University of Nevada, Reno

Use of synthetic sugar analogs to probe plant cell wall function in Arabidopsis thaliana

A dissertation submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy in Biochemistry

by Jose A. Villalobos Jr

Ian S. Wallace / Dissertation Advisor

December 2020 THE GRADUATE• SCHOOL

We recommend that the dissertation prepared under our supervision by

entitled

be accepted in partial fulfillment of the requirements for the degree of

Advisor

Committee Member

Committee Member

Committee Member

Graduate School Representative

David W. Zeh, Ph.D., Dean Graduate School

i

Abstract:

Plants are terrestrial photosynthetic multicellular organisms responsible for producing most of the world’s oxygen. Plants are also responsible for fixating approximately 1011 metric tones of carbon dioxide into reduced forms of carbon that are used for plant metabolism as well as human and animal nutrition. Approximately 70% of plant biomass is derived from plant cell walls. Plant cell walls are polysaccharide-rich extracellular matrices that encapsulate nearly all plant cells and collectively, these polysaccharides are the most abundant biopolymers on the planet. The evolution of cell wall polysaccharides was integral for plants to populate terrestrial environments.

Functionally, cell wall polysaccharides are essential for normal plant growth and development, and compromises to the structure or biosynthesis causes severe developmental defects and are often lethal. The essential nature of cell wall polysaccharides creates challenges in conducting genetic studies to elucidate the biosynthesis and function cell wall polysaccharides.

The goal of this work was to implement a chemical biology strategy to probe cell wall function in a spatial, temporal, and dose-dependent manner. Our strategy was to use semi-rationally designed monosaccharide analogs that may inhibit glycosyltransferases by competitively binding to their active sites. We screened a small library of monosaccharide analogs and found three analogs: 2-deoxy-2-fluoro-L-fucose (2F-Fuc),

2-deoxy-2-fluoro-D-mannose (2F-Man), and N-dodecyldeoxynojirimycin (ND-DNJ) inhibit growth in Arabidopsis thaliana. Only 2F-Fuc repressed growth by inhibiting fucosylation of a pectic cell wall polysaccharide while 2F-Man and ND-DNJ inhibited ii growth in unexpected ways. In 2F-Man, the primary mechanism of inhibition was through the glucose repression pathway in Arabidopsis. The mechanism of 2F-Man toxicity was further supported in yeast as it also inhibited growth through a similar glucose repression pathway. Uniquely, ND-DNJ inhibited glycosylation events in sphingolipids in Arabidopsis and as a result inhibited crystalline cellulose deposition, highlighting a novel connection between cellulose and glycosylated sphingolipids.

Finally, we discovered suberin deposition occurs in response to cellulose biosynthesis inhibition.

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Acknowledgments:

This dissertation would not be possible without the support and advice from everyone around me over the years. First, I would like to thank my advisor Dr. Ian

Wallace for the opportunity to work on various projects and continuously challenging me to think critically and creatively. For believing in me when I didn’t and giving me that extra push when I needed it, none of this would be possible without you.

I would also like to thank my committee members: Dr. Dylan K. Kosma, Jeff F.

Harper, John C. Cushman, and Laina M. Geary. Your advice and questions continuously helped improve the direction of my research and getting me to always think outside the box. Thank you all for taking the time to answer my random questions, providing me insight into techniques I was not familiar with, and helping me grow as a graduate student. I would like to thank Jeff for backcrossing our resistant lines. I would also like to thank Dylan for showing me the ins and outs of operating a GC. I would also like to thank Zackary Wahrenburg for our suberin analysis and your expertise with the GC-MS.

Finally, I would like to thank the Cahoon lab for the sphingolipidomic analysis.

A special thank you to the members of the Wallace lab who have come and gone over the years: Devin Smith, Bret Hart, Edward Cruz, Andrew Larson, Celeste

Rodriquez, Eli Holschbach, Megan Warner, Tori Speicher, Hans Joseph Struffert, Bo Yi,

Timothy Fox, Brett Allen, Sienna Ogawa, Gabriel Aguilar, Jon Lau, KassaDee Herring,

Daniel Jones, Daniel Kinder, and Sarah Pennington. From bouncing ideas back and forth, talking science, teaching me new techniques, to the camaraderie we shared made my

Ph.D. experience that much more fulfilling. I would like to thank Bret for the imaging iv experiments with ND-DNJ and Devin for analyzing the kymographs that led to our mind- blowing result. A special thanks to Andrew Larson for your help with kinase assays. I’m grateful for the team I had the opportunity to work with. v

Table of Contents

List of Tables x

List of Figures xi

Chapter 1: Introduction to plant cell walls and function. 1

I. Plant Cell walls 1

II. Cell walls: Pectin structure and function 2

a. Homogalacturonan 5

b. Rhamnogalacturonan-I 9

c. Rhamnogalacturonan-II 11

III. Cell walls: Hemicellulose structure and function 14

a. Mannan and Glucomannan 14

b. Xyloglucan 17

c. Xylan 19

IV. Cell walls: Cellulose biosynthesis and regulation 20

a. Cellulose production 22

b. CSC accessory proteins 25

c. Regulation of CSCs 28

V. Non-polysaccharide wall components: lignin and suberin 32

a. Lignin 32

b. Suberin 35 vi

VI. Cell walls: Integrity sensing and Cell wall damage 37

a. Cell wall integrity sensing 37

VII. Plant Glycoproteins and Glycolipids: Structure, function and metabolism 40

a. Protein Glycosylation 40

b. Glycosylphosphatidylinositol anchors 45

c. Glucosylceramides 48

d. Glycoinositol phosphoceramides 51

VIII. Use of small molecules, monosaccharide analogs, and monosaccharide 53 mimicking small molecules a. Common cell wall biosynthesis inhibitors 54

b. Monosaccharide analogs as diagnostic tools and inhibitors 57

c. Iminosugars a class of monosaccharide mimicking compounds 60

IX. Screening of monosaccharide analogs as semi-rational inhibitors of plant 61 cell wall biosynthesis References 68

Chapter 2: 2-Deoxy-2-Fluoro-L-Fucose is a metabolically incorporated 114 inhibitor of plant cell wall polysaccharide fucosylation. I. Introduction 114

II. Materials and Methods 117

III. Results 122

IV. Discussion 145

References 149

Chapter 3: 2-Deoxy-2-Fluoro-D-Mannose is an inhibitor of energy 154 metabolism in Arabidopsis thaliana. I. Introduction 154 vii

II. Materials and Methods 158

III. Results 167

IV. Discussion 191

References 197

Chapter 4: 2-Deoxy-2-Fluoro-D-Mannose is a potent inhibitor of fungal 212 growth. I. Introduction 212

II. Materials and Methods 214

III. Results 223

IV. Discussion 244

References 248

Chapter 5: Sphingolipids are required for cellulose deposition and cellulose 258 synthase complex motility. I. Introduction 258

II. Materials and Methods 261

III. Results 265

IV. Discussion 283

References 292

Chapter 6: Cellulose biosynthesis inhibitors induce suberin production in 306 Arabidopsis thaliana as part of the Cell Wall Damage response. I. Introduction 306

II. Materials and Methods 310

III. Results 312

IV. Discussion 318 viii

References 323

Chapter 7: Discussion 330

I. Potential of fluoro fucose resistant lines 330

a. Current state of the ffr lines and their potential 330

b. Other utilities of 2F-Fuc 333

II. Sugar sensing in plants and fungi in relation to glucose and mannose 337

a. Sugar sensing in plants 338

b. Sugar sensing in yeast 342

III. Implications of sphingolipid biosynthesis on cellulose 346

a. Known glycosylated sphingolipid biosynthesis mutants: do they contain 346 less cellulose b. Non-motile observation in ND-DNJ and DCB 347

c. Utility of other iminosugars 350

IV. Cell wall integrity sensing and differential responses to cell wall damage 352

a. THESIUS1 and STRUBBELIG in response to cellulose defects 352

b. Lignin and suberin, in response to cell wall damage 353

V. Possible strategies to isolate novel GTs 355

a. Capture of GTs with photo-reactive crosslinkers 355

b. Isolation of partial polysaccharide glycans as substrates 359

VI. Implications of sugar analogs as herbicides 361

VII. Other monosaccharide analogs to explore in plant growth 363

VIII Conclusions 365 ix

References 367

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List of Tables

Chapter 1

Table 1.1: Monosaccharide analog screen on Arabidopsis. 66

Chapter 2

Table 2.1: Primer sequences used in this chapter. 123

Table 2.2: T-DNA lines used in this chapter. 131

Table 2.3: Analysis of potential fluoro fucose resistant lines. 141

Table 2.4: Segregation Ratios of ffr x Col-0 F2 progeny. 144

Chapter 3

Table 3.1: Retesting fmr lines on 50 µM 2F-Man 185

Chapter 4

Table 4.1: Primer sequences used in this study 219

Table 4.2: IC50 Values of hypersensitive deletion strains. 236

Chapter 5

Table 5.1: Nojirimycin analogs screened against Arabidopsis 268 Col-0. xi

List of Figures

Chapter 1

Figure 1.1: General representation of where cell wall polysaccharides, 4 glycan containing lipid, and proteins are synthesized.

Figure 1.2: Structures of common pectin polymers found in plant cell walls. 6

Figure 1.3: Structures of common Hemicellulose polymers found in plant 15 cell walls.

Figure 1.4: Schematic of cellulose biosynthesis. 21

Figure 1.5: Structures of Lignin and Suberin monomers. 34

Figure 1.6: Structures of N-linked Glycans and GPI-Anchors found in plant 41 cells.

Figure 1.7: Structures of GlcCers and GIPCs found in plant cells. 49

Figure 1.8: Structures of cellulose biosynthesis inhibitors and RG-II 55 biosynthesis inhibitor.

Figure 1.9: Hypothetical SN2 mechanisms of deoxy and fluor/Azido 63 monosaccharide analogs in non-retaining GTs.

Figure 1.10: Monosaccharide analog screen on Arabidopsis thaliana. 65

Chapter 2

Figure 2.1: Effects of fucose analogs on Arabidopsis root growth. 125 xii

Figure 2.2: Effects of short-term 2F-Fuc treatment on Arabidopsis seedlings. 127

Figure 2.3: Matrix polysaccharide analysis of 2F-Fuc treated seedlings. 129

Figure 2.4: Analysis of fucosyltransferase mutants for 2F-Fuc sensitivity. 132

Figure 2.5: Chemical complementation of 2F-Fuc growth defects with 134 Boron.

Figure 2.6: Isolation of fkgp-3 and analysis of resistance to 2F-Fuc. 136

Figure 2.7: Schematic of 2F-Fuc and Ac3 2F-Fuc metabolic incorporation 138 into a plant cell.

Figure 2.8: Analysis of 2F-Fuc resistant fluoro fucose resistant (ffr) lines. 140

Figure 2.9: Mapping of ffr lines allelic to fkgp-3. 143

Chapter 3

Figure 3.1: Effects of 2-deoxy-2-fluoro-D-Mannose on Arabidopsis growth. 169

Figure 3.2: Cell wall analysis of 2F-Man treated seedlings. 172

Figure 3.3: Analysis of N-linked glycosylated membrane protein, GPI- 175 Anchored proteins, and Ascorbate content.

Figure 3.4: Analysis of GIPC headgroup monosaccharide composition. 179

Figure 3.5: Sphingolipid analysis of 7-day-old 2F-Man treated Arabidopsis 181 seedlings. xiii

Figure 3.6: Retesting of potentially resistant lines on 2F-Man. 184

Figure 3.7: Mapping of fmr6 and fmr11 F2 lines. 187

Figure 3.8: Assaying resistance of fmr mutants on 2F-Man and 2F-Glc. 188

Figure 3.9: Assaying glucose repression in fmr6 and fmr11. 190

Figure 3.10: Quantification of ATP and F6P in 2F-Man treated seedlings. 192

Chapter 4

Figure 4.1: Current model of glucose repression mediated by Hxk2. 215

Figure 4.2: Screening of fluoro sugars, iminosugars and known cellulose 225 biosynthesis inhibitors against Saccharomyces cerevisiae strain S288c.

Figure 4.3: Growth of S288c on 2F-Man in liquid cultures and solid media. 227

Figure 4.4: Principle of Bar-Seq 229

Figure 4.5: Growth of resistant strains from Yeast Deletion collection. 233

Figure 4.6: Analysis of hypersensitive strains from Yeast Deletion 235 collection.

Figure 4.7: Current model of Rim101 pathway activation in yeast by alkaline 238 stress.

Figure 4.8: Analysis of other deletion lines involved with glucose repression, 240 Rim101 pathway, and other genes (hxk1Δ and suc2Δ). xiv

Figure 4.9: Analysis of Hxk2 rescue against 2F-Man. S288c, hxk2Δ, and 242 pAG426GDP-EGFP-Hxk2.

Figure 4.10: Hxk2 activity on 2F-Man. 245

Chapter 5

Figure 5.1: Nojirimycin analog screen on Arabidopsis thaliana. 267

Figure 5.2: N-Dodecyl-deoxynojirimycin inhibits growth in Arabidopsis. 269

Figure 5.3: Sphingolipid analysis of ND-DNJ treated Arabidopsis seedlings. 272

Figure 5.4: Sphingolipid analysis of PDMP and DCB treated Arabidopsis 275 seedlings.

Figure 5.5: Root width measurements of ND-DNJ treated seedlings. 277

Figure 5.6: Cell wall composition in seedlings treated with ND-DNJ and 279 PDMP.

Figure 5.7: Callose staining of ND-DNJ treated seedlings. 282

Figure 5.8: Quantification of Cellulose Synthase Complex (CSC) velocities 284 upon ND-DNJ and PDMP treatment.

Chapter 6

Figure 6.1: Investigating suberin deposition in Arabidopsis seedlings on 314 cellulose biosynthesis inhibitors Dichlorobenzonitrile (DCB) and Isoxaben (ISX). Figure 6.2: Time course assay of suberin deposition on cellulose 315 biosynthesis inhibitors. xv

Figure 6.3: Quantification of polymeric suberin from 7-day-old roots. 317

Figure 6.4: Quantification of soluble suberin from 7-day-old roots. 319

Chapter 7

Figure 7.1: Hypothetical Mendelian genetics models for ffr lines with non- 331 recessive recombinant frequencies.

Figure 7.2: Models of glucose repression in Arabidopsis and S. cerevisiae. 341

Figure 7.3: Proposed models of glycosylated sphingolipid role in CSC 349 motility.

Figure 7.4: Proposed model of suberin deposition under cell wall damage. 356

Figure 7.5: Structures of monosaccharide and dual functioning 358 monosaccharide analogs.

Figure 7.6: Structures of alternative fluorinated monosaccharides and 364 Carbasugars.

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Chapter 1: Introduction to plant cell walls and their function.

I. Plant cell walls

Plants are terrestrial photosynthetic multicellular organisms responsible for producing most of the world’s oxygen. Plants are also responsible for the annual fixation of approximately 1011 metric tons of carbon dioxide into more reduced forms of carbon that are used for plant metabolism as well as human and animal nutrition (Beer et al., 2010).

In human society, plants have been utilized as renewable sources of food, animal feedstock, textiles, construction materials, pharmaceuticals, and other industrial feedstocks including bioenergy. Particularly, plant biomass serves as the primary raw material for most of the listed utilities above (Carroll and Somerville, 2009; Somerville,

2006). Approximately 70% of plant biomass is composed of biological material derived from the plant cell wall (Pauly and Keegstra, 2008). Plant cell walls are polysaccharide- rich extracellular matrices that encapsulate nearly all plant cells, and collectively, these polysaccharides are the most abundant biopolymers on the planet.

Functionally, cell wall polysaccharides are critical for normal growth and development in plants by providing structural support, serving as a barrier to external pathogens, maintaining turgor pressure for cell expansion, and determining the shape and size of plant cells (Cosgrove, 2005; Burton et al., 2010).

Plant cell walls can be broadly grouped into primary and secondary cell walls. Nearly all plant cells are surrounded by a primary cell wall, which is synthesized during cellular growth and development. Once a plant cell reaches its mature size, the primary cell wall is remodeled, and the secondary cell wall is synthesized. In the secondary cell wall, 2 additional polysaccharides are deposited such as hemicelluloses and non-polysaccharide polymers such as lignin and suberin (Hao and Mohnen, 2014; Meents et al., 2018;

O’Neill and York, 2018). The evolution of vascular plants would not be possible without the development of secondary cell walls as they provide the strength necessary to withstand the negative pressures generated by transpiration. Secondary cell walls are typically deposited between the primary cell wall and the plasma membrane in three distinct layers depending how cellulose is deposited (Zhong et al., 2019). The S1 layer has a crossed cellulose microfibril organization, the S2 layer is the thickest layer has the cellulose microfibrils layered in parallel to the axis cell elongation, and the S3 layer has cellulose microfibrils in an flat helical orientation surrounding the cell (Timell, 1967).

An overall key role in secondary cell walls is to rigidify the cell wall and provide the necessary structure for vascular tissues.

Structurally, cell wall polysaccharides are heterogenous but, can be grouped into three primary categories: acidic pectins, neutral hemicellulose, and cellulose. Each of these polysaccharide networks will be discussed in more detail below including non- polysaccharide cell walls and other cellular glycans.

II. Cell walls: Pectin structure and function

Pectins collectively represent the most unique and structurally complex group of wall polysaccharides found in plants, constituting approximately 35% of primary cell walls in dicots, ~10% in grasses primary wall, and up to 5% in woody tissues (Mohnen, 2008).

Pectin has been attributed to a number of functional roles, including cell-cell adhesion, ion binding, fruit development, pollen tube growth, seed hydration, cytokinesis, growth, 3 and development (Mohnen, 2008; Ochoa-Villarreal et al., 2012a; Anderson, 2015; Xiao and Anderson, 2013).

Pectins have valuable properties associated with human health, including reducing low density lipoprotein (LDL) cholesterol, where pectins have been commonly referred to as soluble fiber in nutrition (Hara et al., 1999), and regulating glucose serum levels.

Pectin consumption also positively influences the human gut microbiome (Koropatkin et al., 2012; Ndeh et al., 2017), triggers apoptosis in prostate and colon cancer cells, and aids in the efficacy of chemotherapy drugs (Jackson et al., 2007; Tan et al., 2018; Leclere et al., 2013). As a raw material, pectins have been used as gelling agents in food, cosmetics, biodegradable films, adhesives, paper substitutes, surface modifiers for medical devices, and have been explored as drug delivery vehicles for pharmaceuticals

(Mohnen, 2008; Anderson, 2015; Sriamornsak, 2003).

Pectins are synthesized in the lumen of the Golgi apparatus, and given the structural complexity of pectins, it is postulated that over 65 distinct enzymatic activities are required for complete synthesis (Mohnen et al., 2008). Polymers are then trafficked through the Trans Golgi Network (TGN) to the plasma membrane and extruded to the extracellular matrix (Figure 1.1) (Kim et al., 2014). Pectins have been conventionally classified into three primary domains: homogalacturonan (HG), rhamnogalacturonan-I

(RG-I), and rhamnogalacturonan-II (RG-II). Each class will be discussed in below in more detail. It is generally postulated that all three pectin polymers are chemically linked and physically associate with other cell wall polysaccharides particularly with cellulose

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Figure 1.1: General representation of where cell wall polysaccharides, glycan containing lipid, and proteins are synthesized. For each major glycan/polysaccharide shown, an example of a glycosyltransferase associated with each.

5

(Nakamura et al., 2002; Wang et al., 2012; Altartouri et al., 2019; Broxterman and

Schols, 2018; Wang et al., 2015).

One of the most interesting functions of pectins is the interactions it has with cellulose. Multidimensional solid-state nuclear magnetic resonance (SSNMR) of near- native rehydrated cell wall samples from Arabidopsis hypocotyls supplemented with 13C- labeled glucose, revealed some interactions between cellulose and xyloglucan, but a greater number of interactions were detected between cellulose and pectic polysaccharides homogalacturonan and rhamnogalacturonan-I (Wang and Hong, 2015;

Wang et al., 2015). These interactions may come into play in cell wall stiffness during lobe formation in epidermal pavement cells. During lobe formation, changes in homogalacturonan methylation state is changed to reduce stiffness of the wall while the change in cellulose microfibrils realign to increase lobe amplitude (Altartouri et al.,

2019). A key feature of cellulose-pectin interactions may result in strengthening the primary cell wall.

a. Homogalacturonan

Homogalacturonan (HG) is the most abundant of the pectic polysaccharides, comprising approximately 60 – 65% of total pectins. HG consists of a linear α-1→4- linked D-galacturonic acid (GalA) polymer that can be methyl esterified at the C-6 position of various GalA residues (Figure 1.2) (Ochoa-Villarreal et al., 2012a). HG is synthesized in the lumen of the Golgi by several Galacturonosyltransferase (GAUT) enzymes, such as GAUT1 and GAUT7. GAUT1 synthesizes HG in a complex with

GAUT7, which anchors GAUT1 to the Golgi (Atmodjo et al., 2011). Interestingly, 6

Figure 1.2: Structures of common pectin polymers found in plant cell walls. Known GTs associated with the addition of monosaccharides are highlighted in red with an arrow pointing to the specific glycosidic bond. RG-II side chain A (A), side chain B (B), side chain C (C), side chain D (D) are highlighted. In all bonds labeled, assume attachment at 1 position unless stated (β2 = β-(1,2)-linked). Unique features of polysaccharide confirmations are boxed.

7 immunoprecipitation experiments for GAUT1 or GAUT7 followed by tandem mass spectrometry identified QUASIMODO 3 (QUA3), which functions as an S-adenosyl-L- methionine-dependent methyltransferase using HG as a substrate to yield methyl esterified HG, but this interaction was only observed in Arabidopsis thaliana cell- suspension cultures (Miao et al., 2011). Another classically known gene is

QUASIMODO1 (QUA1)/ GAUT8, which encodes a membrane-bound glycosyltransferase

(GT) necessary for proper HG biosynthesis. Two loss-of-function mutants of Arabidopsis

QUA1, qua1-1 and qua1-2, displayed a reduction in GalA compared to wildtype plants and display cell-adhesion defects in hypocotyls (Bouton et al., 2002). In conjunction to

QUA1, QUA2 is a Golgi-localized HG methyltransferase. A nonsense mutation in QUA2, qua2-1, showed 50% less HG without affecting other polysaccharides. Additionally, transcriptomic analyses demonstrated that QUA1 and QUA2 are transcriptionally co- expressed (Mouille et al., 2007). Interestingly, a random mutagenesis screen on qua1-1 and qua2-1 lines revealed a suppressor of the cell-adhesion defect named ESMERALDA 1

(ESMD1). The ESMD gene encodes a Golgi-localized putative protein O- fucosyltransferase controlling the response to the synthesis of HG. The qua2-1 cell adhesion defect is abolished in the esmd1-1 background, although the reduction in cell wall GalA content persists, suggesting that ESMD1 plays a role in cell wall integrity sensing or signals for cell wall modifications that consequently lead to loss of cell- adhesion (Verger et al., 2016).

HG is de-methyl esterified by Pectin Methyl Esterases (PME, E.C. 3.1.1.11) to expose the carboxyl groups of HG. PMEs belong to the Carbohydrate Active enzyme

(CAZy) class 8 of carbohydrate esterases (Lombard et al., 2014) with 66 PME-encoding 8 genes in Arabidopsis thaliana (Sénéchal et al., 2014). There are two classes of PMEs encoded in the Arabidopsis genome, type-I PMEs have a short or absent N-terminal protein precursor (PRO) domain, and type-II PMEs contain multiple PRO domains. It is commonly accepted that the PME PRO domains are cleaved by SUBTILISIN-LIKE

SERINE (SBT) protease in the Golgi in order to be secreted to the cell wall. Structurally,

PRO-domains resemble PME inhibitors (PMEIs) suggesting cleavage of PRO-domains are required for PME activity (Levesque-Tremblay et al., 2015). De-methyl esterification of HG occurs in two distinct patterns, block-wise and random. In the block-wise de- methyl esterification pattern, PMEs remove methyl groups from large stretches of HG creating long negatively charged regions. These areas interact with calcium ions to form egg-box motifs rigidifying the cell wall (Figure 1.2) (Jarvis and Apperley, 1995; Hocq et al., 2017). Random de-methyl esterification occurs under low pH conditions and is often one of the first steps in cell wall loosening and expansion (Hocq et al., 2017;

Wakabayashi et al., 2003). It is still not clear which specific PMEs contribute to each type of HG de-methyl esterification and limited insight has been established for different tissues and seed coats. Some examples of PMEs include PME34 and PME6 that are critical for heat tolerance and stomatal conductance under CO2 respectively (Rui et al.,

2018). Other examples include PME58 in seed coat mucilage, PME5 in shoot development, and PME3 required for root growth (Hocq et al., 2017; Turbant et al.,

2016).

PMEIs are regulators of PMEs and inhibit de-methyl esterification of HGs. The

PMEI family is similarly large compared to the PME family, with the Arabidopsis genome encoding 69 PMEI genes in total, suggesting that for every PME there is a likely 9 a specific PMEI to inhibit it (Scheler et al., 2015). PMEIs bind to the active site of PME and prevent further de-methyl esterification of HG, possibly preventing cell wall degradation or remodeling by the release of oligogalacturonides through the action of endopolygalacturonase (Latarullo et al., 2016; Sénéchal et al., 2014). Interestingly, under biotic stress of necrotrophic pathogens, PMEIs inhibit PMEs excreted by the pathogen

(Levesque-Tremblay et al., 2015; Sénéchal et al., 2014; Gigli-Bisceglia et al., 2020).

Several studies have investigated PMEI overexpressing lines to assess their functions. For example, an AtPMEI2 overexpression line limited vegetative growth, and the corresponding pmei2-1 mutant line displayed larger vegetative growth compared to wildtype. Additionally, the PMEI1 and PMEI2 are critical for pollen tube development

(Raiola et al., 2004; Wolf et al., 2003). Another interesting PMEI is PMEI6 which promotes proper release of seed coat mucilage (Saez-Aguayo et al., 2013). Overall, HG structure is relatively simple yet the fine tuning of HG methylation leads to complex and critical physiological functions including regulating growth, pollen tube development, and pathogen defense (Mravec et al., 2017; Bosch et al., 2005; Gigli-Bisceglia et al.,

2020).

b. Rhamnogalacturonan-I

Rhamnogalacturonan-I (RG-I) is the second most abundant pectic polysaccharide, making up anywhere between 20 – 35% of total pectins in the cell wall (Mohnen, 2008).

RG-I is composed of a backbone with a repeating GalA and L-Rhamnose (Rha): [α-

(1→2)-D-GalA-α-(1→4)-L-Rha], that is partially branched at C-3 and C-4 in Rha with arabinans composed of L-arabinose furanose (Araf) [α-(1→5)-Araf] or galactans 10 composed of D-galactose (Gal) [β-(1→4)-D-Gal] (Ochoa-Villarreal et al., 2012a;

Mohnen et al., 2008). There are also branched arabinogalactan side chains built with a backbone commonly consisting of [β-(1→4)-D-Gal] with branching at C-6 position with

[α-(1→3)-Araf] (O’Neill and York, 2018). Additionally, many of the galactan and arabinan side chains are capped with an α-linked L-fucose (Fuc) (Figure 1.2) (Nakamura et al., 2001).

The biosynthesis of the RG-I backbone is poorly understood, but the RG-I backbone is thought to be synthesized by GAUT11 and GAUT-Like 5 (GATL5) in

Arabidopsis. These enzymes have been implicated in the addition of GalA using UDP-

GalA as a substrate in mucilage RG-I, although each added GalA to an existing oligo polymer indicating both are required the production of RG-I backbone (Caffall et al.,

2009; Kong et al., 2013). The addition of Rha to the RG-I backbone is carried out by RG-

I: RHAMNOSYLTRANSFERASE 1 (RRT1) utilizing UDP-L-Rha as a substrate and an

RG-I oligomer, also implying it does not initiate RG-I biosynthesis. Interestingly, RRT1 is involved in RG-I mucilage biosynthesis (Takenaka et al., 2018). There are four RRTs encoded in the Arabidopsis genome. Only the rrt1 has been described but single mutants of rrt2, rrt3, and rrt4 did not affect RG-I content (Takenaka et al., 2018). To our knowledge a triple knockout of rrt2/rrt3/rrt4 has not been described. Currently there has been little success in elucidating initiators of RG-I biosynthesis in general and no

GAUT/GATLs or RRTs have been identified in other tissues. Arabinan side chains in

RG-I are primarily biosynthesized by Arabinan Deficient 1 (ARAD1), where homozygous knockouts of ARAD1 (arad1-1) display a 54% reduction in Ara content in stems (Harholt et al., 2006, 2012). Galactans are synthesized by a protein complex of 11

Galactan Synthase 1 (GALS1), GALS2, and GALS3 with GALS1 acting as the initiator and the combination of all three elongate a growing Galactan polymer (Ebert et al.,

2018a; Laursen et al., 2018). Fucosyltransferase enzymes involved in the addition of Fuc to the ends of galactan and arabinan side chains have not been elucidated.

The overall function of RG-I is still poorly understood. One hypothesis is that

RG-I functions as a linkage support for other pectic polysaccharides such as HG and

Rhamnogalacturonan-II (see below) (Ochoa-Villarreal et al., 2012a; Caffall and Mohnen,

2009). More interestingly, RG-I is modified in seed-mucilage by removing galactan and arabinan by Mucilage-Modified 2 (MUM2) and Beta-Xylosidase 1 (BXL1) respectively.

Mutations of each perturb the expansion of RG-I in seed-mucilage when exposed to water (Arsovski et al., 2009; Dean et al., 2007). Additionally, the mum2-1 RG-I contained oxidized Gal in branched RG-I galactans, not detected in wildtype.

Interestingly, a mutation in a gene called RUBY-particles in mucilage (ruby-1), encoding a galactose oxidase, suppressed the mum2-1 phenotype and did not display any oxidized

Gal. The ruby-1 mutant did display cell adhesion defects in seed coat epidermal cells

(Šola et al., 2019)

c. Rhamnogalacturonan-II

Rhamnogalacturonan-II (RG-II) is both the most complex polysaccharide in the cell wall, consisting of 12 unique monosaccharides and over 20 unique glycosyl-linkages.

RG-II comprises the smallest portion of pectic polysaccharides, representing approximately 8 – 10% (Mohnen, 2008). The structure of RG-II was first elucidated from red wine residue and its structure has been relatively well conserved among other plants 12

(Pellerin et al., 1996), although there has been a previous report indicating some variations in the larger side chains (Pabst et al., 2013). RG-II consists of a backbone of linear [α-(1→4)-GalA] resembling non-methyl esterified HG with 4 unique side chains.

Side chain A and side chain B branch at the C-2 position in the GalA backbone both beginning with D-Apiose (Api) and Rha [β-(1→2)-Api-β-(1→3)-Rha] but structurally diverge from Rha. Side chain A also contains GalA, L-galactose (L-Gal), D-xylose (Xyl), and L-fucose (Fuc), while side chain B contains L-aceric acid (AceA), D-galactose (Gal),

L-arabinose pyranose (Arap), Araf, and Fuc. Side chains C and D are the simplest in structure consisting of 2 individual monosaccharide both attaching at the C-3 position of the GalA backbone. Side chain C contains both 2-keto-3-doxy-D-manno octulosonic acid

(Kdo) and Rha [α-(2,3)-Kdo-α-(1,5)-Rha], while side chain D contains 2-keto-3-deoxy-

D-lyxo heptulosaric acid (Dha) and Araf [β-(2,3)-Dha-β-(1,5)-Araf] (Figure 1.2)

(Mohnen, 2008; Pellerin et al., 1996; Bar-Peled et al., 2012).

Of all cell wall pectic polysaccharides, RG-II biosynthesis is the least understood.

One of the few characterized glycosyltransferases involved in RG-II biosynthesis is

Rhamnogalacturonan-II xylosyltransferase 1 (RGXT1), and its homologs RGXT2 and

RGXT4. RGXTs transfer Xyl utilizing UDP-Xyl as a substrate to a Fuc acceptor yielding a [α-(1→3)-] linkage found on side chain A in RG-II. Individual knockouts of RGXTs had no discernable phenotype implying a functional redundancy between RGXT1 and

RGXT2 (Egelund et al., 2006, 2008). RGXT4 on the other hand is critical for pollen tube and root growth (Liu et al., 2011). Two Sialyltransfase-like proteins, SIA1 and SIA2, have been hypothesized to catalyze the addition of Dha and Kdo to RG-II side chains, although further investigation is required to confirm their catalytic activity (Dumont et 13 al., 2014). Despite few GTs associated with RG-II biosynthesis, other enzymes associated with RG-II biosynthesis have been identified. One example is MURUS1 (MUR1), which encodes a GDP-D-Mannose-4,6-dehydratase that catalyzes the first step of GDP-Fuc biosynthesis. The mur1-1 mutant has little to no Fuc in all wall polysaccharides

(xyloglucan, RG-I, Arabinogalactans), but it is in RG-II that loss of Fuc results in reduced growth (Bonin et al., 1997). Interestingly, the loss of Fuc (6-Deoxy-L-Galactose) is partially compensated by the structurally similar monosaccharide L-Gal in RG-II

(Reuhs et al., 2004). In Nicotiana benthamiana UDP-Api/UDP-Xyl synthase (NbAXS) has been shown to produce UDP-Api and UDP-Xyl for RG-II biosynthesis, with knockdowns exhibiting reduced growth and premature cell death due to loss of Api (Ahn et al., 2006). Additionally, GOLGI-LOCALIZED NUCLEOTIDE SUGAR

TRANSPORT 3 (GONST3) also known as Golgi GDP-L-Gal transporter (GGLT) transports GDP-L-Gal into the Golgi to the site where a hypothetical L-

Galactosyltransferase incorporates L-Gal into RG-II. Knockdowns of GGLT1

Arabidopsis plants displayed dwarf phenotypes (Sechet et al., 2018). Interestingly, the mur1-1 and GGLT-silenced phenotypes can be partially rescued by exogenous boric acid.

The most unique feature of RG-II is the self-dimerization using boron as a cross-link

(Funakawa and Miwa, 2015; Pellerin et al., 1996). This boron cross-link occurs in side chain A of RG-II through Api and is postulated to be formed during polysaccharide biosynthesis and secretion but, not in the extracellular matrix (Chormova et al., 2014).

RG-II has also been shown to create boron cross-links with Glycoinositol

Phosphoceramides from Rosa cell cultures (Voxeur and Fry, 2014). 14

III. Cell walls: Hemicellulose structure and function

a. Mannan and Glucomannan

Mannan is a homomeric polymer consisting of β-(1→4)-linked D-mannose (Man)

(Figure 1.3) found in lignified secondary cell walls of gymnosperms and in Arabidopsis can be found in thickened secondary cell walls of stem and leaf tissues (Handford et al.,

2003). Earlier studies indicated mannan is synthesized by Cellulose Synthase-Like D

(CSLD) proteins CSLD2, CSLD3, and CSLD5 which are crucial for the transition from early developmental stage to the next in Arabidopsis. Immunodetection using LM21 monoclonal antibody that detects β-(1,4)-linked mannan oligos (Marcus et al., 2010) in the csld2/csld3/csld5 triple mutant had no detectable levels of mannan in stems.

Additionally, the triple mutant heavily accumulated anthocyanins, suggesting loss of mannan biosynthesis leads to stress response (Verhertbruggen et al., 2011). Interestingly, the double mutant csld2/csld3 had identical growth to wildtype in 7-day-old seedlings. In contrast double knockouts of csld2/csld5, csld3/csld5 and the triple knockout had severe reduction in root growth suggesting CSLD5 is required during early developmental stages unlike CSLD2 and CSDL3 (Yin et al., 2011). Additionally, heterologously expressed CSLD5 in tobacco microsomal preparations incorporated Man onto an endogenous acceptor using GDP-[14C]-Man. Suggesting CSLD5 is sufficient for mannan biosynthesis in early developmental stages. Interestingly, microsomes expressing CSLD2 or CSLD3 had no radio labeled Man incorporation activity, but microsomes expressing

CSLD2 and CSLD3 had incorporation activity suggesting CSLD2 and CSLD3 are not redundant and form a complex together to produce mannan (Yin et al., 2011;

Verhertbruggen et al., 2011). Interestingly, mannan biosynthesis is not exclusive to 15

Figure 1.3: Structures of common Hemicellulose polymers found in plant cell walls. Known GTs associated with the addition of monosaccharides in Arabidopsis are highlighted in red with an arrow pointing to the specific glycosidic bond. In all bonds labeled, assume attachment at 1 position unless stated (β2 = β-(1,2)-linked).

16

CSLD proteins, CELLULOSE SYNTHASE-LIKE A (CLSA) proteins have also been shown to produce mannan in a synthetic biology approach (Voiniciuc et al., 2019;

Liepman et al., 2005).CSLAs are associated with glucomannan synthesis, Recombinant expression of CSLA2 and CSLA9 have been shown to produce mannan as well. CSLA9 expressed in Drosophila Schneider 2 cells produced two distinct polymers depending on the radiolabeled nucleotide sugar added. Addition of GDP-[14C]-D-mannose (GDP-[14C]-

Man) produced a β-(1→4)-linked mannan glycan and a β-(1→4)-linked glucan polymer was produced when GDP-[14C]-D-glucose (GDP-[14C]-Glc) (Liepman et al., 2005).

Additionally, CSLA2 expressed in Pichia pastoris produced relatively pure mannan polymer using endogenous GDP-Man (Voiniciuc et al., 2019). Mannan produced by

CSLD and CSLA proteins indicate promiscuous selectivity between GDP-Man and GDP-

Glc. This promiscuity is further implicated in a chimera between CSLD3 and

CELLULOSE SYNTHASE 6 (CESA6). The catalytic region between the second and third transmembrane domain of CSLD3 was fused with the N and C-terminus containing the transmembrane domains of CESA6 yielding CESA6:CSLD3 Catalytic Domain

(CESA6:D3CD). When expressed in the cesa6 mutant, procuste1-1 (prc1-1), rescued the phenotype and cellulose content was restored to wildtype. Kinetic assays of CSLD3 in proteoliposomes demonstrated CSLD3 used UDP-Glc as a substrate (Yang et al., 2020)

No catalytic activity was observed when GDP-Man was used as a substrate, but this may be due to CSLD3 dependence on CSLD2 for mannan synthase activity (Yin et al., 2011;

Verhertbruggen et al., 2011). This result suggest CSLD3, a known for mannan biosynthesis, has a catalytic domain is also capable of synthesizing β-(1→4)-glucans.

Overall mannan biosynthesis is carried out by both CSLD and CSLA proteins. 17

Glucomannan polysaccharides consist of a β-(1→4)-linked Man with β-(1→4)- linked Glc disbursed throughout the backbone (Figure 1.3) (Pauly et al., 2013).

Glucomannan in Arabidopsis is synthesized by Cellulose Synthase-Like A (CSLA) proteins CSLA2, CSLA3, CSLA7 and CSLA9. CSLA7 is a putative glucomannan synthase critical for embryogenesis with csla7 homozygous mutants having arrested growth during embryogenesis (Goubet et al., 2003). Heterologous expression of CSLA9 in Drosophila Schneider 2 cells displayed mannan and glucomannan synthase activity through the incorporation of GDP-[14C]-Man or GDP-[14C]-Glc respectively in reaction mixtures containing non-radiolabeled GDP-Man and GDP-Glc (Liepman et al., 2005).

Stems of Arabidopsis plants homozygous for csla9-1 had substantial labeling intensity with an anti-mannan antibody and displayed little glucomannan in polysaccharide analysis by carbohydrate gel electrophoresis (PACE). Interestingly, stems from the double knockout csla2-1/csla3-2 had similar mannan labeling as wildtype, but the triple mutant csla2-1/csla3-2/csla9-1 had no detectable mannan labeling. Additionally, the double knockouts clsa2-1/csla9-1 and csla3-2/clsa9-1 had less mannan labeling than the clsa9-1. Unexpectedly growth and stem strength was unaffected in any of the mutants

(Goubet et al., 2009). Together these results suggest that CSLA2, CSLA3, and CSLA9 synthesized glucomannan in stems but loss of glucomannan has no observable defects in stems. Glucomannan appears to be an important polymer in embryogenesis, as the csla7 is lethal.

b. Xyloglucan 18

Xyloglucan is one of the most prominent hemicelluloses in dicot plants, comprising 20 – 30% of hemicelluloses in the primary cell wall. It consists of a core β-

(1→4)-Glucan backbone with every Glc residue linked to an α-(1→6)-D-xylose (Xyl) residue. Every other Xyl monosaccharide in xyloglucan is decorated with Gal is in a β-

(1→2)-linkage and occasionally an α-(1→2)-Fuc is added to Gal (Figure 1.3) (Pauly et al., 2013). The core glucan chain backbone is synthesized by Cellulose Synthase-Like C

4 (CSLC4) in Tropaeolum majus seeds. Activity of CSLC4 was assessed in vitro by heterologous expression of TmCSLC4 in Pichia pastoris resulting in the production of a

β-(1,4)-Glucan polymer. This same study also found an Arabidopsis CSLC4 that is co- expressed with other xyloglucan biosynthetic genes (Cocuron et al., 2007). Five xyloglucan xylosyltransferases (XXTs) genes are encoded in the Arabidopsis genome, which are postulated to possess xyloglucan α-(1→6)-Xyl transferase activity onto a β-

(1→4)-Glucan in vitro (Vuttipongchaikij et al., 2012). Of the five, three have been studied in Arabidopsis. Arabidopsis mutants of XXT1 or XXT2 were phenotypically indistinguishable from wildtype, suggesting that XXT1 and XXT2 are genetically redundant. On the other hand, plants carrying the double mutant xxt1/xxt2 grew more slowly and were smaller at maturity. The xxt1/xxt2 double mutant also exhibited shorter root hairs (Cavalier et al., 2008). Analysis of xxt1/xxt2 and wildtype by digesting root cell walls using Driselase, a cocktail of endo- and exoglycanases lacking α-xylosidase activity producing isoprimeverose (Xyl-α-(1→6)-Glc), revealed the xxt1/xxt2 had no detectable isoprimeverose, while wildtype had a detectable peak through HPAEC analysis (Cavalier et al., 2008). XXT5, another putative xyloglucan xylosyltranferase in Arabidopsis, also influences root hair morphology. Unlike xxt1 and xxt2, the xxt5 root hairs were shorter 19 and displayed a bubble-like extrusion at the tip. Overall, the xxt5 plants had reduced xylosylated xyloglucan in whole seedlings (Zabotina et al., 2008). MUR3 is a galactosyltransferase that adds a β-(1→2)-Gal onto Xyl residues. The mur3 mutant had severely altered xyloglucan composition, exhibiting reduced Gal and Fuc xyloglucan content, but the resulting mutant was phenotypically comparable to wildtype in terms of overall growth and inflorescence stem tensile strength (Madson et al., 2003). Finally,

MUR2/ FUCOSYLTRANSFERASE1 (FUT1) transfers Fuc to Gal in xyloglucan to form an α-(1→2)-linkage. Similar to the mur3 mutant, mur2 had a phenotype nearly indistinguishable from wildtype. Additionally, the tensile strength of cell walls was nearly identical. Analysis of xyloglucan in mur2 had no detectable fucosylated xyloglucan (Vanzin et al., 2002).

Xyloglucan was first hypothesized to cross-link with cellulose microfibrils and aid in cellulose’s load-bearing capacity (Pauly et al., 2013). This hypothesis does not seem to be supported based on genetic knockouts of xyloglucan biosynthesis genes described above, although it has been shown that xyloglucan influences cellulose deposition and alters microtubule patterns based on observations in the xxt1/xxt2 double knockout etiolated hypocotyls (Xiao et al., 2016). Interestingly, xyloglucan does assist in longitudinal expansion in stomatal aperture as the xxt1/xxt2 had stomatal apertures were smaller compared to wildtype (Rui and Anderson, 2016).

c. Xylan

Xylan is the most prominent hemicellulose in secondary cell wall. It consists of a linear β-(1→4)-Xyl back bone with regular distribution of α-(1→2)-GlcA, α-(1→2)-Ara, 20

α-(1→2)-Ara- β-(1→2)-Xyl, and occasionally α-(1→2)-Ara-5-ferulate in monocots

(Figure 1.3) (Rennie and Scheller, 2014; Pauly et al., 2013). The backbone of xylan is synthesized by a triad of glycosyltransferases belonging to GT47 and one in GT43 family. These enzymes include Irregular Xylem 9 (IRX9), IRX10, and IRX14. Loss-of- function mutations in any of the three enzymes leads to irregular xylem development in secondary cell walls and compromised growth. Arabidopsis stem microsomal fractions of each mutant showed reduced xylosyltransferase activity when supplemented with an exogenous acceptor β-(1,4)-Xyl6 (Lee et al., 2012, 2007; Brown et al., 2007). In

Arabidopsis, xylan is commonly decorated with α-(1→2)-GlcA mediated by Glucuronic acid substitution of xylan 1 (GUX1) and GUX2. The gux1-2/gux2-1 double knockout had no detectable xylan with substituted GlcA. Stems from the gux1-2/gux2-1 were weaker compared to wildtype, but did grow normally (Mortimer et al., 2010). The weakened stems may be the result of less xylan-lignin cross-links facilitated by GlcA in xylan (Hao and Mohnen, 2014; Kang et al., 2019).

IV. Cell walls: Cellulose biosynthesis and regulation

Cellulose is the most abundant biopolymer on the planet and the main load-bearing component of the cell wall. It consists of a linear polymer of β-(1→4)-Glucose (Glc)

(Figure 1.4A) molecules arranged in a paracrystalline array consisting of 18 cellulose polymers (Figure 1.4B) (Cosgrove, 2005). Cellulose is the primary raw material used in textiles, ruminant animal production, and as feedstock in ethanol production (Robertson et al., 2017; Somerville, 2006). It is also the primary source of insoluble fiber in the human diet. Despite what appears to be its simplicity in structure, cellulose is surprisingly 21

Figure 1.4: Schematic of cellulose biosynthesis. (A) Structure of a single cellulose polysaccharide representing β-(1,4)-Glc. (B) Cross section of a proposed 18-mer crystalline cellulose microfibril displaying an 2:3:4:4:3:2 confirmation. (C) General organization of CSC representing a trimer of CESAs (CESA1: red, CESA3: purple, CESA6: green) in a 1:1:1 ratio organized as a single subunit (gray) part of the hexamer. (D) Organization of CSC and accessory proteins actively synthesizing cellulose trekking along cortical microtubules (blue/purple). (?) Interaction with COB and cellulose to form crystalline cellulose remains as a hypothetical.

22 complex in terms of the structure of its microfibrils, its biosynthesis, and the protein complexes required to produce cellulose.

a. Cellulose production

Cellulose is synthesized at the plasma membrane by the cellulose synthase complex (CSC), which consist of multiple Cellulose Synthase A (CESA) catalytic subunits (Paradez et al., 2006; Purushotham et al., 2016a; Cho et al., 2017). The CESA subunits utilize UDP-Glc as a donor substrate to attach new Glc units to an elongating β-

(1→4)-glucan chain and travel along the plasma membrane following cortical microtubules (Paredez et al., 2006). Two in vitro studies demonstrated catalytic activity of CESA5 and CESA8 from Physcomitrella patens and Populus tremula x tremuloides respectively synthesizing measurable cellulose fibrils in liposomes (Purushotham et al.,

2016b; Cho et al., 2017). There are 10 CESA genes encoded in the Arabidopsis genome each playing specific roles in plant development. The CSC requires CESA1, CESA3, and

CESA6 (or CESA6-like) for primary cell walls (Persson et al., 2007; Desprez et al., 2007).

For secondary cell wall cellulose deposition, CESA4, CESA7, and CESA8 are required

(Turner and Somerville, 1997; Taylor et al., 2003; Persson et al., 2007). The remaining

CESAs (CESA2, CESA5, CESA9, and CESA10) play minor roles in tissue specific processes, such as seed coat cellulose deposition (Somerville, 2006).

Most studies investigating the structural aspects of CESAs have been primarily been based on the crystal structure of bacterial cellulose synthase A (BcsA) shown to produce a single β-(1→4)-glucan chain, has served as a working model for cellulose biosynthesis. This model demonstrated the extrusion of the expanding glucan through the 23 transmembrane region and conformational changes of the catalytic domain (Morgan et al., 2013, 2016). Both the CESA and BcsA proteins contain 8 transmembrane (TM) domains and contain the amino acid motif associated with many processive GTs belonging to the GT-2 superfamily (cazy.org) (Saxena and Brown, 1997; Morgan et al.,

2013; Nagahashi et al., 1995). The CESA proteins carry additional domains that distinguish them from BcsA. Plant CESAs contain a Zinc finger domain between the N- terminus and the first transmembrane domain, and this domain is hypothesized facilitate the oligomerization of CESA proteins (Kurek et al., 2002). Additionally, CESAs contain a Plant-Conserved Region (P-CR) and Class-Specific Region (CSR) sharing an 80% and

40% sequence similarity respectively, in the Arabidopsis genome. The role of these regions remains unclear but, computational modeling of the Gossypium hirsutum CESA1

(GhCESA1) cytosolic domain predicted the P-CR and CSR function as aids in CSC formation (Sethaphong et al., 2013, 2016). Recently, the cryo-electron microscopy (EM) structure of a CESA8 trimer from Populus tremula x tremuloides (Ptt) was described

(Purushotham et al., 2020). PttCESA8 forms a homo-trimeric structure with a 3-fold symmetry down a axis and is perpendicular to the plasma membrane. P-CRs form 2 alpha helical structures and the second helix with a proceeding loop promotes and stabilizes the trimerization. P-CRs stabilization of the homo-trimer (Purushotham et al., 2020). The P-

CR interaction between PttCESA8 partially supports P-CR and CSR promoting complex formation hypothesis proposed by (Sethaphong et al., 2013, 2016) and rejecting the involvement of the Zinc finger domain in stabilizing the complex. The cryo-EM structure demonstrated the CSR does not interact with the CESA monomers, but faces the surface of the complex in the cytosolic region. The authors hypothesized CSR plays a 24 role in the formation of the hexametric complex, interacting with other CESA trimers.

The most peculiar aspect of the CESA8 trimer has to do with the Zinc finger, it self- dimerizes and protrudes out of the protein forming a stock-like structure. The zinc finger composes the tip of the stalk while the adjacent hypervariable region makes up the center of the stock. The overall the resolution of the stock was too low, but it is hypothesized to promote hetero-trimerization with other CESA proteins (Purushotham et al., 2020).

Alternatively, the hypervariable region adjacent to the Zinc finger domain, contains several predicted phosphorylation sites (Jones et al., 2016; Speicher et al., 2018) and may provide a scaffold for regulatory kinases to access their perspective phosphorylation site.

The Zinc finger may be the site where CELLULOSE SYNTHASE INTERACTING 1

(CSI1) interacts with the complex. Great progress has been made in the understanding of

CESA protein structure and with the first heteromeric trimer of CESA8 is the first major step in understanding how the CSC complex forms and how hetero-complexes interact.

Based on freeze fracture electron microscopy, the CSC displays a six-fold symmetry ranging between 25 – 30 nm in diameter. Initial analysis of immunogold labeling indicated the CSC is composed of a 36-mer following a ratio of 1:2:3 producing cellulose microfibrils consisting of 36-glycan polymers (Mueller and Malcolm Brown,

1980; Kimura et al., 1999; Doblin et al., 2002). More recently, increasing evidence suggest the CSC is an 18-mer consisting of a hexamer of trimers following a 1:1:1 ratio based on computational, quantitative immunoblotting, immunoprecipitation mass spectrometry, and 13C-NMR (Figure 1.4C) (Gonneau et al., 2014; Nixon et al., 2016; Hill et al., 2014; Vandavasi et al., 2016; Kubicki et al., 2018; Kumar et al., 2018; Jarvis,

2018). The 18-mer hypothesis is supported by the cryo-EM CESA8 homo-trimer 25 structure (Purushotham et al., 2020). In conclusion, the CSC is composed of a hexamer of trimers likely producing cellulose microfibrils consisting of 18-glycan polymers.

b. CSC accessory proteins

As stated previously, cellulose biosynthesis is carried out by the CSC, yet only the catalytic components of the complex have been discussed. Several CSC-associated proteins are essential for proper cellulose biosynthesis and deposition, including proteins that control the crystallinity of cellulose microfibrils, guide the complex along microtubules, aid in cellulose synthesis, and intracellular trafficking (Figure 1.4D).

One of the first accessory proteins associated with CSCs was Cellulose Synthase

Interacting 1 (CSI1), which was identified in a yeast two-hybrid screen for interactors of

CESAs (Gu et al., 2010). CSI1 facilitates the co-localization of the CSC along cortical microtubules by interacting with the CSC and cortical microtubules (Li et al., 2012; Lei et al., 2012). CSI1 is a 2150 amino acid protein containing 10 armadillo repeats dispersed throughout the protein with a C2-domain near the C-terminus. The C2-domain is required for interaction to cortical microtubules and deletion of the C2-domain results in mislocalization of CSI1 to the cytosol (Bringmann et al., 2012). Loss-of-function csi1 mutants did not reduce cellulose content nor caused any severe defects in vascular bundle, but display a “twisting” morphology in roots, hypocotyls, and leaf rosettes.

Additionally, csi1 mutants also displayed shorter roots, hypocotyls, siliques, and stems.

These phenotypic defects are a result of uncoupled CSCs tracking across the plasma membrane in random directions (Bringmann et al., 2012; Gu and Somerville, 2010).

Furthermore, CSI1 also facilitates de novo exocytosis of CSCs using a plant-specific 26 protein PATROL1 (Zhu et al., 2018) and endocytosis under abiotic stress into small

CESA compartments (SmaCCs) or microtubule associated cellulose synthase compartments (MASCs) (Lei et al., 2015), SmaCCs/MASCs are TGN vesicles housing

CSCs (Crowell et al., 2009; Gutierrez et al., 2009). Interestingly, the forces generated by motile CSCs should make cortical microtubules positionally unstable, yet they remain positionally stable (Paradez et al., 2006). Cellulose-Microtubule Uncoupling (CMU) proteins are localized at the plasma membrane and anchor cortical microtubules which allows them to maintain their position. Disrupting CMU function causes lateral cortical microtubule displacement and lead to the “twisting” morphology similar to csi1 mutants

(Liu et al., 2016).

KORRIGAN1 (KOR1) is a β-(1,4)-endoglucanase that directly interacts with

CESA proteins and is required for proper cellulose deposition (Lane et al., 2001; S et al.,

2001; Dé Ric Nicol et al., 1998). The kor1 mutants exhibit several phenotypic defects associated with cellulose biosynthetic deficiencies, including impaired cytokinesis, reduced CSC velocities, reduced anisotropic cell growth, and reduced overall plant growth (Lane et al., 2001; Dé Ric Nicol et al., 1998; Vain et al., 2014; Lei et al., 2014).

Interestingly, KOR1 is a poly-N-linked glycosylated protein containing 8 different glycosylation sites. Loss of N-linked glycosylation of KOR1 abolishes its β-(1→4)- endoglucanase activity, leading to the phenotypes found in the kor1 mutants (Liebminger et al., 2013; Rips et al., 2014). The β-(1→4)-endoglucanase activity of KOR1 seems to be critical for CSC motility and CSC endocytosis under stress. In kor1-1 mutants treated with cellulose biosynthesis inhibitor CGA-325-615 (CGA), fewer CSCs were found in

MASC/SmaCCs compared to wildtype (Vain et al., 2014), suggesting that KOR1 plays 27 are role in this process. The β-(1→4)-endoglucanase activity of KOR1 is thought to cleave the growing cellulose microfibril, allowing the endocytosis machinery to efficiently remove the CSC from the plasma membrane without the potential steric hinderance from the microfibril, yet this remains to be experimentally demonstrated.

COBRA (COB) is a Glycosylphosphatidylinositol (GPI)-anchored protein localized at to the plasma membrane that plays a vital role in determining the orientation of cell expansion in Arabidopsis (Schindelman et al., 2001; Benfey et al., 1993; Roudier et al., 2002, 2005). Loss of COB leads to severe growth defects, complete loss of anisotropic cell expansion, reduced crystalline cellulose content, altered pectin methylesterification, and elicits an immune response (Schindelman et al., 2001; Ko et al.,

2006; Roudier et al., 2005; Sorek et al., 2015). Interestingly, a suppressor screen of cob in

Arabidopsis found a suppressor named mongoose 1 (mon1) that reduced the phenotypic defects associated with the cob mutant. The mon1 mutant was mapped to MEDIATOR16, a subunit of the transcriptional mediator complex suggesting that the cob phenotypes can be suppressed by disruption of a cell wall integrity sensing mechanism (Sorek et al.,

2015). Other mediators have been described to suppress detrimental phenotypes lignin biosynthetic mutants (Taylor-Teeples et al., 2015; Bonawitz et al., 2014). COBRA-like

(COBL) proteins have been described previously to interact with cellulose microfibrils in vitro, indicating these proteins play a role in cellulose biosynthesis and or aggregation of cellulose (Benfey et al., 1993). A few COBL genes have been described previously to be involved in cellulose biosynthesis in a tissue-specific manner. The rice COBL BRITTLE

CULM1 interacts with cellulose microfibrils and influences crystallinity (Liu et al.,

2013). COBL2 is part of a complex network of interacting pathways in seed coat 28 mucilage deposition that plays a specific role in cellulose deposition in seed coat mucilage, where disruption of the COBL2 gene reduces cellulose rays and increases pectin solubility when seeds are rehydrated (Ben-Tov et al., 2015, 2018). Another interesting COBL protein, COBL10, is critical for male fertility in Arabidopsis. Pollen carrying cobl10 mutations had reduced pollen tube growth and loss of directional sensing leaving them unable to find female ovules. Additionally, cellulose microfibril deposition was altered. These data suggest that proper coordination cell wall biosynthesis is crucial for plant reproduction (Li et al., 2013). Overall, the exact biological function COB and

COBL proteins remains relatively unclear, but it may play a role in crystalline cellulose or proper cellulose microfibril alignment.

The most recently identified accessory proteins of the CSC are Companion of

Cellulose synthase (CC) proteins. CC proteins were shown to directly interact with both

CSCs and cortical microtubules and are integral to the redelivery of CSCs to the plasma membrane after abiotic stress (Endler et al., 2015). CC proteins are unique to plants containing a cytosolic N-terminal domain, a single transmembrane region, and an apoplastic domain of unknown function. Additionally, CC proteins do not act redundantly to CSI1 and their function is independent of CSI1 (Endler et al., 2016).

Interestingly, the N-terminal cytosolic domain resembles a hTau40 architecture that promotes polymerization and bundling of cortical microtubules and has been shown in vitro to promote microtubule bundling in a Tau-like fashion (Kesten et al., 2019).

c. Regulation of CSCs 29

The CSCs activity, expression, and localization are tightly regulated through post- translational modified as well as trafficking through environmental stimuli and cellular cues (Speicher et al., 2018; Kesten et al., 2017; Polko and Kieber, 2019). Here, regulation of CSCs through common post-translational modification prompted by cellular processes and environmental stimuli outside of cell wall integrity sensing, discussed in in section VI. As dynamic as the CSC is and the constant stimuli from the environment such as light, mechanical stimuli, predators, pathogens, and soil conditions; rapid control of

CSCs are vital. The most dynamic form of regulation in CSCs is through post- translational modification, specifically through phosphorylation. Several phosphorproteomic studies have identified a myriad of phosphorylation sites, many of which are conserved across every CESA protein in the N-terminal region. Additionally, several CSC accessory proteins except for COB proteins have been found to be phosphorylated (Nühse et al., 2004; Nakagami et al., 2010; Facette et al., 2013).

The discovery of many CSC phosphorylation sites has encouraged further studies on the impact these phosphorylation sites in CSC regulation and catalytic activity. Initial studies utilized site directed mutagenesis to change Ser/Thr phosphorylation sites to a phosphonull (Ser/Thr to Ala) or phosphomimic (Ser/Thr to Glu). In the case of CESA1, several phosphorylation sites (S162, T165, T166, S167, and S686) were studied in the temperature sensitive CESA1 mutant radial swelling 1 (rsw1) (Chen et al., 2010; Arioli et al., 1998). Transgenic lines expressing CESA1T166E, CESA1S686E, CESA1S688E,

CESA1S162A, CESA1T165A, and CESAS167A fully rescued the rsw1 phenotype. The

CESA1T166A, CESA1S686A, CESA1S688A, CESA1S162E, CESA1T165E, and CESAS167E lines partially rescued the rsw1 phenotype but, still exhibited reduced cell expansion. Live-cell 30 imaging of these lines in the rsw1/prc1: YFP-CESA6 background demonstrated asymmetrical movement of CSCs across the plasma membrane (Chen et al., 2010). A later study of two phosphorylation sites in CESA3 (S211 and T212) the CESA3 mutant je5 background (Chen et al., 2016b). Expression of CESA3S211A negatively impacted hypocotyl and root length but, did not impact root hair length. In contrast, CESA3S211E reduced root hair length but hypocotyls and roots were not significantly impacted. The

CESA3T212E negatively impacted root hair length and hypocotyls but, not in primary roots. CESA3T212A mimicked wildtype root, hypocotyl and root hair lengths. Live-cell imaging of hypocotyls in each line showed CSCs were directionally asymmetric in

CESA3S211A and CESA3T212E, potentially explaining the reduced hypocotyl lengths observed. The CESA3S211E and CESA3T212A lines on the other hand displayed bidirectional velocities similar to wildtype (Chen et al., 2016b). Taken together, these findings suggest CSC activity is fine-tuned by phosphorylation in different tissues.

Phosphorylation and S-acylation of CESA7 have been studied previously and are implicated in protein degradation and delivery to the plasma membrane (Kumar et al.,

2016; Taylor, 2007). In CESA7, a peptide containing a variable region (N-terminal region) from amino acids 175 – 190 were shown to be phosphorylated in vivo. Plant extracts resulted in degradation via a proteasome dependent pathway. This suggest plants can utilized protein degradation as a mechanism for CSC regulation (Taylor, 2007). An alternative post-translational modification of CESA7 is through S-acylation. In CESA7, four cysteine residues in the N-terminus and two at the C-terminus were shown to be S- acylated and mutating these sites resulted in CSCs not being delivered to the plasma 31 membrane. This study predicted over 100 S-acyl groups in a single CSC (Kumar et al.,

2016).

The availability of light has been implicated in the regulation of CSC activity and its composition. For example, procuste 1-1 (prc1-1) is a mutation encoded in CESA6 causing reduced cell elongation of hypocotyls. Implicating deferential expression of

CESA proteins between different tissues (Fagard et al., 2000). Live-cell imaging of GFP-

CESA5/prc1-1 demonstrated reduced CSC velocity. Interestingly, application of microtubule destabilizer, Oryzalin, increased CSC velocity. This indicated the reduced

CSC velocity of cesa6 in the dark is caused some form of indirect interaction with cortical microtubules in CSCs containing CESA5 (Bischoff et al., 2011). This same study also demonstrated PHYTOCROME B (PHYB) increases the velocities of CSCs containing CESA5. Mutating four phosphorylation sites in the N-terminus of CESA5

(S122, S126, S229, and S230) to phosphomimics in the cesa6 background had greater

CSC velocities compared to the phosphonulls and CESA5 control. This implicates the activation of PHYB causes the phosphorylation of CESA5 increasing the activity of

CESA5 containing CSCs (Bischoff et al., 2011).

CSCs can also be regulated by a phytohormone brassinosteroids (BR) through the

BR-signaling pathway. BR-signaling begins when BR binds as a ligand to a lucine-rich receptor-like kinase BRASSINOSTEROID INSENSITIVE 1 (BRI1) and recruits

BRASSINOSTEROID INSENSITIVE 1-ASSOCIATED RECEPTOR KINASE (BAK1) and lead to a cascade of phosphorylation events leading to the phosphorylation of

BRASINOSTEROID INSENSITIVE 2 (BIN2) inhibiting its kinase activity. Substrates of 32

BIN2 are BRASSINAZOLE RESISTANT 1 (BZR1) and BRI1-EMS-SUPRESSOR

(BES1), when phosphorylated are targeted for degradation. (Planas-Riverola et al., 2019).

BZR1 and BES1 are transcription factors for thousands of genes in Arabidopsis including

CESA genes involved in both primary and secondary cellulose biosynthesis (Xie et al.,

2011). Interestingly, BR-signaling also directly regulates active CSCs through BIN2.

Arabidopsis mutants containing the constitutively active BIN2 mutation (bin2-1) exhibit reduced cellulose content in etiolated hypocotyls (Sanchez-Rodriguez et al., 2017).

Additionally, CSC velocities were reduced in the bin2-1 compared to wildtype. This same study also demonstrated BIN2 phosphorylates CESA1 at T157 but only under the condition where S162 is phosphorylated in vitro. This suggest not only is BIN2 a negative regulator of cellulose biosynthesis but also requires the priming of S162. This indicates at least two signaling inputs are required for the downregulation of CSC activity

(Sanchez-Rodriguez et al., 2017).

V. Non-polysaccharide wall components: lignin and suberin

Cell walls are not exclusively composed of polysaccharides. Other non- polysaccharide cell wall components include the phenolic polymer lignin and the lipid- phenolic biopolyester suberin. These polymers are not typically found in the primary cell wall and typically found in other sections. Lignin is typically deposited in secondary cell walls and suberin can be found between the primary cell wall and the plasma membrane commonly referred as suberin-lamellae (Graça, 2015; Terrett and Dupree, 2019)

a. Lignin 33

Lignin is a polyphenolic polymer deposited in secondary cell walls in vascular plants. This polymer is the primary contributor of the inherent recalcitrance in the production lignocellulose-based biofuels (Carroll and Somerville, 2009). Lignin consists of aromatic monolignols produced through the phenylpropanoid pathway using phenylalanine as the starting metabolite. Predominant monolignols within the lignin polymer characterize the type of lignin (H, G, S, and C lignin). Common monolignols are p-coumaryl alcohol (H unit), coniferyl alcohol (G unit), sinapyl alcohol (S unit) and caffeyl alcohol (C unit) (Figure 1.5) (Xie et al., 2018). C-lignin is an uncommon species of lignin found the seed coats of vanilla orchid and in cacti (Chen et al., 2013). A small molecule, p-iodobenzoic acid, is an inhibitor of monolignol biosynthesis targeting

Cinnamate-4-hydroxylase (C4H). This compound reduced overall lignin through reduction in p-coumaric acid (van de Wouwer et al., 2016). Monolignols are covalently linked through combinatorial radical coupling by generating free radicals by a tightly controlled cell wall localized oxidation systems LACCASE/O2 (LAC) and

PEROXIDASE/H2O2 (PRX). The formation of free radicals in monolignols is resonance stabilized providing more than a single point (carbon or oxygen) for radical coupling to occur. This situation also applies to the second monolignol. The potential for more than one possible radical coupling reaction, leads to a highly randomly polymerized lignin that contributes to plant biomass recalcitrance (Tobimatsu and Schuetz, 2019). Lignin is covalently bonded to the plant cell wall through modified xylan in secondary cell walls

(Terrett and Dupree, 2019). In Eudicots, lignin is attached to the carboxyl group in GlcA distributed in xylan polysaccharides. In Arabidopsis, two Glucuronic acid substitution of xylan (GUX) enzymes adds a single GlcA for every 8 xylosyl residues in xylan. The 34

Figure 1.5: Structures of Lignin and Suberin monomers. Monolignols of coniferyl alcohol (G), p-coumaryl alcohol (H), sinapyl alcohol (S), and caffeyl alcohol (C). Commonly found Suberin monomers are highlighted, (1) ω-hydroxy fatty acids (FAs), (2) Dicarbyxylic FAs, (3) Saturated FAs, (4) Primary alkyl alcohols, (5) Ferulic acid, and (6) Tyramine. Alternative functional groups to double bond found in (1) and (2) are boxed (7).

35

GUX1/GUX2 mutants (gux1/2) had drastically reduced recalcitrance to enzymatic cell wall deconstruction, showing increases in the amount of glucose and xylose, 30% and

700% respectively, released during saccharification (Lyczakowski et al., 2017).

Lignin plays several interesting roles in plants as a defense polymer, including possessing antimicrobial properties, acting as a physical barrier against pathogens, and preventing the diffusion of toxins produced by pathogens (Sattler and Funnell-Harris,

2013). Typically, ectopic lignin deposition can be induced upon damage to the cell wall, particularly resulting from cellulose biosynthesis defects. Commonly, cell wall damage caused biotic and abiotic stress induces ectopic lignin deposition (Gigli-Bisceglia et al.,

2018). Plants with compromised cellulose biosynthesis, such as the ectopic lignin1-1 mutant or plants treated with cellulose biosynthesis inhibitors, such as isoxaben, induce ectopic lignin deposition (Caño-Delgado et al., 2003; Denness et al., 2011). Interestingly, cellulose defect-associated ectopic lignin deposition is dependent in cell wall integrity sensing receptor like kinases THESEUS1 (THE1) and STRUBBELIG (SUB). In the the1-1 and sub-1 loss of function mutants still displayed a reduction in cellulose in the prc1-1/the1-1 and sub-1 treated with isoxaben, but had reduced ectopic lignin compared to the prc1-1 or isoxaben treated seedlings (Hématy et al., 2007; Chaudhary et al., 2020).

These observations suggest that lignin not only serves as a structural component in plant cell walls, but also serves as a defensive polymer upon cell wall damage.

b. Suberin

Suberin is a lipid-phenolic biopolyester deposited between the primary cell wall and plasma membrane of root endodermal and peridermal cells. Suberin is impermeable to 36 water and acts as a physical barrier against pathogens and toxins (Nawrath et al., 2013).

Many of the monomers composing suberin are interesting targets as potential replacements for non-renewable petrochemical-derived products for plastic manufacturing (Pinto et al., 2009). This complex polyester consists of several aliphatic and aromatic constituents, including fatty acids (FAs), alkyl primary alcohols, bifunctional ω-hydroxy FAs, Dicarboxy FAs, glycerol, ferulic acid, and tyramine (Figure

1.5) (Vishwanath et al., 2015). These monomers are hypothetically polymerized together based on Transmission Electron Microscopy (TEM) of suberin-lamellae cell walls consisting of light and dark bands. The light bands are hypothesized to primarily consist of aliphatic groups and the dark band consisting primarily of polyaromatics (Graça,

2015). Currently, the synthesis of the polyester remains controversial. A myeloblastosis

(MYB) transcription factor, MYB41, increases the transcription of genes encoding for the proteins necessary to produce suberin. Overexpression of AtMYB41 displayed elevated levels of suberin deposition in atypical tissues like epidermal and mesophyll cells in leaves. Additionally, monolignol content was elevated in the AtMYB41 overexpression lines, suggesting AtMYB41 may also influence lignin deposition. Interestingly,

AtMYB41 expression is increased under salt-stress, suggesting that this transcription factor may play a role in the response to abiotic stress (Kosma et al., 2014).

Similar to lignin, suberin biosynthesis is activated upon cell wall damage, pathogen induced immunity, or under drought stress (Kolattukudy, 2001). In Solanum tuberosum

L., wound periderm accumulates suberin over time preventing pathogen invasion and limiting water loss (Schreiber et al., 2005). Interestingly, an oligomer derived from cellulose, cellobiose, is perceived as a signal molecule similar to known elicitors such as 37 chitin oligomers and oligogalacturonides in Arabidopsis. Treatment with cellobiose both causes pattern-triggered immunity (PTI) and up-regulates AtMYB41 transcription, resulting in increased resistance to P. syringae and increased suberin biosynthetic gene expression respectively (de Azevedo Souza et al., 2017).

VI. Cell walls: Integrity sensing and Cell wall damage

Plants, being sessile organisms, must adapt to their environment to survive. As previously mentioned, the cell wall is the first barrier between a plant cell and its surrounding environment (Voxeur and Höfte, 2016). Plants commonly use receptor-like kinases (RLKs) to sense a myriad of environmental cues. In Arabidopsis, there are over putative 600 RLKs identified based on expression data on multiple abiotic and biotic stresses (Lehti-Shiu et al., 2009). A subfamily of RLKs, Catharantbus roseus Receptor- like kinase 1-like proteins (CrRLK1Ls), consisting of 17 members encoded in the

Arabidopsis genome. All CrRLK1Ls share similar domain structures containing a malectin-like domain, a transmembrane domain, and a cytosolic serine/threonine kinase.

Many of these CrRLK1Ls have diverse functions in cell growth, plant morphogenesis, reproduction, immunity, hormone signaling, and stress responses (Franck et al., 2018).

Here, a few CrRLK1Ls THESEUS1 (THE1), FERONIA (FER), ANXUR1/2,

BUDDHA’s PAPER SEAL1/2, and an atypical leucine-rich repeat receptor kinase

STRUBBELIG (SUB) are directly involved in cell wall integrity (CWI) (Vaahtera et al.,

2019).

a. Cell wall Integrity sensing mechanism 38

CWI sensing is a critical function in maintaining cell wall integrity during biotic or abiotic stress, or developmental. Compromises to CWI are often caused by cell wall damage (CWD), which can be sensed by the release of oligogalacturonides (OGs), cellobiose, or changes in cell wall confirmation (Vaahtera et al., 2019). A common response to CWD involves the increase in reactive-oxygen species (ROS), phytohormone production, deposition of lignin and callose, and arresting of the cell cycle (Gigli-

Bisceglia et al., 2020). A set of CrRLK1Ls, ANXUR1/ ANXUR2 (ANX1/ ANX2) and

BUDDHA’s PAPER SEAL 1/ BUDDHA’s PAPER SEAL 2 (BUPS1/BUPS2), from a complex together and are pollen tube specific receptors required pollen tube integrity sensing. Loss of function mutations in ANX/BUPS causes premature pollen tube bursting and growth arrest. The receptor complex is localized to the apical plasma membrane in pollen tubes (Boisson-Dernier et al., 2009; Li and Yang, 2018; Ge et al., 2017).

THESEUS1 was first identified as a membrane bound CrRLK1L expressed in elongating cells. Loss of function alleles of the1 (the1-1 and the1-2) attenuates the growth defects of CESA6 mutant prc1-1 in Arabidopsis (Hématy et al., 2007). Additionally, the1-

3 crossed with CESA1 (rsw1-10), CESA3 (eli1-1), and CSI (pom1-2) mutants also attenuated the phenotypes associated with each mutant. The phenotypes were also enhanced in mutants carrying THE1-overexpression (Hématy et al., 2007). THE1 is also involved in response to cellulose biosynthesis inhibitor, isoxaben. The the1-1 mutant also attenuated ectopic lignin deposition and ROS accumulation during isoxaben treatment

(Denness et al., 2011). Interestingly, NITRATE REDUCTASE 1 and NITRATE

REDUCTASE 2 (NIA1, NIA2) act downstream of THE1 signaling during CWD and are required for cell cycle repression associated with CWD. The double knockout (nia1/nia2) 39 did not repress cell cycle gene transcripts upon isoxaben treatment. Additionally, nia1/nia2 attenuated ectopic lignin deposition and phytohormone including the triple homozygous line, nia1/nia2/the1-4, the1-4 is a gain of function mutation in THE1 (Gigli-

Bisceglia et al., 2018). Another receptor kinase, STRUBBELIG (SUB), displays a similar phenotype to the1 mutants in terms of reduced ectopic lignin deposition upon isoxaben treatment. Additionally, SUB is required for the growth arrest in the prc1-1 as the sub-

9/prc1-1 partially rescued the prc1-1 phenotype (Chaudhary et al., 2020)

FERONIA is another well-known CrRLK1L most famously known in female fertility

(Capron et al., 2008). Outside of plant reproduction, FER is a positive regulator of plant- triggered immunity (PTI), with loss of function fer-2 and fer-4 mutants displaying mitigated ROS production after treatment with the elicitor molecule elf18. These mutants were also more susceptible to Pseudomonas syringae pv. Tomato DC3000 coronatine- minus strain (Stegmann et al., 2017). Additionally, fer-4 seedlings were hypersensitive to several abiotic stresses, inducing heat, cold, and salt stress. The fer-4 was also hypersensitive to Abscisic acid (ABA)-induced growth inhibition, suggesting FER inhibits the ABA growth inhibition responses (Chen et al., 2016a). The most direct effect of salt stress on the cell wall is the disruption of the egg-box motif of HGs. FER has been demonstrated to be a HG integrity sensor, as the fer-2 and fer-4 hypersensitivity to salt stress stems from elevated cell swelling and cell bursting compared to wildtype.

Additionally, the hypersensitivity of fer-4 mutants to salt stress can be partially rescued by exogenous calcium and boric acid minerals associated with HG and RG-II respectively. Salt stress is postulated to induce a FER-dependent calcium influx that 40

activates a cell wall repair mechanism to reinforce the wall to maintain wall integrity

(Feng et al., 2018).

VII. Plant Glycoproteins and Glycolipids: structure, function, and metabolism

Glycosylation events are critical in multiple biological process outside of cell wall

biosynthesis. (Lairson et al., 2008; Strasser, 2016; Henrissat et al., 2009; Michaelson et

al., 2016; Cheung et al., 2014). The major types of glycosylation processes that are

relevant to the results in this dissertation, such as protein glycosylation,

glycosylphosphatidylinositol-anchors, glucosylceramides, and glycoinositol

phosphoceramides, all have multiple functions, but all have an influence in cell wall

biosynthesis and cell wall integrity sensing.

a. Protein Glycosylation

Protein N-linked glycosylation is the most common co- and post-translational

modification for proteins destined to enter the secretory pathway. N-linked glycosylation

plays a critical role for several biological processes, such as a glycan-dependent quality

control process in the ER, protein stability, protein-protein interactions, plant stress

tolerance, and cellulose biosynthesis (Ebert et al., 2018b; Lukowitz et al., 2001; Strasser,

2016). In eukaryotes, a series of Asparagine (N)-linked glycosylation (AGLs) enzymes

synthesize a precursor glycan using Dolichol-pyrophosphate (Figure 1.6). The assembled

premature glycan is then transferred en bloc to a targeted Asp residue by the

oligosaccharyltransferase complex (Strasser, 2016). The consensus sequence of nascent

polypeptide for en bloc transfer is Asn-X-Ser/Thr (X signifying any amino acid), where

the glycan is attached to the asparagine residue. Interestingly a noncanonical sequence 41

Figure 1.6: Structures of N-linked Glycans and GPI-Anchors found in plant cells. Known GTs associated with the addition of monosaccharides in Arabidopsis are highlighted in red with an arrow pointing to the specific glycosidic bond. In all bonds labeled, assume attachment at 1 position unless stated (β2 = β-(1,2)-linked). Number Man residues are cleaved by Mannosidases.

42

(Asn-X-Cys) has been observed to be N-linked glycosylated in the double-repeat B subunit of Shiga toxin 2e (Matsui et al., 2011).

The completed N-linked glycosylated protein is then transported to the Golgi cisternae for further processing. The first major processing of the immature N-linked glycan is trimmed down removing Glc and several Man residues. The terminal Glc is removed by GLUCOSIDASE I (GCSI), genetic knockouts of GCS1 are embryo lethal as it produced shrunken non-viable seed (Boisson et al., 2001). GCSII appears to exist as a heterodimer consisting of GCSIIα and GCSIIβ that hydrolyze the remaining Glc residues

(Burn et al., 2002; Von Numers et al., 2010). The GCSIIα subunit hydrolyzes the other two Glc restudies. Interestingly, the first described mutation of GCSIIα was a temperature-sensitive mutant labeled as rsw3 as it displayed swollen root phenotype and reduced crystalline cellulose deposition similar to the rsw1-1 mutant. Attempts at obtaining a null-mutant of GCSIIα were not successful, suggesting GCSIIα function is essential (Burn et al., 2002). In contrast, gcsIIβ null mutants displayed no visible phenotypes, but displayed compromised immunity. Treatment of gcsIIβ with elf18, a known elicitor of the immune defense response, did not elicit ROS production or expression of defense genes (Von Numers et al., 2010). The final trimming processes are carried out by three α-MANNOSIDASES (MNS) glycosylhydrolases and GOLGI α-

MANNOSIDASE II (GMII) (Liebminger et al., 2010, 2009; Kang et al., 2008; Schoberer et al., 2019). All three MNS proteins trim the terminal Man residues in the highly branched portion of the N-linked glycan and two Man residues in the linear portion that previously had the Glc residues. MSN1 and MNS2 appear to be functionally redundant and cleave Man 1,2, and 3 (Figure 1.6), but the double knockout displayed reduced root 43 growth. MNS3 specifically cleaves a terminal Man 4 (Figure 1.6) of the N-linked glycan.

The triple mutant has N-linked glycans with all Man residues intact and a result severe growth inhibition (Liebminger et al., 2010, 2009; Schoberer et al., 2019). The last bit of trimming is carried out by GMII, cleaving Man 3 and 5 after MNS1/2/3 removes Man 4 and 6 (Figure 1.6). A null mutation of GMII, hybrid glycosylation 1 (hgl1) displays no visible phenotype under normal conditions but is hypersensitive to salt stress (Kang et al.,

2008). Overall, N-glycan trimming is a critical process for plant immunity, crystalline cellulose deposition, and salt-tolerance.

Post trimming the N-linked glycan is then decorated with several monosaccharides. One of the first monosaccharides added is GlcNAc by N-

AcetylGlucosaminyltransferase I (GNTI) and GNTII. Each GNT1 adds a single GlcNAc to the glycan. Both mutants of GNTI, complex glycan 1 (cgl1), and gntII do not display any observable phenotypes, but cgl1 is hypersensitive to salt stress (Kang et al., 2008;

Yoo et al., 2015). GALACTOSYLTRANSFERASE I (GALT1) catalyzes the addition of

Gal to the glycan and similarly with other N-linked glycan processing enzymes, no visible phenotype was observed in the galt1-1 and RNAi knockdown lines under standard conditions (Strasser et al., 2007). XYLOSYLTRANSFERASE (XYLT) incorporates xylose into the core Man of the N-linked glycan. Similarly to galt1-1, xylt mutants did not display a visible phenotype in Arabidopsis (Strasser et al., 2004). Fuc incorporation is the final monosaccharide to complete the mature N-linked glycan. Fucosylation of N- linked glycans is catalyzed by FUT11, FUT12, and FUT13. Mutations in each FUT gene do not cause any visible phenotypes in Arabidopsis (Strasser et al., 2004; Kang et al.,

2008). Glycosyltransferases in N-linked glycan maturation are summarized in Figure 1.6. 44

Collectively, mature N-linked glycans may not be essential for normal plant function but contribute to salt-tolerance.

Currently, over 2,000 potential N-linked glycosylation sites have been identified in Arabidopsis through the N-glyco-FASP proteomic method and two-dimensional nano-

LC/MS, using the deamination of glycosylated Asn using Protein N-linked Glycosidase

(PNGase) (Song et al., 2013; Zielinska et al., 2012). As mentioned previously, the presence of N-linked glycans on KOR1 is critical for proper function (Liebminger et al.,

2013; Rips et al., 2014). This can be indirectly supported by the loss of N-linked glycosylation in the point mutation in GDP-Mannose Pyrophosphorylase (cyt1-1) mutant.

This mutation exhibits growth inhibition and reduced cellulose content linked to loss of

N-linked glycosylation (Lukowitz et al., 2001). This observation alone indicates that simple N-linked glycosylation is critical for cellulose biosynthesis, but the ability to produce complex N-linked glycans has a genetic interaction with KOR1 mutants. For example, the hgl1-1 mutant lacks α-mannosidase II activity necessary for proper N-linked glycan trimming. Under normal conditions, the hgl1-1 roots grow similarly to wildtype, but this mutant is hypersensitive to salt stress. This phenotype is also observed in the complex glycan 1 (cgl1) mutant, which lacks N-acetyl-glucosaminyl transferase I in the

Golgi, and the staurosporin and temperature sensitive 3a (stt3a) mutant, which encodes a subunit of the oligosaccharyltransferase complex (Frank et al., 2008; Kang et al., 2008;

Kaulfürst-Soboll et al., 2011). A strong negative genetic interaction between the KOR1 mutant (rsw2-1) and stt3a was observed, and the double mutant displayed a severe growth phenotype (Kang et al., 2008). The perturbed oligosaccharyltransferase activity of stt3a-2 coincides with the rationale that KOR1 requires the presence of N-linked glycans, 45 but the complexity of the resulting N-linked glycans does not influence KOR1 activity

(Liebminger et al., 2013). Interestingly, the double mutants containing rsw2-1 and one of the mutants necessary for producing complex N-linked glycans ( rsw2-1/cgl1-3 and rws2-

1/hgl1-1) displayed a severe negative genetic interaction, displaying reduced overall growth (Rips et al., 2014; Kang et al., 2008). These results suggest there may be an interaction(s) between proteins with complex N-glycans that assist with KOR1 function or cooperate with KOR1 during cell wall biosynthesis.

b. Glycosylphosphatidylinositol anchors

Glycosylphosphatidylinositol (GPI)-anchors are glycosylated lipids that attach proteins to the outer leaflet of the plasma membrane (Mamode Cassim et al., 2019). They consist of a highly saturated phospholipid chain attached to a core glycan chain consisting (in order from the phospholipid to the peptide) of inositol, glucosamine, and three Man residues, with the first Man occasionally containing a β-1,4 linked galactose or

N-acetyl glucosamine (Figure 1.6). The peptide is attached to the core glycan by phosphoethanolamine at its C-terminus (Yeats et al., 2018). The biosynthesis of GPIs has been heavily studied in mammalian systems and in S. cerevisiae, but is largely accepted that the biosynthesis of GPI anchors in plants is conserved (Cheung et al., 2014). Several

GPI biosynthetic genes have been isolated in Arabidopsis. SETH1, an ortholog of human

PIG-C and fungal Gpi2p, and SETH2 (PIG-A/Gpi3p) are involved in the addition of glucosamine in GPI synthesis. Heterozygous mutants of SETH1 and SETH2 displayed male-specific fertility defects, including poor pollen tube germination and pollen tube growth in seth1 mutants and seth2 pollen. No homozygous lines of seth1 mutants or 46 seth2-1 were isolated (Lalanne et al., 2004), suggesting that this mutation is gametophytically lethal. The addition of the first Man residue in GPI synthesis is facilitated by PEANUT 1 (PNT1) (PIG-M/Gpi14p). Homozygous pnt1 mutants were seedling lethal and had poor male transmission rates. Interestingly, pnt1 embryos had an altered cell wall composition with decreases in crystalline cellulose content and increases in arabinose containing pectins (Gillmor et al., 2005). ABNORMAL POLLEN TUBE

GUIDANCE 1 (APTG1), is a mannosyltransferase adding the third Man to a GPI anchor.

The aptg1 mutant is embryo lethal and as observed with other GPI biosynthetic mutants, male fertility was negatively impacted. Unlike the previously mentioned mutants, aptg1 had pollen tubes that germinated at similar rates as wildtype and had similar pollen tube lengths. The mislocalization of COBL10 at the apical plasma membrane of the pollen tube tip was also observed in the aptg1 mutant. These observations suggest that GPI anchors also influence an anchored protein’s localization specificity in the plasma membrane (Dai et al., 2014). The completed glycan is transferred en bloc by GPI8 (PIG-

K/Gpi8p) to the targeted peptide. A mutation in GPI8, gpi8-1 has reduced GPI- transamidase activity leading to poor root growth, stomatal formation, flowering, and male fertility defects (Bundy et al., 2016). A C-terminal GPI signal peptide is cleaved by the GPI-transamidase between amino acids indicated as ω + 0 and ω + 1 and the GPI is attached at the carboxy group of the C-terminus to form an amide bond. The consensus sequence is not conserved, and it remains challenging to predict how many GPI anchored proteins are encoded in the Arabidopsis genome (Yeats et al., 2018). One proteomic study attempted to elucidate the number of GPI anchored proteins in Arabidopsis.

Enriched membrane fractions from Arabidopsis callus were treated with a 47 phosphatidylinositol-specific phospholipase C to release GPI-anchored proteins. The released GPI-anchored proteins were then subjected to LC-MS/MS revealing a total of

250 predicted GPI-anchored proteins in Arabidopsis (Borner et al., 2003), a fairly large number compared to the 150 in mammals and 50 in yeast (Cheung et al., 2014). A common theme presented for each mutant with compromised GPI biosynthesis is that they are defective in pollen tube growth or female ovule sensing. As mentioned previously, the CSC accessory protein COBRA is GPI-anchored protein involved in proper cellulose deposition (Schindelman et al., 2001), and COBL10, expressed in pollen tubes, is specifically involved in pollen tube elongation (Li et al., 2013). There are other

GPI anchored proteins in pollen, most notably the LORELEI-like 2 and 3 (LLG2/3). Both

LLG2 and LLG3 interact with receptor-like kinase 1-like (RLK) ANUXR

(ANX)/BUDDHA’s PAPER SEAL (BUPS) and collectively are involved in pollen tube integrity. Briefly, pollen tube growth is a complex and dynamic process, involving fine- tuned rigidification of pollen tube cell walls and loosening at the apical region. Failure to do so either results in growth arrest or premature pollen bursting (Bosch et al., 2005; Li and Yang, 2018). These observations were made in anx1 and anx2 mutants displaying immediate pollen tube bursting in vitro and complete growth arrest in vivo (Li and Yang,

2018). A similar in vitro pollen tube burst and in vivo growth arrest phenotype was observed in RNAi knockdown lines of LLG2/3, suggesting that LLG2/3 are coreceptors of ANX/BUPS (Feng et al., 2019). The GPI-anchored LLG1 has been shown to interact with FERONIA (FER), a homolog of ANX/BUPS, in seedling tissues. Loss-of-function mutations in the llg1-2 and fer-4 have a near identical phenotypes. One interesting observation in llg1-2 and fer-4 was the reduced root hair lengths and root hairs that 48 produced cell walls weak enough to cause cytoplasmic discharge or leaks (Li et al.,

2015). Collectively, these data demonstrate one facet of GPI anchors in male fertility and in growth involves cell wall integrity sensing.

c. Glucosyl Ceramides

Glucosyl Ceramides (GlcCers) are part of a class of lipids called sphingolipids.

Sphingolipids are unique class of lipid structures that play major structural roles in plasma membrane and endomembrane morphology as well as important roles in programed cell death, membrane trafficking, plant pathogen interaction, protein anchoring to the plasma membrane, fertility, and developmental processes (Gronnier et al., 2016; Tartaglio et al., 2017; Ternes et al., 2011). Sphingolipids are composed of a fatty acid (FA) group with acyl chain lengths ranging from 16 to 26 carbons, an 18- carbon sphingosine base, commonly referred to as a long chain base (LCB), and a structurally variable headgroup. Arabidopsis uses the ceramide synthase enzymes

LONGEVITY ASSURANCE GENE ONE HOMOLOG 1 (LOH1), LOH2, and LOH3 to synthesize ceramides utilizing a single FA and LCB as substrates (Figure 1.7A).

Overexpression lines of LOH2 accumulated sphingolipids with short chain FAs (16 carbons), causing impaired growth and induced programmed cell death. In contrast, overexpression of LOH1 or LOH3 resulted in accumulation of sphingolipids with very long chain fatty acids (VLCFA) (18+ carbons) producing larger plants. Together these results demonstrate the importance of acyl chain length in sphingolipids (Luttgeharm et al., 2015a; Ternes et al., 2011). The ceramide synthase biosynthetic reaction can be inhibited by the mycotoxin Fumonisin B1, causing severe growth inhibition and 49

Figure 1.7: Structures of GlcCers and GIPCs found in plant cells. (A) Structures of lipid moiety of FA and Long Chain Base (LCB) and the synthesis of the ceramide component for GlcCers and GIPCs. (B) Structures demonstrating the monosaccharide composition in GlcCers and GIPCs. Known ceramide synthases and GTs associated with the addition of monosaccharides in Arabidopsis are highlighted in red with an arrow pointing to the specific glycosidic bond. In all bonds labeled, assume attachment at 1 position unless stated (β2 = β-(1,2)-linked).

50 programed cell death (Luttgeharm et al., 2016), Ceramide FA moieties can be hydroxylated to form hydroxy ceramides (hCers) by FATTY ACID 2-HYDROXYLASE

1 (FAH1) and FAH2. Both FAH1 and FAH2 are non-redundant and have preferential activity based on ceramide FA lengths. FAH1 mainly 2-hydroxylated VLCFAs, whereas

FAH2 selectively 2-hydroxylated short chain FAs (Nagano et al., 2012). Ceramides and hCers comprise only 1-2% of sphingolipids in plants, suggesting that ceramides and hCers are primarily intermediates to more complex sphingolipids (Michaelson et al.,

2016). Ceramides and hCers are substrates for Glucosylceramide Synthase (GCS), which catalyzes the addition of glucose at the C1 position of the LCB using UDP-Glucose as a donor substrate (Figure 1.7B). The gcs-1 mutant was viable at the seedlings stage with severe growth reduction but failed to develop beyond the seedlings stage. The gcs-1 mutant, similar to GPI biosynthesis defects, exhibited a severe reduction in male fertility, caused by poor pollen tube germination and arrested growth (Msanne et al., 2015).

Additionally, GCS activity can be inhibited by N-[2-hydroxy-1-(4-morpholinymethyl)-2- phenyl ethyl]-decanamide (PDMP). PDMP treatment disrupted vesicular trafficking and caused abnormal endomembrane morphologies (Melser et al., 2010).

Currently, there is no direct evidence that GlcCers are implicated in cell wall biosynthesis, but a relationship between these processes may be inferred based on the gcs-1 mutant having poor fertility similar to all GPI anchor biosynthesis mutants (Msanne et al., 2015; Dai et al., 2014; Lalanne et al., 2004; Gillmor et al., 2005; Bundy et al.,

2016). GlcCers may also play a role in either membrane organization to stabilize GPI anchored proteins and plasma membrane associated proteins, or organizational roles in endomembranes to help provide a stable working platform for GPI biosynthesis. An 51 important aspect of plant membranes to consider here are lipid rafts otherwise known as

Detergent Resistant Membranes (DRMs). The common name of DRMs is derived from their ability to produce insoluble membranes after treatment with a mild detergent, such as Triton X-100. Often in DRMs, there is a high concentration of sphingolipids and proteins compared to non-DRM membrane fractions. This scenario was shown to be the case in Arabidopsis callus membrane extracts (Borner et al., 2005). This same study utilized a modified bacterial phosphinothricin acetyl transferase (PAT) fused with a GPI- anchoring signal to generate a PAT-AtGPI4 to demonstrate it can be GPI-anchored in

Arabidopsis calli and membrane extracts with increasing Triton X-100 concentrations remained in DRM fractions (Borner et al., 2005). Further analysis of DRMs revealed a greater concentration of protein, phytosterols, and sphingolipids compared to solubilized membranes (Borner et al., 2005). Together, these findings suggest that GlcCers play a critical role in plasma membrane organization necessary for GPI-anchored protein stability and organization of membrane bound proteins with a subset involved with cell wall integrity sensing. A small additional piece of evidence that might further indirectly support GlcCer involvement in organization of membrane proteins is the high concentrations of GlcCers and Glycoinositol phosphoceramides (GPICs) in pollen (8- fold) compared to leaves. Additionally, sphingolipid biosynthetic genes, such as GCS, were highly expressed in pollen comparted to whole seedlings (Luttgeharm et al., 2015b).

This simply further supports the role of glycosylated sphingolipids like GlcCers and

GIPCs in male fertility and or pollen tube integrity sensing.

d. Glycoinositol phosphoceramides 52

Glycoinositol phosphoceramides (GIPCs) are a distinct classification of glycosylated sphingolipids unique to plants that represent the largest proportion (64%) of sphingolipids and 25% of total lipids in the plasma membrane (Markham et al., 2006). In plants, GIPCs are implicated in several biological processes, including cell wall anchoring, cell surface recognition, signaling precursors, and immune responses

(Gronnier et al., 2016). Like GlcCers, GIPCs contain a ceramide moiety, followed by a phospho-myo-inositol, and a core glycan group consisting of D-Glucuronic acid (GlcA),

Man, and occasionally other pentose monosaccharides (Figure 1.7B). The addition of phospho-myo-inositol is facilitated by IPC synthase, resulting in the synthesis of inositol phosphoceramides (IPC) (Wang et al., 2008). IPCs are then processed further by the addition of GlcA by INOSITOL PHOSPHORYLCERAMIDE

GLUCORONOSYLTRANSFERASE 1 (IPUT1). Attempts to generate null mutations of

IPUT1 (iput1-1 and iput1-2) resulted in no viable pollen (Tartaglio et al., 2017; Rennie et al., 2014), suggesting that IPUT1 is essential for male fertility and that the resulting mutants are gametophytically lethal. Interestingly, the monocation-induced [Ca2+] increases 1 (moca1) mutant carries a four amino acid deletion in the sixth transmembrane region of IPUT1. Unlike other attempted IPUT mutations, moca1 was viable and demonstrated hypersensitivity to salt stress (Jiang et al., 2019). The unique aspect of moca1 was the dampened calcium influx after initial exposure to high salt conditions compared to wildtype. These observations suggest that GIPCs are actively involved in salt-sensing and calcium influx associated with adaptation responses to salt stress (Jiang et al., 2019). Following the addition of GlcA, Man is added by GIPC

MANNOSYLTRANSFERASE 1 (GMT1) (Fang et al., 2016). Loss-of-function 53

mutations in GMT1 (gmt1-1) causes severe developmental defects in seedlings, limiting

biochemical experiments to be conducted cell cultures.

One of the first pieces of data that demonstrated a role in cellulose biosynthesis was

in the characterization of GOLGI LOCALIZED GDP-MANNOSE TRANSPORTER 1

(GONST1) (Mortimer et al., 2013). GONST1 transports GDP-Man into the Golgi for

GIPC mannosylation. Plants homozygous for gonst1-1 had poor growth and spontaneous

lesions in leaves. Analysis of sphingolipid glycosylation profiles revealed a severe

reduction in complex GIPC head group profiles. The primary monosaccharide deficiency

that contributed to loss of GIPC head group complexity was Man, further supporting the

GDP-Man translocase activity is directly necessary for GIPC head group biosynthesis

(Mortimer et al., 2013). GDP-Man is directly used by GMT1 to add the first Man to

GIPCs in the Golgi. Interestingly, this study revealed that gonst1 mutants exhibit a severe

reduction in crystalline cellulose content, up to 50% in stems (Fang et al., 2016). The

relationship between GIPC head group composition and cellulose biosynthesis is unclear,

but there may be a reasonable association between GIPCs in the plasma membrane and

the membrane bound CSC.

VIII. Use of small molecules, monosaccharide analogs, and monosaccharide mimicking

small molecules

The use of small molecules that interfere with the function or the activity of enzymes

have served as useful tools to probe biological functions, control of pest, diagnostic tools

in healthcare, and as pharmaceuticals (Aktar et al., 2009; MacKinnon and Taunton, 2009;

Kim et al., 2016; Nash et al., 2011). Several small molecules have been used in plants to 54 facilitate the study of numerous biological processes. For example, Oryzalin destabilizes and depolymerizes microtubules in plants. This molecule has assisted the study of numerous microtubule-associated processes, including determining that CSI1 associated the CSC to cortical microtubules, where the trajectories of CSCs in the pom2-4 (csi1) background and Oryzalin treated wildtype, were not linear as in the WT control

(Bringmann et al., 2012). In etiolated hypocotyls, CSCs containing CESA5 were slower compared to CSCs containing CESA6. This was due to phosphorylation events in N- terminus of CESA5 causing a rigid association of cortical microtubules, with velocities that can be restored when Oryzalin is applied (Bischoff et al., 2011).

Small molecules are also used as herbicides in residential areas and in agriculture.

There is a myriad of herbicides available and that target nearly every major biological process in plants. For example, herbicides targeting pigment biosynthesis (Pentoxazone),

NADPH formation (Paraquat), amino acid biosynthesis (Glyphosate, flumetsulam), lipid biosynthesis (metazachlor, sethaxydim), and cell wall biosynthesis inhibitors

(Wakabayashi and Böger, 2004a, 2004b; Smyth et al., 2013). Here, the focus will be in common cellulose biosynthesis inhibitors (Figure 1.8).

a. Common cell wall biosynthesis inhibitors

The use of cellulose biosynthesis inhibitors (CBIs) have become increasingly important in agriculture due to increased rates of herbicide resistant weeds, which account as the primary biotic stress in crops (Douglas and Gaines, 2014; Fickett et al.,

2013; Küpper et al., 2018). CBIs are a structurally diverse group of small molecules that selectively inhibit the production of cellulose. Effects of CBIs include loss of anisotropic 55

Figure 1.8: Structures of cellulose biosynthesis inhibitors and RG-II biosynthesis inhibitor. (1 – 6) Structures of cellulose biosynthesis inhibitors the cause CSC compartmentalization into SmaCCs/MASCs (1) Isoxaben, (2) Thaxtonin A, (3) Acetobixan, (4) Quinoxyphen (5) CGA 325’615, and (6) Triazofenamide. (7) Structure of Dichlorobenzylnitrile (DCB), halts CSC motility. (8 – 9) Structure of (8) KDO, monosaccharide found in RG-II and (9) 2β-deoxy KDO, a known inhibitor of RG-II biosynthesis.

56 growth, reduced growth, and cell death (Sabba and Vaughn, 1999). Most known CBIs can be categorized into two primary groups. The first group of compounds such as

Isoxaben, Thaxtonin A, Acetobixan, Quinoxyphen, CGA 325’615, and Triazofenamide

(Figure 1.4 [1 – 6]) inhibit cellulose biosynthesis by sequestering CSCs into

SmaCCs/MASCs. The second group, such as Dichlorobenzolnitrile (DCB), arrest CSC motility at the plasma membrane (Figure 1.4 [7]) (Tateno et al., 2016; DeBolt et al.,

2007; Xia et al., 2014). There are also a few other compounds that fit in either group 1 or group 2 categories, but their inhibitory mechanisms are unique. The CESA Trafficking

Inhibiter (CESTRIN) is likely a group 2 CBI as it reduces CSC velocities in hypocotyls and causes dissociation of CSI1 from the plasma membrane (Worden et al., 2015). A second CBI, Indaziflam is similar to DCB because it reduces CSC velocities but also decreases colocalization between CSCs and cortical microtubules. This observation suggests that Indaziflam interferes with CSC and cortical microtubule association mediated by CSI1 (Brabham et al., 2014). Arabidopsis plants treated with CBIs commonly exhibit several changes in cellular homeostasis, triggering a series of transcriptional changes including increased transcripts associated with wounding response, DAMP elicitor response, and ectopic lignin deposition (Caño-Delgado et al.,

2003; Duval and Beaudoin, 2009).

Multiple Arabidopsis mutants have been isolated that are resistant to several

CBIs. One of the earliest published resistant mutants, isoxaben resistant A1 (ixr A1) and ixr A2 were 90 to 300 times more resistant to Isoxaben compared to wildtype (Heim et al., 1989). Later, both mutants were renamed ixr1-1 and ixr1-2 respectively as both had genetic lesions in CESA3 (Scheible et al., 2001). A semi-dominant mutant, ixr2-1, has 57 also been isolated corresponding to a point mutation is CESA6 (Desprez et al., 2002).

Four additional ixr1 alleles have been found in the class specific region and in transmembrane regions of various CESA proteins (Shim et al., 2018). Additionally, this same study investigated another CBI, Flupoxam and found several flupoxam resistant

(fxr) mutants in CESA3 (fxr1-1, fxr1-2, and fxr1-3) and CESA1 (fxr2-1, fxr2-2, and fxr2-

3). Interestingly, all of the fxr mutants had missense mutations in loci corresponding to transmembrane regions. Collectively, Isoxaben and Flupoxam resistant mutants also produced cellulose with reduced crystallinity compared to wildtype, suggesting that these mutants would also convey a decrease in recalcitrance to enzymatic cellulose degradation

(Shim et al., 2018). A DCB resistant mutant has been isolated in Arabidopsis labeled as

DH75, which had an 800-fold increase in resistance (Heim et al., 1998). Unfortunately, there has not been follow up in the DH75 mutant. Recently, a novel CBI labeled as C17 has been described and was shown to inhibit growth of Isoxaben resistant mutants ixr1-1, ixr1-2, and ixr2-1. This suggest C17 inhibitory effect mechanism is different from

Isoxaben (Hu et al., 2019). This same study was also able to generate a mutant resistant to C17 in CESA3S983C using CRISPR-Cas9 based editor technology, purposing to use as a positive transformant selection marker (Komor et al., 2017; Hu et al., 2019).

b. Monosaccharide analogs as diagnostic tools and inhibitors

Monosaccharides analogs have served as useful tools to inhibit, chemically label, and metabolically mark cellular glycans. There is a multitude of monosaccharide analogs that inhibit glycosyltransferases described in mammalian systems, fungi, and plants. In mammalian systems several monosaccharide analogs, many of which are fluorinated, 58 have been described to inhibit N-linked glycosylation events, such as 2-deoxy-2-fluoro-

L-fucose (2F-Fuc), 3-deoxy-3-fluoro-axial-D-N-Acetylneuraminic acid, and 4-deoxy-4- fluoro-D-N-acetyl-glucosamine (Rillahan et al., 2013; Barthel et al., 2011). Other monosaccharide analogs have been utilized in tumor suppression in liver (HepG2) with

2F-Fuc and in Hela cells with 2F-Gluc. Both compounds inhibited N-linked glycosylated protein fucosylation and glucose uptake respectively (Zhou et al., 2017; Niccoli et al.,

2017). In fungi, two glucose analogs have been utilized in inhibiting fugal growth. In budding yeast (S. cerevisiae), 2-deoxy-D-glucose (2D-Glc) inhibited growth by activating glucose repression (McCartney et al., 2014). Additionally, 2D-Glc and 2- deoxy-2-fluoro-D-glucose (2F-Glc) have been shown to inhibit GPI biosynthesis in malaria parasite protozoan Plasmodium falciparum (De Macedo et al., 2001). In plants, relatively few monosaccharide analogs have been used as inhibitors of plant growth. One analog, 2β-deoxy-KDO, was a potent inhibitor of RG-II biosynthesis and can be partially rescued by exogenous boric acid. This result implies that the inhibition of KDO transferase in RG-II biosynthesis reduced the efficiency of the boron cross-bridge formation (Smyth et al., 2013). 2F-Fuc has been utilized in rice cell cultures to inhibit fucosylation events of N-linked glycans in recombinant human acid alpha-glucosidase

(rhGAA). The reduction of N-liked glycan fucosylation is critical for the production of therapeutic glycoproteins to reduce immunogenicity-inducing potential (Kim et al.,

2020).

Monosaccharide analogs have also been utilized as probes and diagnostic tools in metabolomics, in crystallo enzymology, and in glycan labeling. One example of monosaccharides in metabolomic labeling is in plant glucose metabolism. Arabidopsis 59 leaves were incubated with 2F-Glc for 4 hours and leaf extract metabolites were quantified. Several fluorine metabolites were identified, including 2-deoxy-2-fluoro-D- gluconic acid, 2F-Glc-6-phosphate, 2-deoxy-2-fluoro-maltose, and UDP-2F-Glc. This observation suggests that 2F-Glc is metabolized into starch and potentially other glycan synthesis utilizing UDP-Glc (Fatangare et al., 2015; Fatangare and Svatoš, 2016). The translocation of a growing glucan and the glycosyltransferase mechanism in bacterial cellulose synthase was elucidated in the crystal structure of BcsA-BcsB. Here 2F-Glc was used to capture the confirmation of catalytic site of BcsA in crystallo (Morgan et al.,

2016).

An important rate-limiting enzyme in cytosolic deoxyribonucleoside salvage pathway in cancer cells is Deoxycytidine kinase, an important target for therapeutics and in positron emission tomography (PET) imaging for detecting cancer. A monosaccharide analog, 2-chloro-2-deoxy-2-[18F]fluoro-9-β-D-arabinofuranosyl-adenine ([18F]CFA), displayed favorable biodistribution in humans and demonstrated favorable accumulation in leukemia cells, detectable in PET imaging (Kim et al., 2016). An interesting imaging technique utilizing monosaccharide analogs is through click-chemistry compatible analogs. Commonly, click chemistry describes the cycloaddition reaction between a metabolically incorporated alkyne analog reacting with an azide containing fluorophore using a copper catalyst (Zhang and Zhang, 2013). Several click-chemistry compatible monosaccharide analogs have been used to label several components in plant polysaccharides. Fucose alkyne was shown to be incorporated and label RG-I polysaccharides in roots (Anderson et al., 2012). One additional analog, 8-azido-8-deoxy-

KDO, was able to incorporate into pectic RG-II in roots (Dumont et al., 2016). Finally, a 60

6-alkynyl-6-deoxy-glucose accumulated in elongating root hairs and were incorporated into callose (McClosky et al., 2016).

c. Iminosugars a class of monosaccharide mimicking compounds

Iminosugars are small molecules that commonly resemble naturally occurring monosaccharides with the oxygen in the pyranose ring substituted with a nitrogen group.

One of the simplest iminosugars available is 1-deoxynojirimycin (DNJ) and its parent compound nojirimycin (Inouye et al., 1966), both of which are naturally occurring.

Sources of DNJ come from Mulberry trees Morus bombycis, M. nigra and from microorganisms such as Bacillus polymyxa and Streptomyces lavendulae (Hughes and

Rudge, 1994). DNJ was first discovered by groups interested in the antidiabetic activity of mulberries, and it became clear that DNJ was a useful inhibitor for a family of enzymes involved in glucosidase inhibition. Soon after, a variety of DNJ analogs were developed including 5 membered rings such as Pyrrolidine, bicyclic compounds like

Casuarin, and N-alkylated compounds like N-butyl-1-deoxynojirimycin (NB-DNJ) and

N-nonyl-1-deoxynojirimycin (NN-DNJ) (Nash et al., 2011).

Alkylated analogs such as NB-DNJ and NN-DNJ have been used as treatments for

Gaucher’s disease, an autosomal recessive glycolipid storage disorder leading to the accumulation of glucosylceramides in macrophages causing dysfunction in multiple organ systems. The disease is caused by mutations in β-glucocerebrosidase gene 1q21

(Elstein et al., 2001). Interestingly, both compounds have been used as treatments for

Gaucher’s disease for glycosidase activity but as either an inhibitor of glucosylceramides or as a chemical chaperone. NB-DNJ is the first oral agent for the treatment of Gaucher’s 61 disease and its primary action is reducing the glucosylceramides by targeting its biosynthesis (Butters et al., 2005; Platt et al., 1994). The second treatment strategy, using a chemical chaperone, involves assisting a partially folded protein to properly fold by using a compound that binds to the active site allowing intramolecular forces to complete proper folding. Both NB-DNJ and NN-DNJ have been utilized as chemical chaperones

(Sawkar et al., 2002; Brumshtein et al., 2007).

IX. Screening of monosaccharide analogs as a semi-rational inhibitors of plant cell

wall biosynthesis.

Although much progress has been made toward understanding of the structure, biosynthesis, and regulation of plant cell wall, many open questions still remain. Many of the GTs involved in wall polysaccharide biosynthesis and many of their functions in other tissues are relatively obscure. Part of the challenge in elucidating the identity of these GTs and other proteins associated with cell wall biosynthesis through forward genetics is that many are gametophytic-lethal (Ochoa-Villarreal et al., 2012b), which often precludes their study. In some studies, such as the rsw2-1 (KOR1 mutant), lethality could be circumvented due to the temperature sensitive nature of rsw2-1 (Lane et al.,

2001). Due to the difficulty to generate and isolate temperature sensitive (they are exceedingly rare) this strategy is a limited approach for isolating cell wall biosynthesis associated proteins (Tan et al., 2009). The use of chemical inhibitors of cell wall biosynthesis to probe for function has almost exclusively revolved around cellulose with few exceptions (Figure 1.4) (Tateno et al., 2016; Smyth et al., 2013; Villalobos et al.,

2015; Dumont et al., 2015). Additionally, the structures of cellulose biosynthesis 62 inhibitors are structurally diverse and do not indicate possible direct mechanisms of inhibition (Figure 1.4) (Tateno et al., 2016).

In order to probe cell wall function to lead to elucidating GTs associated with cell wall biosynthesis, we purchased a small library of monosaccharide analogs containing deoxy, azido, and fluoro substitions. We hypothesized that deoxy-substituted monosaccharide analogs could be incorporated into a growing glucan chain, but the absence of a hydroxyl group could prevent the incorporation of a proceeding monosaccharide (Figure 1.9A and 1.9B). The analogs containing an alternative nucleophile (Azido or fluoro groups) would hypothetically destabilize the oxocarbenium ion-like transition state causing an unfavorable reaction (Figure 1.9C and 1.9D) (Lairson et al., 2008).

Here, Arabidopsis seed were germinated and grown for 7 days in MS media (see materials and methods in chapter 2) supplemented with each tested monosaccharide analog including a solvent control (0.1% DMSO) and a no treatment control. Root lengths from seedlings in each treatment were quantified (Figure 1.10) (Table 1.1).

Overall, four monosaccharide analogs;2F-Gluc, 2-deoxy-2-fluoro-D-mannose (2F-

Man),2F-Fuc, and 1,3,4-triacetal-2-deoxy-2-fluoro-L-fucose (Ac3 2F-Fuc) were found to severely inhibit root growth in Arabidopsis (Figure 1.10) (Table 1.1). The first set of monosaccharide analogs to be characterized in this dissertation, 2F-Fuc and Ac3 2F-Fuc, is based on published findings detailed in (Villalobos et al., 2015) also covered in chapter

2. 63

64

Figure 1.9: Hypothetical SN2 mechanisms of deoxy and fluor/Azido monosaccharide analogs in non-retaining GTs. (A – B) In the scenario of deoxy monosaccharides inhibiting possible extending glycosyltransferases is through incorporation. (A) The native reaction uses the elongating glycan as the acceptor (Green) is deportanated by the nucleophile in the active site (Red) where the deprotonated hydroxyl group targets the alpha-carbon of the donor (Black). This leads to a final product of the new monosaccharide integrated into the glycan. (B) In this scenario, the deoxy monosaccharide is incorporated into the acceptor in the previous reaction and in the next reaction the hydroxyl group necessary to the enzymatic reaction is missing thus, acting a terminal sugar of an incomplete polysaccharide. (C – D) Fluoro/Azido monosaccharides hypothetically inhibits GTs by destabilizing the transition state. (C) In this reaction the same exchange of electrons occurs as in (A) forming a Oxocarbenium ion-like transition state followed by the final product. (D) When a fluoro monosaccharide acts as the donor, the same flow of electrons occurs but, the Oxocarbenium ion-like transition state is unstable due to the beta-carbon’s increased partial charge from the C – F bond.

65

Figure 1.10: Monosaccharide analog screen on Arabidopsis thaliana. Arabidopsis Col-0 seed were germinated and grown in MS media with indicated treatments for 7 days. Root lengths were quantified and represented as a box and whisker plot displaying minimum and minimum, maximum, mean, first, and second interquartile range n = 40 – 267.

66

Table 1.1: Monosaccharide analog screen on Arabidopsis.

Compound Mean SEM Significant n Chapter Ref.

No treatment 25.71 ± 0.3262 n/a 267 n/a

0.1% DMSO 18.61 ± 0.3324 **** 144 n/a

1 mM 3D-Gluc 28.83 ± 0.3214 **** 85 n/a

100 μM 4D-Gluc 25.59 ± 0.3352 ns 85 n/a

100 μM 5T-Gluc 28.42 ± 0.3677 **** 92 n/a

1 mM 2Az-Gluc 28.99 ± 0.4434 **** 83 n/a

100 μM 6Az-Gluc 33.3 ± 0.362 **** 85 n/a

100 μM 2F-Gluc 3.075 ± 0.09313 **** 105 3

100 μM 3F-Gluc 30.08 ± 0.5706 **** 40 n/a

1 mM 4F-Gluc 13.61 ± 0.3959 **** 92 n/a

100 μM 6F-Gluc 31.21 ± 0.3134 **** 92 n/a

1 mM 2D-Gal 13.51 ± 0.4118 **** 90 n/a

100 μM 6Az-Gal 15.66 ± 0.3741 **** 123 n/a

100 μM 2F-Gal 17.91 ± 0.5313 **** 81 n/a

100 μM 3F-Gal 17.64 ± 0.4731 **** 145 n/a

100 μM 4F-Gal 23.92 ± 0.5283 *** 152 n/a

100 μM 2F-Man 3.072 ± 0.0822 **** 151 3 & 4

100 μM 3F-Man 14.25 ± 0.5706 **** 78 n/a

100 μM 4D-Fuc 18.38 ± 0.3322 **** 168 2

100 μM 2F-Fuc 1.491 ± 0.03988 **** 116 2

100 μM Ac3 2F-Fuc 1.546 ± 0.03848 **** 117 2

1 mM 5D-Ara 14.16 ± 0.3026 **** 143 n/a

1 mM D-Rha 19.6 ± 0.2996 **** 136 n/a

Mean and SEM values are reported as mm.

Dunnett's mutliple comparisons (***, p < 0.001; ****, p < 0.0001)

67

In this dissertation novel findings include, 2F-Fuc and Ac3 2F-Fuc inhibit RG-II fucosylation, Chapters 3 and 4 discusses how 2F-Man inhibits growth through the glucose repression pathway in Arabidopsis and in yeast. These results prompted us to also investigate iminosugars in Chapter 5 and discovered N-dodecyldeoxynojirimycin

(ND-DNJ) inhibits glycosylated sphingolipid biosynthesis resulting in reduced crystalline cellulose deposition through reducing CSC velocities. Finally, in Chapter 6 we discovered a novel response to cellulose biosynthesis inhibition with CBIs.

68

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Chapter 2: 2-Deoxy-2-Fluoro-L-Fucose is a metabolically incorporated inhibitor of plant cell wall polysaccharide fucosylation.

i. Introduction

L-Fucose (6-deoxy-L-galactose) is a common monosaccharide found in various plant cell wall components, including xyloglucan, Rhamnogalacturonan-I,

Rhamnogalacturonan-II, and Arabinogalactan proteins. (Vanzin et al., 2002; Mohnen,

2008; Atmodjo et al., 2013; Anderson, 2015; Tryfona et al., 2014). Commonly, fucose is incorporated into the cell wall polysaccharides as a terminal monosaccharide of branching polysaccharide ends. In xyloglucan, the backbone structure is a homopolymer of β-1→4 linked glucose molecules. The glucose chain is decorated with α-1→6-linked xylose residues on three of every four glucose residues (oligomers). In some xyloglucan oligomers, the branching chain extends with a β-1→2 linked galactose and terminated with an α 1→2 linked fucose (Pauly and Keegstra, 2016). Rhamnogalacturonan-I (RG-I) is a complex polymer consisting of a backbone structure of repeating L-Rhamnose-

(1→4)-α-D-Galacturonic acid-(1→2)-α-L-Rhamnose. L-fucose is found at the terminal end of arabinan or galactan side chains (Nakamura et al., 2001). Rhamnogalacturonan-II

(RG-II) is one of the few polysaccharides where L-fucose is not exclusively relegated as a terminal monosaccharide in the branching chain. Rhamnogalacturonan-II backbone consist of a linear α-1→4 linked galacturonic acid with 4 possible side chains referred to as side chains A – D. Fucose can be found on side chain B at a terminal end linked α-

1→2 to D-galactose. Uniquely, side chain A also contains an L-fucose residue not located at the terminal end. Instead it is the third monosaccharide in the growing chain

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(Mohnen, 2008). This is the one example where L-fucose may play a critical role in a cell wall polymer.

L-fucose is not commonly found freely in the cytosol. Generally, all sources of de novo L-fucose occur in the form of its nucleotide sugar GDP-L-fucose. The de novo biosynthetic pathway begins with GDP-D-mannose as a substrate for GDP-Mannose-4-6- dehydrotase (GMD) yielding GDP-4-keto-6-deoxy-D-mannose (Bonin et al., 2003). This intermediate is the substrate to GDP-4-keto-6-deoxy-D-mannose-3,5-epimerase-4- reductase1 (GER1) to yield the final product GDP-L-fucose (Bonin et al., 1997; Bonin and Reiter, 2000). A loss-of-function mutation in the Arabidopsis GMD2, also referred to as murus1 (mur1), exhibits a 98% reduction in cell wall-associated L-fucose, and the majority of L-fucose is replaced with L-galactose in this mutant (Bonin et al., 1997;

Reuhs et al., 2004). Although free L-fucose is relatively uncommon in the plant cytosol, the primary source of free L-fucose is primarily from cell wall degradation from remodeling under the action of FUCOSIDASE1 (FUC1) and ALPHA FUCOSIDASE1

(FXG1) (de la Torre et al., 2002; Léonard et al., 2008). Free L-fucose is converted to

GDP-L-fucose through two subsequent enzymatic reactions. First, L-fucose is phosphorylated yielding L-fucose-1-P followed by coupling to GDP to yield GDP-L- fucose. Both reactions are carried out by the bifunctional enzyme FUCOSE KINASE

GTP PYROPHOSPHORYLASE (FKGP). The Arabidopsis loss-of-function mutations fkgp-1 and fkgp-2 accumulate up to 100-fold more free L-fucose compared to wildtype plants (Kotake et al., 2008).

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Interestingly, there is no evidence to suggest that L-fucose is converted into any other metabolic product, suggesting that this monosaccharide can be considered a “dead end metabolite” (Bar-Peled and O´Neill, 2011). This observation suggests that targeting fucosyltransferases using L-fucose analogs may yield a methodology to probe the function of cell wall-associated fucosylation events. Currently, the fucosyltransferases

(FUTs) responsible for several cell wall fucosylation events have been characterized.

Arabidopsis FUCOSYLTRANSFERASE1 (FUT1) fucosylates D-Galactose residues in the xyloglucan side chain, as demonstrated by the observation that null mutants of FUT1

(mur2) completely lack xyloglucan fucosylation (Vanzin et al., 2002). Arabinogalactan protein fucosylation is carried out by two fucosyltransferases, FUT4 and FUT6 (Liang et al., 2013). The fucosyltransferases responsible for attaching fucose to RG-I and RG-II have not been identified, however, there are multiple FUT genes that may encode a fucosyltransferase that uses a growing RG-I or RG-II polymer as a substrate (FUT2:

At2g03210, FUT3: At1g74420, FUT5: At2g15370, FUT7: At1g14070, FUT8:

At1g14100, FUT9: At1g14110, and FUT10: At2g15350).

Protein N-linked glycosylation is a common co- and post-translational modification of many proteins destined to the plasma membrane. L-fucose is also found in these N- linked glycans and participates in salt tolerance (Strasser, 2016; Kang et al., 2008). These

L-fucose residues are added to a maturing N-glycan by three FUT proteins. FUT11 and

FUT12 catalyze the addition of an α-1→ 3 fucose linkage to the first core N-Acetyl

Glucosamine of the N-linked glycan (Yoo et al., 2015; Both et al., 2011; Kaulfürst-Soboll et al., 2011). FUT13 catalyzes the addition of α-1→ 4 fucose linkage at the terminal ends of N-linked glycans (Rips et al., 2017; Wilson et al., 2001). Interestingly, genetic

117 knockouts of fut11, fut12, fut13, double, and triple knockouts do not display a phenotype grown under normal conditions (Kang et al., 2008).

Previously, fluorinated monosaccharides have been used as monosaccharide- selective inhibitors of glycosyltransferases in mammalian systems (De Macedo et al.,

2001; Kim et al., 2016; Barthel et al., 2011; Zhou et al., 2017). Furthermore, fluorinated

L-fucose analogs have been used as both a treatment to suppress liver cancer HepG2 and reduced N-link glycan fucosylation (Zhou et al., 2017; Rillahan et al., 2012). In this chapter, we report that the fluorinated fucose analog 2-deoxy-2-fluoro-L-fucose (2F-Fuc) is an inhibitor of Arabidopsis growth, resulting from reduced Rhamnogalacturonan-II fucosylation. Additionally, 2F-Fuc must be metabolically converted to GDP-2F-Fuc through the action of FKGP to serve as an inhibitor of cellular glycosyltransferases

(Villalobos et al., 2015).

ii. Materials and Methods

General plant growth and monosaccharide analogs screen: Arabidopsis thaliana Col-0 seeds were sterilized in 30 % [v/v] sodium hypochlorite, 0.1 % [w/v] sodium dodecyl sulfate for 20 minutes at 25 °C. Seeds were washed five times in sterile water and incubated at 4 °C for 48 hours before plating.

Seeds were germinated on MS media (1/2X Murashige and Skoog salts, 10 mM MES-

KOH pH 5.7, 1% [w/v] sucrose, and 1% [w/v] phytoagar) and grown vertically for 7 days under long day conditions (16-hour light/ 8-hour dark) at 22 °C. Arabidopsis mutants harboring T-DNA insertions in xyloglucan fucosyltransferase (FUT1), N-linked glycan fucosyltransferases (FUT11, 12, and 13), and arabinogalactan protein fucosyltransferases

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(FUT4 and FUT6) were previously described elsewhere (Perrin et al., 1999; Liang et al.,

2013; Strasser et al., 2004; Anderson et al., 2012; Vanzin et al., 2002). Monosaccharide analogs were purchased from Carbosynth (Berkshire, UK). Monosaccharide analogs were diluted in DMSO and included in growth media at the indicated concentrations. Plants treated with 0.1% [v/v] DMSO served as negative controls. After 7 days of growth, seedlings were straightened on the treatment plate and scanned with a flatbed scanner.

Root lengths were quantified using ImageJ (imagej.nih.gov/ij).

For short-term treatment experiments, Col-0 seed were sterilized, stratified, and plated on MS medium as described above. These seedlings were grown under long day conditions at 22 ℃ for 4 days. Seedling were then transferred to fresh plates with or without 100 µM 2F-Fuc and were incubated for an additional 5 days under the previously described growth conditions. For the first three days, root lengths were quantified every

12 hours using ImageJ. Five days after transfer, the root tips of the representative seedlings were imaged with Leica EZ4HD video dissecting scope at 35X magnification.

FKGP knockout isolation:

The Arabidopsis FKGP gene was queried against the Salk Institute T-DNA insertional mutant database (Alonso et al., 2003), and several potential T-DNA lines were identified. These seed populations were propagated on MS media as described above and transplanted to soil after 14 days. Transplanted seedlings were maintained in a growth chamber at 22 ℃ under long day conditions (16-hour light/ 8-hour dark) until maturity.

Genomic DNA isolation for PCR genotyping was performed essentially the same as previously describe (Edwards et al., 1991). One leaf from each plant was removed and

119 homogenized in a 1.5 mL Eppendorf tube with a Teflon pestle. Four hundred microliters of Edwards buffer (200 mM Tris-HCl, pH 7.5, 250 mM NaCl, 25 mM EDTA, 0.5% [w/v]

SDS) was added to the homogenate, and the samples were incubated at 25 ℃ for 1 hr.

Samples were centrifuged at 12,000 x g for 10 min, and 300 µL of supernatant was transferred to a new Eppendorf tube containing equal volume of 2-propanol. Samples were vortexed and centrifuged at 12,000 x g for 5 min. The supernatant was removed and

750 µL of 70% [v/v] ethanol was added. Samples were centrifuged for 3 min at 12,000 x g, the supernatant was removed, and the pellet was air dried for 20 min. The dried pellet was resuspended in 100 µL of sterile water.

Each genomic DNA samples was analyzed by genotyping PCR using locus specific primers see Table 2.1, and ExTaq polymerase (Takara Biol, Mountain View,

CA). Reactions were cycled under the following conditions: 95 ℃ initial denaturation for

5 min, 35 cycles of 95 ℃ for 30 sec, 52 ℃ for 30 sec, 72 ℃ for 1.5 min, final extension at 72 ℃ for 7 min. The resulting PCR products were separated on 1% [w/v] agarose gels and documented with a Bio-Rad Gel Doc XR+ Image analysis workstation.

Alditol acetate monosaccharide analysis:

Five-day-old Arabidopsis hypocotyls were placed in a pre-weighed 2 mL screw cap microcentrifuge tube and incubated with 1.5 mL of 70% [v/v] ethanol for 6 hours at

25 ℃ on a nutating shaker. The samples were then centrifuged at 3,000 x g for 10 minutes, and the supernatant was removed. An additional 1.5 mL of 70% ethanol was added, and this wash step was repeated. The seedlings were then washed with 1.5 mL of

1:1 [v/v] chloroform: methanol for 6 hours at 25 ℃. The chloroform methanol wash was

120 removed, and samples were dried under at 50 ℃ heating block. Three steel balls (2 mm diameter) were added to the dried plant material and homogenized in a Retsch MM301 ball mill for 2.5 minutes at 25 Hz. The steel balls were removed and the resulting alcohol insoluble residue (AIR) was weighed.

Five to ten mg of dry plant material was subjected to weak acid hydrolysis by adding 250 µL of 2 M trifluoro acetic acid (TFA) and 100 µg of D-myo-Inositol as an internal standard (MP Biochemicals). Samples were incubated at 121 ℃ for 1.5 hours, and then centrifuged at 12,000 x g for 10 minutes. Two-hundred and fifty microliters of supernatant were transferred to an 8 mL screw cap glass vial, and TFA was evaporated under a stream of N2 gas at 40 ℃. The resulting residue was dissolved in 300 µL 2- propanol and evaporated under a stream of N2 gas at 40 ℃. This process was repeated two additional times. Two hundred microliters of freshly prepared reduction solution (10 mg/mL NaBH4: 1 M NH4OH) was added to each sample and incubated at 25 ℃ for 1.5 hours. The reduction reaction was quenched with 150 µL of glacial acetic acid and samples were evaporated under a stream of N2 gas at 40 ℃. Two-hundred and fifty microliters of 9:1 [v/v] of methanol: acetic acid was added to the residue and evaporated under a stream of N2 gas. This process was repeated two additional times. Samples were then washed with 250 µL of methanol for a total of 4 times. Alditol acetylation was performed by adding 50 µL of pyridine and 50 µL of acetic anhydride to each sample, sealed tightly, and incubated at 121 ℃ for 20 minutes. Samples were placed in ice for 30 minutes and solvent was gently evaporated under N2 gas at 25 ℃. Samples were washed with 200 µL of toluene for a total of three times. Residues containing alditol acetates were resuspended in 1 mL of ethyl acetate and 4 mL of water. Samples were vortexed

121 and centrifuged at 50 x g for 20 minutes. The upper organic phase was transferred into an

8 mL screw cap glass tube and gently evaporated under N2 gas at 25 ℃. Samples were prepared for gas chromatography by resuspending in 300 µL of acetone. One hundred microliters of each samples were transferred to GC vials and diluted with an additional

200 µL of acetone.

Samples were analyzed on an Agilent 7980A gas chromatography instrument

(Agilent Technologies) equipped with a 30 mm x 0.25 mm x 0.25 µm SP-2380 fused silica capillary column (Supelco), under the following conditions: inlet temperature of

250 ℃, initial oven temperature 160 ℃ held for 2 minutes, first temperature ramp at 20

℃/min to 245 ℃, hold at 245 ℃ for 12 minutes, second temperature ramp at 20 ℃/min to 270 ℃. Data were analyzed using GC Open Lab software (Agilent Technologies).

Random mutagenesis screen and isolation of fluoro fucose resistant lines:

Ethyl methyl sulfonate (EMS) mutagenized Arabidopsis seed were sterilized as described above and screened by germinating and growing for 7 days on MS media containing 25 µM 2F-Fuc accompanied with Col-0 wildtype as a negative control.

Potentially resistant candidates were identified by visually screening for plants with longer roots in the presence of 2F-Fuc, and these plants were rescued by transfer to MS media without 2F-Fuc. Resistant mutant candidates were grown for an additional 7 days under long day conditions (16-hour light/ 8-hour dark) at 22 ℃. Potentially resistant candidates were transferred to soil and maintained until maturity. Seed was collected for further testing and designated as fluoro fucose resistant (ffr).

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Seed from each potentially resistant line were rescreened in MS media without treatment (NT), 25 µM 2F-Fuc, and 25 µM Ac3 2F-Fuc for 7 days. Col-0 was used as a negative control and fkgp-3 as a positive control. Root lengths were quantified using

ImageJ. Mature leaves from select ffr lines were collected and genomic DNA was extracted as described above (Edwards et al., 1991). The FKGP gene was PCR amplified using Phusion polymerase in 1000 base pair fragments using primers listed on Table 2.1.

PCR products were separated in a 1% [w/v] agarose gel and gel purified with a Gel extraction kit (Qiagen) following the manufacturer’s instructions. Purified FKGP fragments were cloned into pCRII-Blunt TOPO (Invitrogen) using the manufacturer’s instructions followed by transformation into Top 10 chemically competent cells. The resulting plasmids were sequenced at the Nevada Genomics Center. Sequences were aligned against the full genome sequence available on TAIR (arabidopsis.org) using

Basic Local Alignment Search Tool (BLAST) (blast.ncbi.nlm.nig.gov/Blast.cgi) to identify the presence or absence of EMS-derived mutations in the FKGP gene.

iii. Results

Effects of fluoro-L-fucose analogs on plant growth:

Previously characterized inhibitors of plant cell wall biosynthesis often demonstrate clear phenotypic defects, such as reduced cell expansion or loss of anisotropic cell growth (Xia et al., 2014; DeBolt et al., 2007; Smyth et al., 2013). We reasoned that further investigation of growth inhibition of L-Fuc analogs in the initial screen (Figure 1.2) could potentially demonstrate clear cell wall defects. The three L- fucose analogs, 2-deoxy-2-fluoro-L-fucose (2F-Fuc), 1,3,4-triacetyl-2-deoxy-2-fluoro-L-

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Table 2.1: Primer sequences used in this chapter

Gene Forward or Left primer (5' - 3') Reverse or Right primer (5' - 3') SALK_ GTGCAAGACAAGCTTTCCAAG CTAGTGGGACCTCCCAAAGAC 037697

LBb1.3 ATTTTGCCGATTTCGGAAC FKGP CACCATGTCTAAGCAGAGGAGGA GCCTCCCCAGACTGTTAACAAGA (Set 1) AA AGTTCACCCAAAGGTCTGG FKGP CCAGACCTTTGGGTGAACTTCTTG TCTGCAACTGCTCCCCATACTTTA (Set 2) TTAACAGTCTGGGGAGGC TGCTCCAACTCTATAGCT FKGP AGCTATAGAGTTGGAGCATAAAG AATACAGATGCTCCAGTTGTA (Set 3) TATGGGGAGCAGTTGCAGA

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fucose (Ac3 2F-Fuc), and 4-deoxy-L-Fucose (4D-Fuc) (Figure 2.1A) were assayed against Arabidopsis thaliana. Fluorinated monosaccharide analogs have been previously characterized as inhibitors of glycosylation events in mammalian systems (Barthel et al.,

2011; Rillahan et al., 2012).

Arabidopsis seeds were germinated and grown in the light for 7 days in the presences of these monosaccharide analogs to determine their effects. Seeds were plated on MS media containing media without added monosaccharide (NT), 0.1% DMSO

(Solvent/negative control), 100 µM L-Fucose (Fuc), 100 µM 2F-Fuc, 100 µM Ac3 2F-

Fuc, and 100 µM 4D-Fuc. Seedlings grown on NT, Fuc, and 4D-Fuc media displayed similar phenotypes with similar root lengths. In contrast, 2F-Fuc and Ac3 2F-Fuc treated seedlings had significantly reduced root lengths compared to non-treated seedlings

(Figure 2.1B and 2.1C). We postulated that Ac3 2F-Fuc would display higher permeability through the cell membrane, then the acetyl groups would be removed by non-specific esterases in the cytosol. To test this hypothesis, the potency of 2F-Fuc and

Ac3 2F-Fuc were compared by measuring Arabidopsis root growth at different concentrations after 7 days of growth. Both 2F-Fuc and Ac3 2F-Fuc inhibited root growth in a dose dependent manner with calculated IC50 values of 10 µM (2F-Fuc) and 2 µM

(Ac3 2F-Fuc) (Figure 2.1D). These results indicate both 2-fluoro-L-fucose analogs are effective inhibitors of Arabidopsis growth, and suggest that Ac3 2F-Fuc passively permeates through the plasma membrane causing it to be a more potent inhibitor.

To test the short-term effects caused by 2F-Fuc on Arabidopsis, seedlings were germinated grown on MS media for 4 days in the light. These seedlings were transferred

125

A B

1 2

3 4

C D

**** ****

Figure 2.1: Effects of fucose analogs on Arabidopsis root growth. (A) Structures of L- fucose (1), 2-deoxy-2-fluro-L-fucose (2), 1,3,4-triacetly-2-deoxy-2-fluoro-L-fucose (3), and 4-deoxy-L-fucose (4) are shown. (B) Arabidopsis seedlings were germinated and grown for 7 days on media containing the following additives: No Treatment (NT), 0.1% DMSO (DMSO), 100 µM L-Fucose (Fuc), 100 µM 2-deoxy-2-fluoro-L- fucose (2F-Fuc), 100 µM 1,3,4-triacetly-2-deoxy-2-fluoro-L-fucose (Ac3 2F-Fuc), and 100µM 4-deoxy-L-Fucose (4D-Fuc). Scale bar represents 10 mm. (C) Root lengths of seedlings from each treatments were quantified and represented with a box plot indicating minimum and maximum values. Dunnett’s multiple comparison (****, p < 0.0001) n = 74-111. (D) Arabidopsis seedlings were germinated and grown for 7 days in media with increasing concentrations of 2F-Fuc (blue line)(IC50 = 11 µM) or Ac3

2F-Fuc (red line)(IC50 = 2 µM), and root lengths were quantified. The resulting data were fit to a single-exponential dose response curve. Error bars represent SEM (n = 80).

126 to MS media with or without 100 µM 2F-Fuc and root lengths were quantified every 12 hours for a total of 72 hours. Seedlings were placed under the same conditions between each measurement. The results throughout the 72-hour time period demonstrate that seedlings grown without 2F-Fuc exhibit relatively linear growth. In contrast, seedlings grown in MS media with 100 µM 2F-Fuc ceased growing 12 hours after transfer (Figure

2.2A). Five days after transfer, seedling and root tip morphology were examined (Figure

2.2B and 2.2C). Root lengths of seedlings on 2F-Fuc were remarkably reduced compared to untreated controls. In addition, the root tips of 2F-Fuc treated seedlings exhibited increased root tip diameter and ectopic root hair production. Overall, these results suggest

2F-Fuc displays similar phenotypes observed by known cell wall biosynthesis inhibitors.

Investigating effects of fluoro-L-fucose analogs on cell wall matrix polysaccharides:

L-Fuc is a relatively common component in several cell wall polysaccharides such as pectic Rhamnogalacturonan-I, Rhamnogalacturonan-II, arabinogalactan proteins, and cellular glycans such as N-linked glycans (Anderson et al., 2012; Mohnen, 2008;

Nguema-Ona et al., 2014; Both et al., 2011). 2F-Fuc might serve as an inhibitor of a specific glycan, as a general inhibitor of fucosylation events, or it could be incorporated into a glycan, thus disrupting the function or the continued synthesis of that said glycan.

To examine the inhibition of cell wall polysaccharide fucosylation events in more detail, the matrix polysaccharides of 7-day-old etiolated Arabidopsis seedlings were quantified by alditol acetate assay (Foster et al., 2010).

Arabidopsis seedlings were grown on MS media (NT) or MS media supplemented with 0.1% DMSO, 100 µM Fuc, 100 µM 2F-Fuc, 100 µM Ac3 2F-Fuc, and 100 µM 4D-

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A B NT 2F-Fuc C NT 2F-Fuc

Figure 2.2: Effects of short-term 2F-Fuc treatment on Arabidopsis seedlings. (A) Arabidopsis seedlings were germinated and grown for 4 days in MS media, then were transferred to media with or without 100 µM 2F-Fuc. Root lengths were quantified every 12 hours for 72 hours. Error bars represent SEM (n = 48). (B) Representation of 9-day-old untreated (NT) and treated seedling (2F-Fuc). Scale bars represent 10 mm. (C) Root tips under 35x magnification of 9-day-old representatives. Scale bars represent 0.5 mm.

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Fuc. Seedlings were harvested and alcohol insoluble residue (AIR) was prepared as described in Materials and Methods. The resulting AIR fractions were hydrolyzed in 2 M trifluoroacetic acid to release monosaccharides form matrix polysaccharides. Released monosaccharides were reduced to sugar alcohols, peracetylated to alditol acetates and used for gas chromatography analysis. Seven commonly occurring monosaccharides were quantified in each sample (Figure 2.3). Both 2F-Fuc and Ac3 2F-Fuc treatment caused a

50% reduction in total cell wall matrix polysaccharide-associated fucose content compared to untreated controls. Additionally, solvent control, Fuc and 4D-Fuc treatment had similar fucose contents compared to untreated controls (Figure 2.3A). All other monosaccharides were unchanged in all treatment groups (Figure 2.3B). These results suggest 2F-Fuc specifically inhibits cell wall fucosylation events.

Genetic analysis investigating the incorporation of fluoro-L-fucose analogs into cell wall polysaccharides:

Continuing the assessment of whether the phenotypic defects associated with 2F-

Fuc treatment are the result of inhibition or incorporation, several glycans were investigated with known fucosyltransferases. FUT1 incorporates L-fucose in the D-

Galactose-(1→2)-α-D-Xylose side chain of xyloglucan (Vanzin et al., 2002). FUT4 and

FUT6 are responsible for attaching L-fucose to the terminal end of arabinogalactan proteins (Liang et al., 2013; Knoch et al., 2014; Wu et al., 2010); FUT11, 12, and 13 incorporate L-Fucose into complex N-linked glycans. (Both et al., 2011; Strasser et al.,

129

A

* *

B

Figure 2.3: Matrix polysaccharide analysis of 2F-Fuc treated seedlings. Alcohol Insoluble Residue (AIR) from 5-day-old dark-grown seedlings grown in the indicated treatments were subjected to Alditol Acetate Assay as described in Materials and Methods. (A) Fucose content of Matrix polysaccharides are represented as a percent mole (%mol). Error bars represent SEM (n = 5). Dunnett’s multiple comparisons (*, p < 0.05). (B) Total cell wall monosaccharides of treated seedlings. Error bars represent SEM (n = 5).

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2004). Homozygous T-DNA lines for each gene encoding each fucosyltransferase are listed in Table 2.2 were germinated and grown in MS media with or without 100 µM 2F-

Fuc. Root lengths of each line were quantified after 7 days of growth (Figure 2.4). All

FUT knockouts exhibited similar root lengths to that of wildtype (Col-0) seedlings in MS media with and without 2F-Fuc treatment, suggesting that the incorporation of 2F-Fuc into xyloglucan, arabinogalactan, and N-linked glycans does not contribute to 2F-Fuc- associated phenotypic defects. Furthermore, the similar root lengths of these mutants in normal MS media suggest that these specific fucosylation events in these glycans do not contribute to 2F-Fuc phenotypic defects.

Complementation of 2F-Fuc phenotypic defects with exogenous boric acid:

The Arabidopsis MUR1 gene, which is primarily expressed in aerial tissues, encodes an isoform of GDP-D-mannose-4,6-dehydratase (GMD2), which catalyzes the first reaction in the de novo biosynthesis of GDP-L-fucose, the active substrate for fucosyltransferases (Bonin et al., 1997, 2003). Mutations in this gene causes severe growth defects due to impaired Rhamnogalacturonan-II fucosylation (RG-II). Genetic evidence from fut1, and fut4/fut6 double mutant lack and detectable L-fucose in xyloglucan and arabinogalactans, respectively. These mutants rule out both cell wall polysaccharides as they have no visible phenotype (Vanzin et al., 2002; Liang et al.,

2013). Interestingly, the phenotype can be partially chemically complemented by exogenous L-fucose or boric acid. The supplementation of L-fucose recovers the lack of

L-fucose in the mur1 mutant, providing the missing monosaccharide in mur1 RG-II.

Wildtype RG-II self-dimerizes in an “enzyme-like” fashion using boric acid as a

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Table 2.2: T-DNA lines used in this chapter

Gene AGI designation T-DNA line fkgp-3 At1g01220 SALK_037697 fut1 At2g03220 SALK_139678 fut4/fut6 Atg2g15390:At1g14080 fut11 At3g19280 SALK_134085 fut12 At1g49710 SALK_063355 fut13 At1g71990 SALK_067444

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Figure 2.4: Analysis of fucosyltransferase mutants for 2F-Fuc sensitivity. Col-0 seedlings were germinated and grown on MS media with or without 100µM 2F-Fuc alongside with fut1 (SALK_139678), fut4;6 double knockout, fut11 (SALK_134085), fut12 (SALK_063355), and fut13 (SALK_067444) mutants. Roots lengths were quantified after 7 days and Sidak’s ns ns ns ns ns multiple comparisons test within each treatment group was calulcated. Error bars represent SEM (n = 30).

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“substrate” generating a boron cross bridge between two RG-II polymers. Compromises in RG-II structure reduce self-dimerization, but exogenous boric acid acting as additional

“substrate” increases self-dimerization partially rescuing the mur1 phenotype (O’Neill et al., 2001). RG-II also exclusively contains the unique monosaccharide 3-deoxy-D- manno-oct-2-ulosonic acid (Kdo), and previous work demonstrated that 2β-deoxy-Kdo, an inhibitor of Arabidopsis CMP-Kdo synthase, phenocopied many of the growth defects observed upon 2F-fuc treatment, including root growth inhibition and cell swelling in

Arabidopsis. Similar to the mur1 mutant, 2β-Kdo-induced phenotypic defects could be partially rescued by the application of exogenous boric acid (Smyth et al., 2013).

To test the possibility of 2F-Fuc phenotypic defects is due to inhibition of RG-II biosynthesis, Arabidopsis seedlings were grown in the presence of boric acid to determine if the phenotype can be partially complemented. Arabidopsis seeds were plated on MS media and MS media containing the following additives, 0.1% DMSO, 1.5 mM boric acid, 100 µM 2F-Fuc, 1.5 mM boric acid + 100 µM 2F-Fuc. Seeds were germinated and grown under long day conditions for 7 days, and root lengths were then quantified

(Figure 2.5C). Seedlings grown under boric acid treatment exhibited shorter roots compared to no treatment and solvent control but did not exhibit the same severity of growth inhibition as 2F-Fuc treatment. Seedlings grown in the presence of 2F-Fuc were partially complemented with boric acid (Figure 2.5A and 2.5B), similar to the partial complementation of 2β-deoxy-Kdo, suggesting that 2F-Fuc phenotypic defects are a result RG-II biosynthesis inhibition. The inhibition of RG-II biosynthesis may occur on side chain A and B where fucose is present with interest on side chain A due to L- fucose’s proximity to apiose where the boron cross link occurs.

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A +B NT DMSO +B +2F-Fuc +2F-Fuc C

**** B

Figure 2.5: Chemical complementation of 2F-Fuc growth defects with Boron. (A) Col-0 seed were germinated and grown for 7 days in media with the following treatments: No Treatment (NT), 0.1% DMSO (DMSO), 1.5 mM Boric Acid (+B), 100 µM 2F-Fuc (+2F-Fuc), and 1.5 mM Boric Acid + 100 µM 2F-Fuc (+B +2F-Fuc). Scale bar represents 10 mm. (B) Higher magnification images of root tips from each treatment. Scale bar represents 0.25 mm. (C) Root length quantification for each treatment. Tukey’s multiple comparisons (****, p < 0.0001), error bars represent SEM (n = 70-97).

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Genetic knockout suggest fluoro-L-fucose analogs are metabolically incorporated:

GDP-L-fucose, the active sugar nucleotide substrate for fucosyltransferases can be synthesized through the de novo or salvage pathway. The de novo pathway utilizes

GDP-D-mannose as a precursor, which is converted through a series of enzymatic reactions by GMD to GDP-4-keto-6-deoxy-D-mannose. This intermediate is a substrate for GDP-4-keto-6-deoxy-D-mannose-3,5-epimerase-4-reductase1 (GER1), which yields the final product GDP-L-fucose (Bonin et al., 1997; Bonin and Reiter, 2000). The salvage pathway converts free L-fucose to GDP-L-fucose via a bifunctional enzyme L-Fucose

Kinase GDP-L-fucose pyrophosphorylase (FKGP) in the cytosol (Figure 2.6A) (Kotake et al., 2008). Current evidence suggests that all non-cellulosic cell wall polysaccharides are synthesized in the lumen of the Golgi apparatus (Oikawa et al., 2013; Kim et al.,

2014) and GDP-L-fucose is transported into the Golgi via a GDP-fucose transporter

(GFT1) (Rautengarten et al., 2016), suggesting that 2F-Fuc must be converted to GDP-

2F-Fuc to enter the Golgi and target fucosyltransferases involved in cell wall biosynthesis.

To test this hypothesis, the FKGP gene was queried against the Arabidopsis T-

DNA insertion database (signal.salk.edu/cgi-bin/tdnaexpress), and potential T-DNA insertion lines were obtained from the Arabidopsis biological resource center (ABRC).

Putative T-DNA lines were screened by PCR genotyping and one allele (see Table 2.2) was confirmed to contain a T-DNA insertion in the fourth intron of the FKGP gene

(Figure 2.6B and 2.6C). With two previously described T-DNA lines (Kotake et al.,

2008), this allele was designated fkgp-3. The fkgp-3 null mutant was grown in MS media

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Figure 2.6: Isolation of fkgp-3 and analysis of resistance to 2F-Fuc. (A) Sequence of reactions catalyzed by the bifunctional FKGP enzyme are demonstrated. (B) Arabidopsis FKGP gene (At1g01220) schematic and location of the fkgp-3 T-DNA insertion. The position of LP, LB, and RP primers used for PCR genotyping are also shown. (C) PCR products from wild-type (Col-0) and fkgp-3 mutant from genotyping with LP + RP and RP + LP primers separated on a 1% agarose DNA gel. (D) Col-0 and fkgp-3 seed were germinated and grown for 7 days on MS media with the following treatments: No Treatment (NT), 100 µM L-Fucose (Fuc), 100 µM 2F-Fuc

(2F-Fuc), and 100 µM Ac3 2F-Fuc (Ac3 2F-Fuc). Scale bar represents 10 mm. (E) Root lengths of each experimental group were quantified and represented as a box plot with bars representing minimum and maximum values. Sidak’s multiple comparison (****, p < 0.0001) (n = 43 – 70).

137

and MS media containing 100 µM Fuc, 100 µM 2F-Fuc and 100 µM Ac3 2F-Fuc for 7 days. The fkgp-3 null mutant did exhibit slightly shorter roots compared to wildtype in regular MS media, but fkgp-3 was completely resistant to 2F-Fuc and Ac3 2F-Fuc comparted to Col-0 (Figure 2.6D and 2.6E). These results indicated that FKGP activity is required for 2F-Fuc growth inhibition and suggest that 2F-Fuc is metabolically converted to GDP-2F-Fuc, the active inhibitor to plant growth.

Forward genetics approach to isolate novel fluoro-L-fucose resistance:

For 2F-Fuc to inhibit growth in Arabidopsis, it must be metabolically converted to

GDP-2F-Fuc and enter the Golgi apparatus, where GDP-2F-Fuc can target cellular fucosyltransferases. A proposed incorporation mechanism is outlined in Figure 2.7. There are still a few unknown enzymes involved in this hypothetical model. There is no known

L-fucose transporter that imports free L-fucose into plant cells, and the fucosyltransferase(s) responsible for incorporating fucose into side chain A and B of RG-

II are unknown. A plant carrying null mutation(s) in L-fucose transporters might yield resistance to 2F-Fuc yet remain sensitive to Ac3 2F-Fuc, because our previous results suggest that this compound can passively traverse the plasma membrane. Additionally, any alterations in the enzymes involved in 2F-Fuc metabolism to GDP-2F-fuc could potentially yield changes in the phenotypes of 2F-Fuc treated seedlings. To isolate novel genes encoding enzymes involved in 2F-Fuc toxicity, a forward genetics screen utilizing ethyl methyl sulfonate (EMS) mutagenized Arabidopsis Col-0 seed was performed to identify mutants that are resistant to 2F-Fuc. Additionally, this screen could yield interesting mutations in cell wall

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Cytoplasm

1. 2. Fuc Fuc GDP-Fuc ? FKGP Golgi 2F-Fuc 2F-Fuc GDP-2F-Fuc 3.

GDP-Fuc GFT GDP-2F-Fuc Non-specific 4. esterase ?

RG-II Ac 2F-Fuc Ac3 2F-Fuc 3 Biosynthesis inhibition

Extracellular space

Figure 2.7: Schematic of 2F-Fuc and Ac3 2F-Fuc metabolic incorporation into a plant cell. 1. The incorporation of 2F-Fuc begins through transport into the cytosol by a currently unknown fucosyltransferase at the plasma membrane. Ac3 2F-Fuc may not require a transporter due to its increased permeability. 2. 2F-Fuc in the cytosol is converted to GDP-2F-Fuc by the action of FKGP. 3. GDP-2F-Fuc is likely transported into the Golgi where through a Golgi-localized nucleotide sugar transporter (GONST), GDP-Fucose Transporter (GFT). 4. GDP-2F-Fuc interacts with an unknown fucosyltransferase involved with RG-II biosynthesis and inhibits its function.

139 integrity sensing or other glycosyltransferases that alter the structure of RG-II (Franck et al., 2018).

EMS mutagenized seed were grown in the presence of 25 µM 2F-Fuc for 7 days and visually screened for increased root elongation. Forty-one potentially resistant candidates were rescued, placed in a new MS media plate and grown in the light for an additional 7 days. Candidate mutants, referred to as fluoro fucose resistant (ffr), were transferred to soil and maintained through their life cycle. Seed from each ffr line was collected and retested on MS media or MS media supplemented with 25 µM 2F-Fuc or 25 µM Ac3 2F-

Fuc. After 7 days of growth, root lengths were measured, and the resulting data is summarized in Table 2.3 and Figure 2.8. Several ffr lines did not yield any seed and thus were not included in the rescreening. Detailed information for each ffr line is referenced in Table 2.3. Lines that did not display resistance when rescreened are ffr12, ffr28, and ffr40. Some lines displayed resistance to both fluorinated fucose analogs, for example ffr5, ffr21, and ffr30. Interestingly, some lines exhibited resistance to 2F-Fuc but not to

Ac3 2F-Fuc, such as ffr14, ffr33, and ffr38.

Examining allelism of ffr lines to fkgp-3:

To narrow the number of novel mutants resistant to 2F-Fuc not attributed to a point mutation in fkgp, the FKGP gene was sequenced in a subset of ffr lines. Select ffr lines were plated on MS media containing 25 µM 2F-Fuc and grown for 14 days under long day conditions. Col-0 seed were plated in conjunction with the ffr lines for back crossing.

Seedlings were transferred to soil and after a week of continued growth, mature leaves were collected for Edward’s genomic DNA extraction (Edwards et al., 1991) as described

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Figure 2.8: Analysis of 2F-Fuc resistant fluoro fucose resistant (ffr) lines. Each ffr line was grown on MS media (blue) without supplementation and MS media supplemented with 25 µM 2F-Fuc (red) and 25 µM Ac3 2F-Fuc (green) for 7 days. Col-0 was used as a negative control and fkgp-3 served as a positive control. Root lengths were quantified and represented as an overlaid bar graph. Error bars represent SEM (n = 5 – 39).

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Table 2.3: Analysis of potential fluoro fucose resistant lines Recessive Allelic to or 25 µM Ac 2F-Fuc No Treatment 25 µM 2F-Fuc 3 fkgp-3 Dominant Line Mean SEM Significant n Mean SEM Significant n Mean SEM Significant n -- -- Col-0 26.12 ± 0.7181 n/a 36 6.409 ± 0.2747 n/a 17 2.274 ± 0.06905 n/a 39 -- -- fkgp-3 23.48 ± 0.6105 ns 39 18.53 ± 0.5157 **** 29 24.1 ± 0.5638 **** 32 -- Recessive ffr1 16.88 ± 0.651 **** 24 13.14 ± 0.7993 *** 11 2.816 ± 0.07757 ns 22 n/a ffr5 23.56 ± 1.22 ns 16 23 ± 0.7356 **** 30 22.91 ± 1.781 **** 12 n/a Recessive ffr6 23.79 ± 1.016 ns 18 17.25 ± 1.57 **** 8 15.79 ± 1.621 **** 15 Yes ffr10 15.21 ± 0.4279 **** 11 10.66 ± 0.502 ns 13 2.832 ± 0.1339 ns 16 n/a ffr12 12.43 ± 0.8709 **** 13 2.103 ± 0.1195 ns 10 2.518 ± 0.1408 ns 14 n/a ffr14 22.84 ± 0.7845 * 22 15.07 ± 2.318 **** 9 2.908 ± 0.1127 ns 21 Yes Recessive ffr15 17.97 ± 1.946 **** 17 19.64 ± 2.114 **** 11 11.3 ± 0.5542 **** 15 Yes Recessive ffr17 19.51 ± 0.9613 **** 16 16.52 ± 2.675 **** 7 3.84 ± 0.3404 ns 15 No Dominant ffr20 22.18 ± 0.4501 ** 24 21.47 ± 1.918 **** 13 19.87 ± 0.7961 **** 21 Yes ffr21 20.44 ± 0.7087 **** 24 15.26 ± 0.3866 **** 23 21.46 ± 0.9954 **** 22 No Recessive ffr22 20.66 ± 0.6322 **** 17 21.91 ± 1.023 **** 9 3.489 ± 0.308 ns 22 n/a ffr24 14.87 ± 0.6487 **** 18 18.94 ± 0.7399 **** 29 13.38 ± 0.5428 **** 19 n/a ffr25 23.68 ± 1.484 ns 13 16.17 ± 1.886 **** 14 9.967 ± 0.7204 **** 17 n/a Recessive ffr26 18.93 ± 0.7739 **** 31 6.059 ± 0.3012 ns 13 2.607 ± 0.06006 ns 15 No Dominant ffr27 16.86 ± 1.019 **** 21 10.26 ± 0.43 ns 17 3.066 ± 1.08 ns 18 n/a ffr28 18.93 ± 0.7739 **** 31 4.371 ± 0.9008 ns 4 2.457 ± 0.09969 ns 9 n/a ffr29 26.81 ± 2.207 ns 14 6.999 ± 0.463 ns 16 2.506 ± 0.07962 ns 21 n/a ffr30 25.29 ± 0.8944 ns 20 19.62 ± 1.67 **** 18 24.06 ± 0.966 **** 20 Yes ffr31 21.67 ± 1.912 * 9 6.922 ± 1.007 ns 5 2.625 ± 0.1745 ns 13 n/a ffr32 31.01 ± 0.9829 *** 16 6.549 ± 0.5831 ns 10 2.196 ± 0.1396 ns 14 Yes ffr33 18.21 ± 2.324 **** 16 16.12 ± 1.19 **** 12 3.544 ± 0.1127 ns 16 Yes Recessive ffr34 22.31 ± 0.8609 * 18 8.852 ± 0.9474 ns 9 2.939 ± 0.1121 ns 16 n/a ffr35 16.48 ± 0.8328 **** 16 13.78 ± 1.827 ** 6 3.03 ± 0.1484 ns 18 Yes ffr36 20.97 ± 1.462 *** 16 21.47 ± 0.8684 **** 15 15.41 ± 1.054 **** 16 n/a ffr37 23.23 ± 0.9712 ns 21 6.419 ± 0.448 ns 16 26.57 ± 1.325 **** 19 n/a ffr38 21.18 ± 0.96 *** 15 19.4 ± 1.201 **** 18 2.628 ± 0.1279 ns 14 n/a ffr39 21.68 ± 0.8425 ** 17 7.713 ± 0.4316 ns 15 2.409 ± 0.1178 ns 16 n/a ffr40 15.61 ± 1.073 **** 18 5.593 ± 0.6946 ns 10 2.533 ± 0.1215 ns 16 n/a ffr41 14.82 ± 0.8486 **** 16 22.27 ± 0.2369 **** 5 16.05 ± 1.793 **** 13 Yes Mean root length (Mean) and SEM values are reported as millimeters. n/a indicates not applicable, not tested, or inconclusive results. Dunnett's Multiple Comparisons test to Col-0, (ns, not significant; *, p < 0.05; **, p < 0.01; ***, p < 0.001; ****, p < 0.0001)

142 in Materials in Methods. The FKGP gene from each individual ffr line was amplified in three fragments approximately 1300 – 1500 bp using primers listed on Table 2.1. The resulting PCR products were gel purified, cloned into pCRII-Blunt -TOPO and transformed into E. coli. Resulting colonies containing FKGP gene fragments were submitted for sequencing at the Nevada Genomics Center.

Of the ffr lines sequenced, six ffr lines contained a point mutation in their FKGP gene containing G to A or C to T mutations (Figure 2.9A). Lines containing mutations in exon sequences in FKGP all encoded missense mutations, including ffr6 (V715I), ffr30

(D76N), and ffr35 (D746N). For ffr15 and ffr20, the point mutations encoded silent mutations suggesting the causative mutation for 2F-Fuc toxicity may not result from the mutations in FKGP. Interestingly, ffr33 and ffr41 contain point mutations in the introns of FKGP (Figure 2.9A). Compared to other ffr lines allelic to fkgp-3, ffr33 is resistant to

2F-Fuc but not to Ac3 2F-Fuc, we hypothesized the loci of the ffr33 mutation leads to altered splicing variants of FKGP. These variants may encode FKGP proteins with reduced activity or have full function but are less abundant due to splicing errors. To test the possibility of splicing variants in ffr33, cDNA was synthesized from RNA was extracted from 7-day old ffr33 and Col-0 seedlings selected on 25µM 2F-Fuc. FKGP fragments were amplified using primer set 3 (Table 2.1) with equal concentrations of cDNA and PCR products were separated on a 1% agarose gel. In ffr33, two distinct bands were present compared to a single product in Col-0, suggesting that the ffr33 mutant incorrectly splices a subset of FKGP transcripts. Interestingly, the larger band in ffr33

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ffr20 A G2546A ffr33 ffr41 K534K G2981A FKGP ffr30 G926A ffr35 C628T G2369A D76N D476N

ffr15 ffr6 G2530A G3103A V462V V715I 500 bp

B 1.5Kb

1.0Kb

Figure 2.9: Mapping of ffr lines allelic to fkgp-3. (A) Loci of each ffr line allelic to fkgp-3. Primers used in this study are indicated in red (Set 1), blue (Set 2), and purple (Set 3) sequences listed in Table 2.1. (B) Qualitative analysis of ffr33 FKGP expression. The third section of the FKGP gene from Col-0 and ffr33 were amplified from cDNA extracted from 7-day-old seedlings of Col-0 and ffr33 and the resulting PCR products were separated on a 1.0% agarose gel.

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Table 2.4: Segregation Ratios of ffr x Col-0 F2 progeny.

Lines # R # S Total % resistant x^2 (25%) Significant ffr17 202 103 305 66.2 76 **** ffr21 55 170 225 24.4 56 ns ffr25 70 247 317 22.1 79 ns ffr26 102 95 197 51.8 49 ****

Number of Resistant (#R) and Sensitive (#S) F2 progeny to 2F-Fuc based on total. Chi squared was calculated based on a segregation ratio of 1:3 (****, p < 0.0001).

145 was qualitatively brighter suggesting the 100 bp intron is the major mRNA product

(Figure 2.9B).

Generation of ffr x Col-0 F2 backcrosses for genome resequencing:

To isolate the gene(s) related to 2F-Fuc resistance from the ffr lines not allelic to fkgp-3 (ffr5, ffr14, ffr17, ffr21, ffr25, and ffr26), some additional genetic analysis is required. One such analysis is to investigate if any of these ffr lines follow simple

Mendelian genetics. Therefore, non-allelic fkgp-3 ffr lines were crossed with Col-0 wildtype. This process is also necessary to perform next-generation mapping using illumina genome resequencing (Austin et al., 2011). The resulting F1 progeny were germinated in MS media and transplanted to soil after 14 days of growth. Plants were maintained for the duration of their life cycle and F2 seeds were collected. Segregation ratios of each ffr x Col-0 F2 progeny were determined by growing F2 seed on 25 µM 2F-

Fuc for 7 days. Resistant and sensitive seedlings were counted and chi – squared analysis determining goodness of fit to 1:3 segregation ratios was performed. Overall, ffr21

(24.4% p > 0.05), and ffr25 (22.1%, p > 0.05) displayed a 1:3 segregation ratio resistant to 2F-Fuc. Interestingly, ffr17 (66.2%, p < 0.0001) and ffr26 (51.8%, p < 0.0001) did not display a 1:3 segregation ratio, instead ffr17 displayed a dominant mutation and ffr26 segregation ratios similar to a heterozygous mutant with a reproductive defect self- crossing (Smith et al., 2018) (Table 2.4).

iv. Discussion and Future Directions

In this chapter, we demonstrate that 2F-Fuc and its acetylated analogs Ac3 2F-Fuc cause severe growth defects in Arabidopsis, including reduced root elongation and

146 reduced cell wall L-fucose content (Figure 2.1, 2.2, and 2.3). Because fucose is a common component of cell wall polysaccharides and other cellular glycans, loss-of- function mutants in the fucosyltransferases that target many of the these glycans were examined and did not mimic the phenotype of 2F-Fuc treatment (Figure 2.4) (Perrin et al., 1999; Tryfona et al., 2014; Both et al., 2011; Strasser et al., 2004). Additionally, the similar root lengths of Col-0 and the fut knockouts under 2F-Fuc treatment indicate that

2F-Fuc is not incorporated into these glycans causing the severe growth phenotype.

Interestingly, the suite of 2F-Fuc associated phenotypes can be partially complemented with exogenous supplementation of boric acid (Figure 2.5). This observation suggests the primary cause of 2F-Fuc growth inhibition is through RG-II biosynthesis inhibition

(Dumont et al., 2015; O’Neill et al., 2001). Furthermore, 2F-Fuc must be metabolically incorporated by FKGP to yield the hypothetical GDP-2F-Fuc to inhibit growth. Growing fkgp-3 under 2F-Fuc and Ac3 2F-Fuc showed no discernable growth inhibition compared to Col-0 (Figure 2.6). These data provide a promising foundation to introduce other fluorinated compounds to Arabidopsis as potential cell wall biosynthesis inhibitors discussed in later chapters.

Summarized in figure 2.7, only half of the enzymes involved with 2F-Fuc incorporation are known (FKGP and GFT1) (Kotake et al., 2008; Rautengarten et al.,

2016). Due to this observation, coupled with the potential of other alterations to the cell wall or wall integrity sensing mechanisms involved with RG-II biosynthesis, we were encouraged to perform a random mutagenesis screen against 2F-Fuc. The resulting screen provided 29 potentially resistant candidates to 2F-Fuc named fluoro fucose resistant (ffr)

(Table 2.3) (Figure 2.8). Sequencing of the FKGP gene in multiple lines demonstrates

147 that 7 ffr lines were allelic to fkgp-3 (Figure 2.9A). ffr33 and ffr41 contained point mutations on the edges of the 3’ ends of intron 5 and 1 respectively. In ffr33, the point mutation leads to altered splicing in FKGP transcripts (Figure 2.9B). Back crosses conducted thus far include four lines (ffr5, ffr14, ffr21, and ffr25) that exhibited simple mendelian genetics with a single gene responsible for resistance. In contrast ffr17, displayed an interesting dominant mutation and ffr26 showed 1:1 ratio of resistant to sensitivity 2F-Fuc (Table 2.4).

Future work associated with 2F-Fuc toxicity should involve elucidating the underlying genetic basis of 2F-Fuc resistance associated with ffr lines described in this chapter by genome resequencing of F2 seedlings. It is unclear why a dominant mutation could lead resistance to 2F-Fuc, but it is possible that a mutation in the fucosyltransferase involved in RG-II biosynthesis is able to incorporate GDP-2F-Fuc into a growing RG-II overcoming the unfavorable enzymatic transition state discussed in chapter 1. Another possibility is increased selectivity of L-fucose transport into the cytosol or in GDP-L- fucose transport into the Golgi, causing a reduction in available 2F-Fuc/GDP-2F-Fuc available.

In a similar study, (Dumont et al., 2015) demonstrated that RG-II dimerization is essentially abolished under 2F-Fuc treatment. What made this piece of data interesting is the detection of what appears to be incomplete RG-II monomers. This experiment can potentially be repeated and the incomplete RG-II fraction can be purified and used as a novel substrate for potential fucosyltransferase(s) that add fucose to a growing RG-II monomer.

148

Fluorinated fucose analogs have been used in previous studies of mammalian species

(Zhou et al., 2017; Rillahan et al., 2013) and more recently in transgenic rice cell suspension cultures to generate N-linked glycan without fucose (Kim et al., 2020), but relatively few studies have utilized monosaccharide analogs as a means to probe the cell wall polysaccharide biosynthesis (Villalobos et al., 2015; Dumont et al., 2015; Smyth et al., 2013). A variety of fluorinated monosaccharides have been utilized in other various ways. One study utilized 2-deoxy-2-fluoro-D-glucose (2F-Glc) as a place holder in the active sight of the crystal structure of bacterial cellulose synthase (Morgan et al., 2013).

2F-Glc has also been used in metabolomics studies in Arabidopsis. This study, like 2F-

Fuc, demonstrate a variety of 2F-Glc metabolites (Fatangare and Svatoš, 2016). Finally

2F-Glc has also been used as an imaging probe in Hela cells and found a reduction in glycolysis (Niccoli et al., 2017).

The associated toxicity of 2F-Fuc treatment in Arabidopsis with the fucosylation of a pectic cell wall polymer highlighted the potential of fluorinated monosaccharides to probe cell wall polysaccharide biosynthesis. In conclusion, 2F-Fuc has reinforced the importance of Fuc in RG-II biosynthesis and has provided novel ffr lines resistant to 2F-

Fuc toxicity. The utility of the ffr lines is to provide the possibility of discovering novel proteins involved in fucose metabolism, RG-II fucosyltransferases, and a novel sensing mechanism associated with RG-II integrity.

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Chapter 3: 2-Deoxy-2-Fluoro-D-Mannose is an inhibitor of energy metabolism in

Arabidopsis thaliana.

i. Introduction:

D-mannose (Man) is a component of several cell wall glycans, primarily in hemicellulosic polymers such as mannan and glucomannan (Rose, 2003; Pauly et al.,

2013; Amos and Mohnen, 2019). Mannan is a homopolymer consisting of β-1→4 linked

D-mannose, whereas glucomannan is a polymer made up of D-glucose-β-1→4-D- mannose dimers (da Cruz, 2013). Genetic studies suggest that mannan is synthesized by

CELLULOSE SYNTHASE-LIKE D (CSLD) proteins CSLD2, CSLD3, and CSLD5, and that this polymer is crucial for the transition between developmental stages in

Arabidopsis thaliana. The triple knockout line csld2/csld2/csld5 has little to no detectable mannan in stems (Verhertbruggen et al., 2011). Interestingly, in vitro biochemical reconstitution of a chimeric CSLD3-CELLULOSE SYNTHASE (CESA) produced a β-

1→4-glucan using UDP-D-glucose (UDP-Glc) as a substrate, suggesting CSL proteins may have more than one substrate (Yang et al., 2020). Glucomannan in Arabidopsis is synthesized by CELLULOSE SYNTHASE-LIKE A (CSLA) proteins CSLA2, CSLA3, and CSLA9 with the triple knockout lacks any detectable glucomannan (Goubet et al.,

2009). Interestingly, investigating the catalytic activity of these proteins in Drosophila

Schneider 2 cells produced two distinct polymers depending of the supplied nucleotide sugar substrate. A β-1→4-D-mannose product was formed when GDP-D-mannose

(GDP-Man) was provided as a sugar nucleotide donor, and glucomannan was produced 155 when supplemented with (GDP-Man) and GDP-D-glucose (GDP-Glc), suggesting mannan is also synthesized by CSLA7 (Liepman et al., 2005).

Man is a component of additional plant oligosaccharides, including N-linked glycans,

Glycophosphoinositol (GPI) anchors, and Glycoinositol phosphoceremides (GIPCs).

Each of these glycan types are important in multiple physiological processes, such as signaling, protein anchoring, cell recognition, and protein localization to the plasma membrane (Gronnier et al., 2016; Liu et al., 2018; Li et al., 2015; Voxeur and Fry, 2014).

N-linked glycans are synthesized in the endoplasmic reticulum (ER) beginning at the dolichol phosphate pathway. A series of ASPARAGINE N-LINKED

GLYCOSYLATION (AGL) proteins synthesize the core glycan structure, although not all glycosyl-linkages have been isolated in plants (Strasser, 2016). The entire glycan structure (N-acetyl-D-glucosamine (GlcNAc)-Man9-Glc3) is transferred to an Asn amino acid in the consensus sequence Asn-x-Ser/Thr in an en bloc mechanism then transported to the Golgi for further processing and transported to the outer leaflet of the plasma membrane (Mellquist et al., 1998).

GPI anchors are lipid anchors for many cell surface proteins that are destined for the outer leaflet of the plasma membrane. In Arabidopsis, there are about 250 predicted GPI anchored proteins (Borner et al., 2003). A few examples include SKU5, COBRA (COB),

COBRA-like (COBL), and LORELEI, which are all GPI anchored proteins involved in directional control of growth and cell expansion in roots, pollen tube extension, and pollen tube female gametophyte interaction respectively (Schindelman et al., 2001; Liu et al., 2013; Li et al., 2013; Sedbrook et al., 2002; Li et al., 2015; Wang et al., 2017). GPI 156 anchors consist of a lipid structure resembling a phospholipid and a core glycan structure consisting of myo-D-inositol-GlcN-Man3-phosphate-ethanolamine. The peptide is attached at its C-terminus at the terminal amine group of ethanolamine (Fujita and

Kinoshita, 2012; Cheung et al., 2014). GPI anchors are synthesized in the ER by a series of glycosyltransferases beginning with SETH1 and SETH2 adding glucosamine to a phosphatidylinositol, then Man is added by PEANUT 1 (PNT1), and ABNORMAL

POLLEN TUBE GUIDANCE 1 (APTG1) adds the third Man in GPIs (Lalanne et al.,

2004; Gillmor et al., 2005; Dai et al., 2014). The GPI anchor is transferred en bloc to the target peptide by GPI8 (Bundy et al., 2016), and the completed product is transported to the Golgi and ultimately to the plasma membrane (Kinoshita et al., 2013).

Finally, GIPCs are a distinct class of glycosylated sphingolipids unique to plants and are the most prominent sphingolipid class in plants making up approximately 64% of all sphingolipids and 25% of total plasma membranes (Markham et al., 2006). In plants,

GIPCs are implicated in several biological processes including cell wall anchoring, cell surface recognition, signaling precursors, and immune responses (Gronnier et al., 2016).

GIPCs consist of a hydrophobic end composed of a hydroxylated fatty acid group and a long chain base (LCB) combined to form a ceramide in the ER. The ceramide is transported to the Golgi and a phospho-myo-inositol is added by INOSITOL

PHOSPHORYLCERAMIDE SYNTHASE (IPCS) synthase generating inositol phosphorylceramide (IPC) (Wang et al., 2008). IPC is then processed further by the addition of D-glucuronic acid by INOSITOL PHOSPHORYLCERAMIDE

GLUCURONOSYLTRANSFERASE 1 (IPUT1) (Tartaglio et al., 2017) followed by the addition of Man by GIPC MANNOSYLTRANSFERASE 1 (GMT1) (Fang et al., 2016). 157

Loss-of-function mutations in IPC and IPUT1 are gametophytic lethal, and the gmt1-1 produces severe developmental defects, including severely stunted growth and reproductive defects, limiting biochemical experiments to cell cultures. Collectively, Man is a structural component to a diverse set of glycans involved in cell wall architecture, developmental process, fertility, and immune response.

The mannosyltransferases associated with the glycans listed above utilize the nucleotide sugar GDP-Man as substrates. GDP-Man is synthesized through a de novo pathway beginning with fructose-6-phosphate (F6P) from glycolysis which is converted to Man-6-P by phosphomannose isomerase (Maruta et al., 2008). Phosphomanno mutase converts Man-6-P to Man-1-P, followed by coupling to GDP by GDP-MANNOSE

PYROPHOSPHORYLASE (GMP) to yield GDP-Man (Bar-Peled and O’Neill, 2011;

Hoeberichts et al., 2008). Interestingly, three mutant alleles in GMP have been previously described in Arabidopsis causing severe developmental defects: cytokinesis-deficient 1

(cyt1), vitamin C deficient 1 (vtc1), and hypersensitive to ammonium (hsn1) (Conklin et al., 1999; Nickle and Meinke, 1998; Qin et al., 2008). GDP-Man also serves as a precursor for three other metabolic products; L-fucose, L-galactose, and ascorbate. As mentioned in chapter 2, L-fucose (GDP-L-fucose) is synthesized from GDP-Man by the action of GMD and GER (Bonin et al., 2003; Seifert, 2004). L-galactose (GDP-L- galactose) is an upstream precursor of ascorbate where GDP-Man is converted to GDP-L- galactose by GTP-mannose-3,5-epierimase (GME). GDP-L-galactose is converted to ascorbate using four sequential enzymatic reactions (Giovannoni, 2007). L-galactose can be found in the side chain A of RG-II but it is predominantly used for ascorbate biosynthesis (Mohnen, 2008; Laing et al., 2007). Interestingly, GME has also been found 158 to produce GDP-L-gulose in small amounts. It has previously been proposed that GDP-L- gulose serves as the precursor of ascorbate, but that has yet to be determined (Wolucka and Van Montagu, 2003).

In chapter 2, the characterization of 2F-Fuc was discussed in detail. In this chapter, the characterization of 2-deoxy-2-fluoro-D-mannose (2F-Man) will be discussed. 2F-Man severely affects Arabidopsis, causing reduced root growth, altered root morphology, and increased lateral root formation. Analysis of major Man containing glycans revealed minimal impacts in seedlings treated with 2F-Man. Interestingly, 2F-Man treated seedlings exhibit reduced cellulose content. A forward genetics screen yielded two 2F-

Man resistant lines containing a point mutation in the HEXOKINASE1 (HXK1) gene, encoding a bifunctional protein that serves as both a hexose kinase and as a sugar sensor.

Finally, 2F-Man reduces total ATP content in Arabidopsis seedlings, suggesting that 2F-

Man does not directly inhibit a glycosyltransferase but potentially alters energy metabolism in plants.

ii. Materials and Methods:

General plant growth and maintenance:

Arabidopsis thaliana Columbia (Col-0) seeds were sterilized in seed cleaning solution (30% [v/v] bleach, 0.1% SDS [w/v]) for 20 minutes at 25 ℃. The seed cleaning solution was removed, and seeds were washed 5 times in sterile water followed by incubation at 4 ℃ for 48 hours before use. Seeds were germinated on MS media (1/2 X

Murashige and Skoog salts, 10 mM MES-KOH pH 5.7, 1% Sucrose [w/v], and 1% phytoagar [w/v]). 2-deoxy-2-fluoro-D-mannose and 2-deoxy-2-fluoro-D-glucose were 159 purchased from Carbosyth (Berkshire, UK), suspended in DMSO to yield 100 mM stock solutions and added to MS media plates at the indicated concentrations. Seedlings were grown under long day conditions (16-hour light, 8-hour dark) at 22 ℃ vertically for 7 days. Seedling roots were straightened scanned with a benchtop scanner and roots lengths were quantified using ImageJ.

Alditol acetate analysis:

Alditol acetate analysis of cell wall matrix polysaccharides were performed as previously described in chapter 2 materials and methods (Villalobos et al., 2015). The remaining pellet (Cellulose and Lignin) after TFA hydrolysis was subjected to Saeman hydrolysis as described previously (Sanchez-Rodriguez et al., 2017; Saeman, 1945) and derivatized to alditol acetates as previously described (Villalobos et al., 2015). The remaining pellet after TFA hydrolysis was washed 2 times with 300 µL of 2-propanol then dried under a stream of N2 gas at 40 ℃. One hundred micrograms of D-myo-inositol was added, and the pellet was hydrolyzed in 63 µL of 70% [v/v] sulfuric acid for 1 hour at 25 ℃. The sulfuric acid hydrolysate was diluted with deionized water to 6% [v/v] sulfuric acid and heated to 100℃ for 3 hours. Samples were cooled to 25 ℃ and centrifuged at 12,000 x g. The soluble fraction was transferred to an 8 mL screw cap glass vial and washed with 1 mL of 15% [v/v] N, N-dioctylamine in chloroform. Samples were vortexed vigorously and centrifuged at 50 x g for 5 minutes. The lower organic phase was discarded. Washes were repeated for a total of four times or until the aqueous phase reached pH 8. One milliliter of chloroform was added and vortex vigorously to remove excess N, N-dioctylamine. Samples were centrifuged at 50 x g, and the lower phase was 160 discarded. Washes were repeated for a total of four times or until aqueous phase reached pH 5. Two hundred microliters of aqueous phase were transferred to an 8 mL screw cap glass vial and evaporated under N2 gas at 40 ℃. The resulting residue was used for reduction and acetylation as described in chapter 2.

Concanavalin A immunoblotting:

Seven-day-old Arabidopsis seedlings were grown on MS media as described above under the following treatment conditions: No treatment (NT), 0.1% [v/v] DMSO

(DMSO), 100 µM D-mannose, and 100 µM 2-deoxy-2-fluoro-D-mannose. Seedling tissue was flash frozen in liquid nitrogen and homogenized with a mortar and pestle to a fine powder. The homogenized tissue was re-suspended in 15 mL of protein extraction buffer (50 mM (4-(2-hydroxyethyl)-1-peperazineethanessulfonic acid (HEPES)-NaOH pH 7.0, 150 mM NaCl, 10 mM MgCl2, 10 mM Ascorbic acid, 5 mM β-Mercaptoethanol,

1% [w/v] Polyvinylpyrrolidone-40) containing 1 Pierce™ EDTA-free protease inhibitor cocktail tablet (Thermo Fisher) at 4 ℃. The homogenate was filtered through 3 layers of

Miracloth, and the filtrate was centrifuged at 10,000 x g for 20 minutes at 4 ℃. The supernatant was transferred to an ultra-centrifuge tube and centrifuged again at 100,000 x g for 1 hour at 4 ℃. The soluble protein fraction was discarded, and the microsomal membrane fraction was resuspended in 1 mL of protein extraction buffer. Resuspended membranes were stored at -80 ℃ until further analysis. Protein concentrations were determined using the Pierce™ BCA protein assay kit (Thermo Scientific) according to the manufacturer’s instructions. 161

Five micrograms of total membrane proteins were separated using a mini-

PROTEAN® TGX™ gel 4-20% (Bio-Rad) at 150 V for 1 hour in SDS-PAGE running buffer (12.5 mM Tris, 19.2 mM glycine, 3.47 mM SDS). Gels were then rinsed with deionized water and soaked in western transfer buffer (25 mM Tris, 192 mM Glycine,

20% [v/v] Methanol). Proteins were transferred onto a nitrocellulose membrane at 2.5 A,

25 V for 5 minutes using Trans-Blot® Turbo™ System (Bio-Rad). After transfer, the membrane was washed in Concanavalin A (ConA) buffer (20 mM Tris-HCl pH 7.4, 500 mM NaCl, 1 mM CaCl2, 1 mM MgCl2) for 5 minutes at 25 ℃. The membrane was blocked in ConA buffer supplemented with 3% (w/v) Bovine Serum Albumin (BSA) for

18 hours at 25 ℃. The blocking buffer was discarded, and the membrane was washed with ConA buffer for 10 minutes at 25 ℃. The membrane was then incubated in ConA buffer supplemented with 3% (w/v) BSA and a 1:1000 dilution of biotinylated ConA for

1 hour at 25 ℃. This buffer was removed, and the membrane was washed 3 times for 10 minutes with ConA buffer + 3% (w/v) BSA at 25 ℃. The membrane was then incubated in ConA buffer supplemented with 3% (w/v) BSA and a 1:1000 dilution of Streptavidin-

HRP conjugate for 1 hour at 25 ℃. The buffer was removed, and the membrane was washed 3 times with ConA buffer + 3% (w/v) BSA at 25 ℃. Membranes were visualized using chemiluminescent developer reagent (90 mM Tris-HCl pH 8.5, 185 µM p-coumaric acid, 1.125 mM luminol, and 0.004% [v/v] H2O2). The membrane was incubated for 45 seconds after the addition of developer reagent, and blots were imaged using a Bio-Rad

Chemidoc MP.

Ascorbate content assay: 162

Seven-day-old Arabidopsis seedlings were grown on MS media as described above under the following treatment conditions: No treatment (NT), 0.1% [v/v] DMSO

(DMSO), 100 µM D-mannose, and 100 µM 2-deoxy-2-fluoro-D-mannose. The seedlings were flash frozen in liquid nitrogen and homogenized with a mortar and pestle to a fine powder. Eighty milligrams of tissue were transferred to a new tube and suspended with 1 mL of Ascorbate extraction buffer (6% Metaphosphoric acid [w/v], 1 mM EDTA), vortexed, and centrifuged at 12,000 x g for 25 minutes at 4 ℃. Two hundred and fifty microliters of the extract were transferred to a new tube, and the pH was adjusted to 5.6 by adding 75 µL of 1 M NaOH. In a UV-transparent 96 well plate, 50 µL of adjusted extract was combined with 50 µL of 10 mM Na2PO4 in triplicate. The baseline absorbance at 245 nm was measured, and then 10 µL of 0.35 U/mL Ascorbate oxidase

(MP biomedicals) suspended in 10 mM Na2PO4 + 0.05% BSA was added to each sample followed by incubation at 30 ℃ for 10 minutes. Reactions were terminated by adding 90

µL of 350 mM HCl, and the absorbance at 245nm was measured again. Ascorbate quantification was based on a standard curve of Ascorbate concentrations ranging from 1 mM to 0.1 mM dissolved in Ascorbate extraction buffer adjusted to pH 5.6. Ascorbate contents were measured by calculating the difference in absorbance at 245 nm before and after the addition of Ascorbate oxidase.

Crude GIPC extraction and enrichment:

GIPCs from Arabidopsis seedlings were extracted as previously described

(Markham and Jaworski, 2007). Seven-day-old seedlings were lyophilized, and 60 mg of tissue were homogenized in a mortar and pestle. Five milliliters of the lower phase in 163

Solvent H ([55:20:25] [v/v/v] 2-propanol: hexanes: water) was added to the homogenate, and lipids were extracted at 50 ℃ for 15 minutes. Samples were centrifuged at 50 x g for

10 minutes, and the supernatant was collected in a separate glass vial. The extraction was repeated with an additional 5 mL of Solvent H lower phase and was heated at 60 °C for

15 minutes. The samples were centrifuged at 50 x g for 10 minutes again, and the resulting supernatant was added to the first collected extract. Extracts were evaporated to dryness under a stream of N2 gas at 50 ℃, and lipids were de-esterified by adding 4 mL of 33% [w/v] methylamine in [7:3] [v/v] ethanol: water followed by an incubation at 50

℃ for 1 hour. The de-esterified extracts were centrifuged at 50 x g for 5 minutes. The supernatants were transferred to a new glass tube and evaporated to dryness under a stream of N2 gas at 50 ℃. The resulting residue was re-suspended in 1 mL of GIPC enrichment buffer ([10:60:6:24] [v/v/v/v] chloroform: ethanol: ammonia: water) and extracted in a shaking incubator at 25 °C for 18 hours. The supernatant was transferred to a 1.5 mL tube and centrifuged at 15,000 x g for 30 minutes at 4 ℃.

Anion exchange cartridges (Phenomenex cartridges strata X-AW 33 µM 200 mg/3 mL) were pre-equilibrated in 3 mL of chloroform. Crude sphingolipid extracts were loaded onto the pre-equilibrated columns, which were then washed with 3 mL of the following solvents in sequential order: Chloroform, (2:1) [v/v] Chloroform: methanol

(C:M), (1:1) [v/v] C:M, (1:2) [v/v] C:M, and methanol. Cartridges were dried for 16 hours, and GIPCs were eluted with 3 mL of GIPC enrichment buffer. Enriched GIPCs were evaporated to dryness under N2 gas at 50 ℃ and subjected to alditol acetate monosaccharide analysis as described above. One hundred micrograms of L-rhamnose was used as an internal standard instead of myo-D-inositol. In preparation for gas 164 chromatography analysis, samples were re-suspended in 100 µL of acetone, transferred to

GC vials, and analyzed on an Agilent 7890A gas chromatography instrument (Agilent

Technologies) with column and method described above.

Sphingolipidomic analysis:

Arabidopsis seedlings were germinated in MS media or MS media supplemented with 0.1% [v/v] DMSO, 100 µM D-mannose, 100 µM 2F-Man, and 100 µM D-mannose

+ 100 µM 2F-Man. Seedlings were grown for 7 days under long day conditions as described above, then transferred to a 15 mL tub, flash frozen in liquid nitrogen, and lyophilized using Labconco freeze dry system. Sphingolipid extraction and targeted lipidomic analysis were performed as previously described (Markham and Jaworski,

2007) at the University of Nebraska Center for Plant Science Innovation.

Blotting for GPI-anchored proteins:

Membrane proteins were extracted from 8-day-old SKU5-GFP and SKU5-

GFP/gpi8-1 lines (Bundy et al., 2016) using methods outlined above. Forty micrograms of membrane proteins were separated on a mini-PROTEAN® TGX™ gels 4-20% (Bio-

Rad) and transferred to a Nitrocellulose membrane as described above. The membrane was blocked for 16 hours in Phosphate Buffer Saline (PBS) (10 mM Na2HPO4, 1.8 mM

KH2PO4-HCl pH 7.4, 137 mM NaCl, 2.7 mM KCl) buffer supplemented with 5% [w/v] non-fat dry milk (NFDM). The blocking buffer was discarded, and the membrane was washed with PBS + 0.01% [v/v] Tween-20 (PBST) buffer for 10 minutes at 25 ℃. The membrane was then incubated in PBS buffer supplemented with 5% [w/v] NFDM and a

1:1000 dilution of anti-GFP rabbit IgG for 1 hour at 25 ℃. This buffer was removed, and 165 the membrane was washed 3 times for 10 minutes with PBST + 1% [w/v] NFDM at 25

°C. The membrane was then incubated in PBS buffer supplemented with 1% (w/v)

NFDM and a 1:1000 dilution of stabilized peroxidase-conjugated goat anti-rabbit antibody for 1 hour at 25 ℃. The buffer was removed, and the membrane was washed 3 times with PBST + 1% (w/v) NFDM at 25 °C. Blots were developed as described above.

ATP and Fructose-6-phosphate quantification:

One hundred to 150 mg of ground flash frozen tissue from 7-day-old Arabidopsis seedlings grown on MS media containing the indicated compound additives were used for the following assays. ATP quantification was performed using the ATP

Bioluminescence Assay Kit HS II (Roche) following the manufacturer’s instructions with minor alterations. Ground flash frozen tissue was placed in a 1.5 mL tube, suspended in

700 µL of 100 mM Tris-NaOH pH 7.75 + 4 mM EDTA, and samples were heated to 100

℃ for 2 minutes. Samples were cooled to 25 ℃ and centrifuged at 10,000 x g for 5 minutes at 4 ℃. The supernatant was transferred to a new 1.5 mL tube. Twenty-five microliters of the supernatant were transferred in triplicate into a Corning® white 96-well flat bottom plate (Sigma-Aldrich) and diluted with an equal volume of dilution buffer provided in the kit. Fifty microliters of the provided enzyme mix were added and recorded luminescence on a Spectramax M5 Microplate Reader (Careforde) at 562 nm and integrated over 1000 ms. ATP concentrations for each sample were determined based on ATP standard curve.

Fructose-6-phosphate (F6P) quantification was performed using Fructose-6-

Phosphate Assay Kit (Sigma-Aldrich). Ground tissue was suspended in 700 µL of 1 x 166

PBS (Phosphate Buffered Saline) (137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4-HCl pH 7.4, 1.8 mM KH2PO4) and vortexed vigorously. Samples were centrifuged at 13,000 x g for 10 minutes at 4 ℃. The resulting supernatant was transferred to a Pierce™ PES10 kDa MWCO Concentrator (Thermo Scientific) and centrifuged at 10,000 x g for 10 minutes at 4 ℃. The resulting flow through was transferred to a clean 1.5 mL tube, and

50 µL of each sample was used for F6P quantification following kit instructions.

Fluorescence was read using Spectramax M5 Microplate Reader at 535 nm Excitation and 587 nm Emission. Concentrations were determined using a F6P standard curve.

Bulk segregant genome resequencing of fmr6 and fmr11:

Seed from EMS mutagenized populations were grown for 7 days in MS media supplemented with 50 µM 2F-Man. After 7 days, potential candidates, referred to as fluro mannose resistant (fmr) mutants, which displayed resistance to 2F-Man as evidenced by increased root elongation in the presence of this compound, were rescued and grown to maturity. The resulting progeny were retested on 50 µM 2F-Man, and candidates with root lengths longer than wild type after 7 days of growth were selected for further study.

The candidates fmr6 and fmr11 satisfied these criteria and were back-crossed to wild type

Col-0 plants. The resulting F1 progeny were grown on MS media, propagated to reproductive maturity, and allowed self-fertilize to generate F2 populations.

F2 seed were selected on MS media supplemented with 50 µM 2F-Man for 7 days. Seedlings displaying root lengths similar to that of their parental lines were transferred to normal MS media and allowed to grow for an additional 5 days.

Approximately, one hundred milligrams of fresh weight from twelve-day-old seedlings 167 were homogenized under liquid nitrogen and genomic DNA was extracted using a

DNeasy® Plant Mini Kit (Qiagen). The extraction was carried out following the manufacturer’s instructions with the exception that genomic DNA was eluted in deionized water instead of the provided elution buffer. Fifteen microliters of genomic

DNA were mixed with 3 µL of 6 x loading dye, and these samples were separated on a

1% agarose gel (Thermo Scientific) at 100 V for 20 minutes in TAE buffer (40 mM Tris,

20mM acetate, 1mM EDTA) to ensure high molecular weight DNA was extracted.

Genomic DNA was confirmed by PCR amplification of Actin 7 using forward primer sequence 5’-ATGGGTCAGAAAGATGCTTACGTTGGTGA-3’ and reverse primer sequence 5’-TCAGGACAACGGAATCTCTCAGCTCCGAT-3’. Cycling conditions used: one cycle at 95 ℃ for 5 minutes, 35 cycles of 98 ℃ for 30 seconds, 52

℃ for 30 seconds, 72 ℃ for 1 minute 15 seconds), and final extension cycle 72 ℃ for 7 minutes. A 1% agarose gel containing the PCR products was ran at 100 V for 20 minutes. DNA concentrations were quantified using Pico green assay. Samples were submitted for gDNA processing and Illumina genome resequencing at the Nevada

Genomics Center. Sequencing reads were aligned to the Arabidopsis reference genome and mismatched base reads not corresponding to EMS induced single nucleotide polymorphisms (SNPs) were filtered out. The proportion of EMS induced SNPs reads were calculated and plotted along the reference genome. EMS induced SNPs located within an open reading frame were selected for further study.

iii. Results:

2F-Man inhibits root growth in Arabidopsis: 168

A previously characterized fluorinated monosaccharide 2-deoxy-2-fluoro-L- fucose (Dumont et al., 2015; Villalobos et al., 2015) produced clear cell wall phenotypic defects in Arabidopsis, such as reduced growth, decreased cell elongation, and loss of anisotropic growth as described in Chapter 2. We reasoned that other fluorinated monosaccharides containing 2-fluoro substitutions that were identified as part of the initial chemical screen described in chapter 1 (Figure 1.10) would potentially have similar cell morphology phenotypes. To test this hypothesis, Arabidopsis seedlings were grown under long-day conditions (16-hour light/8-hour dark) in MS media and MS media supplemented with 0.1% (v/v) DMSO (DMSO), 100 µM D-mannose (Man), and 100 µM

2-deoxy-2-fluoro-D-mannose (2F-Man) (Figure 3.1A) for 7 days. Seedlings grown under

DMSO and Man treatment exhibited similar growth phenotypes compared to untreated controls. In contrast, seedlings treated with 2F-Man displayed severe growth defects, including reduced growth and altered root morphology. Interestingly, 2F-Man treated seedlings produced lateral roots earlier than untreated plants (Figure 3.1B). Root lengths of 2F-Man treated seedlings displayed an 85% growth reduction compared to untreated seedlings (Figure 3.1C).

To further understand the effective dose range of 2F-Man treatment against

Arabidopsis, seeds were germinated and grown for 7 days on MS media with a range of

2F-Man concentrations from 0.1 to 100 µM. Root lengths were quantified and a dose response curve was generated, yielding an inhibitory concentration at 50% (IC50) of 12

µM (Figure 3.1D). To assess the short-term effects of 2F-Man treatment, seedlings were grown in MS media for 4 days and transferred to MS media with or without 100 µM 2F-

Man. Root lengths were quantified every hour for a total of 30 hours. Seedlings 169

A B C

(1)

**** (2)

D E

IC50 = 12 µM

Figure 3.1: Effects of 2-deoxy-2-fluoro-D-Mannose on Arabidopsis growth. (A) Structures of D-Mannose (1) and 2-deoxy-2-fluoro-D-Mannose are shown. (B) Arabidopsis Col-0 seed were germinated and grown for 7 days in MS media with the following treatments: No Treatment (NT), 0.1% DMSO (DMSO), 100 µM D- Mannose (Man), and 100 µM 2-deoxy-2-fluoro-D-Mannose (2F-Man). Scale bar represents 10 mm. (C) Root lengths from seedlings of each treatment were quantified and represented as a box plots with error bars demonstrating minimum and maximum values. Tukey’s multiple comparisons (****, p < 0.0001) (n = 67 – 74). (D) Arabidopsis Col-0 seed were germinated and grown for 7 days in MS media with increasing concentrations of 2F-Man. The root lengths were quantified, and the resulting data was fit to a single-exponential dose response curve. Error bars represent SEM n = (57 – 74). (E) Col-0 seed was germinated and grown in MS media for 4 days, then were transferred to media with or without 100 µM 2F-Man (2F-Man). Root lengths were quantified every hour for a 30-hour growth period. Error bars represent SEM (n = 12).

170 transferred to MS media without 2F-Man supplementation served as controls and continued linear root growth during the duration of the experiment. Under 2F-Man treatment, complete root growth inhibition was observed within 13 hours of treatment

(Figure 3.1D). Overall, these results indicate 2F-Man is an effective inhibitor of plant growth in a dose dependent and temporal manner.

Cellulose but not matrix polysaccharides are altered in 2F-Man treated seedlings:

Man is a common component of several hemi-cellulosic polysaccharides, such as mannan and glucomannan (Wang et al., 2013; Pauly et al., 2013; Davis et al., 2010).

Glucomannan is synthesized by three CSLA enzymes CSLA2, CSLA3, and CSLA9. The triple knockout line csla2-1/csla3-2/csl9-1, displays little to no glucomannan in Stems with little to know growth differences in wildtype (Goubet et al., 2009). A mutation in

CSLA7, showed little to no mannan polysaccharides resulting in poor pollen tube elongation and ceased embryogenesis (Goubet et al., 2003). We postulated that 2F-Man could serve as an inhibitor of mannosylation events related to these polymers.

Additionally, D-mannose is the precursor for L-fucose (Fuc), suggesting that 2F-Man may be metabolized to GDP-2F-Fuc, which is the ultimate toxic species (Bonin and

Reiter, 2000; Bonin et al., 2003). The cyt1-1 mutant in the GDP-D-mannose pyrophosphorylase (GMP) displayed reduced GDP-Man, leading to termination of embryogenesis and reduced cellulose content (Lukowitz et al., 2001). The reduction in cellulose may be caused by loss of mannosylated glycans critical for the function accessory proteins involved in cellulose biosynthesis such as N-linked glycans, GPI- anchors, or GIPCs (von Schaewen et al., 2015; Roudier et al., 2005; Mortimer et al., 171

2013). Additionally, the lack of GDP-Man may also reduce the availability of mannan and glucomannan. We postulate 2F-Man functions as an inhibitor of mannosyltransferases targeting many of these glycans leading to altered cell wall polysaccharides including cellulose. To investigate the potential of 2F-Man altering cell wall polysaccharides, 7-day-old dark grown Arabidopsis hypocotyls grown in MS media without sucrose and MS media without sucrose supplemented with 0.1% DMSO, 100 µM

Man, and 100µM 2F-Man were subjected to cell wall analysis via an Alditol Acetate assay.

Alcohol insoluble residue (AIR) from 7-day-old hypocotyls was subjected to weak acid hydrolysis, and the resulting insoluble fraction was saved for Saeman hydrolysis to quantify cellulose content. Soluble fractions containing matrix monosaccharides were reduced to their sugar alcohol derivatives the per-acetylated to yield their alditol acetates for gas chromatography. Man and Fuc content is summarized in in Figure 3.2A. Man content in 2F-Man treated seedlings was marginally increased compared to untreated and solvent controls, but this result was not statistically significant

(Dunnett’s multiple comparisons p = 0.1569, n = 5). Fuc on the other hand, was mildly reduced in 2F-Man treatment compared to no treatment and solvent controls yet statistically not significant compared to no treatment (Dunnett’s multiple comparisons p

= 0.6096, n = 5). All other monosaccharides exhibited no statistically significant alteration in 2F-Man treated seedlings compared to all other controls (Figure 3.2B).

The insoluble fractions that remained after weak acid hydrolysis were subjected to Saeman hydrolysis to liberate glucose (Glc) from crystalline cellulose (Saeman, 1945). 172

A C

ns ns ns ** ns ns ns

B

Figure 3.2: Cell wall monosaccharide analysis of 2F-Man treated seedlings. Alcohol Insoluble Residue (AIR) from Arabidopsis Col-0 seedlings grown in the indicated treatments was subjected to Alditol Acetate assay as described in Materials and Methods, and the resulting unhydrolyzed residue from weak acid hydrolysis was subjected to Saemen Hydrolysis to quantify crystalline cellulose content. (A) Mannose and fucose content of the Matrix polysaccharides are represented as percent mole [%mol]. Error bars represent SEM (n = 5). (B) Total cell wall matrix monosaccharide composition of treated seedlings. Error bars represent SEM (n = 5). (C) Cellulose content of treated seedlings represented as µg Glucose per mg AIR (µgGlc/mg AIR). Dunnett’s multiple comparisons (**, p < 0.01), Error bars represent SEM (n = 11).

173

The resulting released Glc was further processed to alditol acetates and analyzed using gas chromatography. Glc content was quantified and represented as micrograms of Glc per milligram of AIR. Cellulose content in 2F-Man treated seedlings was reduced by 40% compared to untreated controls (Figure 3.2C). Overall, these results indicate that matrix polysaccharides are marginally impacted by 2F-Man treatment, suggesting that the severe growth phenotypes cannot be explained by disruption of Man containing polysaccharides.

Interestingly, the reduction in crystalline cellulose content could potentially explain the significant growth reduction associated with 2F-Man treatment and could result from the

Man-specific glycosyltransferase inhibition in a number of possible targets, such as N- linked glycans, GPI anchors, and Glycoinositol phosphoceremides (GIPCs) (Lukowitz et al., 2001; Borner et al., 2003; Fang et al., 2016; Mortimer et al., 2013).

2F-Man alters other mannose containing glycans but not ascorbate content:

N-linked glycosylation in plants begins through the dolichol phosphate pathway to generate the core glycan beginning with the addition of 2 N-acetylglucosamines to dolichol phosphate followed by the addition of D-mannose in the cytosol. When four D- mannose sugars are added, the growing glycan flips into the lumen of the ER and continues to be synthesized to a final precursor (Banerjee et al., 2017; Jadid et al., 2011).

Korrigan (KOR1) is a β-1→4 endoglucanase required for proper cellulose deposition.

Interestingly, KOR1 contains 8 N-linked glycosylation sites (Lane et al., 2001; Rips et al., 2014; Liebminger et al., 2013). 2F-Man may inhibit protein N-linked glycosylation but instead of targeting a specific protein it targets all N-linked glycans. This provided a rationale to investigate N-linked glycosylation using the Concanavalin A (ConA) lectin, 174 which binds terminal mannose residues, to probe the abundance and composition of core

N-linked glycans (Dwyer and Johnson, 1981).

Arabidopsis seedlings were grown for 7 days in MS media or MS media containing 0.1% DMSO, 100 µM Man, and 100 µM 2F-Man. Proteins were extracted and membrane proteins were separated from crude extracts as described in Materials and

Methods. Equal amounts of membrane proteins were separated by SDS-PAGE and transferred to a nitrocellulose membrane, which was probed with Biotinylated ConA to investigate the extent and composition of N-linked glycosylation in these samples.

Untreated seedlings or seedlings treated with 100µM Man displayed similar signal intensities. Interestingly, DMSO treatment caused a decrease in signal, but this observation could be attributed the unfolded protein response common in DMSO treatment (Kang et al., 2017). Though this effect is seen in mammalian species, this observation is not commonly observed in plants. The most peculiar result appears in 2F-

Man treatment, where the ConA signal intensity is higher compared to all other groups, suggesting that 2F-Man may inhibit mannosidase activity directly or is incorporated into an N-linked glycan that is resistant to trimming by mannosidases (Huttner et al., 2014)

(Figure 3.3A).

Another potential target of 2F-Man growth inhibition are GPI anchors. GPIs are synthesized in the ER beginning in the outer leaflet of the ER membrane. After the addition of an myo-D-inositol and D-glucosamine to the lipid moiety it is translocated into the luminal side of the ER membrane, where 3 Man sugars are added followed by an ethanolamine moiety. Target proteins are attached via their C-terminus to the 175

Figure 3.3: Analysis of N-linked glycosylated membrane protein, GPI-Anchored proteins, and Ascorbate content. (A) Arabidopsis Col-0 seed were germinated and grown for 7 days in MS media containing additives previously described. Protein was extracted from each treatment and membranes were separated from soluble protein extract. Five micrograms of total membrane proteins were separated on an SDS- PAGE, transferred to a nitrocellulose membrane and probed for N-linked glycans using Concanavalin A. (B) SKU5-GFP seed were germinated and grown in media with treatments described previously. Five micrograms of total membrane protein were separated on an SDS-PAGE, transferred to a nitrocellulose membrane and probed with Anti-GFP antibody. Purified CC1-GFP protein was used as a positive control. (C) Col- 0 seed were germinated and grown for 7 days in indicated treatments, ascorbate was extracted from approximately 100 mg of ground flash frozen tissue with 6% hexametaphosphoric acid. Ascorbate content was quantified with Ascorbate Oxidase. Error bars represent SEM (n = 5).

176 ethanolamine nitrogen (Cheung et al., 2014). COBRA (COB), a GPI anchored protein is involved in regulating crystalline cellulose deposition and affects the orientation of cell expansion in roots (Schindelman et al., 2001; Liu et al., 2013), indicating that GPI- anchored proteins are involved in cellulose deposition. If 2F-Man targets GPI anchor biosynthesis, then there should be an overall decrease in GPI anchored proteins, including one such protein SKU5, involved with directional root growth (Sedbrook et al., 2002).

We obtained a GFP-SKU5 transgenic Arabidopsis line (Bundy et al., 2016) and grew it for 7 days on MS media or MS media supplemented with 0.1% DMSO or 100

µM 2F-Man. Proteins were extracted, blotted, and subjected to Western blot using an anti-GFP antibody. Purified CC1-GFP was used as a positive control. Similar to the results observed for N-linked glycosylation, DMSO treated GFP-SKU5 line had a less intense signal compared to no treatment control. Signal intensity of 2F-Man treated seedlings was similar to untreated control, suggesting that 2F-Man does not affect GPI anchor biosynthesis or trafficking (Figure 3.3B).

The most peculiar possible target of 2F-Man growth inhibition is Ascorbate biosynthesis. The Arabidopsis vtc1 mutant contains a point mutation in the GMP gene allelic to the cyt1 mutant. This mutant contained approximately 25% of the free ascorbate compared to wild type plants (Conklin et al., 1999), suggesting that the precursor for

Ascorbate is GDP-Man. GDP-Man undergoes 5 enzymatic reactions to yield Ascorbate

(Giovannoni, 2007). These observations led us to the hypothesis that 2F-Man may affect

Ascorbate biosynthesis. Specifically, the key step may be in the L-Galactose 177 dehydrogenase where the partial positive charge of the adjacent carbon bonded with fluorine will make the formation of the lactone unfavorable or impossible.

To test this hypothesis, ascorbate was extracted with 6% hexametaphosphoric acid from seven-day-old seedlings grown under previously described treatments. Extracts were normalized to a pH of 5.6 with HCl. To determine ascorbate content, the absorbance of each sample was recorded at 245 nm before treatment with Ascorbate Oxidase.

Absorbance was recorded after treatment and content was quantified based on change of absorbance to a standard curve. Ascorbate content for each treatment is summarized in

Figure 3.3C. Overall, ascorbate content is unchanged in 2F-Man treated seedlings compared to controls. These data suggest that 2F-Man does not impact ascorbate biosynthesis and cannot explain the observed stunted growth phenotype.

Overall GIPC monosaccharide content is mildly reduced in 2F-Man treated seedlings:

GIPCs are a unique category of glycosylated ceramides found exclusively in plants that are homologous to gangliosides in mammalian systems (Gronnier et al., 2016).

Genetic and chemical relationships between GIPC and cell wall biosynthetic processes have been proposed. For example, there is evidence to suggest that GIPCs tether RG-II fragments to the plasma membrane through boron cross-linkages (Voxeur and Fry, 2014).

Additionally, genetic evidence suggests that GIPC composition plays a role in cellulose biosynthesis, although the mechanistic basis of this relationship is unclear. In

Arabidopsis, all GIPCs contain a core ceramide attached to a phosphate followed by a myo-D-inositol residue and D-Glucuronic acid. Additional monosaccharides are added to the GIPC head group. For example, Man can be attached to the D-glucuronic acid residue 178 through the action of GMT1 (Fang et al., 2016). Alternatively, GLUCOSAMINE

INOSITOLPHOPHORYLCERAMIDE TRANSFERASE 1 (GINT1) adds an N- acetylglucosamine (GlcNAc). In Arabidopsis, loss of function gint1 mutants did not display any phenotypic defects, but it is lethal in rice where GlcNAc containing GIPCs are the major species (Ishikawa et al., 2018). GMT1 uses GDP-Man as a substrate, which is transported into the Golgi via GONST1 (Mortimer et al., 2013). Importantly, previous work has shown that both gonst1-1 and gmt1-1 mutants exhibit compromised GIPC composition and reduced crystalline cellulose content (Mortimer et al., 2013; Fang et al.,

2016), raising the question of whether 2F-Man negatively impacts GIPC biosynthesis.

To test this hypothesis, GIPCs were extracted from Arabidopsis seedlings grown for 7 days under previously described conditions and GIPCs were enriched as described in Materials and Methods based on an established protocol (Markham et al., 2006).

Enriched GIPCs were subjected to TFA hydrolysis and similar chemistry was utilized in analysis of matrix polysaccharides. In initial experiments, L-rhamnose was not detected in the GC chromatogram, and thus it was used in subsequent experiments as an internal standard. Through this method D-arabinose, D-xylose, Man, D-galactose, Glc, and myo-

D-inositol were detected and quantified (Figure 3.4A). Glc may be the result of glucuronic acid, glucosamine, and Glc. Interestingly, small amounts of D-xylose were detected which has not been previously reported. Overall, most monosaccharides were slightly negatively affected by 2F-Man, specifically Glc and Man had the greatest decrease upon 2F-Man treatment compared to untreated controls. It is difficult to attribute the decrease in Glc and Man due to similar decreases observed in Man treated seedlings 179

Figure 3.4: Analysis of GIPC headgroup monosaccharide composition. (A) Seven- day-old Arabidopsis seedlings grown under previously described treatments were lyophilized, whole sphingolipid extracts were enriched for GIPCs and subjected to Alditol Acetate Assay as described in materials and methods with rhamnose as an internal control. Monosaccharide content was quantified and represented as µg/ mg dry weight. Tukey’s multiple comparisons (*, p < 0.05; **, p < 0.01). Error bars represent SEM (n = 6). (B) Mannose content from enriched GIPC extracts are shown. Tukey’s multiple comparisons (*, p < 0.05; ****, p < 0.0001). Error bars represent SEM (n = 6).

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(Figure 3.4A and 3.4B). Overall, these results and results from N-linked glycosylation demonstrate a modest change compared to untreated controls, suggesting that the primary target of 2F-man growth inhibition is not attributed to any of the tested glycans or ascorbate.

To further investigate the impact of 2F-Man on Glc and Man in GIPCs, targeted sphingolipidomic analysis of 7-day-old Arabidopsis seedlings grown in MS media or MS media supplemented with 0.1% DMSO, 100 µM Man, 100 µM 2F-Man, and 100 µM

Man + 100 µM 2F-Man were lyophilized, and subjected to established sphingolipid extraction procedures described in Materials and Methods. The Man+2F-Man treatment was included in this experiment to investigate the effects of metabolic competition between Man and 2F-Man on GIPCs The resulting extracts were subjected to targeted lipidomic analysis to quantify the abundance or GIPC, but also included other sphingolipids such as Glucosylceramides (GlcCers) and hydroxyceramides (hCers).

Overall, 2F-Man and Man+2F-Man treatment had slight decreases in total GIPCs and

GlcCers but were not statistically significant under Dunnett’s multiple comparisons test

(p > 0.05) (Figure 3.5A and 3.5B). Total hCers were unchanged in all treatment conditions (Figure 3.5C). Interestingly, analysis of t18:0 containing GIPC and GlcCer species were reduced under 2F-Man treatment (Figure 3.5A and 3.5B). The slight decreases in Glc and Man in enriched GIPCs (Figure 3.4) from 2F-Man treated seedlings may be reflected in the small decreases observed in total GIPCs (Figure 3.5A). It is unexpected to observe the slight decrease in GlcCers, a sphingolipid species that does not contain Man, in 2F-Man treated seedlings (Pata et al., 2012). It is possible 2F-Man weakly binds GLUCOSYLCERAMIDE SYNTHASE (GCS) (Melser et al., 2010) 181

Figure 3.5: Sphingolipid analysis of 7-day-old Arabidopsis seedlings grown under MS media with no treated (NT), 0.1% DMSO (DMSO), 100 µM 2F-Man (2F-Man), and 100 µM Man + 100 µM 2F-Man (Man+2F-Man) are shown. Totals of each sphingolipid species are represented as bar graphs. Detailed list of long chain bases (LCB) and hydroxy fatty acids (hFA) are represented as heat maps demonstrating relative amounts of each species to no treatment. (A) GIPC content with LCB and hFA shown bellow. Error bars represent SEM (n = 3 – 4). (B) GlcCer content with LCB and hFA shown bellow. Error bars represent SEM (n = 3 – 4). (C) hCer content with LCB and hFA shown bellow. Error bars represent SEM (n = 3 – 4).

182 slightly inhibiting GlcCer biosynthesis. Alternatively, the changes in GIPCs and GlcCers in 2F-Man treated seedlings may be a secondary effect of a yet unknown primary target of 2F-Man.

Forward genetics approach to isolate 2F-Man resistance:

The limited insight from all glycans analyzed of 2F-Man treated seedlings have not provided a clear mechanism to explain the growth phenotype elicited by 2F-Man treatment. Investigating for possible metabolic incorporation mechanisms, free Man is phosphorylated by a Hexokinase to yield D-mannose-6-phosphate, where the salvage pathway and the de novo pathway merge. D-mannose-6-phosphate is converted D- mannose-1-phosphate by phosphomanno mutase (PMM) (Qian et al., 2007) followed by the coupling to GDP by GMP (Lukowitz et al., 2001). This limited the number of genes to target in a reverse genetic strategy, since many of these genes are essential and homozygous mutants cannot be obtained. To gain further insight into the mechanism of

2F-Man toxicity, we performed a forward genetic screen to identify 2F-Man resistant mutants.

Ethyl methyl sulfonate (EMS) mutagenized seeds were screened on MS media supplemented with 50 µM 2F-Man. EMS seeds and Col-0 (wildtype) were plated and grown for 7 days under long day conditions. After 7 days, mutant seedlings with roots longer than wild type Col-0 seedlings treated under the same conditions were transferred to normal MS media and grown for an additional 7 days. Seedlings were transferred to soil and maintained for the remainder of their life cycle. Seeds were collected from each candidate and referred to as fluoro mannose resistant (fmr). A total of 15 fmr lines were 183 obtained and were then retested on 2F-Man. Each fmr line was grown on MS media supplemented 50 µM 2F-Man including Col-0 as a negative control. Additionally, Col-0 was grown on MS media to serve as a proxy positive control. Seedlings were grown for 7 days and root lengths were quantified (Figure 3.6). Overall, most fmr lines displayed limited resistance to 2F-Man. Interestingly, fmr6 and fmr11 displayed the greatest level of resistance to 2F-Man, yet these mutants were not completely resistant to 2F-Man, with the root lengths of these mutants achieving only 50% of untreated Col-0. Other candidates displayed modest 2F-Man resistance during these assays, such as fmr1, fmr5, and fmr7. Both fmr6 and fmr11 were selected for further study.

Both fmr6 and fmr11 contain point mutations in HXK1:

Prior to genome resequencing and next-generation mapping (Austin et al., 2011), back crosses of fmr6 and fmr11 to Col-0 were generated. Seed from F1 populations were collected and grown in MS media for 14 days. Seedlings were transferred to soil and maintained through their life cycle. F2 seeds were collected and rescreened against 50

µM 2F-Man to asses segregation ratios. Initially, we hypothesized both lines may display a segregation ratio of 1:3 indicating a single recessive mutation causing resistance.

Interestingly, 20.5% of fmr6 seedlings displayed resistant to 2F-Man, lower than what would be expected of a 1:3 ratio of a single recessive gene. This may be due to multiple mutations causing resistance or it could be some mutations detrimental to growth may be linked to the causative mutation. On the other hand, 26.3% of fmr11 seedlings displayed resistance to 2F-Man, conforming to the 1:3 segregation ratio (Table 3.1).

184

Figure 3.6: Growth analysis of 2F-Man resistant lines. All fmr lines were grown on MS media containing 50 µM 2F-Man for 7 days. Root lengths were quantified using ImageJ as describe in Materials and Methods, error bars represent SEM, (n = 6 – 34).

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Table 3.1: Segregation Ratios of fmr x Col-0 F2 progeny

Line #R #S Total %R x^2 (25%) Significant fmr6 137 530 667 20.5 168 ** fmr11 178 498 678 26.3 169 ns

Number of Resistant (#R) and Sensitive (#S) F2 progeny to 2F-Man based on total. Chi squared was calculated based on segregation ratio 1:3 (**, p < 0.01)

186

Resistant seedlings from fmr6 and fmr11 were collected and genomic DNA was extracted. Genomic DNA from both lines were prepared for Illumina genome resequencing at the Nevada Genomics Center and point mutations corresponding to EMS mutations (G/C to A/T) (Greene et al., 2003) were for both lines (Figure 3.7A and 3.7B).

Interestingly, both fmr6 and fmr11 have EMS mutations clustered at higher frequency in chromosome 4. Point mutations were found in the gene HXK1 with fmr6 containing a missense mutation causing a V179I. The increased hydrophobicity of Isoleucine may alter the tertiary structure of HXK1. The fmr11 line contained a mutation located within the 5th intron region potentially causing HXK1 mRNA splicing errors. HXK1 is a bifunctional enzyme that phosphorylates hexose sugars and serves as a glucose sensor.

Under high glucose concentrations (6% in MS media), HXK1 represses growth in

Arabidopsis by entering the nucleus and regulating gene expression (Cho et al., 2006). A mutation in HXK1 referred to as glucose insensitive 2 (gin2) is completely insensitive to glucose repression (Cho et al., 2006; Moore et al., 2003).

Due to the unique function of HXK1 in glucose repression, we considered whether other hexose analogs could inhibit growth via a similar mechanism. To test this hypothesis, one other potent inhibitor 2-deoxy-2-fluoro-D-glucose (2F-Glc), which is also a substrate for HXK1 (Figure 1.10), was assayed against the fmr6 and fmr11 mutants. Both lines and Col-0 were grown in MS media and MS media supplemented with 50 µM 2F-Man or 50 µM 2F-Glc for 7 days. Upon inspection, both fmr6 and fmr11 mutants were resistant to 2F-Glc compared to Col-0 (Figure 3.8A). Quantification of root lengths further demonstrated resistance to 2F-Glc but at 50 µM, Col-0 root lengths were more varied (Figure 3.8). These results suggest that 2F-Glc inhibits growth in 187

Figure 3.7: Mapping of fmr6 and fmr11 F2 lines. Chromosomes are shown with every mutation corresponding to EMS point mutation (G to A) (Red). SNP index corresponds to the proportion of EMS mutations found in Illumina sequencing reads. (A) Map of fmr6 and (B) fmr11.

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Figure 3.8: Assaying resistance of fmr mutants on 2F-Man and 2F-Glc. (A) Wildtype Col-0, fmr6, and fmr11 were germinated and grown on MS media containing the following treatments: No Treatment (NT), 50 µM 2F-Man (2F-Man), and 50 µM 2- deoxy-2-fluoro-D-Glucose (2F-Glc). Scale bar represents 10 mm. (B) Root lengths were quantified and represented as a box and whisker plot displaying minimum and maximum. Tukey’s multiple comparisons (****, p < 0.0001) (n = 15 – 20).

189

Arabidopsis through a mechanism that is dependent on HXK1 function. To further investigate the connection between fmr6 and fmr11 to glucose repression insensitivity, both lines were grown under increasing concentrations of Glc for 7 days. Col-0 seedlings grown under higher concentrations of Glc exhibited severe growth repression. While similar results were observed for the fmr6 and fmr11 mutants, the repression was not as severe as Col-0 (Figure 3.9A). Interestingly, a recurring observation in this experiment was that fmr6 and fmr11 mutants germinated exhibited higher germination rates compared to Col-0 on MS media containing high glucose concentrations. Overall, both lines display higher rates of germination at every concentration except for 4% D-glucose concentration (Figure 3.9B). These results support HXK1’s role in seed germination

(Rolland et al., 2002).

Energy metabolism in 2F-Man treated seedlings:

Resistance to 2F-Man through mutations in HXK1 highlights HXK1’s complex and involved interactions with Golgi (da‐Silva et al., 2001), mitochondria (Galina et al.,

1995), nuclear localization (Cho et al., 2006), and other plastids in plants (Wiese et al.,

1999). Many of these interactions influence Chloroplast development, seed germination, leaf development, root and shoot elongation, aquaporin expression, and programed cell death (Kelly et al., 2017; Kunz et al., 2015; Kushwah and Laxmi, 2017; Dingenen et al.,

2019; Bruggeman et al., 2015). As varied and complex as these interactions and biological activities are, possibly the simplest way to combine all these interactions is direct involvement of HXK1 with energy metabolism. Similar to how the HXK2 protein in yeast plays pivotal role in glucose repression and 2-deoxy-D-glucose 190

Figure 3.9: Assaying glucose repression in fmr6 and fmr11. (A) Root length quantification of Col-0, fmr6, and fmr11 on increasing concentrations of glucose, root lengths are represented as a percentage of No treatment (NT). Error bars represent SEM (n = 15 – 29). (B) Portion of seeds germinated on increasing concentrations of glucose. Seedlings that germinated were counted as a 1 and seed that failed to germinated were designated as 0, then values were averaged. Error bars represent SEM (n = 19 – 30).

191

(Trumbly, 1992; McCartney et al., 2014), energy metabolism could be negatively impacted by monosaccharide analogs such as 2F-Man. To test this hypothesis, ATP and

Fructose-6-phosphate (F6P) content was quantified in 2F-Man treated seedlings. Seven- day-old seedlings grown on MS media or MS media supplemented with 0.1% DMSO,

100 µM Man, and 100 µM 2F-Man were ground in liquid nitrogen to a fine powder and

ATP and F6P was quantified using commercially available kits described in Materials and Methods. Overall ATP content was reduced by approximately 50% in 2F-Man treated seedlings compared to all other controls (Figure 3.10A). In contrast, F6P content was reduced in 2F-Man treated seedlings but after performing a Dunnett’s multiple comparisons, results were not statistically significant when compared to non-treated seedlings (NT) (p = 0.5112) (Figure 3.10B). Overall, these results indicate 2F-Man reduces energy metabolism in Arabidopsis, specifically in ATP production.

iv. Discussion and Future Directions:

In summary, 2F-Man severely reduces growth in Arabidopsis with an IC50 of 12

µM and an inhibitory time of 12 hours (Figure 3.1). The first hypothesis tested that 2F-

Man inhibits mannosylation and or fucosylation events in the cell wall. Man is metabolized to GDP-Man and through downstream enzymatic reactions is converted to

GDP-L-Fuc, a substrate for fucosyltransferases (Bar-Peled and O’Neill, 2011). Analysis of matrix polysaccharides demonstrated little to no change in Man or Fuc content and only slight changes in wall monosaccharides (Figure 3.2A and 3.2B). The remaining pellet after weak acid hydrolysis containing cellulose and lignin was hydrolyzed with

72% H2SO4 to release glucose from cellulose. Interestingly, glucose derivatization to 192

Figure 3.10: Quantification of ATP and F6P in 2F-Man treated seedlings. Seven-day- old Arabidopsis seedlings were grown under the following treatments: No Treatment (NT), 0.1% DMSO (DMSO), 100 µM D-Mannose (Man), and 100 µM 2F-Man (2F- Man). (A) ATP content was quantified in each treatment group, Dunnett’s multiple comparisons (*, p < 0.05). Error bars represent SEM (n = 9). (B) Fructose-6-phosphate (F6P) was quantified. Error bars represent SEM (n = 6)

193 alditol acetates revealed a reduction in crystalline cellulose content in 2F-Man treated seedlings (Figure 3.2). This led to 4 possible targets of 2F-Man toxicity; N-linked glycans, GPI anchors, ascorbate, and GIPCs. Analysis of N-linked glycan, GPI anchors, and ascorbate content were not negatively affected in 2F-Man treatment seedlings (Figure

3.3). In contrast, GIPCs monosaccharide head group content was modestly globally reduced. This observation included monosaccharides that are added into the GIPC head group before Man is added such as myo-D-inositol and Glc (in the form of D-glucuronic acid) (Figure 3.4). These decreases do not mimic the severe monosaccharide decreases found in GIPC mannosylation mutants gonst1-1 and gmt1-1 (Mortimer et al., 2013; Fang et al., 2016). Oddly, Man treatment had a modest negative impact in GIPC head group profiles. This suggests higher concentrations exogenous Man may have a negative impact in GIPC head groups composition.

Due the lack of a clear mechanism of 2F-Man growth inhibition, an EMS screen was performed to identify Arabidopsis mutants that are resistant to 2F-Man. Overall, 15 potential candidates were isolated, and their progeny were retested in 50 µM 2F-Man, with fmr6 and fmr11 displaying the most robust resistance to 2F-Man (Figure 3.5). Both fmr6 and fmr11 were backcrossed to Col-0 and F2’s segregation ratios for each line was assessed (Table 3.1). Resistant seedlings were pooled, and genomic DNA was extracted for whole genome resequencing. EMS induced mutations were mapped out and the highest frequency was found in chromosome 4 with the most likely candidate gene

HXK1 (Figure 3.6). Interestingly, both lines also exhibited resistance to 2F-Glc (Figure

3.7). 194

HXK1 is a bifunctional enzyme that both phosphorylates hexose sugars and acts as a glucose sensor in plants, similar to its yeast homolog HXK2 (Cho et al., 2006; Vega et al., 2016). To summarize fmr6 and fmr11 in relation to HXK1, both lines grown on increasing concentrations of D-glucose were moderately less sensitive to glucose repression and exhibited increased germination ratios compared to Col-0 controls (Figure

3.8). These observations led to the hypothesis that 2F-Man may affect energy metabolism in Arabidopsis, where ATP is decreased in 2F-Man treated seedlings (Figure 3.9). These results collectively suggest that 2F-Man growth inhibition is partially dependent on

HXK1 function causing a reduction in ATP production and limiting energy availability for energetically expensive biological processes such as cell proliferation, growth, and expansion (Dingenen et al., 2019; Moore et al., 2003). Finally, high concentrations of

Man (15 mM) has been reported to inhibit Arabidopsis seed germination in a HXK mediated fashion (Pego et al., 1999). The difference in inhibitory concentrations between

Man and 2F-Man may be due to the potential increase in affinity of 2F-Man to HXK1 from the carbon-fluorine bond’s greater dipole moment compared to carbon-oxygen bond. The increased affinity of 2F-Man may simulate a “high” mannose concentration causing growth repression.

Man can be found in a diverse number of glycans, but in terms of downstream metabolic products, Man can only be converted to L-fucose, L-galactose, and ascorbate

(Bulley and Laing, 2016; Voxeur et al., 2011; Reuhs et al., 2004; Bonin et al., 1997). The caveat to this assumption is that free Man when phosphorylated to Man-6-phosphate can be converted to fructose-6-phosphate through phosphomannose isomerase (PMI) and enter either glycolysis or any other monosaccharide products (Maruta et al., 2008). This 195 pathway can be ruled out in 2F-Man growth inhibition because the 2-carbon cannot be oxidized due to the fluorine substitution. This leaves the metabolic products listed above where 2F-Man can be metabolized. If 2F-Man is metabolized to GDP-2F-Man and subsequently to GDP-2F-Fuc, the primary target would be RG-II fucosylation as previously described (Villalobos et al., 2015; Dumont et al., 2015). Additionally RG-II would be severely inhibited if GDP-2F-Man is metabolized to GDP-L-2F-Galactose as part of ascorbate biosynthesis where it has been reported L-galactose replaces L-fucose in

RG-II in the mur1 mutant (Reuhs et al., 2004; Höller et al., 2015). Evidence provided in this study does not support the inhibition of RG-II biosynthesis, one fucose content in the cell wall was minimally affected and ascorbate content was unaffected under 2F-Man treatment (Figure 3.2A and Figure 3.3C). There is a possibility that 2F-Man has mild effects on all glycans tested and collectively inhibit growth.

Characterizing 2F-Man in Arabidopsis presented an avenue of study that was not previously anticipated. It is clear the biological mechanism of 2F-Man growth reduction is not due to inhibition of any mannose containing glycan. Instead, we observed what appears to be a signaling effect centered around HXK1. Under the model of HXK1 mediated growth inhibition under 2F-Man treatment, 2F-Man binds HXK1 in a similar fashion to Glc or Man under high concentrations is then translocated into the nucleus and transcriptionally regulates gene expression associated with the reduced growth (Cho et al., 2006). It would be interesting to investigate the localization of HXK1 under 2F-Man treatment. Additionally, our screen demonstrated HXK1 as the primary protein involved in 2F-Man toxicity, it would be worth testing 2F-Man against known glucose insensitive

(gin) mutants related to phytohormone signaling such as gin1, gin4, gin5, and gin6 which 196 correspond to abscisic acid (ABA) biosynthesis, ethylene signaling, and ABA signaling

(Rognoni et al., 2007).

As mentioned previously, HXK1 homology in yeast HXK2 is also involved in glucose repression and the hxk2Δ is insensitive to glucose repression as the gin2-1 mutant

(Trumbly, 1992; Moore et al., 2003). In the following chapter we screened 2F-Man and other monosaccharide analogs for growth inhibition in Saccharomyces cerevisiae.

Several iminosugars, known glycosylhydrolase inhibitors, were included in this screen

(Nash et al., 2011).

197

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Chapter 4: 2-Deoxy-2-Fluoro-D-Mannose is a potent inhibitor of fungal growth.

i. Introduction:

As mentioned in chapter 3, 2-deoxy-2-fluoro-D-mannose (2F-Man) in Arabidopsis thaliana targets the sugar sensor HEXOSE KINASE 1 (HXK1). Under high D-glucose

(Glc) conditions, physiological and transcriptional changes occur that inhibit growth, chloroplast development, and germination rate (Dingenen et al., 2019; Kunz et al., 2015;

Cho et al., 2006; Moore et al., 2003). Interestingly, a similar phenomenon has been observed under high D-mannose (Man) concentrations in Arabidopsis (Pego et al.,

1999a). One piece of evidence to suggest HXK involvement in glucose-mediated growth repression is based on mannoheptulose, an inhibitor of HXK activity that mitigates Man- induced germination inhibition (Cho et al., 2006; Pego et al., 1999b). In S. cerevisiae, the homolog to AtHXK1, ScHxk2 has the same sugar sensing mechanism that has been heavily studied. Under high Glc conditions, a similar growth repression response is observed, and ScHxk2 transcriptionally represses of HEXOSE KINASE 1 (ScHxk1) and

GLUCOSE KINASE (ScGlk1) expression while increasing its own expression (Guez et al., 2001). Similar to AtHXK1, ScHxk2 acts a sugar sensor and transcriptionally regulates gene expression (Ahuatzi et al., 2004; Kunz et al., 2015; Dingenen et al., 2019; Herrero et al., 1998). Additionally, ScHXK2 serves as a negative regulatory component in programed cell death (PCD)/necrosis by interacting with mitochondria under normal conditions by resisting Reactive Oxygen Species (ROS) induced necrosis (Amigoni et al.,

2013; Kaeberlein et al., 2005). In contrast, AtHXK1 actively participates in PCD in the myo-inositol 1- (MIPS1) mutant. MIPS1 synthesizes myo-inositol from glucose-6- 213 phosphate. In the mips1 mutant, light-dependent lesions develop in leaves due to PCD.

The mips1/hxk1-2 double knockout suppresses the mips1 PCD in leaves (Bruggeman et al., 2015). Therefore, the mechanistic similarities in Arabidopsis and S. cerevisiae glucose repression represent a potential avenue to further elucidate the mechanistic basis of 2F-Man toxicity, assuming 2F-Man inhibits growth in S. cerevisiae in a similar fashion.

ScHxk2 (referred to as Hxk2 in the body of this chapter) under high Glc conditions, enters the nucleus, recruits Multicopy inhibitor of Gal gene expression 1 (Mig1), Mig2,

Resistant to glucose repression 1 (Reg1), Sucrose non-fermenting 1 (Snf1), Snf4, and

Galactose Metabolism 83 (Gal83) to transcriptionally repress a variety of genes. The current model of repression is based on repression of Sucrose 2 (SUC2) (Vega et al.,

2016). Mig1 and Mig2 are zinc finger transcriptional repressors that directly bind DNA and both interact with Hxk2 in its closed confirmation (Ahuatzi et al., 2004; Vega et al.,

2016; Lutfiyya and Johnston, 1996). Reg1 is a regulatory subunit in Reg1/Glycogen 7

(Glc7) phosphatase complex regulating the localization of Hxk2 maintaining it in the nucleus. Snf7 is a protein kinase that phosphorylates Hxk2 at Ser-14 signaling nuclear export to alleviate Hxk2 mediated repression (Fernańdez-García et al., 2012; Vega et al.,

2016). Snf4 and Gal83 are regulatory subunits of Snf1 kinase that serve both as an activating subunit and to maintain Snf1 in the nucleus, respectively (Corvey et al., 2005)

(Figure 4.1).

In this chapter, S. cerevisiae strain S288c was screened against 2F-Man. We also included several other fluorinated sugars, iminosugars, and known cellulose biosynthesis 214 inhibitors to test if any of these other compounds negatively impact yeast growth. This screen revealed that 2F-Man is a potent inhibitor of S288c growth with 2F-Glc and N- dodecyldeoxynojirimycin (ND-DNJ) (Discussed in Chapter 5) also negatively impacting growth. Utilizing the Yeast Deletion Collection to screen for genes involved with 2F-

Man toxicity, we identified two unique mechanisms that lead to total or partial 2F-Man resistance in S. cereivisiae. The first is glucose repression through Hxk2 and Reg1 gene deletions leading to complete resistance against 2F-Man. This mechanism is homologous to AtHxk1 point mutations found in fmr6 and fmr11 in chapter 3, and further suggests that mechanistic basis of 2F-Man toxicity relies on targeting the glucose repression pathway. The second is the Rim101 pathway, involved in responding to environment alkaline conditions and lipid toxicity (Rockenfeller and Gourlay, 2018; Subramanian et al., 2012; Lamb et al., 2001; Rockenfeller et al., 2018).

ii. Materials and Methods:

Strains used in this study:

Yeast strains containing known deletions were generously provided by Dr. Sue

Liebman: hxk2Δ::KanMX4, reg1Δ::KanMX4, snf1Δ::KanMX4, snf4Δ::KanMX4, gal83Δ::KanMX4, tos3Δ::KanMX4, elm1Δ::KanMX4, hxk1Δ::KanMX4, mig1Δ::KanMX4, mig2Δ::KanMX4, sak1Δ::KanMX4, suc2Δ::KanMX4, rim101Δ::KanMX4, rim21Δ::KanMX4, rim20Δ::KanMX4, rim13Δ::KanMX4, rim9Δ::KanMX4, rim8Δ::KanMX4, snf7Δ::KanMX4, and dfg16Δ::KanMX4. The pooled homozygous diploid Yeast Deletion Collection (YDC) was obtained from ThermoFisher.

The wild-type strain used for initial screens and subsequent controls, S288c were used. 215

Figure 4.1: Current model of glucose repression mediated by Hxk2.

216

Automated growth curve analysis:

Saccharomyces cerevisiae strain S288c or mutants isolated form the yeast deletion collection were propagated in solid YPD(s) media (1% Yeast extract [w/v], 2%

Peptone [w/v], 2% dextrose [w/v], 1.5% agar [w/v]) at 30 ºC. Single colonies were inoculated in 10 mL of liquid YPD (exclude 1.5% agar), which was grown for 16 hours in a 30 ℃-shaking incubator. The resulting saturated culture was diluted 100-fold in 1 mL YPD aliquots. Appropriate treatments were added here at specified concentrations in these 1 mL aliquots. Fluorinated sugars were obtained from Carbosynth (Berkshire,

United Kingdom) and iminosugars were obtained from Toronto Research Chemicals

(TRC) (Ontario, Canada). For automated growth curves, two hundred microliters of each diluted culture were transferred in triplicate to a 100-well Honeycomb™ plate (Growth

Curves USA, Piscatawny, NJ). Growth data was collected using a Bioscreener C2

(Growth Curves USA, piscatawny, NJ) using the following parameters: 30 ℃ incubation, record OD600nm every 5 minutes, shake 30 seconds before reading for a total of 18 hours.

GraphPad Prism v8 was used to analyze data and calculate Area Under the Curve (AUC) values. Known deletion knockouts were assayed in a similar manner but cultured in 5 mL of YPD.

Spot assays for S288c and all other deletion knockouts were propagated as described above and were grown to saturation in 5 mL of YPD. OD600nm was recorded for each culture and diluted to indicated concentrations in 1 mL of YPD. Ten microliters of each culture were spotted in duplicate in solid YPD with indicated treatments. Plates were placed in a 30 ℃ incubator for 18 hours. Plates were scanned on benchtop scanner. 217

Screening Yeast Deletion Collection and Bar-seq preparation:

One hundred milliliters of YPD + G418 was inoculated with a culture of homozygous Yeast Deletion Collection (YDC) and placed in a 30 ℃-shaking incubator for 18 hours. The resulting saturated culture was distributed into two sterile 50 mL tubes and centrifuged at 2,000 x g for 5 minutes. Cell pellets were resuspended in 5 mL of YPD

+ 15% [v/v] glycerol and 200 µL aliquots were distributed to sterile 0.5 mL tubes and stored at -80 ℃. Ten milliliters of YPD + 500 µg/mL G418 were inoculated with a homozygous culture of YDC and grew to late log phase in a 30 ℃-shaking incubator.

One hundred microliters of saturated culture were transferred into three 15 mL tubes containing 10 mL of YPD + 0.1% [v/v] DMSO and three 15 mL tubes containing 10 mL of YPD + 100 µM 2F-Man and grown for a total of 20 generations. Genomic DNA was extracted from 6.5 x 106 cells by centrifuging cells at 2,000 x g for 5 minutes. Cells were resuspended in 100 µL of 0.2 M lithium acetate + 1% [w/v] SDS and heated at 70 ℃ for

5 minutes. Five hundred microliters of 70% ethanol were added and centrifuged at 15,000 x g for 3 minutes. The resulting supernatant was discarded. Pellets were resuspended in

100 µL of sterile water, and cellular debris was removed by centrifugation at 15,000 x g for 30 seconds. The resulting supernatants contained genomic DNA, and 1 µL was used in following PCR reactions. Phusion polymerase (Thermofisher) was used for amplification of Up and Down tags using primers listed in Table 4.1. PCR cycling conditions used are as follows: 1 x [95 ℃ 5 minutes], 35 x [95 ℃, 30 sec; 55 ℃, 30 sec;

72 ℃, 1:30 minutes], 1 x [72 ℃, 6:30 minutes; 4 ℃ ∞]. PCR products of Up and Down tags had lengths of 950 bp and 850 bp respectively, which were gel purified using

QIAquick ® gel extraction kit (Qiagen) using the manufacturer’s instructions. Up and 218

Down tags from each replicate were combined and submitted to the Nevada Genomics center for Bar-Seq.

Molecular cloning:

Ten milliliters of liquid YPD was inoculated with S288c and placed in 30 ℃- shaking incubator for 18 hours. The resulting saturated culture was diluted to 1.0 x 107 cells/mL in 1 mL YPD. Genomic DNA was extracted as described above and Hxk2 was cloned using primers listed in Table 4.1. PCR reactions were performed using cycling conditions as follows: 1 x [95 ℃ 5 minutes], 35 x [95 ℃, 30 sec; 55 ℃, 30 sec; 72 ℃,

2:30 minutes], 1x [72 ℃, 6:30 minutes; 4 ℃ ∞]. PCR products were separated on a 1%

[w/v] agarose gel (ThermoFisher). The band of 1461 bp corresponding to the size Hxk2 gene was excised and purified using QIAquick® Gel Extraction Kit (Qiagen) following manufacturer’s instructions. Purified HXK2 DNA was cloned into pENTR-D-TOPO

(Invitrogen) following manufacturer’s instructions and resulting plasmid was transformed into OneShot® TOP10 Chemically Competent E. coli cells (Invitrogen). Cells were spread on and solid Luria Broth (LB) + 50 µg/mL Kanamycin and placed in a 37 ℃ incubator for 18 hours. Several resulting colonies were picked and inoculated 10 mL of

LB + 50 µg/mL Kanamycin and placed in a 37 ℃ incubator for 18 hours. Cultures were centrifuged at 3,500 x g for 15 minutes at 4 ℃ and extracted Hxk2-pENTR plasmid using

QIAquick® Spin Miniprep kit (Qiagen). The Hxk2-pENTR plasmid was sequence verified by the Nevada Genomics center. Hxk2 was cloned into destination vector pAG426GPD-EGFP-ccdB using an LR clonase (Invitrogen) kit followed by transformation into DH5α and selection on solid LB + 100µg/mL Ampicillin. 219

Table 4.1: Primer sequences used in this study

Primer Forward 5’ to 3’ Reverse 5’ to 3’ Sets

Up-tag GCTATCTGCATTAACCCTCA TATTCATTCGTGATTGCGCC barcode CTAAAGGGGATGTCCACGA GGTCTCT

Down-tag GTTACTCACCACTGCGATCC GCTATCTGCATTAACCCTCA barcode CTAAAGGGCGGTGTCGGTCT CGTAG

HXK2 CACCATGGTTCATTTAGGTC AGCACCGATGATACCAACG CAAAA GA

220

Ten milliliters of LB + 100µg/mL Ampicillin was inoculated with resulting colonies and grown for 18 hours at 37 ℃. Plasmids were extracted from each culture as described above.

Yeast strain hxk2Δ was made competent by propagating on solid YPD media, and a colony was inoculated in 10 mL of liquid YPD and grown to saturation in a 30 ℃- shaking incubator. OD600nm was recorded and diluted to OD600nm of 0.2 in 25 mL of YPD.

The culture was incubated in a 30℃-shaking incubator until an OD600nm of 0.6 was obtained then the culture was centrifuged at 2,000 x g for 5 minutes. The resulting pellet was decanted and washed with 10 mL of sterile water and centrifuged again at 2,000 x g for 5 minutes. The cell pellet was resuspended in 1 mL of 100 mM lithium Acetate + 1 x

TE [10 mM Tris-HCl pH 8.0, 1 mM EDTA] buffer and transferred to a 1.5 mL tube. The cell suspension was then centrifuged again at 2,000 x g for 5 minutes, decanted, and resuspended in 500 µL of 100 mM lithium acetate + 1 x TE buffer. Yeast transformation was carried out by combining 10 µL Salmon sperm DNA (Carrier DNA) (Trevigen), 10

µL of pAG426GDP-EGFP-Hxk2 (5,000 ng), 100 µL competent hxk2Δ, 480 µL 50%

[v/v] sterile Poly Ethylene Glycol (PEG), 60 µL 10 x TE [100 mM Tris-HCl pH 8.0, 10 mM EDTA], 60 µL 1 M lithium Acetate, and 75 µL DMSO. Cells were incubated at 42

℃ for 30 minutes. Cells were then centrifuged at 5,000 x g for 30 seconds and decanted.

Cells were resuspended in 500 µL of 1 x TE and centrifuged at 5,000 x g for 30 seconds.

This process was repeated an additional 2 times. Cells were resuspended to a final volume of 200 µL in 1 x TE and spread on Synthetic dropout (-) uracil (SD-URA) plate

(for 1 L: 1.7 g Yeast Nitrogen Base-NaOH pH 6.15, 5.0 g ammonium sulfate, 20.0 g dextrose, 1.9 g synthetic dropout – uracil amino acid mix (Invitrogen), and 15.0 g Agar). 221

Plates were placed in a 30 ℃ incubator for 72 hours. The resulting colonies were cultured in liquid SD-URA and were tested for sensitivity to 100 µM 2F-Man on the Bioscreener, using wildtype (S288c) and hxk2Δ grown in liquid Synthetic complete (SC) (1 L: 1.7 g

Yeast Nitrogen Base-NaOH pH 6.15, 5.0 g ammonium sulfate, 20.0 g dextrose, 1.9 g synthetic dropout – uracil amino acid mix (Invitrogen), and 20 mg uracil) as controls.

Protein expression, purification, and protein assays:

For protein expression, Hxk2 was cloned into the destination vector pET60-DEST from pENTR-Hxk2 using LR Clonase as described above to produce an Hxk2 variant that was N-terminally tagged with Glutathione S-Transferase (GST) and C-terminally 6X

His-tagged. The resulting pET60-Hxk2 was transformed into E. coli Nico21 (DE3) cells.

One liter of LB + 100 µg/mL Ampicillin was inoculated with DE3 cells harboring the pET60-HXK2 construct and incubated in a shaking incubator at 37 ℃ until the culture reached an OD600nm of 0.6. The culture was then induced with 500 µL of 1 M

Isopropyl β-D-1-thiogalactopyranoside (IPTG) to yield a final concentration of 500 µM.

The culture was then incubated in an 18 ℃-shaking incubator for 18 hours. The culture was pelleted at 3,000 x g for 15 minutes at 4 ℃. Cell pellets were transferred to a sterile

50 mL tube and cells were resuspended in 20 mL of Glutathione Sepharose buffer (GSB)

(25 mM Tris-HCl pH 7.5, 150 mM NaCl, 10 mM MgCl2, 5 mM β-Mercaptoethanol, 1

Pierce protease inhibitor tablet). Cells were lysed by the addition of lysozyme (Sigma-

Aldrich) to a final concentration of 2 mg/mL, vortexed, and incubated on ice for 30 minutes. Samples were then sonicated for 10 seconds, rested on ice for 30 seconds, and repeated this process for a total of six times using a Branson Sonifier 450. Membranes 222 were separated by centrifugation at 100,000 x g for 1 hour at 4 ℃. The resulting supernatant was transferred to a new 50 mL tube and placed in ice.

One and a half milliliters of Glutathione Sepharose (GS) 4 fast flow resin was equilibrated with 10 mL of GSB for 10 minutes, then the excess buffer was removed.

Total protein was added to the resin, incubated for 1 hour at 4 ℃ on an orbital incubator at 25 RPM. In a room maintained at 4 ℃ constantly, a column apparatus was prepared and the total extract + resin was transferred to the column and allowed to flow through into a 50 mL collection tube. Five hundred milliliters of GSB was used to wash the column and 20 µL of the flow through was collected for later use. GST-Hxk2-His was eluted with 15 mL GSB-KOH pH 7.5 + 10 mM Glutathione, and the flow through was collected in a new 50 mL tube. One milliliter of HisPur™ Ni-NTA Superflow Agarose was equilibrated with 10 mL of Protein wash buffer (PWB) (25 mM Tris-HCl pH 7.5,

150 mM NaCl, 10 mM MgCl2, 1 Peirce Protease inhibitor tablet) for 5 minutes and excess buffer was discarded. GST-Hxk2-His was added to the Ni-NTA resin and incubated for 5 minutes. Resin + Protein was transferred to a new column and allowed to flow through. Five hundred milliliters of PWB was used to wash the column and GST-

Hxk2-His was eluted with 15 mL of Ni-NTA Elution buffer (PWB supplemented with

300 mM imidazole) and collected in 1 mL fractions.

All fractions were concentrated in four Pierce Concentrator 10 MWCO centrifuge columns by transferring 500 µL of elution fractions and centrifuged at 12,000 – 15,000 x g for 10 minutes at 4 ℃. Following concentration, the buffer was exchanged with 500 µL protein buffer (25 mM Tris-HCl, pH 7.5, 150 mM NaCl, 10mM MgCl2) followed by 223 centrifugation at 12,000 x g for 10 minutes at 4 ℃. This process was repeated for a total of three times. A final 500 µL of protein buffer was added, and the protein was transferred to a 1.5 mL tube, mixed by inversion, and distributed into 200 µL aliquots in

0.5 mL tubes and placed on ice. The total protein concentration was determined using

Pierce™ BCA Protein Assay Kit following manufacturer’s instructions.

To assess kinase activity of GST-HXK2-His on 2F-Man, the ADP-Glo™ kinase assay kit (Promega) was used with slight modifications. Reactions were carried out in 25

µL: 2.5 µL 10x Kinase Buffer (250 mM MOPS-NaOH, pH 7.0, 100 mM MgCl2), 2.5 µL

10 mM Ultra-pure ATP, 2 µg GST-HXK2-His, 2.5 µL 10 mM 2F-Man or other hexose sugars, and brought up to 25 µL with sterile diH2O. Reactions were incubated at 30 ℃ for 30 minutes, then 25 µL of ADP-Glo Reagent was added and incubated at 25 ℃ for 40 minutes. Fifty microliters of Kinase detection reagent were added and incubated at 25 ℃ for 30 minutes. Samples were transferred to a 96-well white bottom plate, and luminescence readings were measured with a Spextramax M5 Microplate reader using an integration time of 0.5 seconds.

iii. Results:

Identification of 2-fluoro-2-deoxy-D-mannose as an inhibitor of S. cerevisiae growth:

Fluorinated monosaccharides were previously described to inhibit glycosylation events in cancer cells, plants, and in protozoans (Kizuka et al., 2017; Dumont et al., 2015;

De Macedo et al., 2001; Villalobos et al., 2015; Barthel et al., 2011; Zhou et al., 2017).

Additionally, fluorinated monosaccharides have been used in imaging to trace sugar metabolism in plants (Fatangare and Svatoš, 2016; Fatangare et al., 2015; Kim et al., 224

2016). Limited studies have investigated the impact of fluorinated monosaccharides on yeast growth, but these monosaccharide analogs have been used to assay fungal enzymes with fluorinated Glc and galactose analogs (Tan et al., 2014). For example, one unique study utilized an iminosugar, perfluoropropyl-DMDP, that reduced α-glucosidase activity in S. cerevisiae (Massicot et al., 2018). To assess potential growth inhibition of fluorinated monosaccharides in S. cerevisiae, we screened a variety of fluoro sugars against the S288c strain to identify monosaccharide analogs that impacted overall yeast growth. Included in this screen were a select handful of iminosugars as well as two known cellulose biosynthesis inhibitors Dichlorobenzyonitrile (DCB) and Isoxaben

(ISX). Growth curves for each treatment were integrated to calculate area under the curve. Overall, three compounds reduced total S. cerevisiae growth; 2-deoxy-2-fluoro-D-

Glucose (2F-Glc) (-25%), N-dodecyldeoxynojirimycin (-28%) and 2F-Man (-74%)

(Figure 4.2).

To further investigate 2F-Man-induced growth inhibition in S. cerevisiae, 2F-Man was retested at 100 µM with 0.1% DMSO as a solvent control, as well as 100 µM Man and 100 µM 2-deoxy-D-glucose (2D-Glc) for comparison. 2D-Glc was included in this study as it is a known monosaccharide analog that inhibits yeast growth at relatively high concentrations (6.09 mM or 0.10 g/100 mL)(Ralser et al., 2008). Structures of Man and

2F-Man are shown in Figure 4.3A. At 100 µM concentrations, 2F-Man severely inhibited growth in S288c. Interestingly, 2D-Glc did not display any growth inhibition, which can be explained by the fact that in the reported literature, inhibition is reported in the millimolar range (Ralser et al., 2008; McCartney et al., 2014; Biely et al., 1971) (Figure 225

Figure 4.2: Screening of fluoro sugars, iminosugars and known cellulose biosynthesis inhibitors against Saccharomyces cerevisiae strain S288c. Solvent controls DMSO and MeOH were at concentrations of 0.1%, all compounds except Dichlorobenzyonitrile (DCB) and Isoxaben (ISX) were tested at 100 µM. DCB and ISX were ran at 1 µM and 10 nM respectively. Tukey’s multiple comparisons (****, p < 0.0001). Error bars represent SEM n = 3.

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4.3B and 4.3C). However, this result demonstrated that 2F-Man is much more efficient at inhibiting yeast growth than the previously described 2D-Glc. The potency of 2F-Man was assessed against S288c by generating a dose response curve. Saturated cultures were diluted 100-fold in YPD aliquots containing several concentrations of 2F-Man ranging from 100 to 0.1 µM. Growth curves were integrated and AUC values for each growth curve were plotted against log[2F-Man] in molar concentration. An inhibitory concentration of 50% (IC50) was calculated at 52 µM (Figure 4.3D). Overall, these results suggest that 2F-Man is a potent inhibitor of S288c growth and S288c may serve as model organism to further elucidate growth inhibition in Arabidopsis as discussed in chapter 3.

Screening of Yeast Deletion Collection against 2F-Man:

In fungi, Man is primarily found in N-linked, O-linked glycans, and GPI anchors.

N-linked glycans in yeast are commonly poly mannosylated, with chain lengths in some species reaching to >50. O-linked glycosylation events in yeast proteins are exclusively

O-mannosylation events, and this is a fairly common post-translational modification in fungal proteins, targeting over 500 proteins in the yeast proteome (Jigami, 2008; Gemmill and Trimble, 1999; Neubert et al., 2016). With several potential glycan targets of 2F-Man growth inhibition, there may be other possible targets of 2F-Man growth inhibition that we have not anticipated similar to our observations in Arabidopsis (see Chapter 3).

Instead of analyzing every possible glycan individually, we decided to utilize the Yeast

Deletion Collection (YDC) to screen for novel resistant and hypersensitive deletion lines.

The YDC was developed in several strains but the homozygous diploid collection 227

Figure 4.3: Growth of S288c on 2F-Man in liquid cultures and solid media. (A) Structure of D-Mannose (left) and 2-deoxy-2-fluoro-D-Mannose (2F-Man) (B) S288c cultures were diluted to OD600nm of 0.1, 0.01, and 0.001 (left to right) and spotted on YPD(s) media supplemented with No Treatment (NT), 0.1% DMSO (DMSO), 100 µM Mannose (Man), 100 µM 2F-Mannose (2F-Man), and 100 µM 2-deoxy-Glucose (2D-Glc). (C) Area Under the Curve (AUC) measurements from growth curves of cultures grown under conditions listed above. Error bars represent SEM; (n = 2).

Tukey’s multiple comparisons (****, p < 0.0001). (D) The IC50 of 2F-Man was measured by exposing S288c cultures to indicated concentrations of 2F-Man. Error bars represent SEM; (n = 3).

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(strain BY4743) was utilized in this screen. Each deletion was generated by homologous recombination targeting the open reading frame in each gene. The recombination event replaces the open reading frame with a KanMX cassette flanked on both sides by a unique identifying DNA sequence referred to as the up and down tag (Shoemaker et al.,

1996; Brachmann et al., 1998) (Figure 4.4A). All of the deletion strains are pooled together and can be used to screen differential responses to added compounds. The utility of the YDC is that the populations can be grown in the presence of an inhibitor and deletion strains resistant to a particular inhibitor will become more abundant and will represent a larger portion of the population compared to a population grown without the inhibitor. Similarly, hypersensitive deletion lines will be underrepresented compared to untreated populations (Figure 4.4B) (Eason et al., 2004; Giaever et al., 2002). Illumina sequencing can be utilized to sequence the deletion-associated DNA barcodes, and the number of times a particular barcode is identified is used as a proxy for the abundance of a single deletion strain.

In this screen, pooled homozygous diploids were grown in triplicate in 0.1%

DMSO and 100 µM 2F-Man. After 20 generations of growth, genomic DNA was extracted from 6.5 x 106 cells then Up and Down tags were PCR amplified using primers listed on Table 4.1 (Figure 4.4A). PCR products were purified and submitted to the

Nevada Genomics Center for Bar-Seq analysis. Up and Down tags were quantified and traced to their corresponding gene (McMahon et al., 2011; Eason et al., 2004). The log2(Fold Change) in tag counts were plotted against -log10(p-value) in a volcano plot

(Figure 4.4). Gene deletions exhibiting the highest fold-change increase in abundance 229

230

Figure 4.4: Principle of Bar-Seq: (A) For every gene in the yeast genome, a unique construct containing a KanMX4 (G418) gene flanked by an up and down tags sequences with a short sequence corresponding to the target gene. Gene deletion is carried out by homologous recombination. (B) In a Bar-Seq experiment, a population of pooled bar-code deletion strains (individual colored Yeast cells) are grown in the presence or absence of a 2F-Man. Some bar-coded deletion strains may be resistant to the inhibitor, and become more abundant (blue cells), while some strains may be hypersensitive to the inhibitor and die off in the population (gold and light green cells). Genomic DNA from both populations can be extracted and enriched for bar- code sequences by PCR. The resulting DNA is sequenced and each bar code sequence is ratiometrically quantified by Illumina sequencing compared to untreated control populations. Bar code sequences that are more abundant reflect strains that are more resistant to the treatment due to their deletion, while less abundant strains are hypersensitive. (C) Ratiometrically quantified data is displayed as a volcano plot, with log2 (Fold-Change) vs –log10 (p-value). Vertical dotted lines represent the threshold for Fold-Change (FC) (6-fold), horizontal line represents p-value threshold (p < 0.05).

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were Hexose kinase 2 (HXK2) (Mortimer et al., 1989) and REG1 (Matsumoto et al.,

1983). Additionally, several gene deletions exhibiting increased sensitivity to 2F-Man were SNF1, SNF4, GAL83, Target of SBF 3 (TOS3), and Elongated Morphology 1

(ELM1). These results suggest the primary mechanism of 2F-Man growth inhibition is through glucose repression (Vega et al., 2016).

Evaluating resistant and hypersensitive deletion lines:

Hxk2 is a bifunctional enzyme that both phosphorylates hexose monosaccharides and functions as a Glc sensor (Trumbly, 1992). Under high Glc conditions, Hxk2 is dephosphorylated by Reg1/Glk7 and enters the nucleus through α-importin Karyopherin alpha homolog 60 (Kap60) and β-importin (Kap95), then regulates transcription in a

Mig1-dependent manner (Peláez et al., 2012; Ahuatzi et al., 2004; Vega et al., 2016).

Hxk2 is exported from the nucleus when phosphorylated at Serine-14 by the kinase Snf1-

P. Hxk2 is dephosphorylated by Reg1/Glc7, which promotes glucose repression

(Fernańdez-García et al., 2012; Gancedo and Flores, 2008). Genetic knockouts of HXK2 and REG1 have been previously reported to be insensitive to glucose repression

(Trumbly, 1992; Fernańdez-García et al., 2012).

To confirm Hxk2 and Reg1’s role in 2F-Man growth inhibition, the respective individual deletion lines, hxk2Δ and reg1Δ, were assayed against 2F-Man. Saturated cultures of S288c, hxk2Δ, and reg1Δ were diluted to OD600nm of 0.1, 0.01, and 0.001.

Diluted cultures were spotted in YPD(s) (no treatment) and YPD(s) + 100 µM 2F-Man and incubated at 30℃ for 18 hours. Overall, hxk2Δ and reg1Δ had identical growth between no treatment and 2F-Man treatment. In contrast, S288c growth was severely 232 inhibited (Figure 4.5A). To further test resistance, growth curves for S288c, hxk2Δ, and reg1Δ were generated under YPD and YPD + 100 µM 2F-Man. Area under the curve values were calculated and summarized in Figure 4.5B. Growth of reg1Δ was mildly reduced compared to wild-type yeast under no treatment, but its growth was unaffected in

2F-Man treatment. Growth in hxk2Δ was also unaffected by 2F-Man treatment (Figure

4.5B). These results suggest that 2F-Man toxicity is heavily dependent on the glucose repression pathway. These results are similar to observations discussed in chapter 3.

Many of the hypersensitive genes identified in the YDC screen, including SNF1, SNF4,

GAL83, TOS3, and ELM1, are also involved in regulating glucose repression. Snf1 is a protein kinase that is activated by activation-loop phosphorylation by Snf1 activating kinase 1 (Sak1), Elm1, or Tos3. Upon activation, Snf1 can phosphorylate Hxk2, promoting disassociation of the transcriptional repression complex, further signaling for

Hxk2 to be exported from the nucleus (Casamayor et al., 2012). The transcriptional repression complex consists of Mig1 and Mig2 that directly bind to DNA with Hxk2 interacting with Mig1 and Mig2. Snf1 interacts directly with Hxk2 and interacts with 2 regulatory proteins Snf4 and Gal83. Finally, as part of the complex, Reg1 interacts with

Hxk2 in the nucleus maintaining Hxk2 in a dephosphorylated state. These interactions were elucidated in the transcriptional repression of the Suc2 promoter (Ahuatzi et al.,

2004; Castillon et al., 2003; Corvey et al., 2005; Fernańdez-García et al., 2012; Vega et al., 2016; Herrero et al., 1998). Based on the current model of glucose repression, all of the hypersensitive deletions identified here are involved in alleviating glucose repression, suggesting that hypersensitivity to 2F-Man treatment may be due to the inability to reverse glucose repression. 233

234

Figure 4.5: Growth of resistant strains from Yeast Deletion collection. (A) Spot assays of S288c, hxk2Δ, and reg1Δ at OD600nm of 0.1, 0.01, 0.001 (left to right) on YPD(s) with and without 2F-Man. (B) Area Under the Curve (AUC) measurements from growth curves of S288c, hxk2Δ, and reg1Δ cultures. Error bars represent SEM; n = 4 – 5.

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Figure 4.6: Analysis of hypersensitive strains from Yeast Deletion collection. (A) Spot assays of hypersensitive deletion strains on YPD(s) with and without supplementation of 50 µM 2F-Man. (B) The IC50 of 2F-Man from each strain was measured by exposing cultures of each strain at the indicated concentrations of 2F-

Man. Error bars represent SEM; n = 3 – 5. IC50 values are shown in Table 4.2

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Table 4.2: IC50 Values of hypersensitive deletion strains

Strain Gene IC50 (µM)

S288c (WT) N/A 42 snf1Δ YDR477W 37 snf4Δ YGL115W 21 gal83Δ YER027C 38 elm1Δ YKL048C 30 tos3Δ YGL179C 39

237

(Rockenfeller and Gourlay, 2018; Lamb et al., 2001). Alkaline conditions or lipid distress is sensed by Rim21 and the three accessory proteins Rim9, Rim8, and Defective for filamentous growth 16 (Dfg16), causing the proteolytic complex ESCRT-III to form.

Rim21, Rim9, and Dfg16 contain transmembrane domains that are embedded in the plasma membrane. The ESCRT-III complex consist of Rim20, Snf7, and Rim13; the protease that cleaves the C-terminus of Rim 101 to initiate its transcriptional activation.

The activated Rim101 causes transcriptional regulation including the repression of

Negative regulator of glucose-repressed genes 1 (NRG1) (Figure 4.7). Interestingly,

NRG1 also mediates glucose repression and negatively regulates the alkaline pH response (Zhou and Winston, 2001; Park et al., 1999; Lamb and Mitchell, 2003). The

Rim101 pathway serves as a mechanism to respond to alkaline conditions and necrosis

(Sarode et al., 2011; Futai et al., 1999; Maeda, 2012; Nishino et al., 2015; Lamb and

Mitchell, 2003; Lamb et al., 2001; Rockenfeller et al., 2018).

To further expand our understanding of the genes involving glucose repression and the Rim101 pathway, relevant individual deletion strains were obtained and assayed for 2F-Man resistance. Additionally, hxk1Δ and suc2Δ were included in this growth experiment to determine if Hxk1 plays a role in 2F-Man toxicity and to assess loss of

Suc2 function in 2F-Man treatment. Both hxk2Δ and reg1Δ served as positive controls to

2F-Man resistance. Growth curves for each strain were generated under YPD and YPD +

100 µM 2F-Man and AUC values were calculated (Figure 4.8). Interestingly, the hxk1Δ strain exhibited no resistance to 2F-Man, suggesting that Hxk1 is not involved in 2F-Man toxicity. Additionally, suc2Δ displayed a similar growth as S288c under 2F-Man treatment conditions. Other strains tested, mig1Δ, mig2Δ, and sak1Δ did not display 238

Figure 4.7: Current model of Rim101 pathway activation in yeast by alkaline stress

239 resistance to 2F-Man toxicity, although it may be reasonable to hypothesize sak1Δ may be hypersensitive to 2F-Man similar to elm1Δ and tos3Δ. These results suggest there may be additional DNA binding proteins other than Mig1 and Mig2 involved in growth repression under 2F-Man treatment than are suggested in the current model of Hxk2 mediated glucose repression.

In the strains related to the Rim101 pathway, nearly all the deletion lines displayed resistance to 2F-Man toxicity except for rim8Δ. In the alkaline/lipid distress receptors; rim21Δ, rim9Δ, and dfg16Δ displayed nearly double the growth of S288c under

2F-Man treatment. Unexpectedly, the deletion of the cytosolic component of the receptor complex rim8Δ, was as sensitive to 2F-Man treatment as S288c. The tested deletion strains with genes encoding proteins part of the ESCRT-III complex; rim21Δ, rim13Δ, and snf7Δ displayed resistance to 2F-Man toxicity. Lastly, rim101Δ was confirmed to display resistance to 2F-Man treatment (Figure 4.8). Interestingly, all the deletion strains related to the Rim101 pathway displayed only partial resistance to 2F-Man treatment, in contrast to the hxk2Δ or reg1Δ deletion strains, which are components of the glucose repression pathway (Figure 4.5). These data suggest that compromising the function of the Rim101 pathway provides greater tolerance against 2F-Man toxicity compared to wildtype.

HXK2 complementation restores some sensitivity to hxk2Δ:

The results discussed thus far demonstrate that Hxk2 and Reg1 are master regulators of 2F-Man sensitivity (Figure 4.5) (Fernańdez-García et al., 2012; Vega et al.,

2016) . If deletion of Hxk2 leads to complete resistance to 2F-Man, then reintroducing 240

Figure 4.8: Analysis of other deletion lines involved with glucose repression, Rim101 pathway, and other genes (hxk1Δ and suc2Δ). Area Under the Curve (AUC) calculations from growth curves of each deletion strain under No Treatment (NT) and 100 µM 2F-Man (2F-Man). Each strain under 2F-Man were compared to S288c using Dunnett’s multiple comparisons (*, p < 0.05; ****, p < 0.0001). Error bars represent SEM (n = 2 – 4).

241

Hxk2 should restore sensitivity to 2F-Man. Here, the Hxk2 gene was cloned into pAG426GDP-EGFP-ccdB from the Advanced Yeast Gateway Vector kit (Addgene) to yield pAG426GDP-EGFP-HXK2. The hxk2Δ strain was made competent as described in

Materials and Methods and transformed with the Hxk2 construct generating hxk2Δ:pAG426GDP-EGFP-HXK2. Colonies were selected and retested for 2F-Man sensitivity in the Bioscreener as described in Materials and Methods (Figure 4.9).

Unexpectedly, the hxk2Δ:pAG426GDP-EGFP-HXK2 only displayed a mild increase to

2F-Man sensitivity. This may be a result of the EGFP tag linked in the N-terminus of Hxk2, where the Ser-14 regulatory site is located. Based on the observed growth phenotype it is likely that Snf1 can phosphorylate EGFP-Hxk2, but also possible that the phosphatase complex Reg1/Glc7, being a larger dimer, is unable to remove phosphate from Ser-14 due to steric hinderance. Alternatively, EGFP may sterically hinder the Mig1 binding region extending from Lys-6 to Met-15 (Herrero et al., 1998). Therefore, C-terminal fusions of

Hxk2 should be tested in similar assays in the future.

HXK2 phosphorylates 2F-Man:

As a bifunctional enzyme, Hxk2 mutants containing truncations that remove catalytic activity or regulatory function have been generated to investigate physiological activities under different carbon sources and in high glucose conditions. The

478 485 hxk2ΔK A -S304F-without catalytic activity (hxk2Δwca), exhibits no catalytic activity, but still participates in glucose repression as it still interacted with Mig1 in the nucleus under high Glc conditions. On the other hand, hxk2ΔK6M15 without regulatory function

(hxk2Δwrf) has no interaction with Mig1, but still phosphorylates Glc (Peláez et al., 2010). 242

Figure 4.9: Analysis of Hxk2 rescue against 2F-Man. S288c, hxk2Δ, and pAG426GDP-EGFP-Hxk2. Growth measurements were taken for a total of 24 hours. Each strain was grown in No Treatment (NT) and 100 µM 2F-Man (2F-Man). Error bars represent SEM (n = 3 – 6).

243

Glc, the same concentration used in our standard YPD media. This observation suggests that under our experimental procedures, a certain ratio of Hxk2 protein resides within the nucleus and an equilibrium of cytosolic and nuclear localized Hxk2 is maintained. Both hxk2Δwrf and hxk2Δwca were screened against 2D-Glc to elucidate the role of Hxk2 in 2D-

Glc toxicity. The hxk2Δwca displayed resistance to 2D-Glc, but hxk2Δwrf was slightly more hypersensitive to 2D-Glc compared to wildtype (McCartney et al., 2014). These results suggested that the primary mechanism of 2D-Glc is through metabolically inhibiting enzymatic pathways dependent the 2-hydroxyl group for their biochemistry, such as glycolysis and pentose-phosphate pathway (McCartney et al., 2014). The hxk2Δwrf mutant may have increased sensitivity to 2D-Glc due to a larger ratio of cytosolic hxk2Δwrf compared to hxk2Δwca that may have wild-type ratios of cytosolic and nuclear localization. Interestingly, hxk2Δ:pAG426GDP-EGFP-HXK2 may somewhat resemble the hxk2Δwrf mutant as it is only partially sensitive to 2F-Man treatment due to the proximity of EGFP to the regulatory region of Hxk2. One aspect of 2F-Man toxicity yet to be explored is the kinase activity of Hxk2 on 2F-Man. 2F-Man likely inhibits Hxk2 activity or there may be downstream enzymatic process, such as generation of GDP-

Mannose or mannose by a GDP-Mannose Pyrophosphorylase or Phosphomannose isomerase 40 (Pmi40), respectively (Lin et al., 2001; Hashimoto et al., 1997).

To test this hypothesis, Kinase activity was assessed from recombinant Hxk2 purified from E. coli as described in Materials and Methods. Recombinant Hxk2 kinase activity was assayed against Glc and 2F-Man using ADP-Glo™ kit. Two micrograms of recombinant protein were used and 1 mM concentrations of Glc and 2F-Man were used in each reaction. Reaction mixtures were incubated at 30℃ for 30 minutes and kinase 244 activity was quantified based on total ADP generated during the incubation. Overall, kinase activity was observed in both Glc and 2F-Man reactions and no activity was observed in (Figure 4.10). These results suggest 2F-Man does not inhibit Hxk2 activity in vitro. Furthermore, 2F-Man-6-phosphate may act as the small molecule that activates glucose repression.

iv. Discussion and Future Directions

To summarize, 2-deoxy-2-fluoro-D-mannose (2F-Man) is a potent inhibitor of S. cerevisiae growth (Figure 4.3). Additionally, 2F-Man is over 60x more potent than a previously described monosaccharide analog inhibitor 2-deoxy-D-glucose (2D-Glc) with the concentration of 2D-Glc needed to resemble 2F-Man growth inhibition at 100 µM is

6.09 mM (0.10 g/100 mL) (McCartney et al., 2014). Furthermore, the IC50 of 52 µM is possibly the optimal potency of 2F-Man. Unlike 2-deoxy-2-fluoro-L-fucose discussed in chapter 2 (Villalobos et al., 2015), the potency of 2F-Man likely cannot be enhanced by acetylation of hydroxyl groups as observed in 1,3,4-triacetly-2-deoxy-2-fluoro-L-fucose.

This hypothesis is derived from the loss of growth inhibition in S288c between 2F-Glc and Ac4 2F-Glc (Figure 4.2). Using the Yeast Deletion Collection (YDC) to screen for gene deletions that contribute to resistance and hypersensitivity to 2F-Man toxicity revealed two unique processes that play a role in 2F-Man growth inhibition; glucose repression and the Rim101 pathway (Figure 4.3). Glucose repression is induced under high Glc conditions, where Hxk2 is phosphorylated as Ser-14 and enters the nucleus and causing transcriptional regulation and represses growth (Herrero et al., 1998). The

Rim101 pathway is a response to increases in pH or lipid distress to acclimate cells to 245 alkaline conditions by expressing multiple ion pumps or stimulate necrosis respectively.

Rim101 is activated by Rim13-mediated C-terminal cleavage of Rim101, and the truncated Rim101 enters the nucleus and transcriptionally regulates NRG1 and SMP1

(Lamb and Mitchell, 2003; Futai et al., 1999; Lamb et al., 2001; Rockenfeller and

Gourlay, 2018). The Rim101 pathway is also involved in yeast cell wall assembly, and rim21Δ cell walls were compromised leading to hypersensitivity to ion stress, caffeine and zymolyase (Castrejon et al., 2006). Assessing the two resistant deletion knockouts related to glucose repression hxk2Δ and reg1Δ (Guez et al., 2001; Fernańdez-García et al., 2012), displayed complete resistance to 2F-Man toxicity (Figure 4.5). The deletion strains snf1Δ, snf4Δ, gal83Δ, tos3Δ, and elm1Δ corresponding with genes involved with alleviating glucose repression (Celenza et al., 1989b, 1989a; Corvey et al., 2005;

Casamayor et al., 2012; Castillon et al., 2003) were hypersensitive to 2F-Man (Figure

4.6). Furthermore, complementing hxk2Δ with pAG426GDP-EGFP-Hxk2 partially restores sensitivity to 2F-Man (Figure 4.9). These data suggest that the key mechanism to

2F-Man toxicity lies in the glucose repression mechanism in S. cerevisiae.

The role of the Rim101 pathway appears to assist in increasing 2F-Man toxicity. This hypothesis is based on the observation that all Rim101 pathway gene deletion strains except for rim8Δ displayed moderate resistance to 2F-Man (Figure 4.8) and the observation that Hxk2 under high glucose conditions increases its own expression while repressing Hxk1 and Glk1 (Herrero et al., 1995). In this model, Hxk2 localizes to the nucleus to repress Hxk1 and Glk1expression limiting available Hexose kinases available to generate glucose-6-phosphate (G6P) overtime. The decreasing G6P limits glycolysis and ultimately reduces ATP content available to use for growth. The Rim101 pathway 246 comes into play when endogenous H+-ATPases at the plasma membrane begin to run low on available ATP causing the media reach a neutral or alkaline pH. This situation activates the Rim101 pathway and the expression of additional H+-ATPases further depleting available ATP that can be used in growth (Lamb et al., 2001). This scenario may explain why the deletion lines in the Rim101 pathway provide some resistance to

2F-Man toxicity. Interestingly, toxicity of 2D-Glc is not mitigated in the rim20Δ deletion strain as observed under 2F-Man treatment, suggesting that there are two different mechanisms of inhibition between 2D-Glc and 2F-Man beyond simply glucose repression. Furthermore, a study analyzing the catalytic activity or regulatory function of

Hxk2 contributing to 2D-Glc toxicity found the catalytic activity is critical to 2D-Glc toxicity (McCartney et al., 2014). This observation likely suggests that 2D-Glc toxicity is due to competitive inhibition of Phospho-glucose isomerase 1 (Pgi1) in glycolysis (Lin et al., 2001). It is unlikely that 2F-Man significantly inhibits glycolysis, because it does not contain the hydroxyl group at the second carbon to convert it to fructose-6-phophate by

Phosphomannose isomerase 40 (Pmi40) (Smith et al., 1992). In this instance, 2F-Man is unable to block Glc in the media from entering glycolysis. Additionally, the concentrations used in this study may also be too low to inhibit glycolysis compared to

2D-Glc. Alternatively, 2F-Man interaction with Hxk2 influences mitotic cell division, lifespan, and overall energy metabolism (Kaeberlein et al., 2005; Barbosa et al., 2016;

Amigoni et al., 2013) by limiting available Hxk2 in the cytosol and repressing HXK1 and

GLK1 expression. As a result of 2F-Man binding to Hxk2 and its phosphorylation, also limits Hxk2 interacting with mitochondria causing ROS-induced PCD (Kaeberlein et al.,

2005; Amigoni et al., 2013). 247

An interesting opportunity to utilize 2F-Man is in treating fungal diseases. The prevalent use of commercially available fungicides has given rise to Candida species that are multidrug resistant (Lockhart et al., 2017; Spinillo et al., 1999). 2F-Man is even more promising since it utilizes a sugar sensing mechanism necessary for pathogenesis to inhibit growth (Lockhart et al., 2017; Spinillo et al., 1999; Van Ende et al., 2019).

Additionally, Candida albicans dependence in the Rim101 pathway in pathogenesis would be an excellent target to exploit through 2F-Man toxicity by overloading the signaling pathway (Yuan et al., 2010).

248

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Chapter 5: Sphingolipids are required for cellulose deposition and cellulose synthase complex motility.

i. Introduction:

Cellulose is among the most abundant biopolymers on the planet, and this paracrystalline polysaccharide is a major component of plant cell wall extracellular matrices. Cellulose is synthesized at the plasma membrane of plant cells (Paradez et al.,

2006) by the cellulose synthase complex (CSC), which consist of multiple Cellulose

Synthase A (CESA) subunits that serve as the CSC catalytic subunits (Purushotham et al.,

2016; Cho et al., 2017). The CSC also contains accessory subunits such as the

KORRIGAN endoglucanase (von Schaewen et al., 2015; Lei et al., 2014), Cellulose

Synthase Interactive 1 (CSI1) (Gu and Somerville, 2010; Lei et al., 2012), the companion of cellulose synthase (CC) proteins (Endler et al., 2015), and the glycosylphosphatidylinositol (GPI) anchored protein COBRA (Borner et al., 2003; Li et al., 2013; Liu et al., 2013). Together, these subunits form a massive molecular machine that extrudes newly synthesized cellulose into the apoplast (Purushotham et al., 2020).

Live-cell imaging has revealed that the localization of CSCs is dynamic and complex.

Fluorescently labeled CSC subunits reside in the Golgi apparatus, in the Trans-Golgi

Network (TGN), in small vesicular compartments known as microtubule associated cellulose synthase compartments (MASCs) or Small CESA compartments (SmaCCs)

(Crowell et al., 2009; Gutierrez et al., 2009), and also as motile particles at the plasma membrane that move with constant velocities of approximately 250 nm/min (Paradez et al., 2006). Plasma membrane-localized puncta are thought to represent actively 259 synthesizing CSCs, and the rate of CSC motility has been correlated with cellulose biosynthetic output in multiple inhibitor and genetic studies (Paradez et al., 2006; DeBolt et al., 2007; Paredez et al., 2008; Gu and Somerville, 2010). Despite these insights, the mechanistic influences that control CSC motility and activity at the plasma membrane are not fully elucidated.

Recent work has suggested a functional relationship between cellulose biosynthesis and the synthesis of complex glycosylated sphingolipids (Mortimer et al., 2013; Fang et al., 2016). Sphingolipids are a unique group of lipids that play major structural roles in plasma membrane and endomembrane morphology as well as critical roles in programmed cell death, membrane trafficking, plant pathogen interaction, protein anchoring to the plasma membrane, fertility, and developmental processes (Ternes et al.,

2011; Gronnier et al., 2016; Tartaglio et al., 2017). Sphingolipids consist of a fatty acid

(FA) group with an acyl chain ranging from 16 to 26 carbons, an 18-carbon sphingosine base, commonly referred to as a Long Chain Base (LCB), and a structurally variable headgroup. In Arabidopsis, LCBs and FAs are used as substrates for the Ceramide synthase enzymes LONGEVITY ASSURANCE GENE ONE HOMOLOG 1 (LOH1),

LOH2, and LOH3 to synthesize ceramides (Ternes et al., 2011; Luttgeharm et al., 2015).

Ceramide FA moieties can be hydroxylated at the 2-position to form hydroxy ceramides

(hCers), and these sphingolipids often undergo further structural modifications, such as the addition of a glucose head group by the action of Glucosylceramide synthase (GCS)

(Msanne et al., 2015) to form glucosylceramides (GlcCers). Alternatively, hCer headgroups can be modified to generate Glycoinositol phosphoceremides (GIPCs)

(Rennie et al., 2014; Fang et al., 2016; Tartaglio et al., 2017). The elaborate headgroup of 260

GIPCs contain a phosphate, followed by an inositol residue bonded commonly in

Arabidopsis to glucuronic acid then mannose residues to generate a complete GIPC (Fang et al., 2016; Rennie et al., 2014). However, novel GIPC headgroup structures continue to be elucidated in Arabidopsis and other plant species (Ishikawa et al., 2018).

Small molecule inhibitors have served as useful tools to further understand many cellular processes and their intricate functions at the molecular level. Recent work has identified a subset of iminosugar compounds derived from 1-deoxynojirimycin (DNJ) that negatively impact growth and development in Arabidopsis and Eragrostis tef. The primary metabolic defect associated with these inhibitors was a consistent reduction in

GlcCer content, suggesting that this class of small molecule may inhibit the biosynthesis of GlcCers (Rugen et al., 2018). Iminosugars are small molecules that resemble naturally occurring monosaccharides with the oxygen in the pyranose ring substituted with a nitrogen group. Among the structurally simplest iminosugars are DNJ and its parent compound nojirimycin (Inouye et al., 1966), both of which are naturally occurring.

Historically, DNJ and derivatives such as N-butyl-1-deoxynojirimycin (NB-DNJ), N- nonyl-deoxynojirimycin (NN-DNJ) and miglitol have been used as treatment for

Gaucher’s disease and diabetes. Gaucher’s is a lysosomal storage disease causing an accumulation of glucocerebrosides in macrophages leading to multi organ dysfunction.

This disease is caused by an autosomal recessive mutation in β-glucocerebrosidase gene

1q21, where the gene product cleaves off D-glucose from glucocerebrosides, completing the catabolism of glycosylated sphingolipids. (Elstein et al., 2001). NB-DNJ has been utilized as an inhibitor of gangliosides biosynthesis, reducing its catabolic intermediate glucocerebrosides. Additionally, NN-DNJ has been utilized as a chemical chaperone to 261 facilitate proper folding of newly synthesized β-glucocerebrosidase (Butters et al., 2005;

Platt et al., 1994; Sawkar et al., 2002). In diabetes, several iminosugars such as DNJ and miglitol, have been utilized to inhibit amylases in the digestive track lowering the glycemic impact in food (Hughes and Rudge, 1994; Nash et al., 2011; Wardrop and

Waidyarachchi, 2010).

Here, we performed an orthogonal small-scale screen of DNJ analogs on Arabidopsis for their potential to inhibit root growth. The result of the screen revealed N-dodecyl-1- deoxynojirimycin (ND-DNJ) as the most robust inhibitor of root growth.

Sphingolipidomic analysis revealed substantial alterations in the sphingolipid profiles of

ND-DNJ-treated plants, including reductions in total GlcCers and GIPCs, as well as substantial increases in free LCBs commonly associated with programmed cell death

(Alden et al., 2011). Further analysis revealed that these perturbations in sphingolipid content led to the inhibition of CSC velocities and cellulose deposition, suggesting that sphingolipids are required for CSC motility.

ii. Materials and Methods:

General plant growth, maintenance, and drug treatments:

Arabidopsis thaliana Columbia (Col-0) seeds were sterilized in seed cleaning solution (30% [v/v] bleach, 0.1% [w/v] SDS) for 20 minutes at 25 ℃. The seed cleaning solution was removed, and seeds were washed 5 times in sterile water followed by incubation at 4 °C for 48 hours before use. Seeds were germinated on MS media (1/2 X

Murashige and Skoog salts, 10 mM MES-KOH pH 5.7, 1% [w/v] Sucrose, and 1% [w/v] phytoagar). N-Dodecyl-Deoxynojirymycin (ND-DNJ) and all other iminosugar 262 compounds screened in this study were purchased from Toronto Research Chemicals

(TRC) (Ontario, Canada), suspended in methanol to yield 100 mM stock solutions, and added to MS plates at the specified concentrations. N-[2-hydroxy-1-(4- morpholinylmethyl)-2-phenylethyl]-decanamide, monohydrochloride (PDMP) (Cayman

Chemicals) was suspended in DMSO to yield a 50 mM stock solution and added to MS plates at specified concentrations. Seedlings were grown under long day conditions (16- hours light/ 8-hours dark) at 22 ℃ vertically for 7 days. Seedling roots were straightened and scanned with a benchtop scanner, and root lengths were quantified using ImageJ. For assays involving dark grown seedlings, MS media without sucrose was supplemented with ND-DNJ or PDMP at indicated concentrations. Sterilized seeds were plated onto

MS media without sucrose and incubated in the light for 1.5 hours. Plates were then wrapped with aluminum foil and grown vertically under long day conditions at 22 ℃ for

5 days. Etiolated hypocotyl lengths were quantified using ImageJ.

Analysis of plant cell wall polysaccharides:

Alditol acetate analysis of cell wall matrix polysaccharides was performed as previously described (Villalobos et al., 2015) with minor modifications. Alcohol

Insoluble Residue (AIR) (5-10 mg) from 7-day-old light grown seedlings was de-starched by resuspending in 1.5 mL of 0.1 M Sodium Acetate-HCl pH 5.0. Samples were heated to 80 ℃ for 20 minutes, then cooled on ice for 5 minutes. Thirty-five microliters of

Amylase from porcine pancreas (50 µg/mL) (Sigma-Aldrich) and 17 µL of Pullulanase

(Sigma-Aldrich) was added, and samples were incubated for 18 hours in a shaking incubator at 37 ℃. Reactions were terminated by heating at 100 ℃ for 10 minutes. 263

Samples were centrifuged at 15,000 x g for 12 minutes, and the supernatant was decanted. Samples were washed with 1 mL of water, vortexed, centrifuged at 15,000 x g for 12 minutes and decanted 3 times. Finally, the samples were resuspended in 1 mL of

Acetone and evaporated to dryness at 50 ℃ under a stream of N2 gas. The resulting pellet was subjected to Alditol acetate analysis (See Chapter 2).

Crystalline cellulose content was quantified by Updegraff assay (Updegraff,

1969) with modifications. Five hundred microliters of Updegraff reagent ([8:1:2] [v/v/v]

Acetic acid: Nitric acid: water) were added to 10 mg of AIR material and heated at 100

℃ for 30 minutes. Samples were centrifuged at 14,000 x g for 15 minutes, and the supernatant was removed. Pellets were washed with 750 µL of water, the centrifugation was repeated, the supernatant was removed, and the samples were dried under a stream of

N2 at 80 ℃. One hundred and seventy-five microliters of 72% [v/v] H2SO4 was added, and the samples were incubated at 25 ℃ for 30 minutes. Samples were vortexed and incubated at 25 ℃ for an additional 15 minutes, then diluted with 825 µL of water. A 200

µL aliquot of each sample was transferred to a fresh 1.5 mL tube. Two hundred microliters of glucose standards were prepared using 1 mg/mL stock to yield 0, 10, 50,

100, and 200 µg. Four hundred microliters of freshly prepared Anthrone reagent (2%

[w/v] Anthrone in concentrated H2SO4) was added to each sample, followed by sample heating to 80 ℃ for 20 minutes, cooled to 22 ℃ and transferred 75 µL to a 96-well flat bottom plate. The absorbance at 625 nm was measured for each sample, and glucose content was calculated based on linear regression from glucose standards.

264

GlcCer targeted metabolomic analysis:

Arabidopsis seedlings were germinated in MS media without supplementation or containing 0.1% [v/v] methanol, 100 µM ND-DNJ, 0.1% [v/v] DMSO, 50 µM PDMP, or

1 µM DCB. Seedlings were grown for 7 days under long day conditions as described above, then transferred to a 15 mL tube, flash frozen in liquid nitrogen, and lyophilized using Labconco freeze dry system. Sphingolipid extraction and targeted lipidomic analysis were performed as previously described (Markham and Jaworski, 2007) at the

University of Nebraska Center for Plant Science Innovation.

Cellulose synthase complex and microtubule imaging:

Arabidopsis transgenic lines co-expressing GFP-CESA3 under its native promotor and mCherry-TUA5 were previously described (Gutierrez et al., 2009) and were obtained from the Ehrhardt lab at Stanford University. Four-day-old etiolated GFP-CESA3; mCherry-TUA5 seedlings were treated with 200 µL Water (NT), 0.1% [v/v] MeOH, 100

µM ND-DNJ, 0.1% [v/v] DMSO, or 50 µM PDMP (diluted in water). Seedlings were mounted as using vacuum grease supports between the slide and coverslip as previously described (Paredez et al., 2006). Images were taken using a CSU-W1 spinning disk head

(Yokogawa) mounted to an inverted Leica DMI-8 microscope equipped with a 100 x oil immersion objective (Apo total internal reflection fluorescence, N.A. = 1.47;Leica) and an iXon Life EMCCD camera (Andor Technology) controlled by VisiView software

(Visitron Systems). GFP fluorescence was excited with a 488 nm laser and emission was recorded with a 525 nm filter. Fluorescence of mCherry was excited with a 561 nm laser 265 and recorded with a 620 nm long-pass filter. CSC motility assays were performed by recording 5-minute time-lapse movies in 10 second increments.

All time-lapse recordings were processed using identical processing conditions in

Fiji (https://imagej.net/Fiji). Movies were corrected for drift using rigid body alignment with the StackReg plugin (Thévenaz et al., 1998). Time-lapses were further processed by performing a 4-frame walking average, background subtraction of 50 pixels, and then contrast enhancement (Thévenaz et al., 1998). Velocities were analyzed using the

“kymograph evaluation” plug-in of Fiji as previously described (Vellosillo et al., 2015).

Approximately 100 velocity measurements were collected from each cell, and cells from at least 6 – 10 seedlings were used to produce CSC speed distributions.

iii. Results:

N-dodecyl-deoxynojirimycin inhibits cell expansion in Arabidopsis thaliana:

Previous work in a number of organisms, including Arabidopsis and other plant species, has indicated that a select subset of compounds based on the unnatural nojirimycin compounds significantly impact plant growth and development (Rugen et al.,

2018). Based on our interest in identifying rationally designed inhibitors of plant cell wall glycosyltransferases as described in chapter 1 and 2 (Villalobos et al., 2015; Xia et al.,

2014), we screened a small panel of commercially available Deoxynojirimycin (DNJ) analogs that were largely orthogonal to a previous study (Rugen et al., 2018) for their ability to induce cell wall biosynthesis defects, including reduced root elongation and epidermal cell swelling. This screen included compounds listed in Figure 5.1 and Table

5.1 with additional compounds. To evaluate the effects of this inhibitor panel, 266

Arabidopsis seedlings were germinated in MS media supplemented with 100 µM test compound, grown for 7 days as described in materials and methods, and their root lengths were quantified (Figure 5.1 and Table 5.1). These experiments revealed that N- dodecyldeoxynojirimycin (ND-DNJ) (Figure 5.2A) effectively inhibited Arabidopsis root growth, while the remainder of compounds in the screen were largely ineffective in this assay. Compounds that a cause reduction in growth compared to untreated controls include N-alkyl DNJ compounds such as N-methyldeoxynojirimycin (-46%), N- ethyldeoxynojirimycin (-31%), N-butyldeoxynojirimycin (-28%), N- cyclohexylpropyldeoxynojirimycin (31%), and a hydroxy containing N-alkyl DNJ,

Miglitol (-29%).

To investigate the phenotypic defects elicited by ND-DNJ, we performed a series of more detailed growth assays. Consistent with the initial screening assays, 100 µM ND-

DNJ treatment inhibited primary root growth in Arabidopsis primary root growth by nearly 80% (Figure 5.2B and 5.2C) and inhibited Arabidopsis dark-grown hypocotyl elongation by 50% (Figure 5.2D). To measure the effective concentration range of this compound, a dose-response curve was constructed to measure the effect of various ND-

DNJ concentrations on primary root growth. Root lengths were quantified, and a dose response curve was generated (Figure 5.2E). The Inhibitory Concentration at 50% (IC50) for ND-DNJ was 21 µM. Overall, these results suggest ND-DNJ is a unique and effective

N-alkylated iminosugar-based inhibitor of Arabidopsis root growth. 267

Figure 5.1: Nojirimycin analog screen on Arabidopsis thaliana. Arabidopsis seedlings were grown for 7 days with MS media supplemented with 100 µM of each indicated compound. Root lengths were quantified using ImageJ software. No treatments control (Blue), Deoxynojirimycin of hexose sugars (Red), N-alkyl deoxynojirimycin analogs (Green), N-alkyl substituted manno and galacto nojirimycin analogs, all other nojirimycin analogs (Black). Error bars represent SEM; n = 38 – 178.

268

Table 5.1: Nojirimycin analogs screened against Arabidopsis Col-0 Compound M.W. Cat.No Mean SEM Significant n 1-deoxynojirimycin 163.17 D245000 23.05 ± 0.2897 ns 163 1-deoxymannojirimycin HCl 199.63 D240000 24.93 ± 0.5901 ** 38 1- deoxygalactonojirimycin HCl 199.63 D236500 22.24 ± 0.4045 ns 77 1-deoxyfuconojirimycin HCl 183.63 D236000 23.42 ± 0.393 ns 73 N-methyl-1- deoxynojirimycin 177.2 M297000 12.15 ± 0.3707 **** 158 N-ethyl-1- deoxynojirimycin HCl 227.66 E915000 16.03 ± 0.1805 **** 161 N-butyl-1- deoxynojirimyin HCl 255.74 B691000 16.18 ± 0.2435 **** 101 N-(n-nonyl)-1- deoxynojirimycin 289.41 N650300 20.34 ± 0.3891 *** 72 N-dodecyl-1- deoxynojirimycin 331.49 D494550 3.035 ± 0.133 **** 109 N-butyl-1- deoxymannojirimycin HCl 255.74 B690750 20.05 ± 0.3037 **** 101 N-(n-nonyl)-1- deoxygalactonojirimycin 289.41 N649900 24.4 ± 0.5115 ns 40 N-5-Carboxypentyl-1- deoxynojirimycin 277.31 C181200 20.2 ± 0.5276 **** 84 N-cylohexylpropyl-1- deoxynojirimycin 287.4 C988150 15.69 ± 0.2837 **** 87 N-(7-oxa-9,9,9- trifluorononyl)-1- deoxynojirimycin 345.36 O845150 22.36 ± 0.4428 ns 39 N-(7-oxydecyl)-1- deoxynojirimycin 305.41 O845000 18.43 ± 0.4658 **** 47 Miglitol 207.22 M344200 16.01 ± 0.2743 **** 71 D-manno-Y-lactan 177.15 M166000 23.18 ± 0.4998 ns 71 No Treatment control n/a n/a 22.6 ± 0.377 n/a 178 Mean root length (Mean) and SEM values are reported as millimeters. Dunnett's Multiple Comparisons test to No Treatment control (**, p < 0.01; ***, p < 0.001; ****, p < 0.0001). Cat.No correspond to Toronto Research Chemicals (TRC) catalog system. 269

Figure 5.2: N-Dodecyl-deoxynojirimycin inhibits growth in Arabidopsis. (A) The structure of N-dodecyl-deoxynojirimycin (ND-DNJ) is shown. (B) Arabidopsis seedlings were germinated and grown on control MS media (NT) or MS media supplemented with 100 µM ND-DNJ or 0.1% Methanol (MeOH) as a solvent control for 7 days in long day conditions (16 hour light/ 8 hour dark) and 5 days in the dark. Scale bars represent 10 mm. (C) Root lengths of control, 0.1% MeOH, and 100 µM ND-DNJ treated seedlings was quantified in ImageJ as described in Materials and Methods (Error bars represent SEM; n = 88 – 107). Tukey’s multiple comparisons (**** p < 0.0001). Hypocotyl lengths were also quantified for 5-day-old etiolated seedlings grown on MS medium without sucrose as described in Materials and Methods (Error bars represent SEM; n = 100 – 159) Tukey’s multiple comparisons (**** p < 0.0001). (D) Dose response curve of ND-DNJ on light grown seedlings.

The IC50 of ND-DNJ was measured by exposing 7-day old light-grown Arabidopsis seedlings to the indicated concentrations of ND-DNJ (Error bars represent SEM; n = 65 – 81).

270

Sphingolipid profiles are altered upon ND-DNJ treatment:

Previous work has suggested that molecules that are structurally similar to ND-

DNJ bind to the enzyme Glucocerebrosidase and serves as an effective drug treatment for

Gaucher’s disease as a substrate assisted chaperone in human cell lines (Nash et al., 2011;

Stirnemann et al., 2017). Additionally these compounds also act as inhibitors of

Glycosylated ceramide biosynthesis in mammalian systems (Platt et al., 1994; Butters et al., 2005). The ceramide analog N-[2-hydroxy-1-(4-morpholinymethyl)-2-phenyl ethyl]- decanamide (PDMP) has also been widely utilized as a GlcCer biosynthesis inhibitor to study vesicular trafficking defects associated with this molecule (Melser et al., 2010), and a recent screen of nojirimycin-based analogs, which did not include ND-DNJ, also suggested that nojirimycin-based molecules are inhibitors of GlcCer biosynthesis.

Importantly, PDMP and L-ido-AEP-DNJ were shown to produce similar phenotypic defects compared to ND-DNJ (Rugen et al., 2018). Therefore, we postulated that ND-

DNJ may inhibit GlcCer biosynthesis in a similar manner to these compounds.

To test this hypothesis, Arabidopsis seedlings were grown for 7 days in the presence of 100 µM ND-DNJ, lyophilized, and subjected to established sphingolipid extraction procedures described in materials and methods. Untreated seedlings and seedlings grown on 0.1% methanol served as negative controls and were processed in the same fashion. The resulting lipid extracts were then subjected to targeted lipidomic analysis to quantify the abundance of various sphingolipid metabolites, including free

LCBs, GlcCer, GIPCs, and hydroxyceramides (hCers). These experiments revealed ND- 271

DNJ treatment exhibited complex effects on sphingolipid biosynthesis. ND-DNJ treatment resulted in a 30% reduction in total GlcCer content compared to negative controls, with decreases primarily reflected in GlcCer containing d18:0 and d18:2.

Additionally, the hydroxy acyl chains associated with the d18:0 and d18:2 species were reduced globally (Figure 5.3A). In contrast, total GIPC content was minimally impacted by ND-DNJ treatment with the only GIPC species containing d18:1 LCBs were reduced

(Figure 5.3B). Total hCer content was not affected by ND-DNJ treatment, but the proportion of hCers containing d18:0 and d18:1 LCBs decreased and hCers containing t18 LCBs increased. There was a near 3-fold increase in h16:0 and a decrease in h20:1

(Figure 5.3C). ND-DNJ treatment also resulted in a near 10-fold increase in total free

LCBs with major increases in free d18:0, t18:0, and t18:1. Notably, many phosphorylated

LCBs were also elevated with d18:1-P, t18:0-P, and t18:1-P as the most prominent

(Figure 5.3D). These results partially support the hypothesis that ND-DNJ, like several nojirimycin related compounds, is an inhibitor of GlcCer biosynthesis, but also inhibits other glycosylated sphingolipids such as GIPCs (Platt et al., 1994; Nash et al., 2011;

Rugen et al., 2018). The structure of ND-DNJ may bind well to the active site of GCS functioning as a competitive inhibitor (Msanne et al., 2015). In addition, ND-DNJ may also bind INOSITOL PHOSPHOCERAMIDE SYNTHASE (IPCS) one of the first enzymes involved in GIPC biosynthesis (Wang et al., 2008). It appears hCer content in the cell is tightly regulated as the totals are relatively unchanged in ND-DNJ treated seedlings (Figure 5.3C). These data suggest, as excess hCers from the inhibition of

GlcCer and GIPC biosynthesis from ND-DNJ, endogenous ceramidases maintain hCer content relatively stable and the increase in free LCB accumulate (Li et al., 2015; Chen et 272

Figure 5.3: Sphingolipid analysis of ND-DNJ treated Arabidopsis seedlings. Seven- day-old Arabidopsis seedlings grown under MS media with no treated (NT), 0.1% Methanol (MeOH), and 100 µM ND-DNJ (ND-DNJ) were analyzed. Totals of each sphingolipid species are represented as bar graphs. Detailed list of long chain bases (LCB) and hydroxy fatty acids (hFA) are represented as heat maps demonstrating relative amounts of each species. (A) GlcCer content with LCB and hFA shown below. Error bars represent SEM; n = 4 – 5. (B) GIPC content with LCB and hFA shown below. Error bars represent SEM; n = 4 – 5. (C) hCer content with LCB and hFA shown below. Error bars represent SEM; n = 4 – 5. (D) Total free LCB and individual LCB species shown below, data points outside of defined gradient (purple) ND-DNJ; t18:1 = 43.63, t18:0-P = 10.38. Error bars represent SEM; n = 4 – 5.

273 al., 2015; Wu et al., 2015).

Sphingolipidomic profiles of PDMP and DCB treated seedlings:

Given that ND-DNJ and PDMP inhibit the CSC in a manner found in DCB treatment in Arabidopsis, it would be an interesting hypothesis to suggest DCB may have an impact in sphingolipid profiles. Given the limited evidence of how DCB inhibits the motility of CSCs in the plasma membrane (DeBolt et al., 2007; Tateno et al., 2016) including limited success of obtaining DCB resistant lines with one mutant line DH75 displaying a four-fold increase in resistance compared to wildtype (Heim et al., 1998;

Tateno et al., 2016). Based on CSC velocity data described in this chapter, we postulated that DCB may impact sphingolipid profiles in Arabidopsis leading to a mechanistic basis for perturbed CSC motility.

Using similar methods for sphingolipidomic analysis, Arabidopsis seedlings were germinated and grown for 7 days in MS media, MS media supplemented with 50 µM

PDMP, 1 µM DCB, and 0.1% DMSO as a solvent control. Samples were freeze dried and total sphingolipids were extracted (Markham et al., 2006). These resulting sphingolipid extracts were then subjected to targeted lipidomic analysis and sphingolipid species were quantified as in the ND-DNJ treated seedlings. Both PDMP and DCB had similar effects on a variety of sphingolipid species but to varying degrees. In agreement with the literature, PDMP treatment resulted in a 40% decrease in total GlcCer content (Yin et al.,

2010; Krüger et al., 2013). Surprisingly, DCB treatment also caused a 20% decrease in total GlcCer content. Further examination of the LCB and hFA composition of GlcCer species revealed that these species were reduced to similar extents across all LCB and 274 hFA chain lengths (Figure 5.4A). Interestingly, GIPCs were also altered by PDMP treatment, which to our best knowledge has not been reported in plants. PDMP treatment resulted in a 30% reduction in total GIPC content, and t18 species were preferentially reduced compared to d18 species. DCB treatment slightly decreased GIPC content with only t18:1 LCB species affected but, d18:0 species were increased nearly two-fold. These changes were reflected in hFAs with all being negatively affected by PDMP and DCB treatment except for h16:0 hFAs (Figure 5.4B). Total hCer content was not altered by

PDMP and DCB treatment. Upon investigating specific LCB species, PDMP treatment caused modest increases in d18:0 (43%) and t18:0 (43%) but was contrasted with modest decreases in d18:1 and t18:1. DCB had a prominent increase in d18:0. Investigating hCers revealed a near absence of h18:0 and h20:1 in both PDMP and DCB treated seedlings (Figure 5.4C). The largest contrast between PDMP and DCB treated seedlings was in the form of free LCBs. Here only PDMP treated seedlings accumulated free

LCBs, whereas DCB treated seedlings displayed large increases in phosphorylated LCBs such as d18:0-P and t18:0-P, but total LCB content was not affected (Figure 5.4D).

These data indicate that PDMP treatment not only inhibits GlcCer biosynthesis as previously suggested but also indicates that GIPC content and composition are negatively impacted by PDMP treatment. This result suggests PMDP not only acts as a

Glucosylceramide synthase (GCS) inhibitor (Krüger et al., 2013; Msanne et al., 2015), but also inhibits GIPC glycosylation events. Overall, DCB treatment induces changes in glycosylated sphingolipid profiles, specifically the most notable change was in GlcCers

275

Figure 5.4: Sphingolipid analysis of PDMP and DCB treated Arabidopsis seedlings. Seven-day-old seedlings grown under MS media with no treated (NT), 0.1% DMSO (DMSO), 50 µM PDMP (PDMP), and 1 µM DCB (DCB) were analyzed. Totals of each sphingolipid species are represented as bar graphs. Detailed list of long chain bases (LCB) and hydroxy fatty acids (hFA) are represented as heat maps demonstrating relative amounts of each species. (A) GlcCer content with LCB and hFA shown below. Error bars represent SEM; n = 4 – 5. (B) GIPC content with LCB and hFA shown below. Error bars represent SEM; n = 4 – 5. (C) hCer content with LCB and hFA shown below. Error bars represent SEM; n = 4 – 5. (D) Total free LCB and individual LCB species shown below. Error bars represent SEM; n = 4 – 5.

276 containing d18:2. These results suggests a potential connection with d18:2 GlcCers and

CSC motility.

ND-DNJ treatment alters cell wall polysaccharide composition profiles including cellulose and callose deposition:

Investigating the root tip morphology by quantifying the thickness of roots 0.25 mm up from the tip of the root indicated that 100 µM ND-DNJ treatment caused primary roots to swell by 30% compared to untreated and MeOH-treated seedlings (Figure 5.5).

This root swelling phenotype is often observed in conjunction with defects in plant cell wall biosynthesis or deposition, suggesting that the sphingolipid compositional change caused by ND-DNJ treatment may be linked to defects in cell wall polysaccharide deposition or biosynthesis (Brabham et al., 2014; DeBolt et al., 2007; Sabba & Vaughn,

1999; Tateno et al., 2016; Xia et al., 2014). Interestingly, previous reports have suggested that defects in glycosylated sphingolipid biosynthesis, such as the Arabidopsis gmt1 mutant, which is defective in GIPC mannosylation, also cause reductions in crystalline cellulose content, suggesting that cellulose deposition and glycosylated sphingolipid metabolism are functionally linked (Mortimer et al., 2013; Fang et al., 2016).

CSC delivery to the plasma membrane is dependent on MASC/SMACCs

(Crowell et al., 2009; Gutierrez et al., 2009) originating from the TGN. Other matrix polysaccharides, such as pectins and hemicelluloses, are also dependent on the TGN to transport these polysaccharides to the extracellular matrix (Kim et al., 2014). As previously mentioned, GlcCers are vital for Golgi morphology and vesicle trafficking

(Melser et al., 2010; Msanne et al., 2015). Therefore, we were interested in understanding 277

Figure 5.5: Root width measurements of ND-DNJ treated seedlings. Arabidopsis seedlings were germinated and grown in MS media (NT) or MS media supplemented with 0.1% DSMO (DMSO) or 100 µM ND-DNJ (ND-DNJ). (A) Root tips were imaged under a Keyence scope. Scale bars represent 100 µm. (B) Root diameter was quantified 0.5 mm up from the tip of the root using ImageJ, and data is represented as a box and whisker plot. Tukey’s multiple comparisons (****, p < 0.0001) n = 12.

278 whether perturbing GlcCer or other glycosylated sphingolipid synthesis in Arabidopsis would disrupt cellulose biosynthesis and matrix polysaccharide deposition.

To address this question, we analyzed cell wall matrix polysaccharide composition via alditol acetate assay (Villalobos et al., 2015) of Alcohol Insoluble

Residue (AIR) from seven-day-old seedlings treated with 100 µM ND-DNJ as well as seedlings treated with 50 µM PDMP, a known GlcCer inhibitor. This analysis revealed that cell wall-associated L-arabinose content increased nearly 2-fold, while the other five monosaccharides remained relatively unchanged under ND-DNJ treatment compared to untreated control and solvent controls. On the other hand, an increase in L-arabinose was not observed in PDMP treatment (Figure 5.6A). Additionally, AIR from ND-DNJ and

PDMP-treated seven-day-old seedlings was used to quantify crystalline cellulose content via an Updegraff assay (Updegraff, 1969; Bauer and Iba, 2014). This analysis demonstrated that seedlings grown under 100 µM ND-DNJ treatment exhibited a 25% reduction in cellulose content, and similar reductions were also observed in 50 µM

PDMP treated seedlings (Figure 5.6B).

Initially, glucose content was not quantified in the matrix polysaccharide fraction because seedlings were grown in light. Under these conditions, seedlings produce a minimal amount of starch which can contribute to the overall matrix polysaccharide glucose signal. Upon closer inspection, the total matrix polysaccharide glucose contents in ND-DNJ and PDMP treated seedlings were nearly 5-fold greater compared to controls.

This observation prompted further analysis of glucose content in ND-DNJ and PDMP treated seedlings. Using the remaining AIR tissue, starch was removed by treating it with 279

Figure 5.6: Cell wall composition in seedlings treated with ND-DNJ and PDMP. Alcohol Insoluble Residues (AIR) were prepared from Seven-day-old light grown Arabidopsis seedlings grown under non-treated MS media (NT), 0.1% Methanol (MeOH), 100 µM ND-DNJ (ND-DNJ), 0.1% DMSO (DMSO), and 50 µM PDMP (PDMP) as described in materials and methods. (A) AIR tissue was hydrolyzed under weak acid conditions and released monosaccharides were derivatized to Alditol Acetates. Error bars represent SEM; n = 4 – 5 biological replicates, n = 2 technical replicates per biological replicate. Dunnett’s multiple comparisons (**** p < 0.0001). (B) Approximately 10 mg of AIR tissue was subjected to Updegraff assay to quantify cellulose content. Glucose content was quantified using Anthrone reagent using a standard curve. Error bars represent SEM; n = 4 biological replicates, n = 3 technical replicates per biological replicate. Dunnett’s multiple comparisons (*** p < 0.001; **** p < 0.0001). (C) Glucose content from untreated section is derived from GC data obtained in in experiments in panel A. Glucose content was reanalyzed with approximately 10 mg AIR after treatment with amylase and pullulanase. Error bars represent SEM; n = 3 – 5 biological replicates, n = 1 – 2 technical replicates per biological replicate. Dunnett’s multiple comparisons (** p < 0.01; **** p < 0.0001).

280 porcine amylase and pullulanase overnight. Samples were centrifuged and the supernatant containing enzymes and soluble glucose was removed. The remaining material was subjected to alditol acetate assay. Glucose content in de-starched seedlings in ND-DNJ and PDMP seedlings were still elevated compared to untreated and solvent controls (Figure 5.6C). The simplest explanation is that ND-DNJ and PDMP causes a significant increase in starch accumulation by shuttling glucose into starch biosynthesis and reducing available glucose for cellulose biosynthesis. Alternatively, the increase in glucose content of ND-DNJ and PDMP treated seedlings may be due to amorphous cellulose. Typically in higher plants, cellulose microfibrils exist as a highly ordered structures, but can also exist in an amorphous form (Szymańska-Chargot et al., 2011).

Cellulose crystallinity can be altered in cesa mutants such as the temperature sensitive mutant radially swollen 1 (rsw1). The rsw1 corresponds to a mutation in CESA1 gene. A key observation in the rsw1 mutant is the reduced crystalline cellulose and elevated amorphous cellulose which would be digestable under weak acid hydrolysis (Arioli et al.,

1998). The altered sphingolipid profiles of ND-DNJ treated seedlings influences the dynamics of CSCs and CSC accessory proteins that do not alter the overall cellulose content but alters the ratio of crystalline and amorphous cellulose. The increases in glucose content in the matrix polysaccharides may be due to amorphous cellulose at varying sizes, where it is possible some cellulose polymers were short enough to be soluble upon α-Amylase and pullulanase treatment or the products used my not have complete selectivity for α1→4-glucose linkages and have some activity with β1→4- glucose linkages. A different hypothesis is that there may be other glucose containing polymers such as callose from a complex signaling. It is common to see callose 281 deposition increase under immune responses and inhibition of cellulose biosynthesis

(Bundy et al., 2016; Grison et al., 2019; Nickle and Meinke, 1998).

To test the potential of callose deposition as a contributor of glucose in the matrix polysaccharides, Arabidopsis seedlings were germinated and grown for 4 days in MS media. Seedlings were transferred to MS media, MS media supplemented with 0.1% methanol, and MS media with 100 µM ND-DNJ. Seedlings were treated for 24 hours then stained with analine blue (Schenk and Schikora, 2015). After de-staining, seedlings were mounted on a Keyence scope and imaged under a DAPI filter. Untreated and methanol treated seedlings displayed little to no callose deposition in the roots. In contrast, ND-DNJ treated seedlings had several puncta associated with callose deposition

(Figure 5.7). Overall, these results suggest callose deposition contributes to the increase in glucose in de-starched ND-DNJ treated seedlings from eliciting an immune response due to decreases in cellulose content.

ND-DNJ and PDMP treatment reduce CSC velocities:

The CSC actively produces cellulose while moving along the plasma membrane following cortical microtubules trajectories (Lei et al., 2012; Li et al., 2012; Lei et al.,

2013). Common cellulose biosynthesis inhibitors display two unique mechanisms of cellulose biosynthesis inhibition. Both mechanisms target the CSC by either slowing down the movement of complexes as observed with 2,6-dichlorobenzonitrile (DCB) or by removing the CSC from the plasma membrane as demonstrated with Isoxaben (DeBolt et al., 2007; Paredez et al., 2006). The observation of reduced cellulose (Figure 5.6B) and changes in sphingolipidomic profiles (Figure 5.3), suggest a correlation between cellulose 282

Figure 5.7: Callose staining of ND-DNJ treated seedlings. Four-day-old seedlings grown in MS media were transferred to MS media (NT), MS me supplemented with 0.1% methanol (MeOH), and 100 µM ND-DNJ (ND-DNJ) for 24 hours. Seedlings were stained with DAPI, washed and imaged under Keyence scope using a DAPI filter. Red arrow points to a stained callose. Left panels: DAPI filter; Right panels: Overlaid with bright field. Scale bar represents 50 µm.

283 biosynthesis and sphingolipid composition. Therefore, we postulated that sphingolipids play a role in cellulose deposition either by the mechanisms of cellulose biosynthesis inhibitors described above or through a delivery mechanism from the TGN where

GlcCers have been implicated in vesicle transport and organelle structure (Melser et al.,

2010; Msanne et al., 2015).

To test this, etiolated 4-day-old GFP-CESA3: mCh-Tua5 seedlings were treated with 100 µM ND-DNJ and 50 µM PDMP for an hour, followed by live-cell imaging of

CSC localization and motility using spinning disc microscopy. Single frames were taken in intervals of 5 seconds for a total of 5 minutes (Figure 5.8A). Velocities of puncta corresponding CSC containing GFP-CESA3 were quantified by generating Kymographs

(Figure 5.8B). The imaging yielded results indicating both ND-DNJ and PDMP function similar to DCB, where CSC velocities of ND-DNJ and PDMP treated seedlings were reduced by 60% and 40% respectively compared to untreated controls (Figure 5.8C).

These data suggest that the altered sphingolipid profiles of ND-DNJ and PDMP treatment reduces CSC motility as observed in DCB treatment.

iv. Discussion and future directions:

Overall, the results presented above indicate that N-dodecyl-1-deoxynojirimycin

(ND-DNJ) is a potent inhibitor of Arabidopsis thaliana root growth. Growth inhibition is the most severe in light grown seedlings, with an IC50 value of 21 µM compared to hypocotyl growth under 100 µM ND-DNJ treatment (Figure 5.2). Previous reports provided evidence that some alkylated iminosugar analog inhibit GlcCer biosynthesis as observed with N-5-(adamantane-1-yl-ethoxy)pentyl-L-ido-1-deoxynojirimycin or GlcCer 284

Figure 5.8: Quantification of Cellulose Synthase Complex (CSC) velocities upon ND- DNJ and PDMP treatment. (A) Videos were taken of 4-day old GFP- CESA3;mChTua5 seedlings under No Treatment (NT), 1 hour of 0.1% Methanol (MeOH), 1 hour of 100 µM ND-DNJ (ND-DNJ) treatment, 1 hour of 0.1% DMSO (DMSO), and 1 hour of 50 µM PDMP (PDMP). Frames were taken once every 5 seconds for five minutes. Time averages were acquired using ImageJ software using the Z-project (average intensity) plugin. Red lines indicate where kymographs in (B) were drawn. (B) Kymographs of no treatment, MeOH, ND-DNJ, DMSO, and PDMP indicate CESA velocities over the time course described in panel (A). Kymographs were drawn in the locations indicated by red lines in (A). Kymographs were generated using the MultipleKymograph plugin on ImageJ software using a linewidth of 1. (C) Violin plots showing distribution of CSC velocities around the mean under NT, MeOH, ND-DNJ, DMSO, and PDMP treatments. Average velocities (dashed lines) of each treatment are reported above, reported as mean +/- SEM with first and second interquartile range (dotted lines). Tukey’s multiple comparisons (**** P<0.0001), n = 523-590 velocity measurements across 6-8 cells.

285 hydrolase inhibition with NB-DNJ treatment (Rugen et al., 2018; Dai et al., 2020).

Sphingolipidomic analysis of ND-DNJ treated seedlings revealed a complex effect on several major sphingolipid species. GlcCer content was reduced in ND-DNJ treated plants, with GlcCer species containing LCB of d18:0 and d18:2 as the most impacted. In

GIPCs overall content is relatively unchanged but reductions were observed in d18:1 containing species. Most notably, free LCBs were substantially increased with t18:1 and t18:0-P displaying 43-fold and 10-fold increase respectively (Figure 5.3) with phosphorylated free LCBs implicated in programed cell death (Alden et al., 2011;

Ormancey et al., 2019).

Seedlings treated with ND-DNJ displayed root swelling and had roots with larger diameters, potentially indicative of cell wall deposition defects (Figure 5.5). Notably,

ND-DNJ treatment increased L-arabinose content compared to untreated and solvent controls. Updegraff assays to quantify cellulose, indicated a reduction in cellulose content in both ND-DNJ and PDMP treatment. These results suggest a correlation to sphingolipid composition particularly with GlcCers and GIPCs, with GIPCs biosynthesis connected with cellulose production described in previous studies (Fang et al., 2016; Mortimer et al., 2013). Interestingly, de-starched samples had increased glucose content that may be explained by the increase in callose deposition in ND-DNJ treated seedlings (Figure 5.7) or may result in fragmented non-crystalline cellulose. The reduction in cellulose content is likely attributed to reduction in CSC velocities as observed in spinning disk confocal microscopy of GFP-CesA3; mChTua5 etiolated seedlings treated with ND-DNJ and

PDMP (Figure 5.8). Due to ND-DNJ and PDMP resembling Cellulose biosynthesis inhibition of DCB treatment, we hypothesized that DCB may also have an impact directly 286 on sphingolipid metabolism. PDMP and DCB treated seedlings were subjected to a sphingolipidomic analysis (Figure 5.4). As previously described (Yin et al., 2010; Krüger et al., 2013), PDMP reduces GlcCer content, and DCB had a similar but attenuated effect, with d18:2 as the most impacted in both treatments. Both compounds affected GIPCs with decreases in t18:1 containing species. Interestingly, free phosphorylated LCBs were dramatically increased in DCB treatment but only mildly elevated in PDMP. This further suggest a connection between cellulose biosynthesis and sphingolipid composition.

The decreases in cellulose content via decreases in CSC motility upon ND-DNJ treatment was an interesting and unexpected observation. With both ND-DNJ and PDMP resembling the effects of DCB on CSC motility. Similar to DCB, ND-DNJ treated seedlings had increased callose deposition (DeBolt et al., 2007), a common symptom of patterned-triggered immunity (PTI) potentially triggered by Damage-associated molecular patterns (DAMP) (Howard, 1997; Kohorn et al., 2006; Jones and Dangl,

2006). Evidence presented in this chapter highlights the increases in callose deposition upon ND-DNJ treatment (Figure 5.7). Other effects of PTI includes increases in lignin deposition, reactive oxygen species (ROS) production, Jasmonic acid, ethylene, and salicylic acid triggered by MAPK signal cascade (Mine et al., 2017; Tena et al., 2011;

Denness et al., 2011; Gigli-Bisceglia et al., 2018; de Azevedo Souza et al., 2017). PTI can be caused by either Pathogen-associated molecular patterns (PAMPs) or by DAMPs, with DAMPS most likely resulting from ND-DNJ treatment (Franck et al., 2018). The most likely DAMP generated by ND-DNJ may result from cellulose oligomers resulting from decreased cellulose biosynthesis. Cellobiose has been previously described to elicit a signaling cascade that is similar to the well-known DAMP response to 287 oligogalacturonides, yet cellobiose did not elicit ROS production or callose deposition, although it did have a synergistic role with other elicitors, such as the PAMPs flg22 and chitooligomers (de Azevedo Souza et al., 2017). Cellobiose is likely a component of the

ND-DNJ response as it ND-DNJ treatment also elicits cell swelling suggesting a mechanical distortion due to weakened cell walls where THESEUS 1 (THE) plays a significant role (Hématy et al., 2007). It would be interesting to observe how Cell wall integrity (CWI) sensors such as THE1, a receptor like kinase that senses cellulose deficiency first described in the prc1-1 background (Hématy et al., 2007), respond to ND-

DNJ or PDMP. The the1-1 mutant mitigated the reduced growth observed in prc1-1, and has also been shown to be less sensitive to Isoxaben treatment compared to wildtype and deposit less ectopic lignin induced by Isoxaben (Merz et al., 2017; Denness et al., 2011).

An additional sensor for cellulose deficiency is STRUBBELIG (SUB) that functions independently of THE1. Interestingly, sub-9 mitigated the reduced growth phenotype of prc1-1 and had reduced ectopic lignin deposition under Isoxaben treatment (Chaudhary et al., 2020). It would be interesting to see how the the1-1 and sub-9 lines respond to ND-

DNJ or PDMP treatment. Would the loss of sensing mechanism would create the appearance of partial resistance to ND-DNJ treatment and how would this affect callose deposition?

As briefly mentioned above, cellulose crystallinity altered in the temperature sensitive mutant radially swollen 1 (rsw1). The rsw1 corresponds to a mutation in CESA1 gene

(Arioli et al., 1998). This mutant has been observed to produced less crystalline cellulose, but instead produced a glucose containing glycan found in matrix polysaccharides (Arioli et al., 1998). The glucose containing glycan is likely amorphous cellulose, which is more 288 digestable under weak acid hydrolysis. Two cesa mutants, cesa1aegeus and cesa3ixr1-2, contain less crystalline cellulose compared to wildtype (Harris et al., 2012). Interestingly, both mutants contain a point mutation in the C-terminal membrane region, A930V in cesa1aegeus and and T942I in cesa3ixr1-2. As a result of these mutations, have elevated CSC velocities (Harris et al., 2012). It appears there is a correlation with CSC velocities and degree of CSC crystallinity. In (Harris et al., 2012), they describe an elevated CSCs velocities reduces crystallinity, but our data suggest reduced velocities of CSCs in ND-

DNJ and PDMP impact cellulose crystallinity. CSC velocities might be regulated to fine tune the ratio between crystalline and amorphous cellulose. Interestingly, COBRA

(COB), is a CSC accessory protein that influences microfibril deposition and orientation of elongation (Roudier et al., 2002). There is no real direct evidence to suggest COB influences cellulose crystallinity. but cob mutants do display reduced crystalline cellulose and have been thought to influence crystallinity (Roudier et al., 2002). If we postulate

COB influences cellulose crystallinity, it may be dependent on CSC velocities to properly orient CSC microfibrils to crystalline microtubules. If CSC velocities are too high then,

COB cannot properly interact with cellulose microfibrils leaving them in an amorphous state. If the complex velocities are too slow on the other hand, there is not enough cellulose being displaced as the complex does not move across the membrane building up the amount of cellulose that does not interact with COB leading to increased amorphous cellulose. Alternatively, cellulose crystallinity may be self-assembling but requires a CSC velocity sweet spot to create favorable conditions for crystallinity to occur.

Alternatively, the changes in sphingolipid composition may potentially contribute to the PTI response and induce programed cell death (PCD) (Cuvillier, 2002; Alden et al., 289

2011; Gronnier et al., 2016). Previous work indicates that GIPCs are receptors for cytotoxins like Necrosis and ethylene-inducing peptide 1-like (NLP) with binding dependent on specific sugar composition in the GIPC headgroup (Lenar et al., 2017).

Mutants with alterations to GIPC headgroup structure, such as loss of mannose in the head group, had elevated levels salicylic acid, common of plant pathogen defense, along with reduced cellulose (Mortimer et al., 2013). The connection between GIPC composition and cellulose biosynthesis remains unclear but, there may be a relationship with plasma membrane organization and GIPCs that affect CSC motility. GIPCs have the ability, in a phytosterol dependent manner, to increase lipid packing at the plasma membrane (Mamode Cassim et al., 2019; Grosjean et al., 2015). Result presented in this study suggest GIPCs and GlcCers are important for CSC motility and it may be due to membrane organization surrounding the CSC. The CSC moves along the plasma membrane following cortical microtubules (Gutierrez et al., 2009), and it can be implied that the lipid nanodomain surrounding the complex contains higher concentrations of

GIPCs and GlcCers compared to other parts of the plasma membrane (Schneider et al.,

2016). This hypothesis may explain the decreases in cellulose with ND-DNJ and PDMP treatment, the altered sphingolipid profiles may make it too difficult for the complex to move across the plasma membrane. This hypothesis may also provide some reason as to how DCB inhibits cellulose production as well. It would be interesting to observe how other sphingolipid biosynthesis inhibitors such as Fumonisin B1 (FB1) (Luttgeharm et al., 2016) would alter CSC velocities as well as known GIPC biosynthesis mutants monocation-induced [Ca+2] 1 (moca1), gonst1, and gmt1 (Fang et al., 2016; Mortimer et al., 2013; Melser et al., 2010; Jiang et al., 2019). 290

An interesting aspect of a relationship between sphingolipids and cellulose deposition is through auxin homeostasis. Generally, auxin is laterally transported by influx transporters such as AUXIN TRANSPORT PROTEIN 1 (AUX1) or AUXIN

TRANSPORTER-LIKE PROTEIN (LAX) and efflux transporters such as PIN-FORMED

TRANSPORTERS (PIN), which are localized in basal and apical regions of the cell

(Lehman et al., 2017). Both the deposition and status of the cell wall influence PIN localization. For example the cesa3 mutants, je5 and regulator of pin polarity (repp), reversed the localization of PIN1 in the PIN::PIN1-HA;pin2 transgenic line from basal to apical in epidermal root cells. Additionally this study also demonstrated that PIN1 relocalization was also observed under DCB and isoxaben treatment, suggesting cellulose deposition influences PIN localization (Feraru et al., 2011). Furthermore, cellulose and pectin methylation status also regulate diffusion and clustering of PIN3 in membrane nanodomains (McKenna et al., 2019). These domains are classified as distinguishable submicron protein or lipid assemblies between 20 nm to 1 µm in size (Ott, 2017).

Inhibiting cellulose deposition with DCB or inhibiting methyl esterase activity on homogalacturonan with epigallocatechin gallate increased the diffusion of PIN3

(McKenna et al., 2019). Acknowledging DCB likely inhibits sphingolipid biosynthesis

(Figure 5.4), suggest sphingolipids may impact PIN localization. Previous studies demonstrated PIN1 is mislocalized into intracellular bodies in seedlings treated with FB1

(Yang et al., 2013). In addition, AUX1 and PIN1 are mislocalized in the loh1-2+/-/loh3-2-

/- double mutant, further supporting sphingolipid composition playing a role in PIN localization (Markham et al., 2011). The influence of cell walls and sphingolipid on PIN localization can be differentiated as PIN polarity in the plasma membrane is lost in cell 291 wall defects and delivery to the plasma membrane is reduced in sphingolipid defects.

Loss of proper auxin distribution often leads to loss of gravitropism, cell expansion, and cell division (Lehman et al., 2017). These cellular processes require dynamic redistribution of CSCs suggesting and continued production of cellulose (Chen et al.,

2018). One hypothesis suggests the altered sphingolipid composition in ND-DNJ, PDMP, and DCB treatment may disrupt auxin homeostasis leading to the reduction in CSC velocities. This hypothesis is probably not a likely outcome as CSC velocities were quantified in dark-grown hypocotyls, where cell expansion is not dependent on auxin homeostasis (Jensen et al., 1998). Although, we currently cannot completely rule out auxin homeostasis influence on cellulose deposition of CSC velocities.

In conclusion, ND-DNJ behaves as a glycosylated sphingolipid inhibitor and further emphasizes a functional relationship between GlcCers and GIPCs in CSC motility.

Through this investigation, a link between a classically known cellulose biosynthesis inhibitor, DCB, and status of GlcCers and GIPCs were established. One hypothesis to

ND-DNJ toxicity is that it acts as a competitive inhibitor of GCS and IPCS. It would be interesting to observe transgenic lines that over expression lines for GCS and or IPCS under ND-DNJ treatment. Would these overexpression lines have some innate resistance to ND-DNJ treatment? 292

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Chapter 6: Cellulose biosynthesis inhibitors induce suberin production in

Arabidopsis thaliana as part of the Cell Wall Damage response.

i. Introduction:

Cellulose is the most abundant biopolymer on earth, and it is the main load-bearing component of the plant cell wall. It is composed of a linear polymer of β-(1→4)-Glucose

(Glc) (Figure 1.4A) molecules arranged in a paracrystalline array consisting of 18 glucan chains (Figure 1.4B) (Cosgrove, 2005). Cellulose is produced by the cellulose synthase complex (CSC), consisting of multiple Cellulose Synthase A (CESA) proteins. The complex synthesizes cellulose at the plasma membrane and its catalytic activity pushes the complex along the plasma membrane following cortical microtubules as described in greater detail in Chapter 5 (Cho et al., 2017; Paradez et al., 2006; Purushotham et al.,

2016).

Cellulose biosynthesis inhibitors (CBIs) are a broad class of compounds that inhibit cellulose production. Although the molecular targets of these compounds remain unclear,

CBIs commonly impact CSC behavior through two unique mechanisms: a pausing mechanism in which CSC motility at the plasma membrane is arrested, or a removal mechanism in which the CSC is depleted from the plasma membrane upon treatment

(DeBolt et al., 2007; Tateno et al., 2016; Xia et al., 2014). There are a variety of small molecules that inhibit cellulose biosynthesis through one of these mechanisms. Isoxaben

(ISX), is a common CBI used to investigate the physiological, genetic, and biochemical responses of cellulose deficiencies (Heim et al., 1990; Duval and Beaudoin, 2009;

Denness et al., 2011; Hamann et al., 2009; de Azevedo Souza et al., 2017). ISX inhibits 307 cellulose through a removal mechanism, in which CSCs are depleted from the plasma membrane and sequestered into small CESA compartments (SmaCCs) or microtubule associated cellulose synthase compartments (MASCs), Trans-Golgi network vesicles that house CSCs and facilitate re-localization from one area of the plasma membrane to the other (Crowell et al., 2009; Gutierrez et al., 2009; DeBolt et al., 2007).

Dichlorobenzonitrile (DCB) is another common CBI, but unlike ISX, the mechanism of

DCB inhibition is to arrest CSC motility at the plasma membrane (DeBolt et al., 2007).

As the primary load-bearing component of the cell wall, it is highly probable that plants monitor the structural integrity of cellulose as a mechanism to sense cell wall integrity. Common responses to cellulose biosynthetic defects from CBI treatment or in cellulose deficient mutants include growth arrest, loss of anisotropic cell expansion, cell swelling, and ectopic lignin deposition (Caño-Delgado et al., 2003; Ha et al., 2019;

Fagard et al., 2000; Hu et al., 2018). Lignin is a polyphenolic polymer commonly deposited in secondary cell walls in vascular plants, but its deposition can be induced upon cell wall damage (CWD), such as cellulose deficiency (Gigli-Bisceglia et al., 2018;

Caño-Delgado et al., 2003). Lignin possesses several valuable properties as a defense polymer, including antimicrobial properties and preventing the diffusion of toxins and pathogens (Sattler and Funnell-Harris, 2013).

Interestingly, many of these physiological responses to cellulose deficiency, including ectopic lignin deposition, are facilitated by the THESEUS 1 (THE1) receptor-like kinase in Arabidopsis. Genetic lesions in Arabidopsis the1 result in partial phenotypic suppression of growth defects in the cellulose biosynthesis mutants prc1-1, eli1-1 and je5 308

(Hématy et al., 2007; Merz et al., 2017). Additionally, the1 mutants are also partially resistant to ISX treatment and mitigate ectopic lignin deposition (Merz et al., 2017).

CWD signaling mediated by ISX treatment includes a cascade of reactive oxygen species

(ROS) and Ca+2 signaling, both of which are required for ectopic lignin deposition. Co- treatment of Arabidopsis plants with ISX and an irreversible ROS inhibitor, diphenyleneiodonium (DPI), exhibited reduced ectopic lignin deposition. Similarly, co-

2+ treatment with ISX and LaCl3, a Ca influx inhibitor, also had reduced ectopic lignin deposition, suggesting that both ROS and Ca2+ transport are required for the early stages of cell wall damage response signaling (Denness et al., 2011). Interestingly, the connection between CWD induced by cellulose deficiency and lignin deposition may be attributed to mechanical distortion at the plasma membrane, as the turgor pressure inside the cell cannot be held in place causing cells to swell. In line with this hypothesis, the severity of ectopic lignin deposition can be reduced in ISX treated seedlings by co- treatment with an osmotic stressor, such as sorbitol. This treatment reduces the mechanical distortion in the cell and potentially mitigates ROS and Ca2+ signaling

(Hamann et al., 2009).

Suberin is a lipid-phenolic biopolyester that is impermeable to water and acts as a physical barrier against pathogens and toxins. Under normal developmental conditions, suberin is deposited between the primary cell wall and the plasma membrane of root endodermal and peridermal cells (Nawrath et al., 2013). This complex polyester consists of several aliphatic and aromatic constituents, including fatty acids (FAs), alkyl primary alcohols, bifunctional ω-hydroxy FAs, Dicarboxy FAs, glycerol, ferulic acid, and tyramine (Vishwanath et al., 2015). These monomers are polymerized together under the 309 primary cell wall. Based on Transmission Electron Microscopy (TEM) of suberin- lamellae cell walls consist of light and dark bands. The light bands are hypothesized to consist of aliphatic groups and the dark bands consisting of polyaromatics (Graça, 2015).

Currently, the assembly of these monomers remains controversial. Suberin plays several important roles as a barrier limiting water and nutrient transport resulting in tolerance to abiotic stresses such as drought and salinity. Additionally, suberin also serves as a barrier against microbiol pathogens (Franke and Schreiber, 2007; Doblas et al., 2017).

In the context of cell wall damage signaling, oligomers of common cell wall polymers

(e.g. pectin-derived oligogalacturonides) or oligomers of fungal cell wall polymers (e.g. chitin oligomers) are perceived as signaling molecules known as Damage Associated

Molecular Patterns (DAMPs) or Pathogen Associated Molecular Patterns (PAMPs), respectively. Recent work has demonstrated that cellobiose, a glucanase-associated degradation product of cellulose, also serves as a potential DAMP in Arabidopsis (de

Azevedo Souza et al., 2017). Treatment with cellobiose causes both pattern-triggered immunity (PTI) and results in the transcriptional induction of MYB41, a master transcriptional regulator of suberin biosynthetic genes (Kosma et al., 2014). As a result, plants treated with cellobiose exhibit enhanced resistance to P. syringae potentially due to the upregulation of suberin biosynthetic genes, but suberin deposition was not detected

(de Azevedo Souza et al., 2017). Interestingly, cellobiose elicits Ca2+ influx but does not elicit ROS production, suggesting that cellobiose may not elicit ectopic lignin deposition

(de Azevedo Souza et al., 2017). Inhibition of cellulose biosynthesis by CBIs would likely produce incomplete cellulose fragments including cellobiose, which upregulates 310 transcripts associated with suberin biosynthesis leading to the hypothesis that CBIs, such as ISX and DCB, would induce suberin deposition.

Here, we investigated whether treatment of Arabidopsis seedlings with CBIs leads to ectopic suberin deposition as a response to cell wall damage. In line with this hypothesis, the root tips of ISX and DCB treated seedlings displayed increased levels of suberin- associated fluorol yellow 088 staining, which accumulated over time. Additionally, GC-

MS quantification of suberin content and composition in ISX and DCB treated seedlings were nearly doubled compared to untreated controls. These results suggest that ISX and

DCB induce suberin production in response to cellulose biosynthesis inhibition.

ii. Methods and Materials:

General plant growth and maintenance:

Arabidopsis thaliana Columbia (Col-0) seeds were sterilized in seed cleaning solution (30% [v/v] bleach, 0.1% [w/v] SDS) for 20 minutes at 25 ℃. The seed cleaning solution was removed, and seeds were washed 5 times in sterile water followed by incubation at 4 ℃ for 48 hours before use. Seeds were germinated on MS media (1/2 X

Murashige and Skoog salts, 10 mM MES-KOH pH 5.7, 1% [w/v] Sucrose, and 1% [w/v] phytoagar). Dichlorobenzyonitrile (Sigma-Aldrich) was dissolved in DMSO to yield a 1 mM stock. Isoxaben (Sigma-Aldrich) was suspended in methanol to yield a 10 µM stock.

Each compound was added to MS media at the indicated concentrations. Seedlings were grown under long day conditions (16-hour light, 8-hour dark) at 22 ℃ vertically for 7 days. Transfer experiments were conducted by germinating and growing Col-0 seed on 311

MS media for 4 days then transferred onto indicated treatments. Three to four seedlings were selected at designated time points for Suberin staining.

Suberin staining and imaging:

Arabidopsis seedlings were transferred to 1 mL of 0.01% [w/v] Fluorol Yellow

088 (Santa Cruz Biotechnology) in Lactic acid (Sigma-Aldrich). Seedlings were heated to

70 ℃ for 30 minutes and washed three times with deionized water for 5 minutes.

Seedlings were loaded into a glass slide submerged in glycerol and imaged with BZ-X series All-in-one Fluorescence Microscope (Keyence) using a GFP filter cube (Ex. 440-

470 nm, Em. 500-520 nm, DM 495 nm, BA 525-550nm).

Delipidation and suberin quantification:

Seven-day-old Arabidopsis seedlings treated with 1 µM DCB, 10 nM ISX, 0.1%

[v/v] Methanol, 0.1% [v/v] DMSO, or untreated for 24 hours were transferred to an 8 mL screw cap glass vial with PTFE-lined caps. Five milliliters of hot (80 ℃) 2-propanol was added, and the samples were incubated at 80 °C for 10 minutes. Samples were cooled to

25 ℃ and ground in a polytron. 2-propanol was added until the tubes contained a total volume of 6 mL, and the samples were placed in a shaking rotator for 2 hours at 25 ℃.

Samples were centrifuged for 10 minutes at 800 x g, and the 2-propanol was removed. An additional 7 mL of 2-propanol was added, and the samples were placed in a shaking rotator for 16 hours at 25 ℃. Samples were centrifuged at 800 x g, the supernatant was removed, and 7 mL of [2:1] [v/v] Chloroform: Methanol was added. The resulting samples were placed on a shaking rotator for 24 hours at 25 ℃. Samples were centrifuged at 1000 x g, the solvent was removed, and 7 mL of [1:2] [v/v] Chloroform: 312

Methanol was added. The resulting samples were placed on a shaking rotator for 24 hours at 25 ℃. Samples were centrifuged at 1000 x g, the solvent was removed, and 7 mL of methanol was added. The samples were again incubated on a shaking rotator for 24 hours at 25 ℃. The samples were centrifuged at 1000 x g, the solvent was removed, and 7 mL of [1:2] [v/v] Chloroform: Methanol was added. Samples were placed on a shaking rotator for 24 hours at 25 ℃. Finally, the samples were centrifuged at 1000 x g, the solvent was removed, and the samples were dried down completely under a vacuum desiccator. The acid-catalyzed transmethylation of suberin to hydrolyze polymerized suberin, derivatization, and quantified using GC-MS as previously described (Kosma et al., 2012; Molina et al., 2006).

iii. Results:

Isoxaben and Dichlorobenzonitrile induce suberin deposition in Arabidopsis thaliana:

One effect often accompanied by altered cellulose biosynthesis is the activation of cell wall integrity sensing and the resulting ectopic deposition of lignin (Li et al., 2014).

Ectopic lignin deposition has been observed in CESA3 mutations ectopic lignin 1-1 and

1-2 (eli1-1 and eli1-2) utilizing phloroglucinol staining (Caño-Delgado et al., 2003). The same ectopic lignin deposition was observed in ISX treated seedlings (Caño-Delgado et al., 2003; Hamann et al., 2009). This response to the inhibition of cellulose biosynthesis may be a wounding or immune response to cell wall damage. Transcriptional analysis of

ISX and thaxtomin A treated Arabidopsis suspension culture cells showed upregulated genes associated with wounding and pathogen response (Duval and Beaudoin, 2009). A previous study suggested a relationship between the DAMP cellobiose, wounding 313 response elicitation, and the transcriptional induction of MYB41, a transcription factor that activates genes necessary for suberin biosynthesis (de Azevedo Souza et al., 2017).

Additionally, MYB41 also induced gene transcripts associated with lignin biosynthesis

(Kosma et al., 2014). These observations lead to the hypothesis that common cellulose biosynthesis inhibitors (CBIs) not only induce ectopic lignin deposition, but also suberin deposition.

To test this hypothesis, Arabidopsis seedling were germinated and grown for 7 days in ½ x MS media or ½ x MS media supplemented with 1 µM DCB, 0.1% Methanol, and 10 nM ISX. Seedlings were stained with the suberin-specific dye Fluorol Yellow088

(FY088), mounted on a Keyence microscope, and imaged for FY088 staining. Suberin- associated staining was absent in the root tips of untreated and methanol treated seedlings

(Figure 6.1A-C and G-I). In contrast, DCB and ISX treated seedlings displayed elevated levels of suberin staining with FY088 (Figure 6.1 D-F and J-L). These data suggest that treating Arabidopsis seedlings with common CBI induces suberin deposition.

To further investigate the deposition of suberin under ISX and DCB treatment,

Arabidopsis seedlings were germinated and grown in MS media for 4 days then transferred to MS media with no supplementation or 1 µM DCB and 10 nM ISX.

Seedlings were collected every 24 hours starting at the 0 hour and collecting at a final time point of 72 hours. Seedlings grown in MS media without supplementation grew normally and displayed no suberin deposition over the course of the experiment (Figure

6.2A). Seedlings in both treatments demonstrated root tip swelling within 24 hours and in conjunction, suberin deposition was observed. Both root swelling and suberin stain signal 314

Figure 6.1: Investigating suberin deposition in Arabidopsis seedlings on cellulose biosynthesis inhibitors Dichlorobenzonitrile (DCB) and Isoxaben (ISX). Arabidopsis seedlings were germinated and grown on MS media containing the following treatments: No Treatment) (A – C), 1 µM DCB (D – E), 0.1% Methanol (G – I), and 10 nM ISX (J – L). Seedlings were stained with Fluoro Yellow-088 (FY088) and mounted on Keyence microscope. Images were taken in bright field (A, D, G, J), GFP filter (Ex. 440-470 nm, Em. 500-520 nm) (B, E, H, K), and overlaid images (C, F, I, L). Scale bars represent 100 µM.

315

Figure 6.2: Time course assay of suberin deposition on cellulose biosynthesis inhibitors. Arabidopsis seed were germinated and grown for 4 days on MS media. Seedlings were then transferred to MS media with no treatment (A), with 1 µM DCB (B), and 10 nm ISX (C). Initial FY088 staining images (0 hr), were from remaining seedlings in MS media. Seedlings were stained with FY088 every 24 hours after transfer for a total of 72 hours. Bright field (Left), GFP (Center), and Merged (Right) images are displayed. GFP filter (Ex. 440-470 nm, Em. 500-520 nm). Scale bars represent 100 µM.

316 intensity increased under DCB and ISX treatment (Figure 6.2B and 6.2C) during the time course of the experiment. Overall, these results suggest that suberin is continuously deposited during the duration of CBI exposure.

ISX and DCB treated seedling display increases in polymerized and soluble suberin:

The increases in overall suberin deposition of ISX and DCB treated Arabidopsis seedlings, indicated that an additional effect of cellulose deficiency on top of ectopic lignin, wounding response, and immune response. Both ISX and DCB treatment reliably increased suberin deposition in a temporal fashion. Interestingly, both CBI inhibitors arrest cellulose production in unique ways, suggesting that different cellulose synthesis inhibition mechanisms still yield ectopic suberin deposition.

To ensure that the cytologically observed product exhibited the chemical composition of suberin, Arabidopsis seed were germinated and grown in MS media for 6 days on filter paper and then were transferred to untreated MS media or MS media supplemented with

0.1% methanol, 0.1% DMSO, 1 µM DCB, and 10 nM ISX for 24 hours. Root tissues from these samples were harvested and subjected to delipidation as described in Materials and Methods. Polymerized suberin was subjected to acidic transmethylation and analyzed using GC/MS as described previously (Kosma et al., 2012; Molina et al., 2006).

Polymerized suberin content for ISX and DCB treated roots are summarized in Figure

6.3. Overall total suberin content increased nearly 3-fold in ISX and DCB treated roots compared to untreated plants or solvent controls (Figure 6.3A and C). ISX and DCB both displayed specific increases in 20:0 Fatty acids (FAs), 18:1 ω-hydroxy (ω-OH) FAs, and 317

Figure 6.3: Quantification of polymeric suberin from 7-day-old roots. Arabidopsis seed were germinated and grown on filter paper in MS media and grown for 6 days then transferred to MS media with no treatment (NT), 0.1% DMSO (DMSO), 0.1% Methanol (MeOH), 1 µM DCB (DCB), and 10 nM ISX (ISX) for 24 hours. Roots were harvested, delipidated and polymerized suberin was analyzed. (A) Total polymeric suberin in ISX treated seedlings are shown. Dunnett’s multiple comparisons (***, p < 0.001) n = 3. Error bars represent SEM. (B) Individual suberin monomers of ISX treated roots indicating fatty acids (FAs), omega hydroxy fatty acids (ω-OH FAs), Dicarboxylic acids (DCA), primary alcohols (1-Alcs), and phenolic compounds (PP). Error bars represent SEM n = 3. (C) Total polymeric suberin of DCB treated seedlings are shown. Dunnett’s multiple comparisons (**, p < 0.01) n = 3. Error bars represent SEM. (D) Individual suberin monomers of DCB treated roots. Error bars represent SEM n = 3.

318

18:1 dicarboxylic acids (DCA). Furthermore, primary alcohols (1-Alcs) displayed mild increases in abundance, but these results were not statistically significant for ISX and

DCB. Interestingly, a unique phenolic component of suberin, ferulate, only increased in

ISX treated roots but not in DCB (Figure 6.3B and D).

One interesting observation from seedlings germinated on ISX and DCB (Figure 6.1) appear to deposit suberin unequally across tissues. Seedlings observed over time under

ISX and DCB treatment (Figure 6.2) also appear to show an unequal distribution of suberin staining. These observations are potentially explained by uneven staining with

FY088, or this result could be an indication of potentially concentrated puncta of unpolymerized suberin. To test whether ISX and DCB elicit the deposition of unpolymerized suberin, washes from the delipidation process from 7-day-old roots treated for 24 hours with DCB and ISX were collected, concentrated, and suberin specific monomers were quantified using GC/MS. The resulting data are summarized in Figure

6.4 for both ISX and DCB. Total soluble suberin from DCB treated roots was increased by 35%, while ISX treated roots increased nearly 2-fold (Figure 6.4A and C).

Furthermore, increases were primarily observed in 1-Alcs with 22:0, 24:0, and 26:0 in

ISX treated roots, while DCB primarily had increases in 18:0, 20:0, and 22:0.

Additionally, 22:0 and 24:0 FAs were increased as well as ferulate in ISX treated roots but, DCB treated roots did not show increases in FAs and ferulate (Figure 6.4C and D).

These results suggest differential affects of cell wall biosynthesis inhibitors on suberin deposition and composition in polymerized and unpolymerized monomers.

iv. Discussion and Future Directions: 319

Figure 6.4: Quantification of soluble suberin from 7-day-old roots. Arabidopsis seed were germinated and grown on filter paper in MS media and grown for 6 days then transferred to MS media with no treatment (NT), 0.1% DMSO (DMSO), 0.1% Methanol (MeOH), 1 µM DCB (DCB), and 10 nM ISX (ISX) for 24 hours. Roots were harvested, delipidated and delipidated fraction was analyzed for suberin monomers. (A) Total soluble suberin in ISX treated seedlings are shown. Dunnett’s multiple comparisons (**, p < 0.01) n = 3. Error bars represent SEM. (B) Individual soluble suberin monomers of ISX treated roots indicating fatty acids (FAs), omega hydroxy fatty acids (ω-OH FAs), primary alcohols (1-Alcs), and phenolic compounds (PP). Error bars represent SEM n = 3. (C) Total soluble suberin of DCB treated seedlings are shown. Dunnett’s multiple comparisons (*, p < 0.05) n = 3. Error bars represent SEM. (D) Individual soluble suberin monomers of DCB treated roots. Error bars represent SEM n = 3.

320

Exposure to CBIs such as ISX and DCB induced suberin deposition in Arabidopsis seedlings. Seven-day-old Arabidopsis seedlings germinated in MS media containing ISX or DCB and stained for suberin using FY088 exhibited drastically increased fluorescence compared to untreated seedlings (Figure 6.1). Additionally, Arabidopsis seedlings that were transferred to MS containing ISX or DCB began to accumulate suberin within 24 hours of transfer, coincident with root tip swelling (Figure 6.2). Polymerized and soluble suberin content in DCB and ISX treated seedlings was also increased compared to untreated and solvent controls (Figure 6.3A and C). Analyzing suberin monomers demonstrated specific increases in 20:0 FAs, ω-OH FAs, and 18:1 DCAs in both treatments. Interestingly, ISX treated seedlings showed specific increases in ferulate content (Figure 6.3B and D). It is not clear why ISX would elicit ferulate biosynthesis and not DCB. It is possible the differences in the mechanisms of inhibition also attributes to the difference in ferulate observed in ISX treated seedlings. Surprisingly, quantification of soluble suberin, from the delipidated fractions, also had increases in suberin in both DCB and ISX. Larger increases in soluble suberin was observed in ISX treated seedlings (Figure 6.4A and C). Interestingly, soluble suberin monomer composition of ISX treated seedlings were elevated most notably in in primary alcohols

22:0, 24:0 and more pronounced ferulate. Comparatively, DCB treatment only had mild increase in primary alcohols (Figure 6.4B and C).

Overall, suberin deposition was observable in ISX and DCB treated seedlings through both staining and GC-MS. Attempts to assay gene transcripts associated with suberin biosynthesis such as ASFT1, GPAT5, and MYB41 (Molina et al., 2009; Kosma et al.,

2014; Belsson et al., 2007) yielded inconclusive results (Data not shown). Further 321 investigation into the transcriptional regulation of suberin biosynthetic genes should be conducted at varying time points after CBI exposure. Previously investigations of wounding responses in plants typically resulted in suberin deposition (Hawkins and

Boudet, 1996; Franke et al., 2009; Schreiber et al., 2005; Boher et al., 2013; Wei et al.,

2020). CBIs may elicit suberin deposition due to a wounding response. As cellulose biosynthesis inhibition compromised cell wall integrity, cells will burst as their cell walls cannot contain the turgor pressure inside the cell (Voxeur and Höfte, 2016; Gigli-

Bisceglia et al., 2020). It can be hypothesized that cells bursting due to compromised cell wall integrity elicits a wounding response leading to suberin deposition. An interesting angle to explore are compromises to the cell wall through salt-stress. Salinity negatively affects the wall by Na+ ions displacing Ca2+ in homogalacturonan (HG), thus weakening the cell wall. FERONIA (FER) aids in salt-tolerance by sensing disrupted HG and causes a Ca2+ influx leading to cell wall modification (Feng et al., 2018). The fer2 and fer4 mutants are hypersensitive to salt-stress as they are unable to acclimate to high salinity resulting in cell bursting. It would be interesting to observe suberin deposition between wildtype and the fer mutants under salt-stress. This experiment would also indicate if suberin deposition is a result of cellulose inhibition or an overall response to compromises in cell wall integrity.

An interesting observation is the increased levels of ferulate in ISX treated seeding, which was not seen in DCB treated seedlings (Figure 6.2 and Figure 6.3). The ferulate increases may be attributed to ISX mode of inhibition as it inhibits cellulose synthase by removing CSCs from the plasma membrane (DeBolt et al., 2007). It is still not clear why

ISX causes CSCs to be removed from the plasma membrane, but known isoxaben 322 resistant (ixr) mutants suggest a direct connection to CESA3 and CESA6 (Tateno et al.,

2016). Interestingly, several ixr mutants corresponding to mutations in CESA3 such as ixr1-1, ixr1-3, and ixr1-4 cause missense mutations in the 7th transmembrane domain

(TM7). Based on the PttCESA8 Cryo-EM structure, TM7 contributes to the structure of the channel where newly synthesize cellulose is extruded of the adjacent PttCESA8

(Purushotham et al., 2020). It is likely that ISX binds directly to this area of the complex and interferes with the channel leading to inhibition of cellulose biosynthesis and as a response it is removed from the plasma membrane. This response may closely resemble a pathogenic response as observed in a bacterial endophyte secreted compound

Acetobixan, another known CBI (Xia et al., 2014). As ISX and Acetobixan essentially inhibit cellulose biosynthesis as a characteristic of pathogen invasion, it would also stimulate ferulate biosynthesis. In contrast, DCB inhibits cellulose biosynthesis by slowing down CSCs, and as demonstrated in Chapter 5. This effect may be a result of inhibiting glycosylated sphingolipid biosynthesis. As the mode of cellulose biosynthesis inhibition is an indirect effect of DCB toxicity, it may not stimulate ferulate deposition as

ISX would. The differences in ferulate between ISX and DCB treatment suggest ferulate may have antimicrobial properties. The response of cellulose biosynthesis inhibition and suberin deposition is a novel mechanism and provides an additional mechanism to explore in cell wall integrity sensing.

323

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Chapter 7: Discussion

I. The potential of fluoro fucose resistant lines:

As detailed in chapter 2, a suite of fluoro fucose resistant (ffr) lines were isolated from a random mutagenesis screen. After rescreening the progeny from the 29 viable ffr lines, 17 of them demonstrated resistance to 25 µM 2F-Fuc and 11 were resistant to 25

µM Ac3 2F-Fuc (Table 2.3). Currently, seven of the confirmed resistant lines are allelic to fkgp-3, signifying that the remaining 10 lines contain point mutations in unidentified genes. Here, a discussion the current status of each ffr line and possible outcomes of genetic mapping are discussed. Additionally, other utilities of 2F-Fuc will be discussed in this section.

a. Current state of the ffr lines and their potential:

As outlined in chapter 2, there are four ffr lines that have been successfully backcrossed and have available F2 progeny. Two of these lines, ffr21 and ffr25, displayed

24.4% and 22.1% resistance to 25 µM 2F-Fuc, respectively, consistent with a single recessive locus underlying the associated resistance trait. The other remaining lines, ffr17 and ffr26 have non-recessive phenotypes displaying resistance to 25 µM 2F-Fuc at 66.2% and 51.8%, respectively, in the F2 population (Table 2.4). It is interesting to note that ffr26 recombination frequency of 51.8% in F2 progeny raises the question, what mutation or combination of mutations would lead to this observation? One hypothesis to the ffr26

F2 recombination frequency may be similar to a dual functioning protein such as

FERONIA (FER) which is involved in cell wall integrity sensing and in plant reproduction. To elaborate, FER is a receptor-like kinase (RLK) critical for maintaining 331

Figure 7.1: Hypothetical Mendelian genetics models for ffr lines with non-recessive recombinant frequencies. (A) Punnett square model depicting the mutation associated with resistance to 2F-Fuc has a reproductive defect in ffr26. Red highlights are non- viable and blue is resistant to 2F-Fuc. (B) Punnett square model depicting a dominant mutation in ffr17 where a homozygous genotype is lethal. Red highlights are non- viable and blue is resistant to 2F-Fuc (C) Hypothetical mechanism of ffr17 resistance to 2F-Fuc but continued sensitivity to Ac32F-Fuc. In wildtype (Col-0), 2F-Fuc transport is dependent on a fucose transporter to enter the cell and be metabolically incorporated to GDP-2F-Fuc. In contrast, the increased permeability of Ac32F-Fuc may enter the cell without the aid of a transporter and non-specific esterases convert it to 2F-Fuc. In ffr17, the hypothesized cause of resistance to 2F-Fuc is through a lack of a transporter, but Ac32F-Fuc can still enter the cell.

332 cell wall integrity during salt stress and is also critical in female pollen tube perception

(Huck et al., 2003; Feng et al., 2018). The fer mutant displayed a relatively normal growth phenotype compared to wildtype but has impaired female fertility with transmission rates as low as 15% (Huck et al., 2003). Under salt-stress, growth is severely impacted in fer-4 with compromised cell wall integrity leading to cell bursting (Feng et al., 2018). It can be hypothesized that ffr26 carries a mutation in a gene encoding a cell wall integrity sensor expressed in roots and plant fertility. The generation of ffr26 X Col-

0 F2 population was based on an F1 progeny by pollinating ffr26 pistils with Col-0 pollen, postulating ffr26 may have impaired male fertility instead of a female fertility. If ffr26 can be shown to have a male reproductive defect will likely display a 50% transmission rate as observed with other sex-specific reproductive defects (Smith et al.,

2018; Huck et al., 2003; Dai et al., 2014; Li et al., 2013a) (Figure 7.1A). This hypothesis does infer the ffr26 line does contain a dominant mutation, where the gene in question behaves as a dominant-negative or haplo-gene.

The ffr17 line is an interesting dominant mutant as there are very few examples of dominant mutations involved with cell wall biosynthesis. The few examples are related to cellulose biosynthesis are found in CESA3 and CESA7 acting as dominant-negative genes

(Carpita and McCann, 2015). Additionally, mutations in Reversable-glycosylated proteins 1 (RGP1) and RGP2 involved in the interconversion of Arabionofuranose (Araf) to Arabinopyranose (Arap) are dominant. Mutations in RGP1 and RGP2 display a decrease in Ara content in the cell wall between 12 – 31% (Rautengarten et al., 2011).

Another example is the EMBRYO DEFECTIVE (EMB30) gene, which is involved in early seedling development, polarized growth, and pectin secretion. The emb30-3 mutant 333 displays abnormal pectin deposition based on immunostaining (Shevell et al., 2000).

What differentiates ffr17 from the described genes is that ffr17 grows similar to wildtype, whereas the rest displayed negative growth phenotypes suggesting the ffr17 seedlings we grown are heterozygous as homozygous genotype would be embryonic lethal (Figure

7.1B). Interestingly, ffr17, despite being resistant to 2F-Fuc, is sensitive to Ac3 2F-Fuc treatment (Figure 2.8), suggesting that the ffr17 mutation potentially encodes an L-fucose transporter that transports free L-fucose and 2F-Fuc. This hypothesis signifies 2F-Fuc cannot enter the cell, but the acetylated 2F-Fuc may diffuse due to increased membrane permeability compared to 2F-Fuc, bypassing the need for a transporter (Figure 7.1C).

b. Other utilities of 2F-Fuc

Fucose analogs have been used in hepatoma cells to inhibit cell migration and proliferation. In hepatoma (HepG2) cell cultures, 2F-Fuc was shown to reduce cell proliferation, limiting the size of tumor colonies without impacting cell viability in vitro.

Interestingly, in vivo analysis of mice inoculated with HepG2 cells treated with 2F-Fuc had smaller tumors and had reduced tumor growth over time compared to untreated controls. This observation is attributed to the reduced core fucosylation in N-linked glycans in 2F-Fuc treated HepG2 cells. The conclusion of these results is that 2F-Fuc inhibits fucosyltransferases in HepG2 cells. (Zhou et al., 2017). One other fucose analog,

6-Alkyne-L-Fucose (Alk-Fuc), a known click-compatible monosaccharide in Arabidopsis

(Anderson et al., 2012), is also a potent fucosylation inhibitor in mammalian cells. Unlike

2F-Fuc, Alk-Fuc primarily depletes cellular GDP-Fuc instead of inhibiting 334 fucosyltransferases in vivo. Additionally, Alk-Fuc did not inhibit cell proliferation, instead it severely reduced cell migration and cell invasion (Kizuka et al., 2017).

The phenotypic effects of 2F-Fuc in Arabidopsis have been attributed to its inhibitory effects in RG-II fucosylation (Chapter 2) (Villalobos et al., 2015; Dumont et al., 2015).

Although 2F-Fuc serves as a novel inhibitor of RG-II fucosylation events, RG-II only represents about 10% of total pectins, which only make up about 35% of the cell wall

(Mohnen, 2008). Cell wall -associated Fuc content decreased by approximately 50% in

2F-Fuc treated seedlings (Figure 2.3), suggesting that 2F-Fuc is a global inhibitor of fucosylation events. This hypothesis was biochemically confirmed as xyloglucan fucosylation, protein N-linked glycans, and incorporation of a click compatible 6-alkyne-

L-fucose (Alk-Fuc) was abolished in roots (Dumont et al., 2015) upon 2F-Fuc treatment.

Interestingly, the abolishment of α-(1,3)-fucosylation events in plant N-linked glycans is one strategy to “humanize” recombinant proteins generated in non-mammalian systems commonly referred to as therapeutic proteins to treated diseases and minimize cancer progression.

Considerable effort has been placed in engineering protein glycans to study the fundamental processes of cellular glycans, generate glycosylated therapeutic proteins, and elicit or inhibit cellular processes. Several techniques have been utilized to engineer protein glycans through genetic approaches and chemical strategies. Here, I will focus on chemical strategies that have been implemented and how 2F-Fuc could be beneficial toward these goals. Small molecules are used in chemical strategies including the use of deoxy-monosaccharides design to limit incorporation of other monosaccharides, 335 substituting hydroxyl groups with novel functional groups, such as monosaccharides containing thiol groups for example to alter functionality, labeling with click-compatible monosaccharides, and the use of monosaccharide analogs as inhibitors (Griffin and

Hsieh-Wilson, 2016). One area of interest in therapeutic protein production is the precise control of N-linked glycosylation events to maximize the effectiveness of recombinant monoclonal antibodies. The N-linked glycosylation state of recombinant monoclonal antibodies affect the stability, efficiency, and effector function. The presence of fucose or galactose in N-linked glycans are the predominant monosaccharides that negatively impact the efficacy of recombinant monoclonal antibodies (Matsumiya et al., 2007;

Hodoniczky et al., 2005). Currently, most biotechnology producers of therapeutic proteins utilize Chinese hamster ovary (CHO) cells engineered to produce proteins with minimally fucosylated and galatosylated N-linked glycans (Sha et al., 2016). A critical limitation of utilizing CHO cells in therapeutic protein production involves challenges with N-linked glycan processing in CHO cells. These cells are prone to environmental variations that are difficult to control during bioprocessing leading to greater variation in the N-linked glycan structure, and the high cost associated with using CHO cells to produce therapeutic proteins make it very cost prohibitive, especially for patients under enzyme replacement therapy (Chen, 2016).

To circumvent some of these challenges, plant expression systems have become an interesting and viable alternative. Fewer plant N-linked glycan structures have been described compared to mammals with only two predominant N-linked glycan products found in plants. One structure is described in Figure 1.6 and the other predominant N- linked glycan does not have terminal galactose. Use of plant cell cultures have yielded 336 success in producing therapeutic proteins at yields comparable to CHO cells (Jung et al.,

2016; Grabowski et al., 2014; Kim et al., 2020). Despite the smaller variation in plant N- linked glycans and having similar yields to CHO cells, concerns arose of therapeutic proteins produced by plants may trigger immune responses causing adverse side-effects

(Chen, 2016). To circumvent these issues, several strategies have been implemented to

“humanize” plant-produced therapeutic proteins. One particular study utilized 2F-Fuc in rice cell cultures to produce non-fucosylated recombinant human acid alpha-glucosidase

(rhAAG), a therapeutic enzyme to treat the genetic disorder Pompe disease (Kim et al.,

2020), a fatal muscular disorder that shuts down heart and skeletal muscle function by the overaccumulation of glycogen in lysosomes (Winkel et al., 2005). Interestingly, 2F-Fuc does not negatively impact rice cell culture growth, unlike what was observed in

Arabidopsis seedlings, and reliably inhibits fucosylation events in N-linked glycosylation. Despite a reduction in N-linked fucosylation, the presence of fucosylated

N-linked glycans in rhAAG was still detected (39.9% with 0.5% PF-68 surfactant) (Kim et al., 2020). It could be possible completely inhibit fucosylation or at least improve the ratio between non-fucosylated and fucosylated N-linked glycans in recombinant proteins by utilizing Ac3 2F-Fuc in combination with PF-68 in the protein producing rice cell cultures. 2F-Fuc may be a valuable tool for start-up companies invested in generating their own novel protein therapeutics using plants cell cultures. Generally, generating a humanized therapeutic protein in plants is a time consuming and laborious process. For example, using Arabidopsis cell cultures a homozygous triple knockout for all three fucosyltransferases involved in N-linked glycosylation (FUT11, FUT12, and FUT13) must be generated (Both et al., 2011; Wilson et al., 2001; Rips et al., 2017). The triple 337 knockout must then be transformed with a cassette with the therapeutic protein of interest and followed by optimization that may or may not yield success. A start-up company without access to fut11/fut12/fut13 triple knockout is where 2F-Fuc really becomes useful because it can circumvent the need of a fut11/fut12/fut13 triple knockout during optimization of “humanized” therapeutic protein expression. The optimized gene cassette containing the therapeutic coding sequence can be later transformed into the fut11/fut12/fut13 triple knockout. In short, 2F-Fuc allows the simultaneous development of the triple knockout and expression optimization of the “humanized” therapeutic protein.

II. Sugar sensing in plants and fungi in relation to glucose and mannose

Sugar sensing in plants and fungi are critical for normal developmental processes.

Sugar sensing is also important in mammalian cells specifically in human pancreatic β- cells and hepatocytes. Both cell types uniquely express Glucokinase B (GlkB), one of four hexokinases in humans, and this enzyme functions as a glucose sensor. Typically, under high glucose conditions, GlkB signals for sugar metabolism and storage in hepatocytes while signals secretion of insulin pancreatic β-cells (Rolland et al., 2006;

Matschinsky et al., 1998; Van Schaftingen et al., 1994). Unlike plants and yeast, GlkB activity is positively regulated by several regulatory proteins in the presence of high glucose concentrations (Shiraishi et al., 2001; Van Schaftingen et al., 1994; Munoz-

Alonso et al., 2000). One of the most interesting aspects of sugar sensing in yeast and plants is the repression of growth in the presence of high glucose concentrations (Moore et al., 2003; Trumbly, 1992). For example, under high glucose concentrations, growth in 338 yeast is repressed by the action of HXK2p (Guez et al., 2001). Interestingly, the deletion of HXK2, hxk2Δ, in S. cerevisiae causes complete insensitivity to glucose-mediated growth repression, but can be partially re-sensitized by human GlkB (Mayordomo and

Sanz, 2001). In this section, the sugar sensing, primarily glucose and mannose, in relation to growth repression in plants and yeast is discussed.

a. Sugar sensing in plants

Sugars play a pivotal role as signaling molecules having hormone-like functions and controlling vital processes in plant systems. Under high glucose conditions, seed germination, seedling development, cell type differentiation, and photosynthesis are all inhibited. In Arabidopsis, studies have indicated that Hexokinase 1 plays a crucial role as a sugar sensor influencing a variety of physiological processes through acting as a transcriptional regulator when bound to glucose or mannose (Hirsche et al., 2017; Kunz et al., 2015; Cho et al., 2006; Kelly et al., 2017; Rolland et al., 2002; Moore et al., 2003;

Pego et al., 1999). As the presence of high glucose inhibits growth in Arabidopsis, this condition serves as a useful avenue to isolate and study genes involved with glucose repression.

One of the first isolated genes associated with glucose repression in Arabidopsis was GLUCOSE INSENSITIVE 1 (GIN1). Three gin1 alleles were isolated, gin1-1, gin1-2, and gin1-3, which all showed insensitivity to high glucose treatment. Interestingly, genetic mapping of the gin1 mutants corresponded to point mutations in ABSCISIC ADIC

DEFICIENT 2 (ABA2), and further evaluation of aba2 mutants demonstrated that they also displayed insensitivity to high glucose concentrations (Cheng et al., 2002). This 339 result implicates that abscisic acid plays a role in glucose-mediated growth repression.

Other hormones have also been implicated in glucose repression, such as ethylene and gibberellin (GA). The CONSTITUTIVE TRIPLE-RESPONSE 1 (CTR1) mutant, ctr1, is constitutively responsive to ethylene signaling and displays insensitivity to glucose repression. Additionally, ctr1 displayed resistance to high mannose growth inhibition and to the negative effects of paclobutrazol (PAC), an inhibitor of GA biosynthesis (Gibson et al., 2001). A CALCINEURIN B-Like 1 (CBL1) is an internal Ca+2 sensor that is closely involved in GA signaling during seed germination. The T-DNA insertional mutants, cbl1-

1 and cbl1-2, displayed hypersensitivity to high glucose conditions, exhibiting lower germination rates, reduced cotyledon greening, and reduced root growth under high glucose conditions. Additionally, the expression of Gibberellin-3-beta-dioxygenase 1

(GA3ox1) was decreased in the cbl1-1 and cbl1-2. Furthermore, cbl1 mutants were hypersensitive to PAC treatment. The loss of cotyledon greening under high glucose treatment was alleviated with exogenous application of aminocyclopropane carboxylic acid, a precursor of ethylene (Li et al., 2013b). Glucose repression is also alleviated in seedlings carrying the auxin response 3 (axr3) mutation, implicating auxin signaling in glucose repression. Additionally, it appears that exogenous application of a synthetic cytokinin, 6-benxylaminopurine (BAP), increased sensitivity to the glucose response

(Kushwah and Laxmi, 2017). Several genes that encode proteins involved with ABA biosynthesis, ABA transcription factors, ethylene biosynthesis, and ethylene response transcription factors have been identified to be involved in response to high glucose concentration (Rognoni et al., 2007; Rolland et al., 2002). To summarize, several 340 phytohormones both mitigate and increase the potency of glucose mediated growth repression, such as abscisic acid auxin, cytokinin, ethylene, and GA (Figure 7.2A).

The signaling events discussed above that are implicated in glucose repression can arguably be categorized as HXK1-independent responses, although this may not be the case due to the face that gin2-1 and gin2-2 (mutants in the HXK1 gene) are non- responsive to Auxin treatment unless exogenous cytokinin is applied (Moore et al.,

2003). Interestingly, a site-directed mutant of Arabidopsis HXK1 that abolished the catalytic activity exhibited similar glucose repression as the wildtype, suggesting catalytic activity is not crucial to glucose repression, which led to the hypothesis that

HXK1 is a bifunctional enzyme (Moore et al., 2003). HXK1 bifunctional activity was confirmed under high glucose conditions, where HXK1 re-localizes to the nucleus and physically interacts with a vacuolar H+-ATPase B1 (VHA-B1) and 19S regulatory particle of proteasome subunit (RPT5B) (Figure 7.2B). Nuclear HXK1, in complex with

VHA-B1 and RPT5B, negatively regulates photosynthetic gene expression (Cho et al.,

2006). Unsurprisingly, vha-B1 and rpt5b mutants displayed similar insensitivity to high glucose concentrations (Cho et al., 2006). One final note in glucose repression, is in plant reproduction. Pollen tube germination is inhibited by several sugars including glucose, but the most potent inhibitors were galactose and mannose (Hirsche et al., 2017). Glucose and mannose inhibition of pollen tube germination can be alleviated in the gin2-1 mutant.

Furthermore an inhibitor of HXK1, mannoheptulose, also alleviated mannose and glucose repression (Hirsche et al., 2017). Overall, these observations suggest that sugar sensing also critically influences pollen tube germination and may have more implications in plant fertility. 341

Figure 7.2: Models of glucose repression in Arabidopsis and S. cerevisiae. (A) Signaling model of Glucose repression and phytohormones interactions that influence seed germination and early seedling development. Glucose may influence Auxin/Auxin signaling at high concentrations. (B) Transcriptional repression model of HXK1 bound glucose interacting with VHA-B1 and RPT5B., TF are putative transcription factors. (C) Model of glucose repression and de-repression in S. cerevisiae.

342

Overall, the data presented in chapter 3 implicated 2F-Man in a HXK1-based sugar sensing mechanism. Based on the random mutagenesis screen conducted and genetic mapping of fluoro mannose resistant 6 (fmr6) and fmr11 had point mutations in

HXK1 gene. Both mutants also displayed some insensitivity to increasing glucose concentrations (Figure 3.8). Furthermore, resistance to 2F-Glc was also observed in fmr6 and fmr11, suggesting 2F-Glc also inhibits growth in a HXK1-dependent manner. It was quite unexpected to find that 2F-Man-associated growth inhibition was not due to the abolishment of mannosylation events but is potentially the result of activation of a sugar- signaling cascade. This result leaves many new avenues to explore to investigate the relationship between 2F-Man and glucose repression signaling. One avenue would be to test 2F-Man on other glucose insensitive mutants and the phytohormones described above to test for resistance or to chemically rescue the 2F-Man phenotype in Arabidopsis.

To further support the role of HXK1 in 2F-Man-associated growth inhibition, it may be important to investigate whether the HXK1 inhibitor mannoheptulose is able to alleviate the negative growth phenotype. 2F-Man has a unique advantage in future investigations in glucose repression where inhibition can be observed at relatively low concentrations

(IC50 = 12 µM), circumventing the potential of osmotic stress eliminating the need of an osmotic control. Overall, it would be interesting to see the localization of HXK1 under

2F-Man treatment. If HXK1 is transported from the cytosol to the nucleus upon 2F-Man treatment, then this would suggest 2F-Man transcriptionally represses growth through the glucose repression pathway.

b. Sugar sensing in yeast 343

In baker’s yeast (Saccharomyces cerevisiae), the sugar sensing and glucose- repression signaling pathways are remarkably similar to plants. Hexokinase-mediated glucose repression has also observed in yeast, and a more robust understanding of its mechanism has been elucidated. Yeast glucose repression is mediated by the Arabidopsis

AtHXK1 homolog, ScHxk2p (Hxk2p) (Guez et al., 2001; Mortimer et al., 1989). Under high glucose conditions, Hxk2p enters the nucleus, and transcriptionally represses

ScHxk1 as well as Glucokinase (Glk), and upregulates its own transcript levels (Herrero et al., 1995). Additionally, as a glucose sensor, Hxk2p mediates programed cell death/necrosis, and interacts with the mitochondria (Amigoni et al., 2013; Barbosa et al.,

2016).

In the current model, Hxk2p enters the nucleus through an α/β-importin- dependent pathway and recruits Mig1p, Mig2p, Resistant to glucose repression 1 (Reg1p)

(Matsumoto et al., 1983), Sucrose non-fermenting 1 (Snf1p), Snf4p (Celenza et al.,

1989), and Galactose metabolism 83 (Gal83p) (Corvey et al., 2005) to form a protein complex that transcriptionally upregulates a variety of genes involved in fermentation and repressing genes associated with catabolism of non-fermentable carbon sources

(Bisson and Fraenkel, 1983; Gancedo, 1992). The current model is based on repression of

Suc2 (Vega et al., 2016). Mig1p and Mig2p are zinc finger transcription factors that directly bind DNA and both physically interact with Hxk2p in the nucleus (Ahuatzi et al.,

2004; Lutfiyya and Johnston, 1996; Vega et al., 2016). Reg1p is the regulatory subunit in the Reg1p/Glc7p phosphatase complex that de-phosphorylates Hxk2p maintaining its localization in the nucleus. In the other hand, Snf1p is a protein kinase that phosphorylates Hxk2p at Serine-14 signaling nuclear export (Fernańdez-García et al., 344

2012). Snf4p and Gal83p function as positive regulatory subunits of Snf1p kinase and maintains Snf1p localization in the nucleus (Corvey et al., 2005). Cytosolic Snf1p can also be phosphorylated by three protein kinase kinases that target it to the nucleus. These protein kinase kinases include Snf1p-activating-kinase 1 (Sak1p), Elongation morphology 1 (Elm1p), and Target of SBF 3 (Tos3) (Orlova et al., 2010; Blacketer et al.,

1993; Kim et al., 2005). A general model depicting glucose repression displaying Hxk2 localization into the nucleus is summarized in Figure 7.2C.

As covered in chapter 4, 2F-Man-associated growth inhibition is completely abolished in the hxk2Δ deletion line. Additionally, the reg1Δ deletion line was also completely resistant to the toxic effects of 2F-Man (Figure 4.5). Deletion of genes associated with alleviating Hxk2-mediated glucose repression displayed hypersensitivity to 2F-Man (Figure 4.6) (Table 4.3). Additionally, the hxk1Δ was still sensitive to 2F-Man treatment, suggesting the effects of 2F-Man toxicity is not associated with metabolism

(Figure 4.8). Many of the studies investigating glucose repression in S. cerevisiae were conducted in media containing 2% [w/v], the same concentrations used in all our treatment conditions. These conditions suggest glucose repression functions in an equilibrium. This equilibrium may be necessary as Hxk2 also interacts with the mitochondria, regulating reactive oxygen species (ROS), preventing programed cell death

(PCD) (Amigoni et al., 2013). As glucose levels decrease, Hxk2 localization equilibrium shifts to the cytosol, but under 2F-Man treatment Hxk2 remains in the nucleus and continues to repress genes necessary to catabolize ethanol produced from fermentation.

Overall, these data suggest that 2F-Man’s inhibitory capacity is associated with the 345 glucose repression system. This presents a unique opportunity to potentially utilize 2F-

Man as an anti-fungal agent.

Over the years, an emergence of multi drug resistant fungal pathogens have been identified and have become an increasing concern in the healthcare field (Lockhart et al.,

2017; Brown et al., 2012). One group of fungal pathogens, Candida sp., has a similar glucose repression system found in S. cerevisiae (Corvey et al., 2005; Sabina and Brown,

2009; Lagree et al., 2020). Unlike S. cerevisiae where glucose repression shift its metabolism to fermentative growth (Bisson and Fraenkel, 1983), the glucose repression system seems to promote virulence in a Hxk2-dependent manner in Candida sp., and studies investigating Candida sp. virulence tested glucose concentrations found in physiological conditions in blood (0.15 – 1%) (Rodaki et al., 2009; Sabina and Brown,

2009; Van Ende et al., 2019; Laurian et al., 2019). In physiological glucose conditions, transcription of genes associated with alternate carbon metabolism are repressed, but unlike S. cerevisiae protein degradation of protein products associated with alternate carbon metabolism are not degraded (Sandai et al., 2012). It would be interesting to analyze Candida sp. such as Candida albicans in in the presence of a known growth inhibitor that utilized the glucose repression for its toxicity such as, 2-deoxy-D-glucose, as observed in S. cerevisiae (Ralser et al., 2008; McCartney et al., 2014). Additionally, it would be interesting to test 2F-Man against C. albicans for growth inhibition. If 2F-Man has an inhibitory effect on C. albicans growth and or virulence it would be a great candidate as an anti-fungal agent. The unique advantage of 2F-Man over other anti- fungal drugs is how 2F-Man requires Hxk2 for inhibitory effects and may potentially 346 restrict the use of glucose as a carbon source as it would shift the CaHxk2 localization equilibrium to the nucleus similar to S. cerevisiae.

III. Implications of sphingolipid biosynthesis on cellulose

As detailed in chapter 5, N-dodecyldeoxynojirimycin (ND-DNJ) is a potent inhibitor of growth in Arabidopsis, primarily targeting Glycosylated sphingolipid biosynthesis causing a reduction in crystalline cellulose deposition. The primary sphingolipid class affected are GlcCers with small compositional alterations in GIPCs (Figure 5.3).

Surprisingly, we also found that GlcCer and GIPC contents were reduced upon treatment with PDMP and DCB (Figure 5.8). It has become commonly accepted that PDMP inhibits GCS activity in plants (Melser et al., 2010; Krüger et al., 2013; Rugen et al.,

2018; Yin et al., 2010), but the results presented here suggest that PDMP may also inhibit

GIPC-associated glycosyltransferases. It was also unexpected to find that DCB, a compound commonly considered a cellulose biosynthesis inhibitor , altered sphingolipid biosynthesis (DeBolt et al., 2007; Peng et al., 2001; Tran et al., 2018). The data presented are still implicitly in agreement that DCB inhibits cellulose biosynthesis, but it may be a secondary effect of inhibiting glycosylated sphingolipid biosynthesis.

a. Known glycosylated sphingolipid biosynthetic genes: do they contain less

cellulose?

Despite the fact that experimental data analyzing crystalline cellulose content has been exclusive to GIPC mutants gonst1-1 and gmt1-1 (Mortimer et al., 2013; Fang et al.,

2016), it can be reasoned that altered GlcCer content may negatively impact cellulose deposition. The best example would arise from the role that GlcCers play in the secretion 347 of proteins destined for the extracellular space. Inhibition in the biosynthesis of GlcCers using the GLUCOSYLCERAMIDE SYNTHASE (GCS) inhibitor PDMP arrested protein secretion and caused defects in Golgi morphology (Melser et al., 2010). Although, the data presented in chapter 5 demonstrate that PDMP treatment at concentrations used in

(Melser et al., 2010) also decreased GIPC content (Figure 5.8). The role of GlcCers in

Golgi morphology was confirmed in the gcs-1 mutant (Msanne et al., 2015). Thus, the gcs-1 would be an ideal mutant to analyze cellulose content and cross with the transgenic line containing GFP-CESA3:mChTua5 to investigate if CSC velocities are reduced. It will also be worth investigating if CSC velocities are reduced in gmt1-1 and gonst1-1 in the GFP-CESA3:mChTua5.

b. Non-motile observation in ND-DNJ and DCB

In addition to the role of GlcCers in exocytosis of proteins from the Golgi, it would be reasonable to hypothesize that GlcCers also play a role in endocytosis (Bashline et al., 2015, 2013; Fan et al., 2015). Typically, CSCs have a relatively high turnover rate at the plasma membrane, lasting between 7 to 8 minutes. CSCs can be held longer at the plasma membrane upon disruption of actin in the Arabidopsis act2/act7 double mutant

(Sampathkumar et al., 2013). Removal of CSCs from the plasma membrane into CSC- containing compartments would result in CSCs either being trafficked to other locations in the cell or targeted for degradation. Analysis of CSC longevity by treating etiolated hypocotyls with cycloheximide has shown that CESA1, CESA3, and CESA6 persist in the cell either in MASC/SmaCCs or the plasma membrane for up to 48 hours (Hill et al.,

2018). The longevity of CSCs and the relatively high turnover rate at the plasma 348 membrane suggest CSCs are constantly shuttled throughout different regions of the plasma membrane to synthesize cellulose where necessary. The dynamic nature of CSC distribution and redistribution requires dynamic post-translational modifications found in phosphorylation to signal both the reduction in catalytic activity and for endocytosis

(Speicher et al., 2018). Phosphoregulation of CSC catalytic activity has been observed in

CESA5 containing CSCs. In etiolated hypocotyls, CESA5 containing CSCs are immobile at the plasma membrane, but when exposed to light, CSC velocities increased due to phosphorylation of S122, S126, S229, and S230 mediated by PHYTOCHROME B

(Bischoff et al., 2011). Currently, phosphorylation or dephosphorylation events signaling for CSC endocytosis has not been elucidated. Here, the role of glycosylated sphingolipids with the redistribution of CSCs from one location at the plasma membrane to another location would be considered the transport model. In plants with compromised glycosylated sphingolipids biosynthesis, CSCs that are hypothetically phosphorylated signaling for relocation and reduced catalytic activity cannot be endocytosed remaining at the plasma membrane at reduced velocities. The slower complexes allow the endocytosis machinery to “catch” the complexes (Figure 7.3A).

An alternative hypothesis to arrest CSC motility under ND-DNJ, DCB, and

PDMP treatment may be due to alterations in detergent resistant membranes

(DRMs)/lipid rafts. In Arabidopsis, lipid rafts contain the highest concentrations of sphingolipids, phytosterols, and proteins compared to other regions of the plasma membrane (Borner et al., 2005). Current estimates suggest that CSCs are comprised of at least 23 protein subunits (Figure 1.4C and 1.4D). This highly organized conglomerate of proteins can reasonably hypothesize to be embedded in a lipid raft with high 349

Figure 7.3: Proposed models of glycosylated sphingolipid role in CSC motility. (A) The transport model depicts CSCs active at the plasma membrane that are hypothetically phosphorylated (Red spheres), signaling to be endocytosed via a clathrin mediated endocytosis. Glycosylated sphingolipids assist in the membrane organization necessary for endocytosis to transport CSCs to a different location in the cell. Compromised sphingolipid composition may interfere with endocytosis and endomembrane transport. (B) The “icebreaker boat” model depicts glycosylated sphingolipids aiding in lipid raft rigidity aiding the CSC move across the plasma membrane. Compromised sphingolipid composition may create challenges for the CSCs to move across the plasma membrane reducing overall velocities hence, reducing cellulose biosynthesis. Sphingolipids (blue), phospholipids (brown), CSC (gray), Phosphate (red).

350 concentrations of sphingolipids. This hypothesis is further supported in lipid rafts from

Aspen cell suspension cultures produced callose and cellulose synthases (Bessueille et al., 2009). Furthermore, lipid rafts from other species containing polysaccharide biosynthesis proteins have been described (Srivastava et al., 2013; Briolay et al., 2009).

The motility of CSCs may be dependent on GlcCers and GIPCs to maintain a certain rigidity in lipid rafts to move through the plasma membrane. The structure would be analogous to an icebreaker boat that pushes through ice-covered waters. Here, under the momentum of the CSC catalytic activity and the organization of glycosylated sphingolipids and phytosterols allows smoother and unencumbered movement across the plasma membrane (Figure 7.3B).

c. Utility of other iminosugars

In their simplest form, iminosugars typically resemble common pyranose and furanose monosaccharides with the oxygen in the ring substituted with nitrogen. The first naturally occurring iminosugars discovered were 1-deoxynojirimycin (DNJ) and its parent compound nojirimycin (Inouye et al., 1966). Soon after, the first synthetic iminosugars were synthesized (Inouye et al., 1968). Sources of naturally occurring DNJ come from Mulberry trees Morus bombycis and M. niger; additional sources were also discovered in microorganisms such as Bacillus polymyxa and Streptomyces lavendulae

(Hughes and Rudge, 1994). This class of compounds have originally generated interest for its glycosylhydrolase inhibitory properties in mammalian systems. Future studies have utilized DNJ and synthetic derivatives for its potential as therapeutic molecules for a variety of medical conditions, such as genetic disorders in carbohydrate metabolism, 351 cancer, and viral infections (Asano et al., 2000; Nash et al., 2011). A large collection of

DNJ analogs were developed over the years from simple N-alkylated iminosugars, such as ND-DNJ, bicyclic iminosugars (castonaspermin), fluorinated iminosugars and as complex multivalent iminosugars or peptide link iminosugar clusters (Zelli et al., 2016,

2015; Wardrop and Waidyarachchi, 2010; Massicot et al., 2018).

In plants, DNJ and two of its analogs N-5-(adamantane-1-yl-ethoxy)-pentyl-L-ido- deoxynojirimycin (L-ido-AEP-DNJ) and N-butyl-deoxynojirimycin (Andriotis et al.,

2016; Rugen et al., 2018; Dai et al., 2020) were screened for growth inhibitory properties.

DNJ was first screened in barley seed for its ability to inhibit starch catabolism. Barley seed placed in media supplemented with DNJ had poor germination compared to untreated seeds (Andriotis et al., 2016). This same group of researchers demonstrated L- ido-AEP-DNJ severely inhibited root growth in Arabidopsis and Eragrotis tef. The loss of root growth in L-ido-AEP-DNJ treated seedlings was due to decreases in GlcCers, between 3 – 10% in certain GlcCer species (Rugen et al., 2018). This was the first study in plants to demonstrate glycosyltransferase inhibitory properties of iminosugars. GlcCer biosynthesis has been elucidated in Arabidopsis (Msanne et al., 2015), but its catabolism has been poorly understood. Recently, one of the four putative glucosylceramidases

(GCD), GCD3, displayed activity against fluorescently labeled C6-NBD GlcCer with preferential activity towards GlcCers with long acyl chains (24+ carbons). Arabidopsis plants carrying T-DNA insertional mutations, gcd3-1 and gcd3-2, displayed no visible phenotypes. Interestingly, NB-DNJ inhibited GCD activity in vitro (Dai et al., 2020).

This study did not analyze NB-DNJ in vivo, but data generated in chapter 5 demonstrated

NB-DNJ treatment inhibited growth (Figure 5.1) (Table 5.1). Despite the fact that gcd3 352 mutants not display any visible phenotypes, this enzyme is only able to hydrolyze

GlcCers with long acyl chains, which are typically found in sphingolipids associated with cell division and growth (Luttgeharm et al., 2015). It would be interesting to investigate other putative GCD enzymes, as NB-DNJ’s inhibitory effect may target other GCD’s.

IV. Cell wall integrity sensing and differential responses to cell wall damage

Cell wall integrity (CWI) sensing mechanisms function to monitor the status of the cell wall during biotic, abiotic stress, and during developmental processes. Compromises in CWI due to cell wall damage (CWD) increases production of reactive-oxygen species

(ROS) and causes calcium influx leading increased phytohormone production and deposition of lignin and suberin (Gigli-Bisceglia et al., 2020; Vaahtera et al., 2019;

Denness et al., 2011; Gigli-Bisceglia et al., 2018). Of interest, disruption of cellulose biosynthesis invokes a CWD response with deposition of ectopic lignin dependent on receptor-like kinases THESIUS1 (THE1) and STUBBELIG (SUB) (Chaudhary et al.,

2020; Merz et al., 2017).

a. THESIUS1 and STRUBBELIG in response to cellulose defects

The first cellulose integrity sensor to be characterized in Arabidopsis is THE1.

Loss of function mutants of THE1, the1-1 and the1-2 attenuate the negative growth defects of cellulose deficient mutants prc1-1, rsw1-10, eli1-1, and pom1-2 (Hématy et al.,

2007). Interestingly, cellulose content in the1-1/prc1-1 was comparable to prc1-1 indicating despite the1-1 partially rescuing the prc1-1 growth defect, it does not complement cellulose deposition. Furthermore, the1-1/eli1-1 exhibited less ectopic lignin deposition compared to eli1-1 mutant (Hématy et al., 2007). Additionally, multiple the1 353 alleles also displayed reduced ectopic lignin deposition under isoxaben (ISX) treatment

(Merz et al., 2017).

SUB functions in a similar fashion as THE1. Under ISX treatment, sub-1 and sub-

9 had reduced ectopic lignin deposition and the sub-9/prc1-1 partially rescued the prc1-1 phenotype (Chaudhary et al., 2020). A defining difference between the response between

THE1 and SUB is the deposition of callose. Disruption of cellulose biosynthesis with isoxaben causes callose deposition in leaves (DeBolt et al., 2007). In the the1 mutants, callose deposition is unaffected when cellulose biosynthesis is perturbed (Hématy et al.,

2007), but callose deposition was not detected in the sub-9 mutant treated with ISX

(Chaudhary et al., 2020). This may be due to the localization of SUB in the plasmodesmata (Vaddepalli et al., 2014), an organelle where callose is commonly deposited (Chen and Kim, 2009). Interestingly, CWI sensing may be a secondary function of SUB, as its localization with QUIRKY (QKY) at the plasmodesmata and regulates cell morphogenesis, orientation of the division plane, and cell proliferation

(Chevalier et al., 2005; Vaddepalli et al., 2014). QKY is a membrane bound protein containing four C2 domains and a phosphoribosyltranferase C-terminal domain (Fulton et al., 2009). QKY has been suggested to function as a co-signaling component of SUB in regulating cell morphogenesis. Interestingly, the qky-11 allele has a similar abolishment of ectopic lignin deposition as observed in sub-1 and sub-9 when treated with ISX.

Additionally, a reduction in callose deposition was also observed under ISX treatment

(Chaudhary et al., 2020).

b. Lignin and suberin, in response to cell wall damage 354

Upon CWD, a signaling cascade of ROS and Ca+2 mediated by THE1 and SUB leading to transcriptional regulation of defense genes and lignin biosynthetic genes leading to the deposition of ectopic lignin (Hématy et al., 2007; Chaudhary et al., 2020;

Vaahtera et al., 2019; Denness et al., 2011; Duval and Beaudoin, 2009; Merz et al.,

2017). Oddly, CWD cascade of ROS and Ca+2 waves appear to be both opposing and synergistic signals. As a synergistic signal, Arabidopsis plants treated with ISX induces

ROS production and Ca+2 influx. Both signals are required for ectopic lignin deposition, but inhibition of either signal by co-treatment of diphenyleneiodonium (DPI), an

+2 irreversible inhibitor of ROS synthesis, or LaCl3, a Ca influx inhibitor, results in reduced ectopic lignin deposition compared to ISX treated seedlings (Denness et al.,

2011). As antagonistic signals, co-treatment with DPI and ISX results in increases in

Jasmonic acid (JA), and the JA content is reduced in LaCl3 + ISX cotreatment.

Interestingly, exogenous JA treatment also reduced ectopic lignin in ISX treated seedlings (Denness et al., 2011).

In contrast, there is a limited understanding of the molecular mechanisms relating

CWD and ectopic suberin deposition. It appears that wound-induced suberin deposition may not be elicited by mechanical distortion as observed in THE1 and SUB-mediated ectopic lignin deposition under ISX treatment. Lignin deposition can be attenuated by introducing osmotic stressors, such as sorbitol (Hamann et al., 2009). Instead, the damage associated molecular pattern (DAMP), cellobiose, elicits Ca2+ influx that may be associated with suberin deposition. (de Azevedo Souza et al., 2017). Cellobiose up- regulates suberin biosynthetic gene transcripts including a master regulator MYB41 after

25 minutes of cellobiose treatment. Despite increased transcripts of suberin biosynthetic 355 genes, researchers in this study were unable to detect suberin in roots (de Azevedo Souza et al., 2017). The inability to detect suberin may be due to the timing of when the detection assay was conducted, where detectable suberin can be observed at later times, although the researchers did not disclose how they attempted to detect suberin deposition.

Highlighted in chapter 6, ISX and DCB treatment induced suberin deposition in

Arabidopsis seedlings detectable with fluoro yellow 088 staining within 24 hours (Figure

6.3). Together these results suggest that suberin deposition can be induced by CWD possibly dependent on cellulose oligomers from disrupted cellulose biosynthesis (Figure

7.4). Interestingly, a substantial portion of suberin deposition induced by ISX and DCB was non-polymerized. This observation presents a potential opportunity to isolate individual suberin monomers as replacements for non-renewable petrochemical-derived products for plastic manufacturing (Pinto et al., 2009).

V. Possible strategies to isolate novel GTs

One of the initial key goals of utilizing these monosaccharide analogs was to isolate glycosyltransferases involved with primary cell wall biosynthesis. Collectively, the data presented in this dissertation suggest that 2F-Fuc directly inhibits an unknown fucosyltransferase involved in RG-II biosynthesis, and that ND-DNJ and 2F-Man target non-cell wall glycosyltransferases or sugar sensing mechanisms, respectively. These results suggest that 2F-Fuc would be ideal probe molecule to identify fucosyltransferases potentially associated with RG-II biosynthesis. Both ND-DNJ and 2F-Man still have some utility, primarily as suitable compounds to test in vitro enzymatic assays.

a. Capture of GTs with photo-reactive crosslinkers 356

Figure 7.4: Proposed model of suberin deposition under cell wall damage. (A) Upon cell wall damage or inhibition of cellulose biosynthesis, cellulose oligomers are sensed by an unknown cell wall integrity sensor and causes a calcium influx. A signaling cascade occurs and MYB41 expression increases. (B) MYB41 upregulates suberin biosynthetic genes and suberin deposition occurs to reinforce the weakened cell wall.

357

There are a myriad of photo-reactive crosslinkers available, including monosaccharide analogs containing diazirine rings, azido groups, and some iminosugar- based probes. These photo-reactive monosaccharide analogs have been used to covalently modify and capture sugar transporters, glycolipid binding proteins, glycosylhydrolases, enzymes in carbohydrate metabolism, and glycosylhydrolases (Yu et al., 2012; Bond et al., 2011; Yang et al., 2002; Sinnott, 2007). The most flexible and non-specific photo reactive groups are diazirine rings. Diazirine ring-containing probes are relatively stable at room temperature, unreactive with nucleophiles, and stable in acidic or alkaline conditions compared to other photo-reactive crosslinkers that can be activated in a controlled fashion (Dubinsky et al., 2012). Upon UV radiation (350 – 355 nm), diazirines generate carbenes that rapidly forms a covalent bond with the nearest molecule available.

This feature also limits photolabeling yields as the carbenes are rapidly quenched by reacting with nearby water molecules. This situation appears to be a disadvantage, but it is actually an advantage because it minimizes non-specific labeling, as only proteins bound to the label can be covalently linked (Dubinsky et al., 2012).

A potentially interesting strategy to utilize photo-reactive diazirine containing monosaccharide analogs is by exploiting known monosaccharide analogs, such as click compatible monosaccharides Alk-Fuc, 6-Alkyne-6-deoxy-glucose (Alk-Glc), and 8- azido-8-deoxy-KDO (Az-KDO) known to be metabolically incorporate into cell wall polysaccharides (Anderson et al., 2012; Dumont et al., 2016; McClosky et al., 2016)

(Figure 7.5). The utility of a monosaccharide containing both a diazirine ring and an alkyne group will allow both the capture and purification of a glycosyltransferase. For example, a hypothetical 6-alkyne-3-deoxy-3-diazirin-L-fucose (Alk-Dia-Fuc) may be 358

Figure 7.5: Structures of monosaccharides and dual functioning monosaccharide analogs. Peracetylated or native monosaccharides 1,2,3,4-tetraacetyl-L-Fuc (1), 1,2,3,4,6-pentylacetyl-D-Glc (4), and KDO (7). Known click-compatible monosaccharides to label glycans in Arabidopsis 6-alkyne-Fuc (2), 6-alkyne-6-deoxy- Glc (5), and 8-azido-8-deoxy-KDO (9). Hypothetical dual functional monosaccharide analogs containing a diazirine ring and click-compatible functional groups (3, 6, 9).

359 metabolically incorporated, converted to GDP-Alk-Dia-Fuc, and the fucosyltransferase associated with RG-II biosynthesis binds GDP-Alk-Dia-Fuc and RG-II. During this binding event, UV light will photo-capture the fucosyltransferase bound to GDP-Alk-

Dia-Fuc. Protein extraction would follow and click-reaction with the crude extracts and an unconventional azide containing affinity tags such as biotinyl-polyethyleneglycol- azide or β-D-maltosyl azide. The resulting product can be enriched through affinity chromatography and the enrich labeled protein can be analyzed through mass spectrometry. The utility of using Alk-Fuc, Alk-Glc, and Az-KDO is that they function as a scaffold, as the position of their alkyne/azido functional groups are incorporated into cell wall polysaccharide (McClosky et al., 2016; Anderson et al., 2012; Dumont et al.,

2016).

b. Isolation of partial polysaccharide glycans as substrates

A potentially unique strategy to isolate novel glycosyltransferases associated with cell wall biosynthesis is to utilize synthetic monosaccharide analogs to inhibit cell wall biosynthesis to generating glycan fragments to serve as suitable substrates for glycosyltransferase activity assays. For example, 2F-Fuc is a potent inhibitor of fucosylation events, particularly in RG-II fucosylation. Dimerization of RG-II via boron cross-links can be monitored through a modified carbohydrate gel electrophoresis assay

(Chormova et al., 2014; Chormova and Fry, 2015; Voxeur and Fry, 2014). Through this technique, it was demonstrated that 2F-Fuc negatively impacted RG-II dimerization

(Dumont et al., 2015). Additionally, a clearly visible band representing an RG-II fragment that likely does not contain at least one Fucose molecule was observed (Dumont 360 et al., 2015). This assay thus presents a unique opportunity. If the RG-II fragment can be isolated and purified from the endo-polygalacturonase digested cell wall fractions, it would serve as an excellent substrate to test uncharacterized fucosyltransferase (FUT) in vitro. One strategy to isolate RG-II fragments may be from utilizing the gel electrophoresis method developed by (Chormova et al., 2014) to separate the different

RG-II species (dimers, monomers, and fragments), then extract the RG-II fragment by a simple elution method used to extract proteins, essentially excising gel fragments with the desired band, breaking down the gel with plastic pestle, resuspending in a protein buffer, and allowing time for diffusion to occur (Kurien et al., 2019). The limitation of using gel-electrophoresis method lies in the staining method. In order to visualize bands representing RG-II species, silver nitrate is used to stain for RG-II reducing ends, and this method oxidizes RG-II and will potentially make the RG-II fragment unsuitable as a substrate. Alternatively, RG-II dimer and monomer can be separated through size exclusion chromatography (O’Neill et al., 1996). Through some level of optimization, it may be possible to separate dimeric RG-II, monomeric RG-II, and fragmented RG-II; then later confirmed using gel electrophoresis.

Once the RG-II fragment from 2F-Fuc-treated Arabidopsis seedlings is isolated, it can be assayed with recombinant FUT proteins with GDP-Fuc. Out of the 13 known/putative

FUT proteins; FUT2, FUT3, FUT5, FUT7, FUT8, FUT9, and FUT10 have not been characterized (Rocha et al., 2016; Both et al., 2011; Liang et al., 2013b; Sarria et al.,

2001; Perrin et al., 1999; Kaulfürst-Soboll et al., 2011). Detection of fucosyltransferase activity in any recombinant FUT protein will serve as an excellent candidate of RG-II fucosylation. Additionally, it would be interesting to investigate their activity with GDP- 361

L-Gal, as L-Gal serves as an alternative monosaccharide to use in place of Fuc in the mur1 mutant (Bonin et al., 1997; Reuhs et al., 2004).

Despite the complexity of the RG-II structure containing over 20 unique glycosidic linkages, only a handful of the RG-II biosynthetic enzymes have been characterized

(Mohnen, 2008; Egelund et al., 2006, 2008; Dumont et al., 2014). The strategy proposed is limited to two fucosyltransferase, which limits the breath of glycosidic linkages that can be explored, due to only two Fuc residues part of RG-II. Interestingly, a large group of glycosylhydrolases have been isolated from Bacteroides thetaiotaomicron that hydrolyze all but one of the over 20 unique glycosidic linkages (Ndeh et al., 2017). This observation opens the number of possible RG-II fragments that could be generated as partial RG-II substrates.

VI. Implications of monosaccharide analogs as herbicides

The severe growth phenotype observed in 2F-Fuc, 2F-Man, and ND-DNJ treatment in

Arabidopsis highlight a potential framework to develop novel herbicides and fungicides for future applications in agriculture or in pharmaceutical use. There has been an increasing number of herbicide resistant weeds found in agricultural fields, driving the demand for novel methods of weed management. Currently, there is a variety of commercially available herbicides that target 25 biological processes including photosynthesis, biosynthesis of cellulose, lipids, amino acids, and phytohormones such as auxin (Dayan et al., 2019). A potentially unique aspect of 2F-Fuc, 2F-Man and ND-DNJ as potential herbicides is that they target biological processes that commercially available herbicides do not. Additionally, resistance to these monosaccharide analogs may take 362 longer to appear in weeds through target-site resistance (TSR) or nontarget-site resistance

(NTSR) mechanisms, as they closely resemble native monosaccharides. A common TSR mechanism involves a single nucleotide mutation in a gene encoding the target protein resulting in a single amino acid change that disrupts the ability of the herbicide to bind

(Gaines et al., 2020). In 2F-Fuc and 2F-Man TSR is less likely to occur as fluorine van der Waals radius is similar to oxygen (Liang et al., 2013a), where a single amino acid change would also limit the use of Fuc and Man. NTSR mechanisms are generally more complex and varied compared to TSR, these mechanisms can include reduced herbicide absorption, sequestration into vacuoles, or metabolized to an inactivate metabolite

(Gaines et al., 2020). The most susceptible monosaccharide analog is 2F-Fuc as we have essentially described an NTSR mechanism through the fkgp-3 mutant (Villalobos et al.,

2015), It is less likely a NTSR mechanism will rapidly evolve in weeds with 2F-Man as

Man is an essential monosaccharide to a variety of biological processes in plants including in HXK1 glucose sensing as described in chapter 3. Hypothetical resistance mechanisms to ND-DNJ in weeds is unclear as it is likely able to permeate through the cell and a random mutagenesis screen we conducted on ND-DNJ yielded no potentially resistant candidates (data not shown). The potential drawback of 2F-Fuc and 2F-Man is the fluorine atom in their structures. It is unclear how stable these analogs are in the environment nor what their potential breakdown products are. It would appropriate to designate these monosaccharide analogs as starter molecules to explore other functional groups in the 2-position. Overall, the three monosaccharide analogs described in this dissertation are excellent starter molecules to develop the next set of herbicides for weed management strategies. 363

VII. Other monosaccharide analogs to explore in plant growth

Over the course of this dissertation, the primary focus was to utilize monosaccharide analogs to probe for glycosyltransferases related to cell wall polysaccharide biosynthesis by first investigating their capacity to inhibit growth in Arabidopsis. As a result, four monosaccharide analogs 2F-Fuc, 2F-Man, 2F-Glc, and ND-DNJ are potent inhibitors of

Arabidopsis growth. Two of these analogs do have some glycosyltransferase inhibitory properties in RG-II fucosylation and glycosylated sphingolipid biosynthesis.

Unexpectedly 2F-Man and 2F-Glc induce glucose growth repression in Arabidopsis and

S. cerevisiae. The primary class of monosaccharide analogs explored in this piece are fluorinated monosaccharides and iminosugars. There are other classes of monosaccharide analogs to explore and there are other monosaccharides analogs in the fluorinated and iminosugar classes to be explored. It would be interesting to analyze other 2-deoxy-2- fluoro monosaccharides not explored here, such as 2-deoxy-2-fluoro-L-rhamnose, 2- deoxy-2-fluoro-D-xylose, and 2-deoxy-2-fluoro-L-arabinose to probe for RG-I backbone,

RG-II backbone, and RG-II side chains containing D-xylose and L-arabinose (Zilei et al.,

2016; Williams et al., 2006; Wong et al., 1988) (Figure 7.6).

One class of monosaccharide analogs to explore to probe for glycosyltransferases involved in plant cell wall biosynthesis are pseudosugars or more commonly known as carbasugars. Similar to iminosugars, the defining feature of carbasugars is the substitution of the oxygen in the pyranose/furanose ring with a methylene group (Figure

7.5). Commonly, carbasugars are commonly known as glycosylhydrolases inhibitors

(Wardrop and Waidyarachchi, 2010; Roscales and Plumet, 2016). This accepted 364

Figure 7.6: Structures of alternative fluorinated monosaccharides and Carbasugars. Structures of alternative fluorinated monosaccharides 2-deoxy-2-fluoro-L-Rhap (11), 2-deoxy-2-fluoro-Xylp (12), 2-deoxy-2-fluoro-Arap (13), and 2-deoxy-2-fluoro-Araf (14). Structures of native monosaccharide (15) and carbasugar (16). Commercially available carbasugars Carba-α-D-galactopyranose (17), Cyclophellitol (18), MK7607 (19), Streptol (20), (+)-Pericosine A (21), (-)-Gabosine A (22), Validamine (23), Valienamine (24), and Voglibose (25).

365 paradigm should not limit exploration as hypothetical glycosyltransferase inhibitors as iminosugars have been commonly perceived as glycosylhydrolases inhibitors, but have some glycosyltransferase inhibitory properties, as described in chapter 5 and other previous studies (Rugen et al., 2018; Nash et al., 2011). These observations may be attributed to the similar transition states between non-retaining glycosyltransferases and non-retaining glycosylhydrolases (Lairson et al., 2008; McCarter and Stephen Withers,

1994). Few examples of carbasugars have been utilized in plants other than in fungicide application in rice and treatment with validamycin inhibiting trehalase in Medicago trancatula causing accumulation of trehalose leading to salt tolerance (López et al., 2009;

Roscales and Plumet, 2016). There is a large diversity of carbasugars commercially available to explore for growth inhibition in Arabidopsis (Figure 7.6).

VIII. Conclusions

Our initial goals of using monosaccharide analogs were to probe cell wall function, but most of them has led us in unexpected directions. 2F-Fuc was the first promising monosaccharide analog that suggested these molecules can be used to probe cell wall function. Arabidopsis treated with 2F-Fuc displayed severe growth inhibition as a result of RG-II fucosylation inhibition. Additionally, multiple 2F-Fuc resistant lines from an

EMS screen showed promising novel avenues to study Fuc metabolism and RG-II biosynthesis (Chapter 2). Shifting focus to 2F-Man characterization in Arabidopsis was the first molecule to go in an unexpected direction. Instead of inhibiting mannosylation events as we hypothesized, it was through the action of AtHXK1-dependent glucose repression that was the result of growth repression we observed. We also isolated 2F-Man 366 resistant lines that correspond to AtHXK1 (Chapter 3). The dependence of 2F-Man toxicity on the glucose repression model was further reinforced in S. cerevisiae, as 2F-

Man toxicity was reliant in a ScHXK2-dependent glucose repression. Interestingly, the

Rim101 pathway also plays a role in 2F-Man toxicity (Chapter 4). The iminosugar, ND-

DNJ, perturbed glycosylated sphingolipid biosynthesis that resulted in reduced CSC velocities negatively impacting crystalline cellulose deposition. Unexpectedly, further experimentation revealed PDMP and DCB inhibit both GlcCer and GIPC glycosylation resulting in reduced crystalline cellulose deposition (Chapter 5). Finally, investigating perturbed cellulose deposition under the action of DCB and ISX, resulted in monomeric and polymerized suberin deposition not just ectopic lignin (Chapter 6). These results collectively highlighted the potential utility of these monosaccharide analogs and opened a variety of experimental avenues to explore. 367

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