocyanin content, and therefore differ from the previously published genotypic clustering. This may be due to variations in the anthocyanin biosynthetic pathway across these accessions and cultivars of rough

133 bluegrass. Accession 18 did, however, share phenotypic traits with U.S. cultivars including increased flavonoid content and decreased leaf cuticular wax.

Cluster analysis using RAPD markers has also shown that the rough bluegrass

U.S. cultivar Sabre is just as similar to Iranian and Afghani accessions as it is to the U.S. cultivar ProAm (Rajasekar et al., 2006). Based on our results, rough bluegrass U.S. cultivars and accessions from Afghanistan and Iran do not exhibit any similarity on phenotypic data, and U.S cultivars and Afghani accessions are the most dissimilar based on the ability to increase anthocyanin content under constant blue light. Despite the trends identified in our analyses, the cluster analysis presented here does not allow prediction for anthocyanin production based on accession/cultivar geographic origin. For example, based on phenotypic data, accession 6 from Germany clustered with the other

German accessions evaluated (7 and 8), but anthocyanin content of accession 6 was not comparable to accession 8 or any of the cultivars evaluated.

It is well known that leaf cuticular waxes scatter and reflect incident light depending on wax particle size and quantity; potentially decreasing the amount of light the plant perceives (Sinclair et al., 1973; Vogelmann et al., 1996; Pilon et al., 1999;

Holmes and Keiller, 2002). Accession 1 (Afghanistan) produced the largest quantity of cuticular wax, and produced very low concentrations of anthocyanin following blue light treatment, suggesting that the greater quantities of wax present in this accession may be decreasing blue light perception. German accessions (7 and 8) and U.S. cultivars 21-25 produced statistically less cuticular wax compared to accession 1, and produced greater concentrations of anthocyanin when exposed to blue light. However, accession 18

134

(Macedonia) produced the smallest quantity of cuticular wax, and also produced very low concentrations of anthocyanin, indicating that the link between cuticular wax and high anthocyanin levels produced under blue light requires further investigation.

Results from the PCA, however, further supported an association between anthocyanin concentration and leaf cuticular wax. Regression of principle component 2 and anthocyanin concentration data showed the relationship between these data; increased quantities of leaf cuticular wax lead to a decrease in blue light induced anthocyanin accumulation. These results suggest that accessions exhibiting larger quantities of cuticular wax will only accumulate anthocyanins in small quantities or not at all when exposed to constant blue light. Cuticular waxes not only function in reflecting/scattering specific wavelengths of light, but increasing quantities of waxes are also known to decrease PPF within the leaf (Hull et al., 1975; Barnes et al., 1996).

However, the impact of leaf cuticular wax on the regulation of anthocyanin production through alteration of optical properties has not been investigated, and it is unknown whether or not waxy accessions exhibit a change in blue light reflection.

It has been demonstrated previously that using rough bluegrass cultivar Havana, anthocyanins can be induced in large quantities, harvested, and the process can be repeated using the same stand of plants after additional growth (Petrella et al., 2016).

Here, we have shown that this process is also repeatable using other U.S. cultivars as well as German accessions of rough bluegrass. We have also shown that phenotypes associated with anthocyanin content may be used for rapid screening of other turfgrass to help screen for species that could increase anthocyanin content following light treatment.

135

Further, U.S. cultivars, other than ‘Havana’, and German accessions may exhibit other benefits that could increase seasonal anthocyanin production including, but not limited to, faster growth following tissue harvest (allowing a greater number of seasonal harvests), improved nutrient use efficiency, or greater resistance to disease and insect pests.

Additionally, the variability seen for light inducible anthocyanin biosynthesis in rough bluegrass presented here should help to enable development of breeding strategies, through both traditional (i.e., crossing based on phenotype) and/or genomic selection methods, designed to increase anthocyanin production in rough bluegrass.

As a turfgrass species rough bluegrass is not used to a great extent, and is only used in certain situations (cool, damp, and shaded environments). This is in the most part due to low stress tolerance, primarily low tolerance to heat and drought stresses, but it can also be seen by the species’ low tolerance to high light. The production of flavonoids, like anthocyanins, serve as a bio-indicators of stress tolerance. As rough bluegrass has been shown to rapidly produce flavonoids during stress, this turfgrass may serve as model plant for evaluating the production of bio-markers for stress tolerance.

Interestingly, for golf courses, rough bluegrass is used in the over-seeding of dormant Bermudagrass greens, especially in the U.S. desert southwest and in Florida.

Golf courses in this region rely heavily on re-claimed irrigation water that has higher than average salt content, and rough bluegrass is considered to be one of the most salt intolerant turfgrasses (Harivandi, 2004). Flavonoid production has been shown to increase under salt stress (Eryilmaz, 2006; Van Oosten et al., 2013; Yan et al., 2014).

While little is known on what role flavonoids may have in the salt stress response, it is

136 well known that flavonoids interact with the metabolism and transport of the phytohormone auxin (Murphy et al., 2000). Therefore, rough bluegrass may provide insight into the function of flavonoid accumulation during salt stress.

137

Reference

Barnes, J.D., K.E. Percy, N.D. Paul, P. Jones, C.K. McLaughlin, P.M. Mullineaux, G. Creissen, and A. R. Wellburn. 1996. The influence of UV-B radiation on the physiochemical nature of tobacco (Nicotinia tabacum L.) leaf surfaces. J. of Exp. Bot. 47(294): 99-109.

Close, D.C. and C.L. Beadle. 2003. The ecophysiology of foliar anthocyanin. The Bot. Rev. 69(2): 149-161. doi:10.1663/0006-8101(2003)069[0149:TEOFA]2.0.CO;2

Dédaldéchamp, F. and C. Uhel. 1999. Induction of anthocyanin synthesis in nonpigmented grape cell suspensions by acting on DFR substrate availability or precursors level. Enzyme and Microbial Tech. 25: 316-321.

Dooner, H.K., T.P. Robbins, and R.A. Jorgensen. 1991. Genetic and developmental control of anthocyanin biosynthesis. Ann. Rev. Genetics. 25: 173-199.

Ebercon, A., Blum, A., and Jordan, W.R. A rapid colormetric method for epicuticular wax content of sorghum leaves. Crop Sci. 17: 179-180.

Evans, J.R. and H. Poorter. 2001. Photosynthetic acclimation of plants to growth irradiance: the relative importance of specific leaf area and nitrogen partitioning in maximizing carbon gain. Plant, Cell, and Environment. 24: 755-767.

Forkmann, G. 1977. Precursors and genetic control of anthocyanin synthesis in Matthiola incanta R. Rr. Planta. 137: 159-163.

Fossen, T., R. Slimestad, D.O. Øvstedal D, and Ø.M. Anderson. 2002. Anthocyanins of grasses. Biochem. Systematics and Ecol. 30: 855-864.

138

Garnier, E., B. Shipley, C. Roumet, and G. Laurent. 2001. A standardized protocol for the determination of specific leaf area and leaf dry matter content. Functional Ecology. 15: 688-695.

Giusti, M., and R.E. Wrolstad. 2001. Characterization and Measurement of Anthocyanins by UV-Visible Spectroscopy. Current Protocols in Food Anal. Chem. 2: 1-13.

Harivandi, M.A. 2004. Evaluating recycled waters for golf course irrigation. USGA green section record. November – December.

He, J. and M. Giusti. 2010. Anthocyanin: Natural colorants with health-promoting properties. Annu. Rev. Food Sci. Technol. 1: 163-187.

Hogewoning, S.W., G. Trouwburst, H. Maljaars, H. Poorter, W. van Leperen, and J. Harbinson. 2010. Blue light-dose responses of leaf photosynthesis, morphology, and chemical composition of Cucumis sativus grown under different combination of red and blue light. J. of Exp. Bot. doi:10.1093/jxb/erq132. 1-11.

Holmes, M.G. and D.R. Keiller. 2002. Effects of pubescence and waxes on the reflectance of leaves in the ultraviolet and photosynthetic wavebands: a comparison of a range of species. Plant, Cell and Environment. 25: 85-93.

Hull, H.M., H.L. Morton, and J.R. Wharrie. 1975. Environmental influences on cuticle development and resultant foliar penetration. The Botanical Rev. 41(4): 421-452.

Hurley, R. 2010. Rough Bluegrass (Poa trivialis L.), p. 67-73. In: Casler, M.D. and R.R. Duncan (eds.). Turfgrass Biology, Genetics, and Breeding. Wiley, New York, NY.

Kalt, W., J.E. McDonald, R.D. Ricker, and X. Lu. 1999. Anthocyanin content and profile within and among blueberry species. Canadian J. of Plant Sci. 79: 617- 623.

Lachman, J., K. Harmouz, M. Šulc, M. Orsák, V. Pivec, A. Hejtmánková, P. Dvořák, and J. Čepl. 2009. Cultivar differences of total anthocyanins and anthocyanidins in red and purple-fleshed potatoes and their relation to antioxidant activity. Food Chem. 114: 836–843.

Long, L.M., H. P. Patel, W.C. Cory, and A. E. Stapleton. 2003. The maize epicuticular wax layer provides UV protection. Functional Plant Biology. 30: 75-81.

139

Luczkiewcz, M. and W. Cisowski. 2001. Optimisation of the second phase of a two phase growth system for anthocyanin accumulation in callus of cultures of Rudbeckia hirta. Plant Cell, Tissue, and Organ Culture. 65: 57-68.

Mansour, R., B. Ezzili, and M. Farouk. 2013. Dyeing properties of wool fabrics dyes with Vitis vinifera L. (Black Grenache) leaves extract. Fibers and Polymers. 14(5): 786-792.

McCormick, S. 1978. Pigment synthesis in maize aleurone from precursors fed to anthocyanin mutants. Biochemical Gene. 16(7/8): 777-784.

Mullineaux, P. and S. Karpinski. 2002. Signal transduction in response to excess light: Getting out of the chloroplast. Curr. Op. in Plant Biol. 5(1): 43-48. doi.org/10.1016/S1369-5266(01)00226-6

Murphy, A.S., W.A. Peer, and Taiz, L. 2000. Regulation of auxin transport by aminopeptidases and endogenous flavonoids. Planta. 211: 315-324.

Narayan, M.R. 2012. Review: Dye sensitized solar cells based on natural photosensitizers. Renewable and Sustainable Energy Reviews. 16: 208– 215.

Petrella, D.P., J.D. Metzger, J.J. Blakeslee, E.J. Nangle, and D.S. Gardner. 2016. Anthocyanin production using rough bluegrass treated with high intensity light. HortSci. 5(9): 1-10

Pilon. J.J., H. Lambers, W. Baas, M. Tosserams, J. Rozema, and O.K. Atkin. 1999. Leaf waxes of slow-growing alpine and fast-growing lowland Poa species: inherent differences in responses to UV-B radiation. Phytochemistry. 50: 571-580.

Rajasekar, S., S. Fei, and N.E. Christians. 2006. Analysis of Genetic Diversity in Rough Bluegrass Determined by RAPD Markers. Crop Sci. 46: 162-167.

R Core Team. 2013. R: A language and environment for statistical computing. Release 3.0.2. R Foundation for Statistical Computing, Vienna, Austria.

Reicosky, D.A. and J.W. Hanover. 1978. Physiological effects of surface waxes: I. Light reflectance for glaucous and nonglaucous Picea pungens. Plant Physiol. 62: 101- 104.

140

Sæbø, A., T. Krekling, and M. Appelgren. 1995. Light quality affects photosynthesis and leaf anatomy of birch plantlets in vitro. Plant Cell, Tissue, and Organ Culture. 41: 177-185.

SAS Institute. The SAS System for Windows. Release 9.2. SAS Institute, Inc., Cary, NC.

Sinclair, T.R., M.M. Schreiber, and R.M. Hoffer. 1973. Diffuse reflectance hypothesis for the pathway of solar radiation through leaves. Agr. J. 66: 276 -283.

Steyn, W.J., S.J.E., Wand, D.M., Holcroft, and G. Jacobs. 2002. Anthocyanins in vegetative tissues: a proposed unified function in photoprotection. New Phytologist. 155: 349-361

Van Oosten MJ, Sharkhuu A, Batelli G, Bressan RA, Maggio A. 2013. The Arabidopsis thaliana mutant air1 implicates SOS3 in the regulation of anthocyanins under salt stress. Plant Mol Biol 83, 405-415.

Vile, D., E. Garnier, B. Shipley, G. Laurent, M. Navas, C. Roumet, S. Lavorel, S. Diaz, J.G. Hodgson, F. Lloret, G.F. Midgley, H. Poorter, M.C. Rutherford, P.J. Wilson, and I.J. Wright. 2005. Specific leaf area and dry matter content estimate thickness of laminar leaves. Annals of Botany. 96: 1129-1136.

Vogelmann, T.C., and L. Olof Björn. 1986. Plants as light traps. Physiol. Plantarum. 68: 704-708.

Vogelmann, T.C. 1993. Plant tissue optics. Ann. Rev. Plant Mol. Biology. 44: 231-251.

Vogelmann, T.C., J.N. Nishio, and W.K. Smith. 1996. Leaves and light capture: Light propagation and gradients of carbon fixation within leaves. Trends in Plant Sci. 1: 65-70.

141

Chapter 4: Modulation of Halotropic Growth in Rough Bluegrass by Blue and Red Light

Abstract

Turfgrasses are said to cover approximately 16 million hectares in the continental U.S. and have been postulated to be largest irrigated crop in the country. Global climate change and increasing demands for fresh water have resulted in decreased availability and increases in the cost for freshwater irrigation. Use of re-claimed water as an alternate irrigation strategy that is on the rise. However, these sources of water can have large concentrations of salt. As less irrigation is being applied, due to restrictions, salts will also aggregate in the soil to a greater degree. Currently, salt exclusion and sequestration are the most understood mechanisms of salt tolerance. However, a tropic growth response has also been recently defined in Arabidopsis, tomato, and sorghum.

Halotropism is a form of adaptive growth where upon exposure to salt, roots will bend/grow away from the area of high salt concentration due to changes in auxin transport and metabolism. Flavonoids have also been shown to modulate the metabolism and transport of auxin; therefore, flavonoids may regulate halotropism. The objective of this study was to determine if the turfgrass species rough bluegrass (Poa trivialis L.) exhibits halotropism, and, secondly, to determine the impact of flavonoid accumulation

142 on halotropism through irradiation with red and blue light. Rough bluegrass exhibited halotropic growth at 350 mM. NaCl, while concentrations of NaCl below 300 mM lead to increased root growth into NaCl containing media. Blue light enhances halotropic growth compared to white light, and halotropism is lost under red light. However, the combinations of blue red light restored halotropic growth in rough bluegrass. Treatment of rough bluegrass under dark conditions also did not lead to halotropism. Further analysis of rough bluegrass roots showed that flavonoids only accumulated in quantifiable quantities under blue and blue/red light treatment, and their concentrations showed a relative increase under salt treatment compared to 0 mM. Following 24 hours of salt treatment, however, plants under white light also increased flavonoid production to quantifiable amounts. Based on these data, light is required for halotropism in rough bluegrass, and more specifically blue light is required to modulate halotropism, potentially through the production of flavonoids.

Introduction

Turfgrasses cover approximately 1.9% of land in the continental United States, and may be one of the largest irrigated crops in the country (Milesi et al., 2005;

Breuninger et al., 2013). The turfgrass industry (representing golf turf, sports surfaces, parks, home lawns, and roadsides) was estimated to have a total value of $40 billion dollars as of 2009, the industry produces more income than many other agriculture commodities, and turfgrasses are known to have a positive benefit on human mental and physical health (Breuninger et al., 2013; Tyrvainen et al., 2014; Seabra et al., 2016).

143

According to the United States Geological Survey (USGS), over 13 trillion liters of water was withdrawn daily across the U.S. in 2010, and approximately 30% was used for irrigation (Maupin et al., 2014). The increasing demands for fresh water have led to overall decreases in turfgrass irrigation along with greater use of re-claimed water as an irrigation source. Treated sewage water (effluent, re-claimed, or re-cycled water), however, often contains NaCl concentrations of 30 mM or higher (Hayes et al., 1990;

Mancino and Pepper, 1992; Harivandi, 2004). Similarly, the physical decrease in irrigation will also lead to soluble salt aggregating within the turfgrass root zone.

The perched water table found within many constructed golf greens can also exacerbate issues associated with poor quality irrigation water or increased salt aggregation (Hummel, 1993; Ben-Hur et al., 2001; Schlossberg and Karnok, 2002;

Obear, 2014). Increased exposure to NaCl causes considerable stress to turfgrass plants, resulting in decreased recovery, reductions in quality, and potential decline. Many turfgrass species are not especially tolerant to salt stress, and the primary method used to decrease salt stress in turfgrasses, soil flushing, may not be a viable option in the future

(Harivandi et al., 1992; Prettyman and McCoy, 2000; Munns and Tester, 2008).

Accumulation of NaCl results in both osmotic and ionic stresses in plants

(Blumwald, 2000; Munns and Tester, 2008). The osmotic stress generated by increased levels of NaCl is a results of changes in soil and apoplastic water potential, and is considered to be the first phase of salt stress (Munns and Termaat, 1986; Munns, 1993,

2002; Tester and Davenport, 2003). As salinity stress progresses over time, Na+ rapidly

144 accumulates in the symplast, and results in ionic specific stress (Munns and Termaat,

1986; Munns, 1993, 2002; Tester and Davenport, 2003).

The uptake of Na+ results in the rapid depolarization of the plasma membrane, followed by the passive efflux of K+ to restore membrane potential, and decreases in the rate of K+ influx can also be observed (Blumwald, 2000; Shabala and Cuin, 2007). The increased Na+: K+ ratio generated by the symplastic accumulation of Na+ initiates the production of reactive oxygen species (ROS), resulting in enzyme inhibition, reductions in stomatal conductance, and decreased rates of photosynthesis (Maathuis and Antmann,

1999; James et al., 2002; Tester and Davenport, 2003).

Responses to NaCl stress fall into three categories: 1) removal of NaCl from the symplast, through sequestration in the vacuole (or transport back into the apoplast); 2) exclusion of NaCl from shoots; and 3) directional, escape-like, growth of roots away areas of high NaCl in the soil. Na+ can be transported directly into the vacuole for sequestration by Na+/H+ Exchanger 1 (NHX1), a vacuolar localized Na+/H+ antiporter which exhibits increased expression in the presence of salt (Apse et al., 1999; Shi and

Zhu, 2002; Apse et al., 2003; Zhu, 2003). In addition to sequestration in the vacuole, Na+ can also be removed from the symplast by transporting Na+ back into the apoplast, a process mediated by the salt-overly sensitive (SOS) family of proteins (Liu and Zhu,

1998; Zhu, 2002; Qiu et al., 2003; Julkowska and Testerink, 2015).

For turfgrasses, the majority salt stress response research has focused on osmotic stress associated with NaCl along with ROS production during salt stress. In both

Bermudagrass (Cynodon dactylon L.) and perennial ryegrass (Lolium perenne L.) the

145 expression of antioxidant enzymes (Superoxide Dismutase (SOD), Catalase (CAT),

Ascorbate Peroxidase (APX), etc.) has been shown to increase, and the activity of these enzymes has also been shown to increase when exposed to 200 and 400 mM NaCl, aiding in decreasing oxidative damage (Hu et al., 2011; Hu et al., 2012a). Glycine betaine, an osmolytic polyamine, application was shown to decrease salt stress in perennial ryegrass

Kentucky bluegrass (Poa pratensis L.), and creeping bentgrass (Agrostis stolonifera L.), and may be due to decreases in oxidative stress as well as potentially through balancing cellular water potential (Hu et al., 2012a; Yang et al., 2012).

Osmotic adjustment through the accumulation of osmolytes is considered to be a primary means of NaCl exclusion. Salt tolerant accessions of Bermudagrass have been shown to rapidly increase production of osmolytic compounds like proline as well as soluble sugar content when compared to salt-intolerant lines (Hameed and Ashraf, 2008).

For turfgrasses, exclusion of Na+ and Cl- from leaf tissue is also considered one the most prominent mechanisms of salinity tolerance (Marcum and Pessarakli, 2006). In zoysiagrass (Zoysia japonica L.) and Bermudagrass in particular, NaCl exclusion occurs through salt gland based secretion, and greater rates of secretion have been correlated with increased salt tolerance (Marcum and Murdoch, 1994; Marcum et al., 1998; Marcum and Pessarakli, 2006).

The maintenance of a high K+/Na+ ratio has been shown to decrease salt stress in

Bermudagrass; therefore, the transport and sequestration of Na+ is also an important salt tolerance mechanism in turfgrasses (Hu et al., 2012b). The zoysiagrass Na+/H+ antiporter

(ZjNHX1) was shown to exhibit Na+/H+ transport in yeast, and this transporter also

146 increases expression in zoysiagrass during salt stress (Du et al, 2009). In red fescue

(Festuca rubra ssp. litoralis L.) a vacuolar ATPase (FrVHA-B) and the Na+/H+ antiporter, NHX1 (FrNHX1), were both shown to increase expression following treatment with 500 mM NaCl in the endodermis and in vascular tissues, indicating the importance of NaCl sequestration in cool season turfrgasses (Diedhiou et al., 2009).

Phytohormones may also have an important role in turfgrass salt stress (Krishnan and Merewitz, 2015). Comparing the salt intolerant creeping bentgrass cultivar

‘Penncross’ to the more tolerant cultivar ‘Mariner’ has shown that the maintenance of higher absicic acid (ABA), jasmonic acid (JA), salicylic acid (SA), and indole 3-acetic acid (IAA) content in root tissue correlates with increased salt tolerance (Krishnan and

Merewitz, 2015). Increased production of, or maintained production of, phytohormones in roots during salt stress can help to maintain root growth under excessive NaCl conditions, or it can also lead to changes in root architecture that aid in decreasing NaCl uptake. For example, during salt stress, abscisic acid (ABA) functions to alter root architecture, upregulate expression of stress-related genes, promote stomatal closure, and decrease shoot growth (Grill and Himmelbach, 1998; Xiong et al., 2002; Duan et al.,

2013; Julkowska and Testerink, 2015).

The phytohormone auxin also regulates root architecture and development during salt stress (Julkowska and Testerink, 2015). Modulation of auxin metabolism and transport is essential for roots to adapt and respond to changing conditions, particularly the initiation of lateral roots and root tropic growth under stress (Eapen et al., 2005;

Julkowska and Testerink, 2015). Halotropism is a recently defined tropic growth

147 phenomena in which, upon exposure to a salt gradient, roots exhibit directional growth away from areas of high NaCl concentrations (Galvan-Ampudia et al., 2013; Rosquete and Kleine-Vehn, 2013). Halotropism is dependent on the asymmetrical re-distribution of auxin from the side of the root nearest to the salt, to opposite side of the root, and this change in auxin distribution results in root tip growth away from NaCl. (Galvan-

Ampudia et al., 2013; Rosquete and Kleine-Vehn, 2013).

As previously stated, the production of antioxidant enzymes and antioxidant metabolites increases salt tolerance in turfgrasses. Interestingly, phenolic antioxidants, flavonoids and anthocyanins, have also been shown to aid in decreasing salt stress; however their involvement may extend beyond merely reducing oxidative stress

(Eryilmaz, 2006; Yan et al., 2013). Flavonoids, in particular, have been shown to be involved in normal root gravitropism through the modulation of auxin metabolism and transport (Murphy et al., 2000).

Indole-3-acetic (IAA) acid is rapidly oxidized as it is transported into the symplast, and oxIAA, the product of IAA oxidation, is not an active auxin signaling molecule; therefore, the antioxidant capacity of flavonoids is thought to increase the total quantity of active auxin in the symplast through ROS quenching (Peer et al., 2013). The

Arabidopsis tt4 mutant is deficient in chalcone synthase (CHS - the first committed enzyme in flavonoid biosynthesis), and does not synthesize flavonoids (Peer et al., 2001).

This mutation results in an increase in auxin transport from shoot-to-root as well as slowed gravitropic bending, and results have shown that flavonoids modulate auxin transport both directly and indirectly through their interaction with auxin efflux proteins

148

(PIN and ABCB transporters) (Brown et al., 2001; Buer et al., 2004; Peer et al., 2004;

Williams et al., 2004; Geisler et al., 2005; Geisler and Murphy, 2006).

The turfgrass species rough bluegrass (Poa trivialis L.) has been shown to hyper- accumulate flavonoids and anthocyanins, and synthesis of flavonoids in this turfgrass can also be controlled in this species through specific light treatments (Petrella et al., 2016a,

2016b). Interestingly, this grass is also known to be salt sensitive relative to other turfgrasses, and rough bluegrass is used to overseed dormant Bermudagrass golf greens in the southern United States that may contain higher than normal levels of NaCl

(Camberato and Martin, 2004; Harivandi, 2004).

The establishment of rough bluegrass during overseeding is critical; therefore, it is necessary to evaluate the response of seedling rough bluegrass roots to NaCl. As halotropism has been shown to occur in seedlings of Arabidopsis, tomato, and sorghum, this tropic growth phenomena may also be important in the development of rough bluegrass seedlings under NaCl stress (Galvan-Ampudia et al., 2013). Based on the above, the objectives of this research were to A) determine if rough bluegrass seedlings exhibit halotropism, and B) to determine if flavonoids modulate halotropism in rough bluegrass through the application of light treatments that either promote or repress flavonoid accumulation.

149

Materials and Methods

LED light box construction

Light tight LED light boxes were constructed in two pieces using aluminum boxes

(Bud industries Inc., Willoughby, OH, USA). The upper half (LED enclosure) was an aluminum box with dimensions 30.5 x 30.5 x 7.6 cm, and the lower box (treatment enclosure) was an aluminum box with dimensions 30.5 x 30.5 x 25.4 cm (Appendix U and V). Both boxes were modified to allow ventilation, and heat sink fans (5.5 x 5.5 cm) were also attached to the upper side of the LED enclosure using thermal adhesive (Arctic

Silver® Inc. Visalia, CA, USA) to help cool the LEDs. On the LED enclosure, two square holes (4 x 3 cm) were cut using a dremmel tool and aluminum cutting wheel on opposing sides of the box. On the treatment enclosure one square hole was cut in a lower corner of the box, and four ventilation holes (5 cm diameter) were cut into the bottom of this box. Brushless centrifugal fans (12V; 75 x 75 x 30 mm) were attached to all three square holes using silicon caulking and aluminum tape, and were fans wired directly to a

12 V AC-DC switching power supply (LED Wholesalers® Hayward, CA, USA)

(Appendix W).

To prevent light from escaping through the centrifugal fans or bottom ventilation holes, cardboard baffles were constructed and attached (Appendix W). Cardboard tubes

(7.62 cm diameter) were cut into lengths of 8.89 cm, and were divided into two halves.

For each baffle, four cardboard circles (7.62 cm in diameter) were cut into unequal halves, and two halves were glued onto each side of the tube at unequal distances to

150 simultaneously allow air flow through the baffle while stopping light from escaping

(Appendix W).

Blue, red, cool white, and warm white (Appendix J and K) 5050 LEDs (5.0 x 5.0 mm) were purchased pre-mounted on flexible metal-core printed circuit strips - 60 LEDs m-1 (Torchstar Inc. ® La Puente, CA, USA). LED strips were cut into lengths of 25.4 cm and were attached side by side to the inner side of the LED enclosure using 3M® 8805 thermally conductive tape (3M® Medina, OH, USA). LED strips were wired in parallel, all wiring was passed through a hole drilled into the side of the LED enclosure (sealed with caulking), and the LEDs were powered using a 12 V AC-DC switching power supply. For experiments combining red/blue or cool/warm white LED light, LED strips were wired independently, and were connected to separate dimming units (LED

Wholesalers® Hayward, CA, USA) to allow for adjustment in spectral quality. For halotropism experiments, the top of the petri plates were 14 cm from the LEDs.

Irradiance was adjusted to 200 µmol m-2 s-1 using the attached dimmers, and simultaneous measurement using a LI-COR quantum sensor and LI-2189 light meter (LI-

COR® Biosciences, Lincoln NE USA). When applying light treatments, the LED and treatment enclosures were sealed together using aluminum tape.

LED array spectral distribution and quantum flux were measured using a

151 peak wavelength of 635 nm (4.61% of the spectrum) (Appendix E and F). Combined cool and warm white LEDs produced a bandwidth from 400-700 nm, and peak wavelengths were 447 nm (0.64% of the spectrum) and 585 nm (1.29% of the spectrum)

(Appendix J and K).

The environment within the LED light boxes was maintained through the four ventilation holes, attached centrifugal fans, and ventilation fans (425 m3 h-1 or greater) were also placed above the LED light boxes to apply a constant flow of air over the light boxes. The temperature and relative humidity within the chamber was monitored using a data logger (Watchdog microstation 1000 series, Spectrum Technologies, Aurora, IL,

USA). With all components running except for the LED lights, temperature within the box was 21.5°C and relative humidity was 30% on average. When the LED lights were turned on, temperature was 24.5°C and relative humidity did not change.

Seed sterilization

Rough bluegrass (cv. ‘Havana’) seed was surface sterilized using methods modified from Ke and Lee (1998). A 1 mL aliquot of seed was placed in a 15 mL centrifuge tube, and approximately 5 mL of 70% (v/v) ethanol was added. The tube was shaken for 30 seconds, and the ethanol was removed. The seed was then washed three times with ddH2O. Following being rinsed with water, 5 mL of a 2.6% sodium hypochlorite (Clorox splash-less bleach, The Clorox Company, Oakland, CA, USA) solution containing 0.2% SDS was added. Tubes were placed on a shaker and allowed to incubate for 45 minutes. The bleach solution was decanted, and the seed was washed

152 three times with ddH2O. To the rinsed seed, 5 mL of a 1.3% sodium hypochlorite with

0.2% SDS was added, and the tubes were placed on a shaker and allowed to incubate for

30 more minutes. Tubes were transferred to a laminar flow hood, seed was vacuum filtered, and the seed was rinsed with autoclaved ddH2O 6-8 times to remove any remaining bleach solution. The seed was allowed to dry within the laminar flow hood, was placed within a petri plate, and was sealed with micropore medical tape (3M®

Medina, OH, USA).

Plant growth conditions

Sterilized rough bluegrass seeds were placed on square petri plates (10 x 10 cm) containing 45 mL ¼ MS media (2.56 mM 2-(N-morpholino) ethanesulfonic acid (MES),

0.8% Phytoblend agar (Caisson Laboratories, Smithfield, UT USA), and pH 5.8). On the agar plates, seeds were placed on two horizontal lines (25 seeds per line) approximately 4 cm apart from one another. Plates were sealed with micropore medical tape, and were placed vertically within a growth chamber (24 hour day length, 60-80 µmol m-2 s-1, 22°C) for 6 days. On average seeds germinated within 2-3 days following placement, and seedlings 3-4 days post germination (DPG) were used in halotropism experiments.

Halotropism experimental methods

Halotropism agar-media was prepared as follows (Appendix Y) (methods modified from Galvan-Ampudia et al., 2013). Square petri plates (10 x 10 cm) were lined with marker on the bottom of the plate to divide the plates into two halves separated

153 by a 0.5 cm border. The petri plates were filled with 30 mL ¼ MS media (2.56 mM

MES, 0.8% Phytoblend, and pH 5.8), and were allowed to set for 1 hour. The lower half of the media was cut out, and 12.5 mL of media containing NaCl was poured into this space. Plates were allowed to set for 1 hour, were sealed with cling wrap, and were placed in the dark at 4°C for 8-12 hours prior to use.

Seedlings 3-4 DPG were transferred to the halotropism media by grasping plants at the base of the shoot using forceps. The root tip was placed on the media in proximity to where the upper most marker line appeared on the petri plate, and the entire plant was adjusted to ensure the root was straight. A total of ten plants were placed on each plate, and the plates were sealed with micropore medical tape. Plates were scanned at this “0 hour”, and plates were scanned at respective intervals to monitor root movement.

Following being scanned, a marker dot was placed on the back of the petri plate in proximity of where the root tip was located.

To determine if rough bluegrass exhibited halotropism, a halotropism experiment was designed using 0, 200, 250, 300, 350, and 400 mM NaCl. For each salt concertation, nine replicates (nine petri plates) with ten plants per plate were used, and the experiment was repeated two independent times. Following seedling placement, plates were vertically placed in a growth chamber (24 hour day length, 60-80 µmol m-2 s-1, 22°C).

Plates were scanned at 0, 12, 24, and 48 hours post transfer, and root angle was measured image J software (ImageJ developers, 2009).

154

Halotropism light box experiments

To determine if light quality modulates halotropism through changes in flavonoid metabolism, plants were exposed to a white LED (50:50 cool to warm white) light control, 100% blue LED light, 100% red LED light, or 70% blue - 30% red LED light at a an intensity of 200 µmol m-2 s-1 constantly. Plants were placed on media without salt (0 mM), or were placed on media containing 350 mM NaCl. For each light treatment, nine replicates with ten plants per plate were made for both 0 mM and 350 mM NaCl (18 total plates per light box). Halotropism media and plant placement followed the same methods as above, however, petri plates were placed in LED light boxes after being placed on their respective petri plates.

Plates were placed vertically within the described LED light boxes, the boxes were sealed with aluminum tape, and all electronics (fans etc.) were turned on except for the LED lights. For all LED treatments, plants were dark adapted for three hours, and following this period lights were turned on for the designated time period. The duration of the light box experiments were run for 12 hours (3 hours dark + 9 hours light) or for 24 hours (3 hours dark + 21 hours light). Plants were also analyzed for the response to treatment under darkness. In a similar fashion to LED light treatments, 0 and 350 mM plates were placed in light boxes with all components running except for the LED lights, and were treated for 24 hours only. Light box experiments were repeated three independent times. For dark treatment experiments, plants were placed in three separate light boxes, and were analyzed at the same time along with a white light control.

155

Following halotropism light box experiments, root tissue was harvested. Roots were cut at the root-shoot transition zone, and roots were placed into pre-weighed microcentrifuge tubes. Tubes were weighed, flash frozen in liquid nitrogen, and were stored at -80°C until tissue was extracted for flavonoid analysis.

NaCl measurements

To determine the quantity of NaCl that roots were being exposed to, the agar in proximity to the roots was analyzed using electrical conductivity (EC). Halotropism plates were made using the same methods as above using 350 mM NaCl. Plates were analyzed at 0 hours (directly post plant placement), 6, 12, 24, and 48 hours following placement onto the halotropism plates. For each time point five plates were analyzed, and a standard curve was made using standard agar plates with known quantities of NaCl.

At the specified time point, the agar between where the new media was poured, and the agar where the roots would be placed was removed. The agar section was placed into a

15 mL centrifuge tube and was mixed using a spatula. The electrical conductivity of the agar was measured using a field scout direct soil EC meter (Spectrum Technologies,

Aurora, IL, USA). Salt concentrations were calculated using a standard curve generated by measuring the EC of agar slabs containing known concentrations of salt.

Tissue extraction

To increase the quantity of tissue for high-performance liquid chromatography

(HPLC) analysis, root tissue within independent experiments was pooled, producing three

156 replicates per treatment (i.e. 3 samples for blue light 0 mM NaCl and 3 samples for blue light 350 mM NaCl, etc.). The internal standard, chrysin (97.5 µg mL-1), was first added, tissue was then triple ground in liquid nitrogen within the microcentrifuge tubes, and the ground tissue was suspended in 0.1% (v/v) acidified (HCl) methanol. Tubes were vortexed for 15 seconds, and were placed in a sonication bath (UCS-10, Jeio Tech Co.,

LTD., Seoul, Korea) set at high for 30 minutes (25-35°C) (. Following sonication, tubes were vortexed a second time, and were then centrifuged at 15,000 g for 10 minutes. The resultant extract was filtered into amber auto sampler vials using 13 mm 0.2 µm PTFE filters (Waters Acrodisc® syringe filter, Waters Corporation Milford, MA, USA).

Samples were stored in the dark at 4°C until analysis.

High Performance Liquid Chromatography (HPLC)

For HPLC analysis, a Beckman Coulter (Beckman Coulter Life Sciences, Brea,

CA, USA) autosampler, a Beckman Coulter system gold® 126 solvent module, an Alltech model 631 column heater, and a Beckman Coulter system gold® 168 photodiode array

(PDA) detector were used for sample analysis. For all samples, the column was maintained at 30°C, and samples were injected at 30 µL. Flavonoids and anthocyanins were analyzed using two independent methods.

Flavonoids were analyzed using a Phenomenex Gemini 5µm C6 phenyl column

(250 x 4.6 mm, 110 Å) (Phenomenex company, Torrance, CA, USA), and 340 and 520 nm were monitored simultaneously (methods modified from Paudel et al., 2013). The mobile phases consisted of (A) 0.2% acetic acid and (B) acetonitrile with a flow rate of

157

0.7 mL minute-1. The following gradient was used for separation: 5% B 1-10 minutes,

22% B at 20 minutes, 28% B at 45 minutes, 80% B at 53 minutes and held through 59 minutes, and 5% B at 72 minutes. Standard curves were produced for the internal standard, chrysin, rutin (a flavonoid), and caffeic acid (a hydroxycinammic acid)

Flavonoid and hydroxycinammic acid content was determined using internal standard calibration (Venton, 2017). The limit of quantification (LOQ) for flavonoids and hydroxycinammic acids was 1 µg mL-1.

Anthocyanins were analyzed using a Phenomenex Prodigy 5µm ODS-3 C18 column (250 x 4.6 mm 110 Å), and 254 and 520 nm were monitored simultaneously

(methods modified from Tulio et al., 2008). The mobile phases consisted of (A) 5% formic acid and (B) methanol with a flow rate of 0.8 mL minute-1. The following gradient was used for separation: 5% B 1-2 minutes, 60% B at 32 minutes and held through 37 minutes, 100% B at 42 minutes and held until 44 minutes, and 5% B at 52 minutes. A standard curve was produced for cyanidin, and anthocyanin content was determined using external standard calibration. All chromatographic data was analyzed using 32 Karat software (Beckman Coulter Life Sciences).

Anthocyanins and non-anthocyanin phenolics were identified using an Agilent

6495 LC-MS/MS (Agilent Technologies, Santa Clara, CA USA), and a Phenomenex

Gemini 5µm C6 phenyl column (250 x 4.6 mm, 110 Å). A mobile phase consisting of

(A) 1% formic acid and (B) acetonitrile was used to simultaneously separate flavonoids and anthocyanin using the flavonoid solvent gradient from above, a flow rate of 1.3 mL min-1 was used, and samples were injected at 10µL. Samples were analyzed using

158 electrospray ionization (ESI) in both negative and positive modes with an Agilent Jet

Stream (350°C, 3500 V capillary, 1000 V nozzle, 18 L m-1 drying gas, 12 L m-1 sheath gas, and 30 PSI nebulizing gas), and MS/MS fragmentation was carried out at a collision voltage of 25 eV.

Greenhouse experiment

To determine if the described LED light treatments altered rough bluegrass seedling establishment in saline soil, the following greenhouse experiment was performed using Lumigrow adjustable LED arrays (Lumigrow Pro 650, Lumigrow,

Emeryville, CA, USA). Plastic 10 x 10 cm pots were filled with soilless media (Metro-

® mix 360, Sungro Horticulture, Agawam, MA, USA), soil was saturated with ddH20 or with a 25 mM NaCl solution, and pots were allowed to drain for 24 hours prior to the start of the experiment. Rough bluegrass (‘Havana’) was seeded at a rate of 9.75 g m-2, and seedlings were exposed to the following supplemental light treatments (50 µmol m-2 s-1): white LED light (Appendix R and S), blue LED light (Appendix N and O), red LED light (Appendix P and Q), or a no supplemental light control. Four replicates of both salt treatments (0 and 25 mM NaCl) were included under each light treatment. Pots were bottom watered with their respective salt treatment (0 mM or 25 mM NaCl) every 4 days for 1 hour.

Light treatments were on individual greenhouse benches that were separated by two layers of black polyester cloth to minimize light contamination, and pots were placed on raised benches that set them 91 cm from the LED array(s). The experiment was run

159 for 18 days, and was repeated two independent times (Nov. 11th – 19th 2016 and Dec. 2nd to Dec. 20th 2016). The saturated electrical conductivity (EC) was measured prior to the start of the experiment, and the saturated EC was also measured at end of the experiment using a field scout direct soil EC meter. Shoot tissue was harvested at the conclusion of both experiments. Shoots were clipped at the soil surface, weighed, and were dried in an oven at 50°C for 3 days. Following drying, tissue was re-weighed to determine dry weight and water content.

Data analysis

Halotropism data was analyzed by measuring the root angle using Image J software (ImageJ developers, 2009). Two measurements were made per plant. The first measurement was the starting angle, or the angle the root was at when placed on the media, and the second measurement was the angle between where the root tip started and where the root tip ended at. A corrected angle was calculated for each plant by subtracting the initial measurement from the second measurement. Corrected angles were sorted, ranked, and grouped in 10° increments, and this data was used to create degree of root bending figures. Ranked corrected data was used to make treatment comparisons with respective control group(s) using Student’s T test (P = 0.05) to determine if a given treatment(s) was bending further away from 90° (Table 4.1).

Ranks were also used to determine what percentage of roots per petri plate were

A) gravitropic B) bending into the new media, or C) bending away from the new media.

Roots with corrected angles between -5 to 5° were categorized as gravitropic (90°

160 growth), roots between -85 to -5° were categorized as bending into the newly poured media, and roots between 85 to 5° were categorized as bending away from the new media

(Table 4.2). Within a single treatment the percent of roots bending in A) B) or C) directions were compared amongst each other using analysis of variance (ANOVA) (P =

0.05). Treatments were also compared back to their respective control(s) using Student’s

T test (P = 0.05). There was no statistical difference between experimental runs of all halotropism experiments; therefore, data for experiments was pooled for statistical analysis.

Corrected angle° Assigned rank Corrected angle° Assigned rank Greater than -105 1 6 to 15 13 -105 to -96 2 16 to 25 14 -95 to -85 3 26 to 35 15 -85 to -76 4 36 to 45 16 -76 to -66 5 46 to 55 17 -65 to -56 6 56 to 65 18 -55 to -46 7 66 to 75 19 -45 to -36 8 76 to 85 20 -36 to -26 9 86 to 95 21 -25 to -16 10 96 to 106 22 -15 to -6 11 Greater than 106 23 -5 to 5 12 Table 4.1: Categories for ranking data for degree of bending statistics

Corrected angle° Assigned rank -105 to -6 1 -5 to 5 2 6 to 106 3 Table 4.2: Categories for ranking data for percent of roots bending statistics

161

For metabolite quantitation, treatments were compared using Analysis of

Variance (ANOVA), and means were separated using Tukey’s Honest Significant

Difference (HSD) (P = 0.05). Data for the greenhouse experiment was analyzed as a percent of 0 mM NaCl control, and light treatments were compared using ANOVA, and means were separated using Tukey’s HSD (P = 0.05). All experiments were analyzed using a completely randomized design (CRD) and data were analyzed using Minitab software (Minitab 17 Statistical Software, 2010).

Results

Halotropism salt concentration test

To determine if rough bluegrass roots exhibited tropic growth following exposure to a gradient of salt, seedlings were exposed to 0, 200, 250, 300, 350, or 400 mM NaCl over a 48 hour period (Appendix Z). After 12 hours of treatment, seedlings treated 250,

350, and 450 mM NaCl exhibited a greater percentage of roots bending away from the salt compared to 0 mM (Table 4.3). At concentrations of 250, 350, and 400 mM NaCl, there was also a significantly lower percentage of roots growing into the salt compared to

0 mM NaCl.

162

Percent of Roots mM NaCl - 45° to 90°y 90°x 90° to 45°w 0 22.21 Bz 62.71 A 14.53 B 200 18.45 B 58.32 A 22.54 B 250 12.74* C 56.63 A 30.07** B 300 14.10 B 59.81 A 26.09 B 350 7.1** C 57.26 A 35.63*** B 400 3.46*** C 63.84 A 34.66*** B * Within column comparison back to 0 mM using Student’s T test (P=0.05) * = P ≤ 0.05; ** = P ≤ 0.01; *** = P ≤ 0.001 z Capital letters represent comparison within rows using ANOVA (P=0.05) y The percent of roots growing into the salt or newly poured media x The percent of roots growing towards the vector of gravity w The percent of roots growing away from the salt or newly poured media

Table 4.3: Salt concentration test. 12 hours post treatment

At 24 hours post NaCl treatment, only seedlings treated with 400 mM NaCl exhibited a greater percentage of roots bending away from the salt relative to 0 mM, but

300, 350, and 400 mM treatments exhibited a smaller percent or roots growing into the salt compared to 0 mM (Table 4.4). Treatment with 350 and 400 mM NaCl did not result in a significant difference between gravitropic and halotropic roots, however, both treatments exhibited a significantly smaller percentage of roots bending into the salt

(Table 4.4).

Following 48 hours of salt treatment, only 400 mM NaCl treatment resulted in a significantly greater percentage of halotropic roots compared to 0 mM (Table 4.5).

However, for 300 and 350 mM treatments, there was still a significantly greater percentage of roots bending away from the salt compared to bending into the salt.

163

Interestingly, when treated with 200 and 250 mM NaCl, there was a greater percentage of roots bending into the salt compared to 0 mM (Table 4.5).

Percent of Roots mM NaCl - 45° to 90°y 90°x 90° to 45°w 0 25.94 Bz 47.72 A 26.33 B 200 23.33 B 52.09 A 24.58 B 250 18.30 B 53.05 A 28.64 B 300 15.21** C 50.39 A 34.41 B 350 9.68*** B 52.76 A 37.00 A 400 5.29*** B 44.96 A 51.71*** A * Within column comparison back to 0 mM using Student’s T test (P=0.05) * = P ≤ 0.05; ** = P ≤ 0.01; *** = P ≤ 0.001 z Capital letters represent comparison within rows using ANOVA (P=0.05) y The percent of roots growing into the salt or newly poured media x The percent of roots growing towards the vector of gravity w The percent of roots growing away from the salt or newly poured media

Table 4.4: Salt concentration test. 24 hours post treatment

The degree to which roots were bending under salt conditions was also determined (Fig. 4.1). At 12 hours 250 - 400 mM NaCl treatments resulted in roots growing away from the newly poured media to a larger degree compared to roots treated with 0 mM NaCl. Following 24 hours, roots treated with 300 – 400 mM NaCl exhibited a larger degree of halotropic growth compared to 0 mM, and at 48 hours only 400 mM led to a greater degree of halotropic bending (Fig. 4.1). Also, both 200 and 250 mM

NaCl treatments led to a significantly smaller degree of bending compared to 0 mM.

Plants under control conditions, 0 mM, naturally exhibited a greater degree of bending into the newly poured media when comparing 12 and 48 hours post treatment (Fig. 4.1).

164

Percent of Roots mM NaCl - 45° to 90°y 90°x 90° to 45°w 0 9.46 Cz 61.98 A 28.56 B 200 24.3** B 55.54 A 20.15 B 250 33.57*** AB 44.74** A 21.70 B 300 17.66* C 50.32 A 32.02 B 350 11.25 C 58.16 A 30.60 B 400 5.82 B 54.78 A 41.36* A * Within column comparison back to 0 mM using Student’s T test (P=0.05) * = P ≤ 0.05; ** = P ≤ 0.01; *** = P ≤ 0.001 z Capital letters represent comparison within rows using ANOVA (P=0.05) y The percent of roots growing into the salt or newly poured media x The percent of roots growing towards the vector of gravity w The percent of roots growing away from the salt or newly poured media

Table 4.5: Salt concentration test. 48 hours post treatment

Determination of the salt concentration in proximity to rough bluegrass roots

The previous experiment demonstrated that treatment with 350 mM NaCl resulted in consistent halotropism in rough bluegrass without being detrimental to plant growth.

However, in this NaCl gradient system, NaCl is diffusing slowly towards the roots, and roots are being exposed to increasing concentrations of NaCl over time. Because of this, it was necessary to determine what concertation of NaCl roots were being exposed to over the course of the 48 hour experiment. Results show that at the time when rough bluegrass plants are placed on the halotropism gradient petri plates (0 hours), roots are exposed to approximately 100 mM NaCl on average (Fig. 4.2). Following 6 hours of salt treatment roots are exposed to 105 mM NaCl, 112 mM at 12 hours, 118 mM at 24 hours, and 130 mM at 48 hours (Fig. 4.2).

165

* Within row comparison back to 0 mM using Student’s T test (P=0.05) † Within column comparison back to the 12 hour time point using Student’s T test (P=0.05) *(†) = P ≤ 0.05; **(††) = P ≤ 0.01; ***(†††) = P ≤ 0.001

Figure 4.1: The degree of root bending following 12, 24. And 48 hours of treatment with various concentrations of NaCl (mM). Numbers within the center of the circles are the total number of plants represented by that treatment. The blue bar represents the percent of roots growing gravitropically. Bars to the right of the blue bar represent roots growing away from the new media (i.e. salt), and bars to the left of the blue bar represent roots growing into the new media (i.e. salt).

166

Modulation of halotropic growth by light

Results show that rough bluegrass exhibits halotropism when exposed to a gradient of salt. To determine what impact flavonoid accumulation had on halotropic growth, plants were treated with white, blue, red, or a combination of red and blue

(70:30) LED light to promote or repress flavonoid synthesis. Following 12 hours of treatment, results show that treatment with white light and 350 mM NaCl resulted in a larger percent of roots bending away from the salt, relative to plants treated with 0 mM

NaCl (Table 4.6). When treated with blue light, there was also larger percent of roots bending away from the salt, and there was also a smaller percent of roots growing into the newly poured media when comparing 350 and 0 mM NaCl treatments.

For red light exposure, however, 350 mM NaCl treatment did not result in an increase in the percentage of roots bending away from the salt, and plants treated with

350 mM NaCl here also exhibited a smaller percent of roots bending away compared to white light. When blue light was added back to spectrum at 70% of the total photosynthetic photon flux (PPF), root distribution under 0 and 350 mM NaCl was similar to blue light treated plants. Compared to white light, plants treated with blue light and salt exhibited a smaller percent of roots bending into the salt and a lager percent of roots bending away (Table 4.6).

167

Figure 4.2: The concentration of salt in close proximity to the roots of rough bluegrass following placement (0 hours) onto the salt gradient plates and up to 48 hours following placement.

Evaluating the degree to which roots bent during light treatment showed that all light treatments resulted in some degree of halotropism (Fig. 4.3). White, blue, and blue/red light treatments led to roots bending to a further in the 90° to 45 direction (away from the salt) compared to 0 mM treated plants. Red light treated plants under 350 mM conditions did also bend to a statistically greater degree compared to 0 mM plants, but the degree to which they bent was also significantly less than white light (Fig. 4.3). Blue light and the combination of blue and red light, however, resulted in roots that bent to a greater degree away from the salt relative to white light, unlike red light treatments.

168

Percent of Roots Light mM NaCl - 45° to 90°x 90°w 90° to 45°v 0 17.41 BZay 71.73 Aa 13.33 Ba White 350 12.25 Ca 56.09 Ab 31.66 Bb 0 16.91 Ba 69.05 Aa 14.03 Ba Blue 350 5.68** Cb 53.09 Ab 41.23* Bb 0 20.91 Ba 66.30 Aa 12.43 Ba Red 350 16.15 Ba 65.59* Aa 18.26*** Ba Blue + 0 15.47 Ba 73.62 Aa 10.91 Ba Red 350 9.01 Cb 57.82 Ab 33.17 Bb * Within column comparison back to 0 mM using Student’s T test (P=0.05) * = P ≤ 0.05; ** = P ≤ 0.01; *** = P ≤ 0.001 z Capital letters represent comparison within rows using ANOVA (P=0.05) y Lower case letters represent comparison between 0 and 350 mM NaCl within a given light treatment x The percent of roots growing into the salt or newly poured media w The percent of roots growing towards the vector of gravity v The percent of roots growing away from the salt or newly poured media

Table 4.6: 12 hour light box experiment; percent of root distribution.

Following 24 hours of treatment with white light and 350 mM NaCl, a larger percent of roots bent away from the salt, relative to plants treated with 0 mM NaCl, similar to 12 hours of treatment (Table 4.7). When treated with blue light, there was a larger percent of roots bending away from the salt, and there was also a smaller percent of roots growing into the newly poured media when comparing 350 and 0 mM NaCl treatments (Table 4.7). For plants treated with red light and 350 mM NaCl, there was a greater percent of roots bending away from the salt when compared to 0 mM, and there was also significantly greater percent of roots bending into the salt for this treatment

(Table 4.7). However, when examining the percent of root movement of plants treated with 350 mM NaCl under red light, it was evident that there was no difference in the

169 percent of roots bending towards or away from the salt, indicating that there was an overall increase in non-directional root bending (Table 4.7). Adding blue light back to the monochromatic red light spectrum led to a restoration of the blue light phenotype, similar to the results of 12 hours of treatment.

Plants were also treated under dark conditions, in the same environment as the

LED treatments, to evaluate the requirement of light in the halotropic response. Dark treatment did not change the percentage of root movement when comparing 0 and 350 mM NaCl treatment (Table 4.7). However, treatment of plants in the dark led to an increase in the roots bending in the - 45° to 90° direction (into the salt/new media) compared to white light, and there was also a significant decrease in the percent of roots bending away from the 350 mM NaCl treatment compared to white light (Table 4.7).

Blue light treated increased the percent of roots bending away from the salt relative to white light. Red light treated plants exhibited a smaller percent of roots bending away from the salt compared white light, and these plants also exhibited a larger percent of roots bending into the salt compared to white light (Table 4.7). White, blue, and blue/red light treatments resulted in a significantly greater degree of root bending when exposed to

350 mM NaCl compared to 0 mM NaCl (Fig. 4.4). Under red light or dark conditions, however, there was no significant difference in the degree of root bending between 0 and

350 mM (Fig. 4.4). Both blue light and the combination of blue and red light exhibited a greater degree of root bending under 350 mM NaCl compared to white light. On the other hand, both red light and dark treatments led to a significant decrease in the degree to which roots bent from the salt when compared to white light (Fig. 4.4).

170

* Within row comparison back to white light using Student’s T test (P=0.05) † Within column comparison between 0 and 350 mM NaCl in a given light treatment using Student’s T test (P=0.05) *(†) = P ≤ 0.05; **(††) = P ≤ 0.01; ***(†††) = P ≤ 0.001

Figure 4.3: The degree of root bending following 12 hours of treatment with 0 or 350 mM NaCl in combination with white light, blue light, red light, or the combination of blue and red light. Numbers within the center of the circles are the total number of plants represented by that treatment. The blue bar represents the percent of roots growing gravitropically. Bars to the right of the blue bar represent roots growing away from the new media (i.e. salt), and bars to the left of the blue bar represent roots growing into the new media (i.e. salt). 171

Percent of Roots Light mM NaCl - 45° to 90°x 90°w 90° to 45°v 0 13.37 Bzay 74.42 Aa 12.21 Ba White 350 14.05 Ca 54.06 Ab 31.89 Bb 0 15.03 Ba 72.89 Aa 12.08 Ba Blue 350 9.06* Cb 56.65 Ab 38.28* Bb 0 13.54 Ba 71.17 Aa 15.29 Ba Red 350 25.98** Bb 51.23 Ab 22.42** Bb Blue + 0 13.07 Ba 75.06 Aa 11.87 Ba Red 350 10.20 Ca 55.01 Ab 34.79 Bb 0 19.26* Ba 64.44** Aa 15.93 Ba Dark 350 21.40** Ba 59.18 Aa 19.42*** Ba * Within column comparison back to 0 mM using Student’s T test (P=0.05) * = P ≤ 0.05; ** = P ≤ 0.01; *** = P ≤ 0.001 z Capital letters represent comparison within rows using ANOVA (P=0.05) y Lower case letters represent comparison between 0 and 350 mM NaCl within a given light treatment x The percent of roots growing into the salt or newly poured media w The percent of roots growing towards the vector of gravity v The percent of roots growing away from the salt or newly poured media

Table 4.7: 24 hour light box experiment; percent of root distribution.

172

Figure 4.4: The degree of root bending following 24 hours of treatment with 0 or 350 mM NaCl in combination with white light, blue light, red light, or the combination of blue and red light. Numbers within the center of the circles are the total number of plants represented by that treatment. The blue bar represents the percent of roots growing gravitropically. Bars to the right of the blue bar represent roots growing away from the new media (i.e. salt), and bars to the left of the blue bar represent roots growing into the new media (i.e. salt). 173

Metabolite analysis of rough bluegrass roots

To evaluate changes in the phenolic profile of rough bluegrass roots exposed to both salt and light treatment, root samples were extracted and analyzed by high- performance liquid chromatography diode array detection (HPLC-DAD). After being treated for 24 hours, roots exposed to blue light and the combination of blue and red light accumulated anthocyanins between 1.25 – 3.11 mg 100 g-1 FW, but there was no statistical difference between treatments (Table 4.8). Anthocyanins were below the limit of detection (LOD) for all other treatments, and at 12 hours.

Light mM NaCl Cyanidin eq. (mg 100 g-1 FW z) 0 Below LODz White 350 Below LOD 0 3.11 ± 1.55 A Blue 350 2.57 ± 0.46 A 0 Below LOD Red 350 Below LOD 0 1.25 ± 0.43 A Blue + Red 350 2.48 ± 0.48 A z FW = Fresh Weight y LOD = Limit of detection

Table 4.8: Anthocyanin concentration of roots treated with 24 hours of light and 0 or 350 mM NaCl. Treatments are compared using ANOVA (P = 0.05) and Tukey’s Honest

Significant Difference (HD). Different letters indicate a significant statistical difference

Results are presented as the average concentration ± standard error (SE).

174

Prior to halotropism experiments flavonoids were below the limit of quantification (LOQ), and the primary phenolic compounds present were hydroxycinammic acids at an average concentration of 286.11 µg g-1 FW. At 12 hours post treatment, only roots treated with blue light and the combination of blue and red light exhibited flavonoid concentrations above the LOQ (Fig. 4.5) (Appendix AA).

There were no significant differences in flavonoid concentration between light treatments or between plants treated with 0 or 350 mM NaCl; however, plants treated with salt exhibited a trend of increasing flavonoid concentration. Following 24 hours of treatment, blue light and blue/red light treated plants produced greater quantities of flavonoids relative to 12 hours, and plants treated with white light and 350 mM exhibited detectable quantities of flavonoids that were statically less than blue and blue/red treated plants (Fig. 4.5) (Appendix BB). Flavonoids were below the LOQ for all other treatments at 24 hours. At both 12 and 24 hours following treatment there was no statistical difference in hydroxycinammic acid content between all treatments, but concentrations did increase slightly over the 12 hour period (Fig. 4.6)

175

Figure 4.5: Quantification of flavonoids accumulated in rough bluegrass roots following

A) 12 hours of treatment and B) 24 hours of treatment. Prior to treatments, flavonoids were below LOQ. Error bars represent treatment standard error (SE), and letters represent mean separation using Tukey’s HSD (n = 3). Different letters indicate a significant statistical difference (P = .05). LOQ = Limit of quantification (1µg mL -1).

176

Figure 4.6: Quantification of hydroxycinammic acids accumulated in rough bluegrass roots following A) 12 hours of treatment and B) 24 hours of treatment. Prior to treatment, hydroxycinammic acids were 286.11 µg g-1 FW. Error bars represent treatment standard error (SE), and letters represent mean separation using Tukey’s HSD

(n = 3). Different letters indicate a significant statistical difference (P = .05). LOQ =

Limit of quantification (1µg mL -1).

177

Phenolic Identification

Three anthocyanin peaks were present in rough bluegrass roots following treatment with 350 mM NaCl and blue light (Fig. 4.7). In negative and positive ion modes, peak 1 exhibited a molecular ion of 447/449, and a mass fragment of 285/287, indicating a cyanidin agyclone (Table 4.9). Here, a neutral loss of 162 is indicative of a hexoside (- H2O), and peak 1 can be tentatively identified as cyanidin 3-glucoside based on previous literature (Petrella et al., 2016). Peak 2 was similar to Peak 1, however the molecular ion was 489/535. In positive mode, a molecular ion of 535 and a fragment of

287 indicate a neutral loss of 162 for hexose as well as loss of 86 for malonyl (- H2O).

The third anthocyanin present exhibited a molecular ion of 301, indicating the presence of a peonidin agyclone (Table 4.9). In positive mode, a molecular ion of 549 and a fragment of 301 indicate a neutral loss of 162 for hexose as well as loss of 86 for malonyl

(-H2O).

178

Figure 4.7: Chromatogram (520 nm) of anthocyanins induced in rough bluegrass roots exposed to 350 mM NaCl and blue light. 1 = Cyanidin 3-glucosisde; 2 = Cyanidin 3- malonylglucoside; 3 = Peonidin 3-malonylglucoside; INTSD = Chrysin internal standard

[M+H]- [M+H]+ RT [M+H]- Fragments Fragments UV-vis Peak (min.)z [M+H]+y (m/z)x (m/z)w (nm)v I.D.u 1 21 447/449 285 287 520 Cy 3-g 2 25.2 489/535 285 287 520 Cy 3-mg 3 27.1 547/549 285,447 301 520 Pn 3-mg z Retention time (minutes) y Molecular ion in negative and positive modes x Mass fragments in negative ion mode w Mass fragments in positive ion mode v Peak UV-vis absorption u Compounds identification Cy 3-g = cyanidin 3-glucoside Cy 3-mg = cyanidin 3-malonylglucoside Pn 3-mg = peonidin 3-malonylglucoside

Table 4.9: Mass spectrometry data for anthocyanins induced in rough bluegrass roots exposed to 350 mM NaCl under blue light.

179

The non-anthocyanin phenolic metabolites aligned for rough bluegrass leaf and root tissue treated with 350 mM NaCl under blue light conditions (Fig. 4.8). Results therefore indicate that phenolics found within both tissues would be similar. Rutin and luteolin 7-glucosude eluted from the column at similar times relative to the flavonoids present in rough bluegrass (Fig. 4.8). The UV-vis absorption spectra of flavonoids in rough bluegrass roots (257, 270, and 350 nm peak) was more similar to luteolin (255,

267, and 349 nm peaks) than to rutin (257 and 356 nm peaks) (Fig. 4.9). Peaks 5 and 6 present in rough bluegrass roots exhibited a similar UV-vis spectra compared to caffeic acid; however, retention time here was not similar to caffeic acid (Fig 4.8 and 4.9).

Evaluating mass spectromery data for non-anthocyanin phenolics shows that for peaks 1-3 a mass fragment of 285 and 287 can be found in negative and postive ion modes (Table 4.10). Mass fragments of this size indicate the presence of either luteolin or kampferol. However grasses in general are considered to accumulate flavone agyclones and glycosides like luteiolin rather than flavonols (Dong et al., 2014). For peak 1, neutral loss of 264 from the molecular ion of 609 and the agyclone indicates two hexoses of 162 (-H2O). The moleucular ion found in peak 2, 579/581, yields 294 when subracted from 285/287, and this indictes the presence of a pentose, 132 (-H2O) and a hexose (-H2O). Peaks 5 and 6 represent hydroxycinammic acids based on the UV-vis absportion spectra, and the mass fragments present (Table 4.10). A mass fragment of 161 is indicative of caffeic acid (-H2O). The data indicate a tentative identification of a caffeic acid hexoside dimer; however the increased mass observed in peak may also

+ - incidate the presence of Na , Cl , or even H2O adducts (Table 4.10).

180

Figure 4.8: Chromatograms (340 nm) of A) Standard phenolic mix, B) Luteolin 7- glucoside standard, C) Rough bluegrass non-anthocyanin phenolic leaf extract, and C)

Rough bluegrass non-anthocyanin phenolic. 1) Luteolin dihexoside, 2) Luteolin pentosylhexoside, 3) Luteolin hexoside 4) Unknown Luteolin derivative, 5) Caffeic acid hexoside dimer, 6) Caffeic acid hexoside dimer, and INTSD = chrysin internal standard.

181

182

RT [M+H]- [M+H]- Fragments [M+H]+ Fragments UV-vis Peak I.D.u (min.)z [M+H]+y (m/z)x (m/z)w (nm)v 1 26.6 609/611 285,298,327,357,489 287,299,329,353 350 L-di-Hex

285,298,309, 327, L-Pent- 2 26.9 579/581 287,299,329,353 350 357, 459 Hex

3 27.6 447/449 285,297,327,339,357 287,299,329,353 350 L-Hex

Caffeic acid 5 30.1 623/642 161,461 N/A 290/330 Hexoside dimer Caffeic acid 6 36.1 665/684 161,461 N/A 280/330 Hexoside dimer z Retention time (minutes) y Molecular ion in negative and positive modes x Mass fragments in negative ion mode w Mass fragments in positive ion mode v Peak UV-vis absorption u Compounds identification L-di-Hex = luteolin dihexoside L-Pent-Hex = luteolin pentosylhexoside L-Hex = luteolin hexoside

Table 4.10: Mass spectrometry data for non-anthocyanin phenolics induced in rough bluegrass roots exposed to 350 mM NaCl under blue light.

Greenhouse experiment

To determine if the application of specific light treatments could improve establishment of rough bluegrass seedlings through modulating flavonoid metabolism, seedlings were treated with 0 or 25 mM NaCl in soilless media and were exposed to no supplemental light, white, blue, or red supplemental light (50 µmol m-2 s-1) for 18 days.

Successive applications of 25 mM NaCl increased the electrical conductivity (EC; SEC =

183 saturated EC method) of the soilless media upwards of 3.2, and control media had an average SEC of 1.6. There was no statistical difference for the SEC among any and all treatments.

For percent control data, there was no statistical difference between experiments 1 and 2, therefore data was pooled. All light treatments exhibited a decline in biomass following NaCl treatment, as seen with the percent of control data (Figure 4.10). There was no difference among light treatments in their ability to decrease NaCl stress, but there was a trend for red light to decrease stress compared to the other treatments.

Figure 4.10: Biomass percent of control (0 mM NaCl) for rough bluegrass treated with

25 mM NaCl. Error bars represent treatment standard error (SE), and letters represent mean separation using Tukey’s HSD (n = 3). Different letters indicate a significant statistical difference (P = .05). 184

Discussion

Halotropic growth in rough bluegrass

When treated with a gradient of NaCl over the course of a 48 hour period, rough bluegrass exhibited consistent halotropic growth when using 350 mM NaCl, and to date rough bluegrass is only the third monocot shown to exhibit halotropic growth following exposure to salt gradients (Galvan-Ampudia et al., 2013; Han et al., 2017 unpublished).

While concentrations below 350 mM did exhibit changes in the vector of root growth, results were variable between 12 and 48 hours. In particular, 300 mM NaCl treatment resulted in an increase in halotropism at 24 and 48 hours. Here, however, the percentage of halotropic roots was never greater than 0 mM, and at 12 hours 300 mM treatment did not lead to the halotropic growth which was seen in lower concentrations. These results suggest that concentrations lower than 350 mM result in root movement that may be due to other outside variables; in particular, because 0 mM treatment also increased bending into the newly poured media over time. Interestingly, at the lower concentrations of NaCl evaluated (200 and 250 mM NaCl) root movement into the salt increased over time relative to 0 mM, suggesting that a series of alternate NaCl interactions may occur in rough bluegrass roots.

Modulation of halotropic growth by spectral composition

To evaluate if the observed halotropic response was modulated by flavonoids, rough bluegrass was treated with flavonoid-inducing light conditions. Over both 12 and

24 hour periods blue light promoted and was required for halotropism in rough bluegrass.

185

Under 24 hour dark conditions, halotropism did not occur relative to 0 mM or white light control. These results suggest that in rough bluegrass, light is required for halotropic growth. This may be due to changes in root growth rates, however, previous research in

Arabidopsis has shown that under dark conditions halotropism is increased, and roots are more sensitive to lower concentrations of NaCl in darkness compared to roots that are exposed to light (Yokawa et al., 2014). Further, under dark treatment there is a greater percent of roots bending into the salt and a smaller percent of roots bending away from the salt compared to white light. These data were similar to what was seen when rough bluegrass was treated with 200 or 250 mM NaCl, and suggest that in darkness rough bluegrass roots are less sensitive to salt and increase foraging in multiple directions.

When treated with red light for 12 hours, roots exposed to 350 mM NaCl bent away from the salt to a greater degree compared to 0 mM treated plants. However, the degree to which red light treated plants bent was significantly less compared to white light, and a significantly smaller percent of roots bent away from the salt under red light compared to white light. At 24 hours post salt treatment, red light did not increase the degree to which roots bent away from the NaCl containing media, and red light led to a significant decline in halotropic growth compared to white light. When evaluating the percent of root distribution for red light treated plants, however, there was a significant increase in the percent of roots bending into the newly poured media when comparing 0 and 350 mM. Further examination shows that this is primarily due to the randomization of root growth under red light, in that there was also a significant increase in the percent of roots bending into the newly poured media when comparing 0 and 350 mM. Under

186 red light and salt treatment, there is a decrease in root gravitropism and a subsequent increase in randomized root growth, while treatment with blue light led to focused directional growth.

Following 12 and 24 hour treatment of roots with 350 mM NaCl and exposed to blue light led to an increase in the percent of roots bending away from the salt and a decrease in the percent of roots bending into the salt compared to white light. The shift in the direction of root movement suggests focused directional growth away from high concentrations of NaCl rather than the randomized root growth pattern seen with red light treatment. Blue light increased halotropic growth compared to white light as early as 12 hours; therefore, a greater proportion and increasing quantity of blue light may be promoting and enhancing halotropic growth through a faster/earlier response. When blue light was combined with red light in a high proportion (70:30), halotropic growth was restored to what was observed with white light and 100% blue light.

Our data shows that halotropism does not occur under dark conditions, halotropism is lost under continuous red light treatment, blue and red light application restore halotropism, and given that blue light enhances halotropic growth relative to white light all suggest that A) light is required and B) more specifically blue light is required for halotropism in rough bluegrass.

Phenolic metabolite accumulation in rough bluegrass roots

I hypothesized that blue light modulate halotropic growth through alterations in flavonoid metabolism, and the results presented above show that specific light treatments

187 modulate halotropism as well as increase flavonoid content in rough bluegrass. Only treatment with blue or blue/red light for 24 hours increased anthocyanin content to detectable levels, and anthocyanins present are similar to what has been found in rough bluegrass and creeping bentgrass (Agrostis stolonifera L.) (Nangle et al., 2015; Petrella et al., 2016a) leaves. Greater concentrations of anthocyanin in leaves has been linked to increased salt stress tolerance, but their contribution to root salt tolerance has not been investigated (Appendix DD) (Van Oosten et al., 2013).

When evaluating the non-anthocyanin phenolic profile of rough bluegrass roots, five major peaks were visible in plants that were treated under blue light and salt; and, most importantly, only plants treated with blue, blue/red, and white light exhibited flavonoid peaks that were above the LOQ (Appendix AA, BB, and DD). Peaks 1-3 exhibited UV-vis spectra and retention times similar to luteolin and rutin flavonoids. The mass fragmentation profile from all flavonoid peaks, however, indicate the presence of either luteolin or kaempferol due to the presence of a fragment of 285/287 (301 or 303 would indicate a quercetin agyclone). The non-anthocyanin phenolic metabolite profile of grass roots has not been evaluated to great extent in previous literature, but it has been reported that grasses accumulate flavone glycosides in leaves, suggesting the agyclone here is luteolin and not kaempferol (Cavaliere et al., 2005; Wojakowka et al., 2012; Dong et al., 2014). Based on this we can hypothesize that in rough bluegrass roots, luteolin is the primary agyclone present due to rough bluegrass leaf tissue exhibiting peaks with similar elution order and retention times.

188

Peaks 5 and 6 exhibited UV-vis spectra unique for hydroxycinammic acids, and mass spectra indicate the presence of caffeic acid hexoside dimers. Prior to any light or salt treatment, only peaks associated with caffeic acid hexoside dimers were above the

LOD, and following treatment these compounds did not show significant change between any treatment (Appendix AA, BB, and FF). Overall this suggests that while caffeic acid hexoside dimers are main constituents of rough bluegrass roots, their presence does not alter stress tolerance, and only changes in flavonoid content may be linked to difference in halotropic root growth observed here.

As halotropism occurred in white light, and was enhanced under blue light, the metabolite data presented here suggest that blue light modulation of halotropic growth may be through increases in flavonoid content specifically. It is well known that flavonoids modulate tropic growth and auxin related responses through ROS quenching, and flavonoids have been shown to increase auxin retention by decreasing auxin efflux through the modulation of PIN protein trafficking and direct inhibition of ABCB transporters (Peer et al., 2001; Peer et al., 2004; Williams et al., 2004; Peer et al., 2004;

Peer and Murphy 2007; Peer et al., 2013; Buer and Muday, 2014).

The Arabidopsis tt4 (transparent test 4) mutant is deficient in chalcone synthase

(CHS), the first committed step in flavonoid biosynthesis, does not synthesize flavonoids, and previous results have shown that this mutant displays delayed gravitropism due to increased auxin transport which could be recovered through flavonoid supplementation

(Brown et al., 2001; Peer et al., 2001; Buer and Muday, 2014). Similar results have been shown in the tomato are (anthocyanin reduced) mutant which is deficient in flavonoid 3

189 hydroxylase (F3H), the enzyme that produces flavonols (Maloney et al., 2014). The tomato are mutant displays enhanced rates of auxin transport from the shoot-to-root, and this mutant exhibited increased auxin content at the root tip relative to actively maturing root zones (Maloney et al., 2014). Similar to auxin transport inhibitors 1-N- naphthylphthalamic acid (NPA) and 2,3,5- triiodobenzoic acid (TIBA), the application of flavonols has also been shown to rescue other auxin transport mutations, including those mutants who do not initiate rhizobia nodulation (Pin Ng et al., 2015).

Aglycone flavonoids have primarily been indicated in the modulation of auxin transport, and flavonols, in particular, have been shown to inhibit auxin transport due to

Arabidopsis being unable to synthesis flavones (Brown et al., 2001; Peer et al., 2001;

Dong et al., 2014). My data do not indicate the presence of flavonols or agylcone flavonoids. However, the flavone apigenin has been shown to inhibit auxin transport similar to quercetin; therefore, as luteolin and apigenin share many common structural features, luteolin may exhibit an inhibitory response similar to apigenin (Jacobs and

Rubery, 1988). Whether or not only agylcones inhibit auxin transport is also questionable. Recently, the flavonoid sugar conjugate kaempferol 3,7-rhamnoside has also been shown to inhibit auxin transport (Yin et al., 2014). Similar to agylcones, glycosylated flavonoids can also be found within the cell wall, indicating that their presence in tissue extracts here are important in modulating auxin transport during halotropism (Strack et al., 1988; Winkel-Shirley, 2001; Agati et al., 2012).

Blue light may also modulate halotropism through other mechanisms. The auxin efflux transporter, PIN3, has been shown to be polarized under unilateral blue light

190 illumination of roots, leading to the re-direction of auxin transport, and resulting in root movement away from blue light (Zhang et al., 2013). The blue light photoreceptor

PHOT1 (Phototropin 1) has been shown to modulate root growth under drought conditions, but PHOT1 function may also only be limited to upper more mature regions of the root where it has been shown to accumulate to a greater degree (Sakamoto et al.,

2002; Galen et al., 2006). While blue light and PHOT1 lead to changes in auxin transporter localization, these responses have been show to occur in mature regions of the root, and PHOT1 expression is limited or absent in regions of the root that are involved in halotropic growth (Sakamoto and Briggs, 2002).

Flavonoids also play a role in regulating redox metabolism, and when compared to other flavonoids, luteolin has been shown to have a more planar structure resulting in a low activation energy requirement for H+ donation and electron transfer reactions (van

Acker et al., 1996; Leoplddini et al., 2004). Luteolin has also been shown to have a relatively high antioxidant capacity in vivo and in vitro (Wolfe and Liu, 2008). The production of reactive oxygen (ROS) species is known to rapidly increase during salt stress, and the inability to balance cellular redox potential is linked to NaCl related toxicity (Zhu et al., 2007; Miller et al., 2010; Bose et al., 2014). Greater quantities of

ROS are also detrimental for auxin signaling as IAA is rapidly oxidized and rendered inactive (Peer et al., 2013).

The presence of increased quantities of luteolin glycosides in halotropic plants treated with blue light suggest that luteolin may be increasing salt tolerance through the modulation of auxin transport and metabolism, along with the mediation of salt induced

191 changes in cellular redox. The quercetin glycoside, rutin, has also been shown to confer salt tolerance in halophytic species through direct ROS scavenging, and indirectly through decreasing K+ efflux through ROS-K+ activated channels (Ksouri et al., 2007;

Ismail et al., 2016). Therefore, halotropism that was observed in rough bluegrass following blue light treatment could be due to flavonoid modulation of auxin transport and ROS induced auxin catabolism, and together this would lead to greater asymmetrical auxin distribution and enhanced halotropic bending.

ROS production due to blue light exposure may also act as a signal. With the blue light photoreceptor, cryptochrome, ROS production can take place through the re-

‧ oxidation of FADH to fully oxidized FAD, producing H2O2, in the presence of oxygen

(Muller and Ahmad, 2011; Ahmad, 2016). This process generally occurs following return to darkness, which did not occur in blue light treatments here, but re-oxidation of

FADH‧ is known to be a competitive reaction even in light (Muller and Ahamd, 2011).

Increasing ROS content is damaging, as mentioned above, but ROS can also serve as signaling molecules (Miao et al., 2006; Maathuis, 2014). Blue light induced production of ROS may serve as an early signal of salt stress, as ROS production is thought to be a signal for NaCl accumulation; therefore, early halotropic bending that was observed in rough bluegrass could be due to advanced production of ROS prior to salt induced ROS production (Consentino et al., 2015; Jourdan et al., 2015. Many blue and red light signaling cascades take place indirectly through the nuclear localized Constitutive

Photomorphogenic/De-etiolated/Fusca protein complex (COP/DET/FUS), in which this protein complex degrades light induced transcription factors like HY5 (Long Hypocotyl)

192 and HYH (HY5 Homolog) (Lau and Deng., 2012). While photoregulation is mostly associated with shoots, both HY5 and HYH have also been shown to regulate auxin transport and signaling in roots (Oyama et al., 1997; Cluis et al., 2004; Sibout et al.,

2006). Together our results show that blue light regulates halotropic growth in rough bluegrass, and the data indicates that this is through increased flavonoid production.

However, due to blue light regulating through alternate pathways, the changes in root tropic growth seen here could also be due to ROS signaling as well as signaling through transcription factors like HY5 and HYH. Overall, however, it is questionable as to how important light is for modulating root growth. Greenhouse experiments here show that light application may improve rough bluegrass established under salt stress, but more controlled experiments need performed to verify this.

Light has been shown to regulate gravitropism, but it may still be a stretch to say that light, blue light in particular, is required for halotropism and salt tolerance (Feldman and Briggs, 1987; Poppe et al., 1996) in the field or greenhouse setting. However, shoot derived signals following light perception could be translocated to the root system, allowing for the photoregulation of root responses. Flavonoids in particular have been shown to be transported long distances, from shoot-to-root or vice-a-versa (Buer et al.,

2007). Therefore, flavonoids may be produced in rough bluegrass shoots and transported to the roots. However, experiments blocking root light exposure need to be performed to determine if increases in shoot derived flavonoids through blue light application can increase flavonoid content in the roots. HY5 also been shown to be transported from the shoots to the roots (Chen et al., 2016; Palme et al., 2016). Again, this suggests that light

193 induced transcription factors like HY5 and HYH could act on roots when light conditions are favorable above ground. Furthermore, light can be conducted through shoots to roots, and in seedlings, like the ones tested here, the relatively short distance between the shoot and roots may increase light propagation to below ground tissue (Sun et al., 2005).

In conclusion, our data show that light is required for halotropism in rough bluegrass due to a loss in halotropic growth under dark conditions, and the data suggest that flavonoids modulate halotropism. Results indicate that the application of blue light enhances halotropic growth through further increases in flavonoid levels, and we have shown that this response is lost under red light treatment. Together this indicates that blue light is required for halotropism, however, because blue light regulates growth and development through multiple pathways, flavonoids may only be one piece of the puzzle.

194

Reference

Agati, G., E. Azzarello, S. Pollastri, and M. Tattini. 2012. Flavonoids as antioxidants in plants: Location and functional significance. Plant Sci. 196: 67-76.

Ahamd, M. 2016. Photocycle and signaling mechanism of plant cryptochrome. Curr op. in Plant Biol. 33: 108-115.

Apse, M.P., G.S. Aharon, W.A. Snedden and E. Blumwald. 1999. Salt tolerance conferred by overexpression of a vacuolar Na+/H+ antiporter in Arabidopsis. Science. 285.

Apse, M.P., J.B. Scottosanto and E. Blumwald. 2003. Vacoular cation/H+ exchange, ion homeostasis, and leaf development are altered in a T-DNA insertional mutant of AtNHX1, the Arabidopsis vacoular Na+/H+ antiporter. The Plant Journal. 36: 229-239.

Ben-Hur, M., F.H. Li, R. Keren, and G. Shalit. 2001. Water and salt distribution in a field irrigation with marginal under high water table conditions. Soil Sci. Soc. of Am. J. 65: 191-198.

Blumwald, E. 2000. Sodium transport and salt tolerance in plants. Curr. Op. in Cell Biol. 12: 431-434.

Bose, J., A. Rodrigo-Moreno, and S. Shabala. 2014. ROS homeostasis in halophytes in the context of salinity stress tolerance. J. Exp. Bot. 65(5): 1241-1257.

Brown, D.E., A.M. Rashotte, A.S. Murphy, J. Normanly, B.W. Tague, W.A. peer, L. Taiz, and G.K. Muday. 2001. Flavonoids act as negative regulators of auxin transport in vivo in Arabidopsis. Plant Physiol. 126: 524-535.

195

Buer, C.S., G.K. Muday, and M.A. Djordjevic. 2007. Flavonoids are differentially taken up and transported long distances in arabidopsis. Plant Physiol. 145: 478-790.

Camberato, J.J. and S.B. Martin. 2004. Salinity slows germination of rough bluegrass. HortSci. 39(2): 394-397.

Cavaliere, C., P. Foglia, E. Pastorini, R. Samperi, and A. Lagana. 2005. Identification and mass spectrometric characterization of glycosylated flavonoids in Triticum durum plants by high-performance liquid chromatography with tandem mass spectrometry. Rapid Commun. Mass Spectrom. 19: 3143-3158.

Chen, X., Q. Yao, Z. Gao, C. Jiang, N.P. Harberd, and X. Fu. 2016. Shoot-to-root mobile transcription factor HY5 coordinates plant carbon and nitrogen acquisition. Curr. Biol. 26: 640-646.

Cluis, C.P., C.F. Mouchel, and C.S. Hardtke. 2004. The Arabidopsis transcription factor HY5 integrates light and hormone signaling pathways. The Plant Journal. 38: 332-347.

Consentino, L., S. Lambert, C. Martino, N. Jourdan, P.E. Bouchet, J. Witczack, P. Castello, M. El-Esawi, F. Corbineau, A. d’Harlingue, and M. Ahamd. 2015. Blue- light dependent reactive oxygen species formation by Arabidopsis cryptochrome may define a novel evolutionarily conserved signaling mechanism. New Phytol. 206: 1450-1462.

Dong, X., W. Chen, W. Wang, H. Zhang, X Liu, and J. Luo. 2014. Comprehensive profiling and natural variation of flavonoids in rice. J. Integrative Plant Biol. 56(9): 876-886.

196

Du, Y., Q. Hei, Y. Liu, H. Zhang, K. Xu, and T. Xia. 2010. Isolation and characterization of a putative vacuolar Na+/H+ antiporter gene from Zoysia japonica L. J. Plant Biol. 53(4): 251-258.

Duan, L.N., D. Dietrich, C.H. Ng, P.M.Y. Chan, R. Bhalerao, M.J. Bennett, et al. 2013. Endodermal ABA Signaling Promotes Lateral Root Quiescence during Salt Stress in Arabidopsis Seedlings. Plant Cell. 25: 324-341. doi:10.1105/tpc.112.107227.

Eapen, D., M.L. Barroso, G. Ponce, M.E. Campos, and G. I. Cassab. 2005. Hydrotropism: Root growth responses to water. Trends in Plant Sci. 10(1): 44-50.

Eryilmaz, F. 2006. The relationships between salt stress and anthocyanin accumulation in higher plants. Biotech. And Biotech. Equip. 20(1): 47-52.

Feldman, L.J. and W.R. Briggs. 1987. Light-regulated gravitropism in seedling roots of maize. Plant Physiol. 83: 241-243.

Friml J. 2003. Auxin transport - shaping the plant. Curr. Op. in Plant Biol. 6: 7-12.

Galen, C., J.J. Rabenold, and E. Liscum. 2006. Functional ecology of a blue light photoreceptor: effects of phototropin-1 on root growth enhance drought tolerance in Arabidopsis thaliana. New Phytol. 173: 91-99.

Galvan-Ampudia, C.S., M.M Julkowska, E. Darwish, J. Gandullo, R.A. Korver, G. Brunoud, M.A. Haring, T. Munnik, T. Vernoux, and C. Testerink. 2013. Halotropism is a response of plant roots to avoid a saline environment. Curr. Biol. 23: 2044-2050.

Geisler, M., J.J. Blakeslee, R. Bouchard, O.R. Lee, V. Vincenzetti, A. Bandyopadhyay. B. Titapiwatanakun, W.A. Peer, A. Bailly, E.L. Richards, K.F.K. Ejendal, A.P. Smith, C. Baroux, U. Grossniklaus, A. Müller, C.A. Hrycyna, R. Dudler, A.S. Murphy, and E. Martinoia. 2005. Cellular efflux of auxin catalyzed by the Arabidopsis MDR/PGP transporter AtPGP1. The Plant Journal. 44: 179-194.

Geisler, M. and A.S. Murphy. 2006. The ABC of auxin transport: The role of p- glycoproteins in plant development. FEBS. 530: 1094-1102.

Grill, E. and A. Himmelbach. 1998. ABA signal transduction. Current Opinion in Plant Biology 1: 412-418. doi:Doi 10.1016/S1369-5266(98)80265-3.

Hameed, M. and M. Ashraf. 2008. Physiological and biochemical adaptations of Cynodon dactylon (L.) Pers. from the Salt Range (Pakistan) to salinity stress. Flora. 203: 683-694.

197

Harivandi, M.A. 2004. Evaluating recycled waters for golf course irrigation. USGA green section record. Nov.-Dec.

Hayes, A.R., C.F. Mancino, and I.L. Pepper. 1990. Irrigation of turfgrass with secondary sewage effluent: I. Soil and leachate water quality. Agron. J. 82: 939-943.

Hu, L., T. Hu, Z. Zhang, H. Pang, and J. Fu. 2012a. Exogenous glycine betaine ameliorates the adverse effect of salt stress on perennial ryegrass. J. Amer. Soc. Hort. Sci. 137(1): 38-46.

Hummel, N.W. 1993. Rationale for the revisions of the USGA green construction specifications. USGA green section record. March-April.

Jacobs, M. and P.H. Rubery. 1988. Naturally occurring auxin transport regulators. Science. 241: 346-349.

James, R.A., A.R. Rivelli, R. Munns and S.v. Caemmerer. 2002. Factors affecting CO2 assimilation, leaf injury and growth in salt-stressed durum wheat. Functional Plant Biol. 29: 1393. doi:10.1071/fp02069.

198

Jourdan, N., C.F. Martino. M. El-Esawi, P.E. Bouchet, A. d’Harlingue, and M. Ahmad. 2015. Blue-light dependent ROS formation by Arabidopsis cryptochrome-2 may contribute toward its signaling role. Plant Signaling and Behavior. 10(8). e1042647.

Julkowska, M.M. and C. Testernik. 2015. Turning plant signaling and growth to survive salt stress. Trends in Plant Sci. 20: 586-594.

Ke, S. and C.W. Lee. 1996. Plant regeneration in Kentucky bluegrass (Poa pratensis L.) via coleoptile tissue culture. Plant Cell Reports. 15: 882-887.

Krishnan, S. and E.B. Merewitz. 2015. Phytohormone responses and cell viability during salinity stress in two creeping bentgrass cultivars differing in salt tolerance. J. Amer. Soc. HortSci. 140(4): 346-355.

Ksouri, R., W. Megdiche, A. Debez, H. Falleh, C. Grignon, and C. Abdelly. 2007. Salinity effects on polyphenol content and antioxidant activities in leaves of the halophyte Cakile maritima. Plant Physiol. and Biochem. 45: 244-249.

Lau, O.S. and X.W. Deng. 2012. The photomorphogenic repressors COP1 and DET1: 20 years later. Trends in Plant Sci. 17(10): 584-593.

Leopoldini, M., I.P. Pitarch, N. Russo, and M. Toscano. 2004. Structure, Conformation, and Electronic Properties of Apigenin, Luteolin, and Taxifolin Antioxidants. A First Principle Theoretical Study. J. Phys. Chem. 108: 92-96.

Liu, J. and J.K. Zhu. 1998. A calcium sensor homolog required for plant salt tolerance. Science 280: 1943-1945. doi:10.1126/science.280.5371.1943.

Maathuis, F.J. and A. Antmann. 1999. K+ nutrition and Na+ toxicity: the basis of cellular K+/Na+ ratios. Annals of Botany. 84: 123-133.

Maathuis, F.J. 2014. Sodium in plants: perception, signalling, and regulation of sodium fluxes. J. Exp. Bot. 65: 849-858. doi:10.1093/jxb/ert326.

Mancino, C.F., and I.L. Pepper. 1992. Irrigation of turfgrass with secondary sewage effluent: Soil quality. Agron. J. 84: 650-654.

Marcum, K.B. and C.L. Murdoch. 1994. Salinity tolerance mechanisms of six C4 turfgrasses. J. Amer. Soc. Hort. Sci. 119(4): 779-784.

199

Marcum, K.B., S.J. Anderson, and M.C. Engelke. 1998. Salt gland ion excretion: A salinity tolerance mechanism among five zoysiagrass species. Crop Sci. 38: 806- 810.

Marcum, K.B. and M. Pessarakli. 2006. Salinity tolerance and salt gland excretion efficiency of Bermudagrass turf cultivars. Crop Sci. 46: 2571-2574.

Maupin, M.A., J.F. Kenny, S.S. Hutson, J.K. Lovelace, N.L. Barber and K.S. Linsey. 2014. Estimated use of water in the United States in 2010. U.S. Geological Survey Circular 1405: 1-64.

Milesi, C., S.W. Running, C.D. Elvidge, J.B. Dietz, B.J. Tuttle, and R.R. Nemani. 2005. Environmental Management. 32 (3): 426-438.

Miller, G., N. Suzuki, S. Ciftci-Yilmaz and R. Mittler. 2010. Reactive oxygen species homeostasis and signalling during drought and salinity stresses. Plant Cell Environ 33: 453-467. doi:10.1111/j.1365-3040.2009.02041.x.

Minitab 17 Statistical Software (2010). [Computer software]. State College, PA: Minitab, Inc. (www.minitab.com)

Munns, R. and M. Tester. 2008. Mechanisms of salinity tolerance. Annu. Rev. Plant Biol. 59: 651-81.

Muller, P. and M. Ahmad. 2011. Light activated cryptochrome reacts with molecular oxygen to form a flavin-superoxide radical pair consistent with magnetoperception. J. Biol. Chem. 286(34): 21033-21040.

Munns, R. and A. Termaat. 1986. Whole-plant responses to salinity. Australian J. Plant Physiol. 13: 143-160.

Munns, R. 1993. Physiological processes limiting plant growth in saline soils: some dogmas and hypotheses. Plant Cell and Environment 16: 15-24.

Munns, R. 2002. Comparative physiology of salt and water stress. Plant Cell and Environment 25: 239-250. doi:DOI 10.1046/j.0016-8025.2001.00808.x.

Murphy, A.S., W.A. Peer, and Taiz, L. 2000. Regulation of auxin transport by aminopeptidases and endogenous flavonoids. Planta. 211: 315-324.

200

Obear, G.R., A.E. Hartemink, and D.J. Soldat. Soils with iron-cemented layers on golf courses in the USA. Geoderma. 232-234: 198-207.

Oyama, T., Y. Shimura, and K. Okada. The Arabidopsis HY5 gene encodes a bZIP protein that regulates stimulus-induced development of root and hypocotyl. Genes and Dev. 11: 2983-2995.

Palme, K., W. Teale, and A. Dovzhenko. 2016. Plant signaling: HY5 synchronizes resource supply. Curr. Biol. 26: 319-337.

Paudel, L., F.J. Wyzgoski, J.C. Scheerens, A.M. Chanon, R.N. Reese, D. Smiljanic, C. Wesdemiotis, J.J. Blakeslee, K. M. Riedl, and P.L. Rinaldi. 2013. Nonanthocyanin secondary metabolite of black raspberry (Rubus occidentalis L.) fruits: Identification by HPLC-DAD, NMR, HPLC-ESI-MS, and ESI-MS/MS analyses. J. Agr. Food Chem. 61: 12032-12043.

Peer, W.A. 2013. From perception to attenuation: Auxin signaling and responses. Curr. Op. in Plant Biol. 16: 561-568.

Peer, W.A., A. Bandyopadhyay, J.J. Blakeslee, S. N. Makam, R.J. Chen, P.H. Masson, and A.S. Murphy. 2004. Variation in expression and protein localization of the PIN family of auxin efflux facilitator proteins in flavonoid mutants with altered auxin transport in Arabidopsis thaliana. Plant Cell. 16: 1898-1911.

Peer, W.A. and A.S. Murphy. 2007. Flavonoids and auxin transport: Modulators or regulators? Trends in Plant Sci. 12(12): 556-563.

Peer, W.A., D.E. Brown, B.W. Tague, G.K. Muday, L. Taiz, and A.S. Murphy. 2001. Flavonoid accumulation patterns of transparent testa mutants of Arabidopsis. Plant Physiol. 126: 536-548.

Peer, W.A., J.J. Blakeslee, H. Yang, and A.S. Murphy. 2011. Seven things we think we know about auxin transport. Mol. Plant. 4: 487-504.

Petrella, D.P., Metzger, J.D., J.J. Blakeslee, E.J. Nangle, and D.S. Gardner. 2016a. Anthocyanin production using rough bluegrass treated with high intensity light. HortSci. 51(9) 1111-1120. 2016. doi:10.21273/HORTSCI10878-16.

Petrella, D.P., Metzger, J.D., J.J. Blakeslee, E.J. Nangle, and D.S. Gardner. 2016b. Effects of blue light and phenotype on anthocyanin accumulation in accessions and cultivars of rough bluegrass. Crop Sci. doi:10.2135/cropsci2016.05.0438.

201

Pin Ng, J.L., S. Hassan, T.T. Truong, C.H. Hocart, C. Laffont, F. Frugier, and U. Mathesius. 2015. Flavonoids and auxin transport inhibitors rescue symbiotic nodulation in the Medicago trunculata cytokinin receptor mutant cre1. Plant Cell. 27: 2210-2226.

Poppe, C., R.P. Hangarter, R.A. Sharrock, F. Nagy, and E. Schafer. 1996. The light- induced reduction of the gravitropic growth-orientation of seedlings of Arabidopsis thaliana (L.) Heynh. is a photomorphogenic response mediated synergistically by the far-red-absorbing forms of phytochromes A and B. Planta. 199: 511-541.

Prettyman, G. and E. McCoy. 2000. Charging and flushing high-sand greens. Golf Course Management. May: 49-53.

Qiu, Q.S., B.J. Barkla, R. Vera-Estrella, J.K. Zhu and K.S. Schumaker. 2003. Na+/H+ exchange activity in the plasma membrane of Arabidopsis. Plant Physiol 132: 1041-1052. doi:10.1104/pp.102.010421.

Rosquete, M.R. and J. Kleine-Vehn. 2013. Halotropism: turning down the salty date. Curr. Biol. 23: 927-929.

Ruiz-Sola, M.A., V. Arbona, A. Gomez-Cadenas, M. Rodriguez-Concepcion and A. Rodriguez-Villalon. 2014. A Root Specific Induction of Carotenoid Biosynthesis Contributes to ABA Production upon Salt Stress in Arabidopsis. Plos One 9. doi:ARTN e90765

Saito, K., K. Yonekura-Sakaibara, R. Nakabayashi, Y. Higashi, M. Yamazaki, T. Tohge, and A.R. Fernie. 2013. The flavonoid biosynthetic pathway in Arabidopsis: Structural and genetic diversity. Plant Physiol. and Biochem. 72: 21-34.

Schlossberg, M.L. and K.J. Karnok. 2002. Use of transparent columns for demonstrating water movement in golf green root zones. J. Nat. Resour. Life Sci. Educ. 31: 1-4.

Shabala, S. and T.A. Cuin. 2007. Potassium transport and plant salt tolerance. Physiologia Plantarum. 133: 651-669.

Shi, H. and J.K. Zhu. 2002. Regulation of expression of the vacuolar Na+/H+ antiporter gene AtNHX1 by salt stress and abscisic acid. Plant Mol. Biol. 50: 543-550.

202

Sibout, R., P. Sukumar, C. Hettiarachchi, M. Holm, G.K. Muday, and C.S. Hardtke. 2006. Opposite root growth phenotypes of hy5 versus hy5 hyh mutant correlate with increased constitutive auxin signaling. PLOS Genetics. 2(11): e202.

Strack, D., J. Heilemann, M. Momken, and V. Wray. 1988. Cell wall-conjugated phenolics from coniferae leaves. Phytochem. 27(11): 3517-3521.

Rasband, W.S., ImageJ, U. S. National Institutes of Health, Bethesda, Maryland, USA, http://imagej.nih.gov/ij/, 1997-2016.

Tester, M. and R.J. Davenport. 2003. Na+ Tolerance and Na+ Transport in Higher Plants. Annals of Botany 91: 503-527. doi:10.1093/aob/mcg058

Tognetti, V.B., P. Mühlenbock, and F. Van Breusegem. 2012. Stress homeostasis – the redox and auxin perspective. Plant Cell and Envrion. 35: 321-333.

Tulio, A.Z., R.N. Reese, F.J. Wyzgozki, P.L. Rinaldi, R. Fu., J.C. Scheerens, and A.R. Miller. Cyanidin 3-rutinoside and cyanidin 3-xylosylturinoside as primary phenolic antioxidant of black raspberry. J. Agr. Food Chem. 56: 1880-1888.

Tyrvainen, L, A. Ojala, K. Korpela, T. Lanki, Y. Tsunetsugu, and T. Kagawa. 2014. The influence of urban green environments on stress relief measures: A field experiment. J. Environ. Psychology. 38: 1-9.

Van Oosten, M., A. Sharkhu, G. Batelli, and R.A. Bressan. 2013. The Arabidopsis thaliana mutant air1 implicates SOS2 in the regulation of anthocyanins under salt stress. Plant Mol. Biol. 83: 405-415. van Acker, S.A.B.E., M. J. de Groot, D. van den Berg, M.N.J.L, Tromp, G.D. den Kelder, W.J.F. van der Vijgh, and A Bast. 1996. A Quantum Chemical Explanation of the Antioxidant Activity of Flavonoids. Chem. Res. Toxical. 9: 1305-1312.

Venton, B.J. 2017. Essentials of analytical chemistry: Internal standards. JoVE. doi: 10.3791/10225.

Winkel-Shirley, B. 2001. Flavonoid biosynthesis. A colorful model for genetics, biochemistry, cell biology, and biotechnology. Plant Phsyiol. 126: 485-493.

Wojakowska, A., J. Perkowksi, T. Goral, and M. Stobiecki. 2012. Structural characterization of flavonoid glycosides from leaves of wheat (Triticum aestivum L.) using LC/MS/MS profiling of the target compounds. J. Mass Spectrom. 45: 329-339.

203

Wolfe, J.L. and R.H. Liu. 2008. Structure-activity relationships of flavonoids in the cellular antioxidant activity assay. K. Agr. Food. Chem. 56: 8404-5411.

Yan, J., B. Wang, J. Jiang, L. Cheng, and T. Wu. 2014. GmFnSll-Contolled soybean flavone metabolism reponds to abiotic stresses and regulates plant salt tolerance. Plant Cell Physiol. 55(1): 74-86.

Yang, Z., J. Yu, E. Merewitz, and B. Huang. 2012. Differential effects of absicic acid and glycine betaine on physiological responses to drought and salinity stress for two perennial grass species. J. Amer. Soc. Hort. Sci. 137(2): 96-106.

Yin, R., K. Han, W. Heller, A. Albert, P.I. Dobrev, E. Zazimalova, and A.R. Schaffner. kaempferol 3-O-rhamnoside-7-O-rhamnoside is an endogenous flavonol inhibitor of polar auxin transport in Arabidopsis shoots. New Phytol. 201: 446-475.

Yokawa, K., R. Fasano, T. Kagenishi, and F. Baluska. 2014. Light as a stress factor in plant roots – case of root halotropism. Frontiers in Plant Sci. 5(718). doi: 10.3389/fpls.2014.00718

Xiong, L., K.S. Schumaker and J.K. Zhu. 2002. Cell signaling during cold, drought, and salt stress. The Plant Cell: 165-183. doi:10.1105/tpc.000596.

Zhang, K., H. Xu, T. Yuan, L. Zhang, and Y. Lu. 2013. Blue-light-induced PIN3 polarization for root negative phototropic response in Arabidopsis. The Plant Journal. 76: 308-321.

Zhu, J.-K. 2003. Regulation of ion homeostasis under salt stress. Curr. Op. in Plant Biol. 6: 441-445. doi:10.1016/s1369-5266(03)00085-2.

Zhu, J.K. 2002. Salt and drought stress signal transduction in plants. Annu Rev Plant Biol 53: 247-273. doi:10.1146/annurev.arplant.53.091401.143329.

Zhu, J., Z. Fu, Y.D. Koo, J.K. Zhu, D.E. Jenney, M.W.W.Adams, Y. Zhu, H. Shi, D. Yun, P.M. Hasegawa, and R.A. Bressan. 2007. An enhancer mutant of Arabidopsis salt overally sensitive 3 mediates both ion homeostasis and the oxidative stress response. Mol. and Cell. Biol. 27(14): 5214-5224.

204

Chapter 5: Overall Conclusions

Abstract

This research has shown that rough bluegrass (Poa trivialis L.) can increase levels of flavonoids and anthocyanins to quantities that are similar to plants used as industrial sources of phenolic metabolites. Results indicate that the application of blue light alone or in combination with red light can increase anthocyanin content compared to white light alone. Rough bluegrass also exhibits natural variation for light-induced anthocyanin accumulation, and this variation can be linked to leaf morphology. Being that rough bluegrass exhibits a high level of stress-induced flavonoid accumulation, this turfgrass may also provide further insight into flavonoid function. Data show that flavonoid upregulation, through the application of blue light, leads to enhanced and sustained halotropism for 24 hours. Under red light, however, halotropism is lost. Overall conclusions and thoughts on the role of flavonoids in turfgrass biology are presented within.

205

Discussion

The application of high intensity white light to rough bluegrass for a prolonged period of time drastically increased anthocyanin content compared to untreated plants

(Appendix M). Calculations show that over an entire growing season, rough bluegrass could out-perform current plant based anthocyanin sources when treated with light based protocols presented here. To increase the efficiency of light treatment, plants were treated with narrow band LED light treatments. Results show that blue light and combinations of blue and red light increase anthocyanin content compared to white light alone, and that photosynthetic stress may further increase anthocyanin content of young plants in particular. However, even though blue light increased anthocyanin content compared to control, blue LED treatment never increased anthocyanin content to levels seen when treated with high-intensity discharge (HID) metal halide lamps.

LED treatments allowed for the separation of temperature stress from light- induced regulation. However, the increased temperature from HID light treatment may also be contributing to anthocyanin regulation. Experiments evaluating the impact of temperature on anthocyanin synthesis should be performed to further optimize the protocols presented here. HID light treatment also increases the amount of ultra-violet

(UV)-A light that plants are exposed to. While we focused on the effects of visible light application, it is known that UV-A light is sensed and responded to in a similar fashion as blue light. Preliminary experiments tested the efficiency of UV-B light application, and while the lamps used contained UV-A wavelengths, the distribution and intensity of UV-

A or the combination of UV-A/B did not increase anthocyanin content at all. However,

206 newer technology has led to the development of UV-A specific LED lights. Future experiments should evaluate the impact of UV-A light alone and in combination with blue light to help further optimize anthocyanin production in turfgrasses.

Preliminary experiments evaluated the effect of high intensity white light application on anthocyanin production in multiple C3 and C4 turfgrasses. Only rough bluegrass exhibited the unique ability to upregulate anthocyanin synthesis; however, initial experiments only evaluated a single cultivar, ‘Havana’. To truly optimize a biological production system a breeding program would be necessary to increase system efficiency. When evaluating accessions and cultivars of rough bluegrass, it was evident that only specific accessions increased anthocyanin content under blue light treatment, in particular accessions from Germany and production cultivars of rough bluegrass.

Compared to other accessions tested, these samples exhibited finer leaf tissue with significantly less cuticular wax. Experiments analyzing the reflective capacity of the cuticle from these accessions and cultivars should be analyzed, and this data may also indicate the potential effectiveness of UV-A light application for anthocyanin induction.

Further analyses should be done to evaluate differences in cuticular waxes of these rough bluegrass accessions and cultivars. As the composition and structure of the plant cuticle is very diverse, analysis of the cuticle here may provide further insight into the function of the cuticle in turfgrass light perception. On a side note, the cuticle is also very important for tolerance to biotic and abiotic stress, and research on the cuticle of rough bluegrass may increase our understanding of the cuticle’s importance in turfgrass stress tolerance.

207

Rough bluegrass is not utilized to a great extent as a turfgrass due to low tolerance to environmental stress. However, this species is used to overseed dormant

Bermudagrass in certain parts of the United States where effluent, saline, water is used for irrigation, and soils tend to have greater levels of NaCl. Rough bluegrass is considered to be intolerant to salt stress, but this species is still used in these situations.

Data has shown that rough bluegrass hyper-accumulates flavonoids in a short period of time; therefore, we investigated what impact flavonoid accumulation had on root tolerance to NaCl stress. Results show that greater quantities of flavonoids in the root increase the movement of the primary root of rough bluegrass seedlings away from areas containing high NaCl concentrations. Flavonoid synthesis was increased using light treatment. Results showed that halotropism was lost under darkness and red light supplementation, and flavonoid concentrations in these treatments was below the limit of quantification.

Blue light application, however, increase flavonoid concentration and resulted in enhanced halotropic growth, but blue light dose response tests should also be done to determine the quantity of blue light required to modulate root tropic growth. Blue light may impact halotropism on its own through pathways other than flavonoid synthesis.

Therefore, experiments supplementing flavonoids into the agar based media should be performed under darkness and/or red light to help determine if halotropism observed here was due to flavonoid mediated changes alone. Supplementation experiments have their own drawbacks including the efficiency of uptake and transport of the compound of interest, but these experiments could still help provide valuable data.

208

Interestingly, when testing various concentrations of salt for halotropism experiments rough bluegrass roots exhibited increased movement into salt containing media at low NaCl concentrations. These results indicate the presence of other root sensing mechanisms, and here NaCl may be serving as an indirect signal at low concentrations. Experiments evaluating the nutrient sensing ability of rough bluegrass seedlings with and without low concentrations of NaCl should be performed to determine if roots of monocots move towards gradients of nutrients. Data would also help to determine if this potential sensing mechanism is dependent on sensing the presence of secondary ions like Na+. Monocots and dicots produce very different types of root systems, and even in the early stages of development dicot plants will produce lateral roots. While these lateral roots scavenge for water and nutrients in various directions, many monocots do not produce lateral roots or produce a very limited number of lateral roots; therefore the primary and/or adventitious roots of monocots may scavenge to a greater degree due to the lack in lateral rooting.

Whether or not halotropism is an important mechanism for increasing salt tolerance is still not known, and little data has been presented on the role of halotropism in maintaining fitness under salt stress. Only seedlings and/or young plants have been evaluated for halotropic growth, and while the establishment of seedlings in saline soils is important, other known mechanisms have been directly linked to salt tolerance of young plants. Halotropism experiments should be performed using plants in various developmental stages to help determine what impact halotropism has on plant growth and development.

209

Rough bluegrass exhibits significant changes in morphology with increasing plant age that may be linked to salt stress tolerance and halotropism. Seedlings and young plants only spread through tiller production, and rooting is adequate. However within one or two months, rough bluegrass begins to produce stolons, tillering is reduced and/or absent, and rooting is greatly diminished. The mat type prostrate growth exhibited by mature rough bluegrass results in decreased stress tolerance, and may be linked to a loss in rooting. This growth habit, however, actually promotes stress tolerance in one sense through avoidance, and stolons act as vegetative propagules and carbohydrate reserves for when environmental conditions are more hospitable. The switch from intravaginal

(tiller) to extravaginal (stolon) axillary buds may be regulated developmentally or by stress, and rough bluegrass may serve as a model plant to investigate this phenomenon due to its poor stress tolerance. Results could help breed turfgrasses with improved stress tolerance through alterations in normal developmental patterns that alter plant growth.

Rough bluegrass is a turfgrass species, and plant in general, that has received little attention. Here we have shown that rough bluegrass possess the ability serve as a bio- production system for secondary metabolites, and we have shown that light treatment and flavonoid concentration lead to alterations in root growth during NaCl stress. This species could be a model monocot in future research, and its low tolerance to stress may provide greater insight into stress tolerance of grasses.

210

Complete Reference

Agati, G., E. Azzarello, S. Pollastri, and M. Tattini. 2012. Flavonoids as antioxidants in plants: Location and functional significance. Plant Sci. 196: 67-76.

Ahmad, M., J.A. Jarillo, O. Smirmova, and A. R. Cashmore. 1998. Cryptochrome blue- light photoreceptors of Arabidopsis implicated in phototropism. Nature. 392: 720- 723.

Ahmad, M. 2016. Photocycle and signaling mechanism of plant cryptochrome. Curr. op. in Plant Biol. 33: 108-115.

American Meteorological Society. 2012. Meteorology glossary. http://glossary.ametsoc.org/wiki/Freshwater.

Apse, M.P., G.S. Aharon., W.A. Snedden, and E. Blumwald. 1999. Salt tolerance conferred by overexpression of a vacuolar Na+/H+ antiporter in Arabidopsis. Science. 285.

Baker, D.A. 2000. Vascular transport of auxins and cytokinins in Ricinus. Plant Growth Regulation. 32: 157-160.

Bandyopadhyay, A., J.J. Blakeslee, O.R. Lee, J. Mravec, M. Sauer, B. Titapiwatanakun, S.N. Makam, R. Bouchard, M. Geisler, E. Martinoia, J. Friml, W.A. Peer, and

211

A.S. Murphy. 2007. Interactions of PIN and PGP auxin transport mechanisms. Biochem. Soc. Trans. 35: 137-141.

Bao A-K, Wang S-M, Wu G-Q, Xi J-J, Zhang J-L, Wang C-M. 2009. Overexpression of the Arabidopsis H+-PPase enhanced resistance to salt and drought stress in transgenic alfalfa (Medicago sativa L.). Plant Sci. 176, 232-240.

Bartel, B., and G.R. Fink.. 1995. Ilr1, an Amidohydrolase That Releases Active Indole-3- Acetic-Acid from Conjugates. Science. 268: 1745-1748.

Beard, J.B. 1973. Chapter 2: Growth and development pgs. 25-53. In. Turfgrass: Science and culture. J.B. Beard. Prentice-Hall. Edgewood Cliffs, NJ. USA.

Bechtold, T., A. Mahmud-Ali, and R. Mussak. 2007. Anthocyanin dyes extracted from grape pomace for the purpose of textile dyeing. J. Sci. Food Agr. 87: 2589-2595.

Beffa, R., H.V. Martin, and P.E. Pilet. 1990. In vitro oxidation of indoleacetic acid by soluble auxin-oxidases and peroxidases from maize roots. Plant Physiol. 94: 485- 491.

Ben-Hur, M., F.H. Li, R. Keren, and G. Shalit. 2001. Water and salt distribution in a field irrigation with marginal under high water table conditions. Soil Sci. Soc. of Am. J. 65: 191-198.

Bennett, S.R.M, J. Alvarez, G. Bossinger, and D.R. Smyth. 1995. Morphogenesis in pinoid mutants of Arabidopsis thaliana. The Plant Journal. 8: 505-520.

212

Blakeslee, J,J., W.A. Peer, and A.S. Murphy. 2005. Auxin transport. Curr. Op. Plant Biol. 8: 494-500.

Blumwald E. 2000. Sodium transport and salt tolerance in plants. Curr. Op. in Cell Biol. 12: 431-434.

Boerjan, W., J. Ralph, and M. Baucher. 2003. Lignin biosynthesis. Ann. Rev. Plant Biol. 54: 519-546.

Boldt, J.K. 2013. Foliar anthocyanins in coleus and ornamental grasses: Accumulation, localization, and function. Univ. of Minnesota, Minneapolis, MN. Ph.D. Diss.

Boldt, J.K., M.H. Meyer, and J.E. Erwin. 2014. Foliar anthocyanins: A horticultural review, p. 209-251. In: J. Janick (ed.). Hort. Rev. Wiley, New York, NY.

Bouly, J. E. Scheicher, M. Dionisio-Sese, F. Vandenbussche, D. Van Der Straeten, N. Bakrim, S. Meier, A. Batschaeur, P. Galland, R. Bittl, and M. Ahmad. 2007. Cryptochrome blue light photoreceptors are activated through interconversion of flavin redox states. J. of Biol. Chem. 282(13): 9389-9391.

213

Burgie, E.S., A.N. Bussell, J.M. Walker, J. Dubiel, and R. D. Vierstra. 2014. PNAS. 111(28): 10179-10184.

Butler, W.L., S. Hendricks, and H.W. Siegelman. 1964. Action spectra of phytochrome in vitro. Phytochem. and Photobiol. 3: 521-528.

Camberato, J.J. and S.B. Martin. 2004. Salinity slows germination of rough bluegrass. HortSci. 39(2): 394-397.

Casal, J.J., R.A. Sanchez, and M.J. Yanovsky. 1997. The function of phytochrome A. Plant Cell and Environ. 20: 813-819.

Casal, J.J. 2013. Photoreceptor signaling networks in plant responses to shade. Ann. Rev. Plant Biol. 64: 403-427.

214

Castandea-Ovando, A., L. Pacheco-Hernandez, E. Paez-Hernandez, J.A. Rodriguez, and C.A. Galan-Vidal. 2009. Chemical studies of anthocyanin: A review. Food Chem. 113: 859-871.

Chai, C., and P.K. Subudhi. 2016. Comprehensive Analysis and Expression Profiling of the OsLAX and OsABCB Auxin Transporter Gene Families in Rice (Oryza sativa) under Phytohormone Stimuli and Abiotic Stresses. Front Plant Sci. 7, 593.

Chalker-Scott, L. 1999. Environmental significance of anthocyanins in plant stress responses. Photochem. and Photobiol. 70(1): 1-9.

Chefdor F, Benedetti H, Depierreux C, Delmotte F, Morabito D, Carpin S. 2006. Osmotic stress sensing in Populus: components identification of a phosphorelay system. FEBS Lett. 580, 77-81.

Chen, M., J. Chory, and C. Frankhauser. 2004. Light signal transduction in higher plants. Ann. Rev. Genet. 38: 84-117.

Chen, X., Q. Yao, Z. Gao, C. Jiang, N.P. Harberd, and X. Fu. 2016. Shoot-to-root mobile transcription factor HY5 coordinates plant carbon and nitrogen acquisition. Curr. Biol. 26: 640-646.

Cho, M., S.H. Lee, and H.T. Cho. 2007. P-glycoprotein4 displays auxin efflux transporter-like action in Arabidopsis root hair cells and tobacco cells. Plant Cell. 19: 3930-3943.

Christians, N. E. 2011. Chapter 2: Introduction to the grasses, p. 9-31. In. N. E. Christians (ed.). Fundamentals of Turfgrass Management – 4th edition. Wiley, New York, NY.

215