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Investigating the role of sucrose phosphate synthase and hexokinase in carbon sink strength. Lazarova, Sofiya 2015

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INVESTIGATING THE ROLE OF SUCROSE PHOSPHATE SYNTHASE AND HEXOKINASE IN CARBON SINK STRENGTH by  Sofiya Lazarova  B.Sc. (Hons.), The University of Toronto, 2012  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  MASTER OF SCIENCE in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Forestry)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  October 2015  © Sofiya Lazarova, 2015   ii Abstract  The production, transport and assimilation of organic carbon ultimately drive the growth of plants. In this work, two enzymes, Sucrose Phosphate Synthase (SPS) and Hexokinase (HXK), prominent in their role of carbon production in the form of sucrose at the source, have been examined for their role at the sink, where carbon is assimilated. It has been postulated that the presence of sucrose-forming enzymes in the sink serves a function to reform sucrose from apoplastic cleavage or partake in a “futile” cycle of sucrose cleavage and such that small changes in metabolite enable large changes in sink carbon strength. In order to determine if SPS is involved in carbon sink strength, A. thaliana TDNA insertional lines and P. trichocarpa RNAi stem and developing xylem with decreased SPS transcript expression were analyzed. It was determined that loss of SPS transcript generally increases soluble sugars: sucrose, glucose and fructose, in the leaf and stem as well as starch in the leaf. Structural carbohydrates were generally unaffected and Klason soluble lignin decreased. Similarly, A. thaliana TDNA insertional lines with decreased HXK transcript expression were utilized to determine the role of HXK using stem tissue as a carbon sink model. Soluble sugars mainly increased in the leaf of athxk3 TDNA insertional line whereas starch increased in both leaf and stem of the same line. Interestingly, structural carbohydrate levels of the cell wall were perturbed in HXK TDNA insertional lines. The results were found to be consistent with the postulated roles of SPS and HXK that predict a function in sucrose formation from apoplastic cleavage, which allows for fine-tuning of major intracellular metabolites and adjustment of sink strength.    iii Preface  This thesis is the original, unpublished and independent work by the author, S. Lazarova.   iv Table of Contents  Abstract .......................................................................................................................................... ii!Preface ........................................................................................................................................... iii!Table of Contents ......................................................................................................................... iv!List of Tables .............................................................................................................................. viii!List of Figures ............................................................................................................................... ix!Acknowledgements ...................................................................................................................... xi!Dedication .................................................................................................................................... xii!Chapter 1: Introduction ................................................................................................................1!Chapter 2: Methodology ..............................................................................................................11!2.1! Selection of A. thaliana genes and P. trichocarpa orthologs .......................................... 11!2.2! Phylogenetic analysis ....................................................................................................... 12!2.3! Arabidopsis TDNA mutants ............................................................................................ 13!2.4! DNA screening ................................................................................................................. 14!2.5! RNAi down-regulation construct development ............................................................... 15!2.5.1! RNAi constructs targeting a single gene ...................................................................15!2.5.2! RNAi constructs targeting multiple genes .................................................................17!2.6! Agrobacterium transformation ......................................................................................... 23!2.7! Hybrid poplar transformation .......................................................................................... 24!2.8! Plant growth and phenotyping ......................................................................................... 25!2.8.1! A. thaliana TDNA insertional lines ...........................................................................25!2.8.2! Hybrid poplar hpRNAi lines ......................................................................................25!  v 2.9! RNA isolation and cDNA production .............................................................................. 26!2.10! Semi-quantitative and quantitative RT-PCR ................................................................. 27!2.11! Non-structural sugars and starch determination ............................................................ 28!2.12! Cell wall chemical analysis ............................................................................................ 30!2.13! High-performance liquid chromatography .................................................................... 30!2.14! Statistical analysis .......................................................................................................... 31!Chapter 3: Results ........................................................................................................................32!3.1! Sucrose phosphate synthase ............................................................................................. 32!3.1.1! Phylogenetic analysis of the A. thaliana and P. trichocarpa protein families ..........32!3.1.2! Endogenous expression of SPS genes in A. thaliana and P. trichocarpa in sink and source tissues ........................................................................................................................ 34!! A. thaliana .......................................................................................................... 34!! P. trichocarpa .................................................................................................... 35!3.1.3! atsps1-1, atsps1-2 and atsps2 A. thaliana TDNA insertional mutants ......................36!! Relative expression of AtSPS1 and AtSPS2 in TDNA insertional mutants ....... 36!! Phenotypic analysis of plant growth .................................................................. 38!! Non-structural carbohydrates in leaf and stem .................................................. 40!! Starch content in leaf and stem .......................................................................... 43!! Structural carbohydrates and lignin of the stem ................................................ 44!3.1.4! RNAi hybrid poplar down-regulation ........................................................................46!! Relative expression of PtSPS1 and PtSPS3 ....................................................... 46!! Phenotypic analysis of plant growth .................................................................. 49!! Non-structural carbohydrates in cambium and source leaf ................................ 51!  vi! Starch content in cambium and source leaf ....................................................... 54!3.2! Hexokinase ....................................................................................................................... 55!3.2.1! Phylogenetic analysis of A. thaliana and P. trichocarpa Hexokinase protein families…… .......................................................................................................................... 55!3.2.2! Endogenous expression .............................................................................................59!! A. thaliana .......................................................................................................... 59!! P. trichocarpa .................................................................................................... 60!3.2.3! athxk1, athxk2 and athxk3 TDNA insertional mutants ..............................................62!! Relative expression of AtHXK1, AtHXK2 and AtHXK3 .................................... 62!! Phenotypic analysis of plant growth .................................................................. 64!! Non-structural carbohydrate in leaf and stem .................................................... 68!! Starch content in leaf and stem .......................................................................... 70!! Structural carbohydrates and lignin in the stem ................................................. 71!Chapter 4: Discussion ..................................................................................................................75!4.1! Effects of loss of SPS in A. thaliana and P. trichocarpa ................................................. 75!4.1.1! Phylogenetic relationships between plant SPS genes and putative function of A. thaliana and P. trichocarpa enzymes based on endogenous expression. ............................. 78!4.1.2! Loss of photosynthetic SPS .......................................................................................81!4.2! Effects of loss of HXK in A. thaliana .............................................................................. 84!4.2.1! Phylogenetic relationships between A. thaliana and P. trichocarpa HXKs ..............93!4.2.2! Effects of loss of AtHXK1, AtHXK2 and AtHXK3 functional enzymes in A. thaliana....... .......................................................................................................................... 94!Chapter 5: Conclusion ...............................................................................................................100!  vii 5.1! Proposed function of SPS enzyme ................................................................................. 100!5.2! Proposed function of HXK enzyme ............................................................................... 102!5.3! An integrated view of sucrose metabolism .................................................................... 104!References: ..................................................................................................................................107!   viii List of Tables  Table 1: List of primers and their uses. ......................................................................................... 18!Table 2: hpRNAi vectors and the coding sequence base pair position of the genes targeted. ...... 23!Table 3: Average quantity of sugar per mass of sample (ug/mg) in atsps1-1, atsps1-2 and atsps2 main stems. ................................................................................................................................... 45!Table 4: Average percent insoluble, soluble and total lignin in the main stem of atsps1-1, atsps1-2 and atsps2................................................................................................................................... 46!Table 5: Average quantity of sugar per mass sample (ug/mg) in athxk1, athxk2 and athxk3 primary stems. ............................................................................................................................... 73!Table 6: Average percent insoluble, soluble and total lignin in the main stem of athxk1, athxk2 and athxk3. .................................................................................................................................... 74!   ix List of Figures  Figure 1: Construction of hpRNAi using the pHELLSGATE12 Gateway cloning system. ........ 16!Figure 2: Condensed phylogenetic tree of plant SPSs. ................................................................. 33!Figure 3: Endogenous expression of SPS genes; AtSPS1 (red dot), AtSPS2 (blue dot), AtSPS3 (green dot) and AtSPS4 (brown dot) in select A. thaliana tissues. ............................................... 35!Figure 4: Endogenous expression of SPS genes in P. trichocarpa cambium and source leaf. ..... 36!Figure 5: atsps1-1, atsps1-2 and atsps2 are TDNA insertional mutants. ..................................... 37!Figure 6: Rosette diameter and stem height of atsps1-1, atsps1-2 and atsps2. ............................ 39!Figure 7: Non-structural carbohydrate contents in stem and leaf of atsps1-1, atsps1-2 and atsps2 harvested at midday. ..................................................................................................................... 42!Figure 8: Percent starch in the stem and leaf of atsps1-1, atsps1-2 and atsps2 harvested at midday. .......................................................................................................................................... 44!Figure 9: Quantitative PCR ........................................................................................................... 49!Figure 10: Stem growth parameters of PtSPS1/3i and PtSPS1/3::PtSPS2i RNAi down regulated hybrid poplar following four months of growth in a greenhouse. ................................................ 51!Figure 11: Non-structural carbohydrates in cambium and source leaf of PtSPS1/3i and PtSPS1/3::PtSPS2i RNAi down-regulated hybrid following four months of growth in a greenhouse and harvested at midday. ........................................................................................... 54!Figure 12: Percent starch content in the cambium and source leaf of PtSPS1/3i and PtSPS1/3::PtSPS2i RNAi down regulated hybrid following four months of growth in a greenhouse and harvested at midday. ........................................................................................... 55!Figure 13: Condensed phylogenetic tree of plant Hexokinase proteins. ...................................... 58!  x Figure 14: Multiple sequence alignment of AtHXK and PtHXK family of proteins using MUSCLE algorithm. ..................................................................................................................... 59!Figure 15: Endogenous expression of the HXK family of genes in select A. thaliana tissues. .... 60!Figure 16: Endogenous expression of HXK transcripts in cambium and source leaf of P. trichocarpa. ................................................................................................................................... 61!Figure 17: athxk1, athxk2 and athxk3 are TDNA insertional mutants. ......................................... 63!Figure 18: athxk1, athxk2 and athxk3 TDNA seedlings on control media, 2-D-glucose; a non-metabolizable glucose analog, or 6% glucose after 7 DAG. ........................................................ 64!Figure 19: Rosette diameter at 21 DAG of athxk1, athxk2 and athxk3 TDNA insertional mutant lines. .............................................................................................................................................. 66!Figure 20: Stem height of athxk1, athxk2 and athxk3 TDNA mutant lines. ................................. 67!Figure 21: Non-structural carbohydrate in the stem and leaf of athxk1, athxk2 and athxk3 TDNA insertional lines harvested at midday. ........................................................................................... 70!Figure 22: Percent starch in the stem and leaf of athxk1, athxk2 and athxk3 TDNA mutant lines harvested at midday. ..................................................................................................................... 71!Figure 23: An integrated view of sink cell carbohydrate metabolism. ....................................... 105!     xi Acknowledgements  I would like to extend my gratitude to my supervisor, Dr. S. Mansfield for providing me with the opportunity to conduct this research, being patient with my mistakes and offering scientific guidance throughout my time in his laboratory.   I would like to thank all past and present laboratory members for taking time out of their day to help me with any questions, for being extremely patient with me, and most importantly, for being excellent friends both in and outside the lab – it was a pleasure to work with you every single day.  Finally I would like to acknowledge the greater Faculty of Forestry community including all the graduate students who made my time here all the more enjoyable.    xii Dedication  I dedicate this work to my mother, Mariyana and my father Nedialko, who have always been the most supportive and loving parents I could ever wish for. To my sister Yanna and my grandparents who are always inspiring me to continue my education onward. I also dedicate this to my partner Brendan, who keeps me grounded when nothing else will. Thank you for believing in me.    1 Chapter 1: Introduction  The building block of plants, organic carbon, is the result of photosynthetic carbon fixation. Amospheric CO2 acquired through plant stomata diffuses into the chloroplasts of leaf mesophyll tissue. There, it is incorporated into organic molecules through the Calvin-Benson cycle to produce triose-phosphates (Triose-P). Triose-P may remain in the chloroplast or be exported to the cytoplasm. In either case, two triose-P are combined to form fructose-1-6-bisphosphate (F1,6-BP) that is consecutively converted to fructose-6-phosphate (F6P), glucose-6-phosphate (G6P) and glucose-1-phosphate (G1P). In the chloroplast, ATP and G1P yields ADP-glucose (ADP-G), which can be used for starch synthesis, a way to store excess photosynthate as transitory starch. In the cytoplasm, G1P can react with a UTP molecule via UDP-glucose pyrophosphorylase (UGP) to form UDP-glucose (UDP-G) (Munch-Petersen, Kalckar, Cutolo, & Smith, 1953). Previously produced F6P (or isomerized G6P) and UDP-G combine to form sucrose-6-phosphate (S6P) in a reversible reaction catalyzed by sucrose phosphate synthase (SPS): UDP-G + F6P !" S6P + UDP  The phosphate residue can thereafter be quickly cleaved by sucrose phosphate phosphatase (SPP) making the reaction essentially irreversible. It is believed that SPS and SPP form an in vivo complex to form a metabolic channel (Maloney, Park, Unda, & Mansfield, 2015). These sucrose molecules can be imported into the vacuole by tonoplast monosaccharide transporters (TMTs) and vacuolar glucose transporters (VGTs)  (Thompson & van Bel, 2012)  or exported out of the cell either symplastically through plasmodesmata, or apoplastically through a sucrose exporter. Alternatively, UDP-G may combine with G6P in a reaction catalyzed by trehalose   2 phosphate synthase (TPS) to form trehalose-6-phosphate (T6P). Similar to S6P, trehalose phosphate phosphatase (TPP) cleaves the phosphate to form trehalose. Trehalose, however, exists in a comparatively small concentration in A. thaliana (Paul, Primavesi, Jhurreea, & Zhang, 2008).    In the dark, the starch carbohydrate reserve stored in the chloroplast is broken down into the disaccharide maltose, glucose or G1P to provide energy when Triose-Ps are not available. Maltose and glucose are exported to the cytoplasm through specific carriers, where maltose is broken down into two glucose molecules (S. Zeeman, Smith, & Smith, 2007). In order to be used for metabolic reactions, such as glycolysis and respiration, glucose is phosphorylated by hexokinase (HXK) to form G6P: hexose + ATP " hexose-6-phosphate + ADP  Stored glucose can also be derived from the vacuole where sucrose is broken down to glucose and fructose by vacuolar invertase (VINV), and the breakdown products passively exported through carriers of the EDR6-like subfamily from the monosaccharide transporters (MST) superfamily (Slewinski, 2011). Sucrose may also be released during the day from the vacuole to balance cytosolic sucrose levels (Thompson & van Bel, 2012). In addition, sucrose may be resynthesized from these monosaccharides, after phosphorylation and interconversion, and again exported symplastically or apoplastically to support the metabolism of sink organs (Granot, David-Schwartz, & Kelly, 2013) .   Sucrose concentrations in the cytosol of mesophyll cells, at any time during the diurnal cycle, drive the passive export of carbohydrate through plasmodesmata to the bundle sheath/phloem   3 parenchyma cells (BSC/PCC). Energy-coupled carriers retrieve any leaked sucrose from the surrounding apoplasm. From there, symplastic loading can continue passively down the concentration gradient into companion cells (CC) and finally into the sieve tube of phloem. Alternatively, the synthesis of raffinose family oligosaccharides (RFO) from sucrose in intermediary cells (modified companion cells) can actively trap these carbohydrates, which are released to the sieve element through large pore plasmodesmata (Unda, Canam, Preston, & Mansfield, 2012). Apoplasmic loading of photosynthate includes the release of sucrose, amino acids and polyols through putative efflux carriers in the BSC/PCC and subsequent uptake by energy dependent carriers on the companion cell-sieve element (CC/SE) complex. This uptake also includes the exchange of water, K+ and amino acids derived from the xylem (Thompson & van Bel, 2012) .   Bulk flow has been accepted as the principle method of solute translocation through the transport phloem, as passive diffusion is unable to sustain the observed flux through sieve tubes. The theoretical aspect of bulk flow is encapsulated by the Münch pressure flow hypothesis (Munch, 1930), which states that the phloem transport rate is the product of the sieve tube cross-sectional area, phloem sap concentration and transport velocity for a certain nutrient species. Transport velocity on the other hand, is determined by the hydrostatic pressure difference between the source and the sink. Hydrostatic pressure is created at the source by the loading of major osmotic species into the phloem sap, of which sugars account for at least 50% (Thompson & van Bel, 2012). Sucrose loading can, therefore, be said to drive bulk flow from source to sink in plants where sucrose is the main transport carbohydrate.     4 Unloading of phloem sap nutrients takes place at sinks along the axial phloem path, often in arranged in series. Various sink types inherently exist including vegetative apices (root apical meristem, shoot apical meristem), stem elongation zones, mature stem zones, terminal vegetative storage organs (tubers), and terminal reproductive storage sinks (fleshy fruit, developing seed) (Patrick, 1997). Unloading may occur only apoplastically such as in meristematic apical zones, mostly symplastically such as in high source to sink ratios in elongating or mature stem/root, or mostly apoplastically in low source to sink scenarios of the elongating or mature stem/root. Irrespective of the unloading pathway, some sucrose leaks from the SE and is transported back in through turgor-regulated transporters located on SE-CC plasma membranes. In areas other than vegetative meristems, phloem sap arrives at the adjoining vascular parenchyma cells (VPC) symplastically through bulk flow in plasmodesmata from the CC. From there sucrose may be transported again symplastically directly into sink cells or apoplastically through carrier mediated transport, with the two possibilities not being mutually exclusive. Apoplastic loading, however, allows for the accumulation of nutrients against the concentration gradient, as it is an energy-expending process (Turgeon, 2006).   Once in the sink cell, sucrose is rapidly broken down and resynthesized. When sucrose was labelled asymmetrically in tomato fruit, 5-10% of the label was detected symmetrically after 1 hour of in vitro chase and following 2 hours, 60-90% was detected symmetrically (N'tchobo, Dali, Nguyen-Quoc, Foyer, & Yelle, 1999). In order to be metabolized, stored in vacuoles, or plastids as starch, sucrose must be broken down by one of sucrose synthase (SuSy) or cytosolic invertase (INV). SuSy cleaves sucrose in the presence of UDP to form UDP-G and a fructose molecule that can be utilized for cellulose synthesis, glycosylation reactions and resynthesis of   5 S6P by SPS (Bar-Peled & O'Neill, 2011; Dennis & Blakeley, 2000). In addition to UDP, it has been suggested that SuSy may use ADP to form ADP-glucose (ADP-G) that can then be utilized for starch synthesis (Baroja-Fernandez et al., 2009). SuSy is a cytoplasmic protein but has also been found to be associated with the plasma membrane suggesting a role in UDP-G channelling for cell wall synthesis. Indeed, overexpression of SuSy in poplar increased cellulose production and wood density in developing xylem (Coleman, Yan, & Mansfield, 2009). INV on the other hand, uses no energy to produce glucose and fructose monomers, but those monomers need to be phosphorylated by HXK and fructokinase (FRK), respectively, for further metabolism. Glucose and fructose monomers may also be created by the action of cwINV from apoplastically derived sucrose (Canam et al., 2006; Canam, Mak, & Mansfield, 2008; Canam, Unda, & Mansfield, 2008). Apoplastic sucrose, glucose and fructose are imported into the cell through sucrose or hexose carriers (Lalonde, Wipf, & Frommer, 2004).   The allocation of source supplies to various sinks is determined by the ability to sense sucrose and hexoses so that appropriate mechanistic changes to the system are made (Bihmidine, Hunter III, Johns, Koch, & Braun, 2013). When sink partitioning is low, photosynthesis is upregulated in the source, and conversely, when photosynthate is abundant sinks are upregulated in enzymes which cleave, store and import sucrose (Kang, Price, Lin, Hong, & Jang, 2010; Li, Xing, Gianfagna, & Janes, 2002; Ramon, Rolland, & Sheen, 2008; Smidansky et al., 2002). Such signals are believed to include sugar sensing by TPS, which produces T6P. T6P exerts its effect by repressing Suc-non-fermenting related kinase-1 (SnRK1), itself a repressor of plant growth, to promote cell wall biosynthesis, starch production and growth. In the developing seed and   6 seedling, G-protein coupled receptors are involved in the sensing of extracellular glucose (Chen 2003). Moreover, hexokinase-1 (HXK1), usually associated with the mitochondrial membrane where it regulates sugar partitioning to the glucose metabolome, has been show to translocate to the nucleus and control the degradation and transcription of the EIN3 transcription factor involved in ethelyne signaling (Yanagisawa, Yoo, & Sheen, 2003). These are just some of the mechanisms associated with cellular sucrose homeostasis, but consistently, in many organisms it has been found that in the sink, SuSy, INV and SPS exert the biggest mechanistic effects on sucrose metabolism. In crops such as sugarcane, SuSy protein is largely regarded as a marker for sucrose accumulation and sink strength (Grof et al., 2007). INV activity in tomato fruit has been associated with late stage accumulation of soluble sugars (Wang, Sanz, Brenner, & Smith, 1993), and when SPS was overexpressed in tobacco, sink sucrose pools were substantially elevated compared to the source (Park, Canam, Kang, Ellis, & Mansfield, 2008) .   The prevailing view is that various sink types compete for nutrients by way of increasing sink strength or maximizing the ability to assimilate the carbon through growth, storage, or respiration. However, carbon may be compartmentalized by various phloem connections, thus making phloem an additional pool for which sinks compete. Thus, it has been suggested that plants actively adjust allocation to sources and sinks rather than sources and sinks passively adjusting allocation (Dietze et al., 2014). On the molecular level, this sugar responsiveness is exhibited on the transcriptional, translational and protein turnover/allosteric regulation levels in order to adjust for the allocation of source supplies to the sink (K. Koch, 1996; K. E. Koch, Ying, Wu, & Avigne, 2000; Ramon et al., 2008; Ruan, 2014; Tiessen & Padilla-Chacon, 2012; Xiong   7 et al., 2013) . The genes responsible include those of sucrose import and use, with INV, SuSy and SPS found to be major players (Nguyen-Quoc & Foyer, 2001) .   It has long been recognized that “futile” cycles of sucrose degradation and resynthesis occur in a variety of sink organs (Geigenberger & Stitt, 1991) where net sucrose degradation occurs. In general, the two enzymes responsible for sucrose degradation, INV and SuSy, have been correlated with hexose accumulation and cellulose synthesis and overall growth (Canam et al., 2008; Coleman et al., 2009). SPS has been found to be expressed in a variety of sink organs - both sucrose accumulating such as sugarcane (Grof et al., 2007), and non-sucrose or hexose accumulating species such as tobacco (Chen, Hajirezaei, & Bornke, 2005; Park et al., 2008; Park, Canam, Kang, Unda, & Mansfield, 2009) - and functions to synthesize sucrose in those organs. Generally, the seemingly wasteful breakdown and resynthesis of sucrose is considered to be necessary following apoplastic unloading so that further storage or intercellular transport can occur (Ruan, 2014), since sucrose is more stable than hexoses due to a lack of a reducing end. Alternatively, sucrose resynthesis may be a mechanism to directly regulate the carbohydrate flux from source to sink tissue. This involves the production of sucrose or the degradation of starch at the source and resynthesis at the sink. Thus, small changes in sucrose concentration can result in large fluxes between the two (Geigenberger et al., 1997) and the plant can therefore quickly respond to changing biotic or abiotic factors. These two alternative hypotheses for the role of SPS in non-photosynthetic tissue have not been adequately teased apart.     8 In tomato fruit, several such futile cycles have been identified: 1) continuous and rapid degradation of sucrose by SuSy in the cytosol and resynthesis by SuSy or SPS; 2) hydrolysis of sucrose in the apoplast or vacuole, import of apoplastic hexoses and subsequent resynthesis in the cytosol; and, 3) rapid synthesis and breakdown of starch in the amyloplast (Nguyen-Quoc & Foyer, 2001). In the initial rapid growth phase of tomato, SuSy predominates in unloading of sucrose symplastically as plants with decreased SuSy activity had a substantially lower sucrose unloading capacity (D'Aoust, Yelle, & Nguyen-Quoc, 1999). Subsequent sucrose resynthesis may be catalyzed by SPS but also by SuSy, which functions in the synthesis direction of the reaction when intracellular hexose concentrations are high (Geigenberger & Stitt, 1993). Intracellular concentrations of sucrose vary widely during tomato fruit development. Once sucrose is imported into the vacuole it is constantly degraded into hexoses by acid invertase (aINV) to maintain sucrose to hexose ratios in that compartment (Ohyama et al., 1995). Glucose and fructose are exported and resynthesized to sucrose by SPS. Finally, starch is continually broken down and resynthesized in the amyloplast as it has been shown that newly accumulated starch is always proportional to previously synthesized starch and the amount of newly unloaded sucrose used for starch is always a constant twenty five percent (N'tchobo et al., 1999). For the inclusion of glucose and fructose into the futile cycles of degradation, whether released by the action of cwINV or aINV, HXK and FRK phosphorylation is required. These interconversion reactions essentially regulate the major metabolites; UDP-G, G1P, F6P, G6P, glucose, fructose and sucrose, that are responsible for cellulose synthesis, starch accumulation, respiration, storage in the vacuole and others (Nguyen-Quoc & Foyer, 2001) .     9 SPS and HXK are two enzymes whose importance in sucrose metabolism has clearly been demonstrated in photosynthetic tissue. Recent evidence has established a role for both in non-phosynthetic tissue as well. For SPS that role could be to synthesize sucrose from apoplastic cleavage products of cwINV action or to play a role in “futile” sucrose cycling along with other enzymes, so that it promotes large changes in carbon flux to the sink. In order to investigate the role of SPS in the sink, using A. thaliana stem and P. trichocarpa developing xylem as models, experiments in both organisms were undertaken. Firstly, known A. thaliana SPS isozymes were used to identify P. trichocarpa orthologs and all genes were examined for level of expression in the defined source and sink tissue. In A. thaliana, a quick growing and widely used plant model organism, homozygous TDNA insertional lines were utilized to study the chosen genes. Those lines were phenotyped visually using growth measurements and chemically through measurement of nonstructural sugars and starch in the leaf and stem, and structural sugars and lignin content of the stem. In an effort to corroborate evidence gathered in A. thaliana, hybrid poplar was genetically manipulated using RNA interference (RNAi) technology to decrease SPS transcript abundance. Genetically manipulated hybrid poplar was also visually and chemically phenotyped using growth parameters, nonstructural carbohydrates and starch of the source leaf and developing xylem. No matter the origin, in order for glucose and other monosaccharides to be used in sink cell metabolism they must be phosphorylated. FRK activity has been shown to be strongly correlated with xylem sink activity in tomato (Damari-Weissler et al., 2009) and hybrid poplar (Roach et al., 2012).   In order to determine whether HXK has a similar effect in carbon flux to the stem, A. thaliana homozygous TDNA insertional lines where measured for a change in growth phenotype during   10 development. In addition, chemical phenotyping was undertaken of nonstructural carbohydrates and starch in the leaf and stem as well as structural carbohydrates and lignin content of the stem. The following chapters describe the findings of the aforementioned experiments, provide interpretation of the data with respect to previous findings and attempt an explanation of the results with room for further experimentation.    11 Chapter 2: Methodology  2.1 Selection of A. thaliana genes and P. trichocarpa orthologs Annotated sequences of four known sucrose phosphate synthase (SPS) genes; AT5G20280 (AtSPS1), AT5G11110 (AtSPS2), AT1G04920 (AtSPS3), AT4G10120 (AtSPS4) in Arabidopsis thaliana were obtained from The Arabidopsis Information Resource (Lamesch et al., 2012) and BLAST-ed against the Populus trichocarpa genome in Phytozome V10 (Goodstein et al., 2012) to identify the corresponding orthologs; Potri.006G064300 (PtSPS1), Potri.018G026100 (PtSPS2), Potri.0018G124700 (PtSPS3), Potri.013G095500 (PtSPS4), Potri.001G317600 (PtSPS5), and Potri.017G057800 (PtSPS6). Of these, three genes, Potri.006G064300 (PtSPS1), Potri.018G026100 (PtSPS2) and Potri.0018G124700 (PtSPS3), were found to be the most highly expressed in the developing poplar xylem versus leaf tissue based on transcriptome sequencing data of 20 field-grown trees. Therefore, three genes were chosen for further investigation (Figure 4) (Hefer, Mizrachi, Myburg, Douglas, & Mansfield, 2015). The orthologs of Potri.006G064300 (PtSPS1), Potri.018G026100 (PtSPS2), Potri.0018G124700 (PtSPS3); AT5G20280 (AtSPS1) and AT5G11110 (AtSPS2), were also found to be the most highly expressed SPS genes in A. thaliana xylem, based on the Arabidopsis eFP browser (D. Winter et al., 2007) and Genevestigator (Hruz et al., 2008).   A similar approach was used to identify P. trichocarpa hexokinase (HXK) orthologs, where the six known HXK and HXK-like (HXKL) A. thaliana genes - AT4G29130 (AtHXK1), AT2G19860 (AtHXK2), AT1G47840 (AtHXK3), AT1G50460 (AtHXKL1), AT3G20040 (AtHXKL2) and AT4G37840 (AtHXKL3) - were BLAST-ed in Phytozome V10 (Goodstein et al., 2012). Six   12 corresponding P. trichocarpa HXK and HXKL genes were identified: Potri.018G088300 (PtHXK1), Potri.001G190400 (PtHXK2), Potri.005G238600 (PtHXK3), Potri.009G050000 (PtHXKL1), Potri.001G254800 (PtHXKL2), and Potri.007G009300 (PtHXKL3). Based on an assessment of poplar transcriptome sequencing data (Hefer et al., 2015) it appears that Potri.001G190400 (PtHXK2) and Potri.005G238600 (PtHXK3) were more highly expressed in developing xylem versus leaf tissue and were therefore chosen for further investigation (Figure 16). The A. thaliana ortholog AT4G29130 (AtHXK1) was also found to be the most highly expressed gene in xylem followed by AT1G50460 (AtHXKL1) and AT2G19860 (AtHXK2). However, AT4G29130 (AtHXK1), AT2G19860 (AtHXK2), and AT1G47840 (AtHXK3) were selected for investigation since AT1G50460 (AtHXKL1) is believed to be a catabolically inactive protein (Granot et al., 2013).  2.2 Phylogenetic analysis In order to determine the evolutionary relationship between the P. trichocarpa SPS genes, their A. thaliana homologs and annotated SPS genes from other relevant organisms, a protein-based un-rooted Neighbor-Joining (NJ) phylogenetic tree was constructed. Annotated A. thaliana protein reference sequences were extracted from the National Center for Biotechnology Information (NCBI) RefSeq Database (Pruitt et al., 2014) and BLAST-ed in Phytozome V10 (Goodstein et al., 2012) to identify and compare the SPS sequences of the organisms of interest. The sequence information of other organisms of interest unavailable on Phytozome were extracted from Uniprot (UniProt Consortium, 2008) using the same search parameters. Sequences were aligned in MEGA6 (Tamura et al., 2011) using MUSCLE multiple sequence alignment, excess sequence trimmed and duplicates eliminated. A NJ test for phylogeny was   13 executed with the Jones-Taylor-Thornton model, and a bootstrap of 2000 replicates; gaps missing were considered complete deletions. The same approach was employed to construct a NJ tree of Hexokinase protein sequences.  2.3 Arabidopsis TDNA mutants Two Transfer DNA (TDNA) A. thaliana mutant seed lines were obtained from the ABRC stock catalogue through TAIR (Lamesch et al., 2012) for AtSPS1: SAIL_764_B04, from here on referred to as atsps1-1, and SALK_148643.41.05.x, from here on referred to as atsps1-2. atsps1-1 contains an insertion in the first exon, while atsps1-2 possesses an insertion in the eleventh exon of AtSPS1 (Figure 5, A). Similarly, SALK_064922.49.80.x a TDNA insertion in AtSPS2, from here on referred to as atsps2, was obtained and contains an insertion in the fifth exon of AtSPS2. A. thaliana mutant seed lines for AtHXK1; SALK_034233, termed athxk1, AtHXK2; WiscDsLox289_292O3, termed athxk2 and AtHXK3; SALK_030722, termed athxk3 contain TDNA insertions in the 3! untranslated region (UTR), ninth exon and second exon, respectively.   To germinate, seeds were sterilized with 70% EtOH for 5 minutes, followed by 10% sodium hypochlorite for 15 minutes and then washed three times with sterile water. Seeds were suspended in 0.1% w/v agar and immediately plated on ½ strength MS medium (Murashige & Skoog, 1962) supplemented with 50 mg/L kanamycin sulphate (Life Technologies Inc., Burlington, ON) as a selection for SALK lines (pROK vector) or 25 mg/L glufosinate ammonium (Sigma-Aldrich Co. LLC., Oakville, ON) for Wisconsin lines (pDs-Lox vector). Seeds were vernalized for 2 days in the dark at 4°C, followed by incubation for 7 days in the light (16h light/8h dark) at 21°C. Approximately ten seedlings from each line which germinated   14 on glufosinate ammonium (25 mg/L) or appeared green and healthy on kanamycin sulphate (50 mg/L) were selected for further analysis.   2.4 DNA screening In order to generate A. thaliana lines homozygous for the respective TDNA insertions, genomic DNA (gDNA) was extracted from plants that were growing in the growth chamber for 2 weeks.  One leaf was ground with 400 µl of DNA extraction buffer (200 mM Tris HCl, 250 mM NaCl, 25 mM NaEDTA, 0.5% SDS) and centrifuged at 13000 RPM for 3 minutes. The supernatant was transferred to a new tube, mixed with 300 µl of isopropanol and incubated at room temperature for 10 minutes. The samples were then centrifuged at 13000 RPM for 5 minutes and the supernatant discarded. To the pellet, 500 µl 70% ethanol was added and samples vortexed. A final spin for one minute was used to pellet the samples again, ethanol was discarded and tubes allowed to air dry for 30 minutes before resuspending DNA in double distilled water. A similar protocol was used to extract DNA from tissue culture-growing hybrid poplar hpRNAi lines. PCR was used to screen individuals for presence of a TDNA insertion in A. thaliana. Gene specific primers were designed using the TDNA primer design website hosted by the SALK Institute Genomic Analysis Lab (SIGNAL). Left border TDNA-specific primer for the pROK vector was self-designed using Primer Designer Software (no longer available) and the left border proximal primer LB1 for pDs-Lox vector was obtained from TAIR (Table 2-1). PCR was also used to screen hybrid poplar for the genomic insertion of the hpRNAi construct using primers amplifying the 35S promoter (Table 2-1). All PCR screening reactions were performed using New England BioLabs Taq DNA Polymerase with ThermoPol Buffer (New England BioLabs Ltd., Whitby, ON) using the manufactures instructions for routine PCR except that the number   15 of cycles was decreased to 25 and reaction volumes decreased to 20 µl.   2.5 RNAi down-regulation construct development In order to clone the full-length coding sequences (CDS) of PtSPS1, PtSPS2, PtSPS3, PtHXK2 and PtHXK3, RNA was extracted using the CTAB method from the developing cambium tissue of P. trichocarpa. Residual gDNA was removed using Abion DNA-free DNA Removal Kit (Life Technologies Inc., Burlington, ON) and first strand cDNA synthesis was preformed using iScript cDNA Synthesis Kit (Bio-Rad Laboratories Ltd., Mississauga, ON) with Oligo(dT) primers. CDS Forward and Reverse primers, listed in Table 2-1, were used for PCR on the cDNA template, and the resulting fragments run on a gel and purified using an in-house gel purification kit. The fragments were then cloned into pCR-Blunt II-TOPO vector using Invitrogen Zero Blunt TOPO PCR Cloning Kit (Life Technologies Inc., Burlington, ON).  Positive colonies were then selected on 50 mg/l kanamycin sulphate and vectors sequenced to confirm identity.   2.5.1 RNAi constructs targeting a single gene For RNAi constructs targeting a single gene, e.g. PtSPS1/3i and PtSPS2i, an approximately 500bp region was chosen that is specific to the target of interest. In addition, PtSPS1 and PtSPS3 share high sequence homology and since it was desirable to down-regulate both genes simultaneously, a region highly similar to both sequences was chosen that contained many regions of 20 bp identical sequence. Other parameters for the efficient design of RNAi constructs were adopted as described in Helliwell & Waterhouse (2003). Target regions were amplified from the CDS using primers listed in Table 2-1, which contained the attB1 on the forward, and attB2 on the reverse primer to facilitate cloning into the pDONR221 vector following a BP   16 reaction using Gateway BP Clonase II Enzyme Mix (Life Technologies Inc., Burlington, ON). pDONR221 constructs were transformed into chemically competent E. coli and positive colonies sequenced to confirm identity (NAPS Unit, UBC). To create a hairpin RNAi construct, Gateway LR Clonase II Enzyme Mix Kit was (Life Technologies Inc., Burlington, ON) used to directionally clone the fragment of interest into the pHELLSGATE12 expression vector (Helliwell & Waterhouse, 2003) driven by the 35S promoter (Figure 1, A). The resulting constructs were sequenced to confirm identity and correct orientation (forward, intron, reverse) of the gene fragment.   Figure 1: Construction of hpRNAi using the pHELLSGATE12 Gateway cloning system. A) Cloning of a fragment targeting a single gene. B) Cloning of fragments targeting two genes.   17 2.5.2 RNAi constructs targeting multiple genes In order to target multiple genes with a single hpRNAi construct, an approximately 250 bp region of highly conserved sequence from each gene of interest was chosen (Table 2-2). Again, in the case of PtSPS1 and PtSPS3, one highly conserved region of 250 bp was selected to represent both genes (Ex:PtSPS1/3::PtSPS2i) and another 250 bp region was chosen towards the 5! end of the CDS to target PtSPS2. A similar approach was take when constructing PtHXK2::PtHXK3i. In order to create a single fragment, a two-step ligation PCR was used where the first reaction amplifies the single gene target from the pCR-Blunt II-TOPO vector, using primer pairs attB1_PtSPS2_F and PtSPS2_R; PtSPS3_F and attB2_PtSPS3_R. In this reaction, the primers create sticky attB ends and sticky ends that match the other fragment. In the second reaction, half of the products of the first reaction are mixed together along with only primers attB1_PtSPS2_F and attB2_PtSPS3_R. In this reaction, the two fragment-specific sticky ends bind together and one, two-gene fragment was created with attB1 and attB2 sticky ends (Figure 1, B). The reactions following the two-step ligation PCR were identical to those described for the design of a single gene hpRNAi construct  18 Table 1: List of primers and their uses. Primer Name Sequence (5’ to 3’)  Use PtSPS1_CDS_F PtSPS1_CDS_R CTTGATGTAGGTCCTGGCTTAG GCCTCTATCCGACGCATTAT Cloning Cloning PtSPS2_CDS_F PtSPS2_CDS_R GATGAGGAGTACGCAAGAGA CTCCATCGTGTGGAGTGATA Cloning Cloning PtSPS3_CDS_F PtSPS3_CDS_R PtHXK2_CDS_F GTCCTGGCCTAGATGACAA GAAGCGCATGATCTCTATCC TTGCCGCAACCACTTCAC Cloning Cloning Cloning PtHXK2_CDS_R PtHXK3_CDS_F GCTAGGAGAGCAGCTCCAA AGAAGGTGGTGGTAGCTGTT Cloning Cloning PtHXK3_CDS_R CAAGGCCTGAGCCATCATT Cloning attB1_PtSPS2_F GGGGACAAGTTTGTACAAAAAAGCAGGCTAACGCCACCTTGAACGTGAA Cloning attB2_PtSPS2_R attB1_PtSPS3_F attB2_PtSPS3_R GGGGACCACTTTGTACAAGAAAGCTGGGTGCAACAGGCCAGACTGGATT GGGGACAAGTTTGTACAAAAAAGCAGGCTGCAGTGGCTGATATGTCTGA GGGGACCACTTTGTACAAGAAAGCTGGGTGATGTGAGGCCACAGAAGTT Cloning Cloning Cloning   19 attB1_PtSPS2_F PtSPS2_R PtSPS3_F attB2_PtSPS3_R attB1_PtHXK2_F-1 PtHXK2_R-1 PtHXK3_F-1 attB2_PtHXK3_R-1 attB1_PtHXK2_F-2 PtHXK2_R-2 PtHXK3_F-2 attB2_PtHXK3_R-2 M13_F M13_R 35S_F 35S_R GGGGACAAGTTTGTACAAAAAAGCAGGCTAACGCCACCTTGAACG ACGACCAAGCTCCATGTT TCCTGGCCTAGATGACA GGGGACCACTTTGTACAAGAAAGCTGGGTTCTTCTGCCACGTGTT GGGGACAAGTTTGTACAAAAAAGCAGGCTTGGCGGATTCCATG AGCGTGCATCTCAACAACCATGGCATCATCAGATGTACCAAACATA CCTCAAGGGCTTATGTTTGGTACATCTGATGATGCCATGGTTGTT GGGGACCACTTTGTACAAGAAAGCTGGGTTGCATCTGAAGTCCC GGGGACAAGTTTGTACAAAAAAGCAGGCTGCTTGTCTGAATGAAG CCACCACATCTTGGCCAACTGCTACCATCTGAAAACGC GAATGGGGAGCGTTTTCAGATGGTAGCAGTTGGCCAAGA GGGGACCACTTTGTACAAGAAAGCTGGGTACTCCATGTTTATAACCA GTAAAACGACGGCCAG CAGGAAACAGCTATGAC AATGCAGCTGGCACGACAGGTTT TGTGTACGCGTCAGCTGCTGCTCT Cloning Cloning Cloning Cloning Cloning Cloning Cloning Cloning Cloning Cloning Cloning Cloning Sequencing Sequencing Screening Screening   20 pROK2_LB p745-Ws atsps1-1_LP atsps1-1_RP atsps1-2_LP atsps1-2_RP atsps2_LP atsps2_RP athxk1_LP athxk1_RP athxk2_LP athxk2_RP athxk3_LP athxk3_RP AtEF1α_RT_F AtEF1α_RT_R GTGGACCGCTTGCTGCAACTCTC AACGTCCGCAATGTGTTATTAAGTTGTC AGCCTCTGTGCTTCTTTTTCC TGACAAAAGATCATTAGTGGTGG TTCTAATAATGTTGCGCCTGG CACCGGGTATGTGTTTTTACG TGCAAGACTTACAAGGTTCGC CCAGCTACTCTGAACCGTCTG GATTTCTACTGGTGCTGCTGG CTCTGCTGCTGGAATCTATGG ATGGATGAGGTGGACAAAACC ATTATGTTTGCGGTATGCAGG TTCTTCTCGTAAAACCCAACAAG AACTCGCATATCGAGTCCATG AGTTCTCGATTGCCACACCTCT ACCATACCAGCGTCACCATTCTTC Screening Screening Screening Screening Screening Screening Screening Screening Screening Screening Screening Screening Screening Screening Semi-q PCR Semi-q PCR   21 AtUBQ5_RT_F AtUBQ5_RT_R AtSPS1_CDS_F AtSPS1_CDS_R AtSPS2_CDS_F AtSPS2_CDS_R AtHXK1_CDS_F AtHXK1_CDS_R AtHXK2_CDS_F AtHXK2_CDS_R AtHXK3_CDS_F AtHXK3_CDS_R PtEF1β_RT_F PtEF1β_RT_R PtSPS1_RT_F PtSPS1_RT_R CAGCTCCACAGGTTGCGTTA CAAGCCGAAGAAGATCAAGCACAAG ATGGCCGGGAACGATTGGGTAAAC TCAGTCCTTGAGAAGCTCTAATTTCTTC ATGGTGGGAAACGACTGGGTGAATAG TCAAGGCTTGAGGAGACTGATCCC ATGGGTAAAGTAGCTGTTGGAGCG TTAAGAGTCTTCAAGGTAGAGAGAGTGA ATGGGTAAAGTGGCAGTTGCAACG TTAACTTGTTTCAGAGTCATCTTCAAGTTC ATGTCACTCATGTTTTCTTCCCCTGTC CTAGTAAATGGAGTTAGTGGCCGCC CCTGCTTCAAGCTTCCCAGG CCTCTGCTGCCTTCTTGTCC TCGCTGGGCATCTTCAGTTTCTG GCTCCTTAACAGGTGGAACCAATC Semi-q PCR Semi-q PCR Semi-q PCR Semi-q PCR Semi-q PCR Semi-q PCR Semi-q PCR Semi-q PCR Semi-q PCR Semi-q PCR Semi-q PCR Semi-q PCR q-PCR q-PCR q-PCR q-PCR   22 PtSPS2_RT_F PtSPS2_RT_R PtSPS3_RT_F PtSPS3_RT_R PtHXK2_RT_F PtHXK2_RT_R PtHXK3_RT_F PtHXK3_RT_R TGGACATTCATCGGGTGCTT CCCACAGCTGCTTATCGGAA CTGTTGAAAAGGAACGGGTGG TGGCCAAGGACGTTGACAATA CCCTTCATACTGAGGACGCC TGTTGCAAAGCTCAACCACG GCGAGATGTCGTTGCTTGTC CATTAACCAGGGCAGACACAC q-PCR q-PCR q-PCR q-PCR q-PCR q-PCR q-PCR q-PCR      23 Table 2: hpRNAi vectors and the coding sequence base pair position of the genes targeted. hpRNAi vector  Gene(s) targeted Base pair position PtSPS1/3i  PtSPS2i PtSPS1 PtSPS3 PtSPS2 _ 382-942 305-926 PtSPS1/3i::PtSPS2i  PtSPS1 PtSPS3 PtSPS2 _ 53-273 271-491 PtHXK2::PtHXK3i-1  PtHXK2 PtHXK3 176-425 180-429 PtHXK2::PtHXK3i-2 PtHXK2 PtHXK3 628-876 607-876  2.6 Agrobacterium transformation In order to insert RNAi vector sequences into the P. trichocarpa genome, Agrobacterium tumefaciens strain EHA105, was transformed with pHELLLSGATE12 hpRNAi expression vectors. Bacteria were transformed using the freeze-thaw method, where 1 µl of purified vector DNA was incubated with a 50-µl aliquot of EHA105 cells and allowed to sit on ice for 30 minutes. Cells were heat-shocked at 37 °C for 5 minutes, placed back on ice and 1 ml of liquid YEP media added to each tube. Tubes were then placed in a shaking incubator for 2 hours at 220 RPM at 28 °C to recover. To plate, the cells where centrifuged at 13000 RPM on a bench-top centrifuge for 1 minute and the supernatant discarded. Cells were then gently resuspended in 100   24 µl of YEP media and spread on YEP-agar plates containing 50 mg/l rifampicin and 100 mg/l spectinomycin. Selection plates with potential transformants were incubated at 27 °C for 2 days and colonies screened using colony-PCR with Forward and Reverse primers amplifying the 35S promoter (Table 2-1). Positive colonies were either directly grown in large-batch liquid media for plant co-inoculation or stored as glycerol stocks at -80 °C.   2.7 Hybrid poplar transformation To generate transgenic hpRNAi expressing plants, hybrid poplar (P. alba x P. grandidentata; P39), plantlets were propagated in sterile woody plant media (WPM)-agar until 4 weeks old. Three hundred and seventy five leaf discs were cut from P39 leaves with a size-4 cork borer for each construct (125 discs served as various antibiotic controls). Leaf discs were incubated overnight at room temperature in liquid WPM. Concurrently, construct-transformed A. tumefaciens colonies were grown in 40 ml liquid WPM culture with 100 µM acetosyringone at 28 °C and 200 RPM. The following day, the O.D. of the A. tumefaciens cultures was measured and adjusted to 0.1 O.D. with sterile WPM. For each WPM-agar petri plate, 25 leaf discs were then inoculated in 10 ml of liquid A. tumefaciens culture and allowed to co-incubate for 30 minutes at 28 °C and 200 RPM. Following which, the culture was discarded and all leaf discs were dried of excess liquid and placed on WPM-agar petri plates supplemented with 0.1 mM TDZ 0.1 mM NAA 0.1 mM BA. Plates were sealed and placed in the dark at 21 °C for two days. On the third day, A. tumefaciens was killed by moving discs to 0.1 mM TDZ 0.1 mM NAA 0.1 mM BA WPM plates supplemented with 500 mg/L carbenicillin, 250 mg/L cefotoxamine and 25 mg/L kanamycin. After five weeks, leaf discs possessing calli were transferred to 0.01 mM BA WPM with antibiotics. At nine weeks any formed shoots were transferred to rooting media   25 containing 0.01 mM NAA WPM and antibiotics. At thirteen weeks, green and elongated shoots that had rooted were transferred off antibiotic media and grown in tissue culture for further analysis.   2.8 Plant growth and phenotyping 2.8.1 A. thaliana TDNA insertional lines Germinated A. thaliana seedlings were planted in autoclaved potting soil (Westcreek Farms Ltd., Fort Langely, BC), watered and sealed in an opaque autoclave bag for 1 week to prevent desiccation. Following potting, the pots were placed in a growth chamber with long-day cycles (16h light/ 8h dark), 99% relative humidity (RH) and 21°C. After one week the plants were watered with tap water every two days, or as needed. Rosette diameter was measured with a ruler at 21 DAG (days after germination). Where applicable, whole rosettes and stems (main and side without flowers) were collected in aluminum foil and immediately frozen in liquid nitrogen.  Watering ceased at 42 DAG, after which stem height was measured and the plants were removed from the growth chamber to desiccate in a cool, dark place. Seeds were collected and, where desirable, the bottom 10 cm of the main stem was collected for cell wall chemical analysis.  2.8.2 Hybrid poplar hpRNAi lines  Positively transformed hybrid poplar clones were bulked up and grown in WPM-agar tissue culture for 5 weeks of growth or until suitable root formation was present. Ten clones from each positive line were planted in 2 L pots of perennial mix (Westcreek Farms Ltd., Fort Langley, BC), shoots were sprayed with tap water and covered with clear plastic cups to prevent desiccation for the first two weeks of growth in the greenhouse. Pots were watered as needed on   26 the greenhouse watering table with nutrient solution having the following molecular composition: pH 7.6, E.C. 1.6, NO3 4.9 mM/l, Cl 0.3 mM/l, S <0.2 mM/l, HCO3 3.2 mM/l, P 2.76 mM/l, NH4 8.0 mM/l, K 4.1 mM/l, Na 0.3 mM/l, Ca 0.3 mM/l, Mg <0.2 mM/l, Si <0.10 mM/l, Fe 17.1 µM/l, Mn 7.7 µM/l, Zn 8.2 µM/l, B 33 µM/l, Cu 8.4 µM/l, Mo <0.2 µM/l. Plant pots were randomly rotated once during the growth period. Tree height was measured every month of growth with a meter stick and plants were photographed at approximately two months of growth.   The trees were harvested for analysis after four months of growth beginning at 9:30 am for approximately two hours. Final height was measured and stem diameter taken at the cut site, which was 10 cm from the soil. The largest unfurled leaf was designated PI=0 and the fifth leaf down, PI=5, collected for analysis as a source leaf and immediately frozen in liquid nitrogen. A 60 cm piece of stem was measured from the cut site, bark stripped and developing xylem (from hereon referred to a cambium) scraped and immediately frozen in liquid nitrogen. Scraped stems were allowed to dry in a cool, dark place for further analysis.   2.9 RNA isolation and cDNA production To isolate RNA from flash frozen A. thaliana, leaves and stems were ground in a liquid nitrogen pre-cooled pestle and mortar thoroughly cleaned with 70% ethanol and treated with Sigma RNaseZAP (Sigma-Aldrich Co. LLC., Oakville, ON). Fifty microliters of tissue volume was transferred to a liquid nitrogen pre-cooled 2 ml safe-lock tube and RNA extracted with TRIzol Reagent (Life Technologies Inc., Burlington, ON) using the manufacturer’s directions. RNA was dissolved in 20 µl of nuclease-free water, tested for purity and concentration on a Themo   27 Scientific NanoDrop Lite Spectrophotometer (Thermo Fisher Scientific Inc., Ottawa, ON) and used immediately for further reactions or stored at -80 °C. A modified approach was used to isolate RNA from developing xylem and source leaf of hybrid poplar (P. alba x P. grandidentata) where tissues were ground using a SPEX 2010 Geno/Grinder (SPEX Sample Prep, Metuchen, NJ). Tissue was placed in vials with two steel ball bearings, pre-cool in liquid nitrogen and then homogenized for 1 minute at 15000 RPM. This process was repeated three times for developing cambium and twice for leaf tissue with sufficient cooling in liquid nitrogen between grinding steps to assure tissue did not thaw. Five hundred microliters of ground tissue was used for RNA extraction and the remaining tissue freeze dried in a Labconco FreeZone 4.5 Benchtop Freezedryer (Labconco Corp., Kansas City, MO) overnight at -50 °C and 0.032 Torr for non-structural sugar and starch analysis. The extraction method used was a modified TRIzol procedure, where a step to pellet polysaccharides was added: following homogenization of the tissue and extraction of the aqueous phase from TRIzol and chloroform to a new tube, 1/30th volume of NaOAc and 1/30th volume of 100% ethanol. Samples were chilled on ice for 10 minutes and centrifuged at 13000 RPM, after which the manufaturer’s protocol was followed and RNA resuspended in 20 µl of nuclease-free water. First strand cDNA was synthesized using iScript cDNA Synthesis Kit (Bio-Rad Laboratories Ltd., Mississauga, ON) with Oligo(dT) primer for both A. thaliana and hybrid poplar lines.  2.10 Semi-quantitative and quantitative RT-PCR In order to determine whether transcription of genes was occurring in A. thaliana homozygous TDNA insertional lines, semi-quantitative reverse transcription (semi-q RT) PCR was used. The primers used for these reactions amplified the full CDS of the genes and are listed in Table 2-1.   28 Control primers testing the integrity of the template amplified AtEF1α for samples of lines atsps1-1, atsps1-2 and atsps2 and AtUBQ5 for athxk1, athxk2 and athxk3. Semi-q RT-PCR reactions were carried out using New England BioLabs Taq DNA Polymerase with ThermoPol Buffer (New England BioLabs Ltd., Whitby, ON). The manufacturer’s instructions for conventional PCR were followed except that the reaction volume was decreased to 20 µl and the first strand cDNA template was undiluted.   To assess the amount of transcriptional down-regulation in hybrid poplar hpRNAi construct lines, quantitative RT-PCR (qRT-PCR) was used. To this end, a 5-point serial dilution series to test primer efficiency was created by mixing 1 µl of all cDNA templates and running each primer pair in a technical triplicate with a respective cDNA dilution. For all hybrid poplar experiments PtEF1β was used as a reference gene (Table 2-1). Finally, qRT-PCR was performed using Bio-Rad CFX96 Touch Real-Time PCR Detection System using Bio-Rad SsoFast EvaGreen Supermix (Bio-Rad Laboratories Ltd., Mississauga, ON) and annealing/extension temperature of 60 °C. For each hybrid poplar line, three biological samples were obtained from developing xylem and source leaf, and were run in technical triplicates. qRT-PCR analysis was performed using Bio-Rad CFX Manager Software v3.1 by importing plate files into a gene study file.  2.11 Non-structural sugars and starch determination Non-structural sugars and starch were quantified using the same protocol for A. thaliana TDNA insertional lines as well as hybrid poplar hpRNAi lines, however, due to the small amounts of tissue available from A. thaliana, the protocol was scaled down to 1/5th the reaction volume. For the un-scaled reaction, 50 mg of freeze-dried tissue was weighed into a 15 ml polypropylene   29 centrifuge tube and 50 µl of galactitol stock (10mg/ml) was added as an internal standard to each tube. Four milliliters of 12 methanol:5 chloroform:3 water (M:C:W) was then added, samples vortexed and allowed to extract overnight at 4 °C. On the following day, tubes were centrifuged at 4000 RPM, supernatant moved to a new 50 ml centrifuge tube and residual sugars re-extracted twice from the pellet in the same manner with 4ml M:C:W, finally achieving 12 ml of supernatant. The pellet was dried in 55 °C oven overnight and used for starch analysis. The aqueous phase was partitioned from the M:C:W by vortexing the samples with 4 ml of water and centrifuging at 4000 RPM for 4 minutes. The top phase was collected and 2 ml of it was vacuum dried overnight. The residue was resuspended in 1 ml of distilled water and filtered through a 0.45 µm filter into an HPLC vial.  For starch determination, the residual pellet was weighed in a 10 ml glass culture tube and 5 ml of 4% H2SO4 added. Samples were gently vortexed and autoclaved for 3.5 minutes at 121 °C. Cooled samples were centrifuged at 500 RPM for 5 minutes and the supernatant collected. HPLC vials were prepared by the addition of 950 µl of sample and 50 µl of fucose stock (10mg/ml) as an internal standard, by weight and passed through a 0.45 µm filter.   For both non-structural soluble sugars and starch, the area under the curve of each chromatograph was manually determined. The slope and intercept were found using the known concentrations of the sucrose/glucose/fructose/galactitol and glucose/fucose standards and used to integrate the concentrations of the samples in MS Excel.     30 2.12 Cell wall chemical analysis A Klason micromethod was used to determine the amount of structural carbohydrate in A. thaliana TDNA insertional lines. To this end, the bottom 10 cm of A. thaliana stems from 10 biological replicates of each line were pooled together and ground using a SPEX 2010 Geno/Grinder (SPEX Sample Prep, Metuchen, NJ) in a vial with two steel ball bearings. Before grinding, vials with sample were pre-cooled in liquid nitrogen then ground three times at 15000 RPM in 1-minute intervals with liquid nitrogen cooling in between. Ground tissue was acetone-extracted with a Soxhlet apparatus overnight in filter-paper pockets. Acetone was allowed to evaporate and tissue was re-dried at 100 °C overnight. Ten milligrams of tissue were weighed into 2 ml autoclave screw-cap tubes and immediately vortexed following the addition of 100 µl of 72% H2SO4 to each sample. Samples were incubated at 30 °C for 1 hour with 500 RPM shaking. Following, 2 ml of double distilled water was added, tubes thoroughly vortexed to resuspend the sample and autoclaved at 140 °C for 75 minutes. After allowing the samples to cool completely, the supernatant was removed by centrifuging the tubes at 15000 RPM for 10 minutes. One thousand eight hundred microliters of sample was removed and filtered through a 0.45 µm filter to HPLC vials.   2.13 High-performance liquid chromatography All non-structural, structural and starch samples were detected using a high-performance liquid chromatography (HPLC) system, Dionex Dx-600 (Thermo Fisher Scientific Inc., Bannockburn, IL) via anion exchange chromatography. The column used for non-structural carbohydrate separation was Agilent HiPlex Ca2+ (Agilent Technologies Inc., Seattle, WA) where the column compartment was kept at 70 °C.  The eluent was HPLC grade water. For the starch and structural   31 sugars a Dionex CarboPac PA1 was employed, and the carbohydrates were eluted using water at a flow rate of 1 mL/min. The machine was equipped with a Dionex ASI-100 auto-injector and a pulsed amperometric detector with a gold electrode where detection was achieved by the post-column addition of 100 mM NaOH at a flow rate of 1 ml/min.   2.14 Statistical analysis To determine whether a value was different from the wild-type plants, a Student’s t-test function in MS Excel was employed for the following measurements: plant height, rosette diameter, stem diameter, non-structural sugars, structural sugars, starch and X-ray crystallinity. Variances were assumed equal; tests were performed with two tails and α = 0.05 for all tests. The familywise error rate was corrected for using Bonferroni correction where each hypothesis was tested at a critical value α/m, where m is the total number of hypothesis. Dispersion of the data was measured using standard deviation of the sample.      32 Chapter 3: Results  3.1 Sucrose phosphate synthase  3.1.1 Phylogenetic analysis of the A. thaliana and P. trichocarpa protein families  In order to investigate the number of sucrose phosphate synthase (SPS) proteins in A. thaliana and P. trichocarpa, and the nature of any orthologous relationships between the two species, a 2000-bootstrap phylogenetic Neighbor-Joining tree was constructed (Figure 2). The analysis clearly shows that A. thaliana contains four and P. trichocarpa six SPS protein coding sequences. PtSPS1 and PtSPS3 are orthologous to AtSPS1 and fall within group 2d, as based on a designation established by Lutfiyya et al. (2007). PtSPS2 is orthologous to AtSPS2, and also falls within group 2d. Potri.013G095500 was found to be orthologous to AtSPS4 in group 4 and is from here on referred to as PtSPS4. Potri.001G317600 and Potri.017G057800 were orthologous to AtSPS3 and are from here on referred to as PtSPS5 and PtSPS6; they fall within group 3d. The presence of two additional, highly homologous SPS enzymes in poplar suggests that PtSPS1 & PtSPS2 and PtSPS5 & PtSPS6 may have arisen due to the recent chromosomal duplication (Tuskan et al., 2006).     33  Figure 2: Condensed phylogenetic tree of plant SPSs. The Neighbor-Joining tree was constructed using predicted and aligned protein sequences from SPS genes using 2000-boostraps. Black square: P. trichocarpa family, black triangle: A. thaliana family. Group designations based on Lutfiyya et al. (2007); d = dicots, m = monocots.   Potri.001G317600 | PtSPS Potri.017G057800 | PtSPS Gorai.011G155900 | GrSPS PGSC0003DMG400026428 | StSPSB Q3HLN2 | NtSPSB Solyc09g092130.2 | SlSPS Eucgr.H00041 | EgSPS Glyma.14G029100 | GmSPS Glyma.08G308600 | GmSPS Glyma.18G108100 | GmSPS AT1G04920 | AtSPS3d LOC Os01g69030 | OsSPS GRMZM2G140107 | ZmSPS GRMZM5G875238 | ZmSPSm Phpat.009G095500 | PpSPS Phpat.015G092800 | PpSPS3 GRMZM2G008507 | ZmSPS LOC Os11g12810 | OsSPS Eucgr.F02931 | EgSPS PGSC0003DMG402019060 | StSPSC Solyc11g045110.1 | SlSPS Q3HLN3 | NtSPSC AT4G10120 | AtSPS4 Potri.013G095500 | PtSPS Gorai.009G262100 | GrSPS Glyma.04G110200 | GmSPS Glyma.06G323700 | GmSPS4 GRMZM2G049076 | ZmSPS GRMZM2G471083 | ZmSPS LOC Os06g43630 | OsSPS GRMZM2G462613 | ZmSPS LOC Os02g09170 | OsSPS GRMZM2G055331 | ZmSPS LOC Os08g20660 | OsSPS2m PGSC0003DMG400029892 | StSPS Solyc08g042000.2 | SlSPS AT5G11110 | AtSPS2 P31928 | SoSPS Eucgr.C01715 |EgSPS Potri.018G025100 | PtSPS2 Gorai.004G292800 | GrSPS Gorai.010G116100 | GrSPS PGSC0003DMG400027936 | StSPS Solyc07g007790.2 | SlSPS Q9SNY7 | NtSPSA Glyma.13G161600 | GmSPS Glyma.17G109700 | GmSPS Eucgr.E03524 | EgSPS Eucgr.E03528 | EgSPS AT5G20280 | AtSPS1 Gorai.013G235900 | GrSPS Potri.006G064300 | PtSPS1 Potri.018G124700 | PtSPS32d1009 91009 01001001001001006 99 91001008 81001001009 98 99 81009 99 61001009 71001001009 91008 55 26 87 89 65 47 3  34 3.1.2 Endogenous expression of SPS genes in A. thaliana and P. trichocarpa in sink and source tissues A. thaliana It was found that A. thaliana’s genome contains four SPS coding sequences. To examine which are most highly expressed in leaf and stem, endogenous expression data of A. thaliana SPS homologs were examined in sink and source tissues (data not shown) using the Arabidopsis eFP browser (D. Winter et al., 2007) and Genevestigator (Hruz et al., 2008) (Figure 3). AtSPS1 was found to be the most highly expressed gene in all tissues, ranging from 12.5 to 15 relative units. AtSPS2 was the second most highly expressed gene in xylem and stem with approximately 12.5 and 11.25 units, respectively. In adult leaf and whole rosette tissue, AtSPS4 was second most highly expressed at 11.75 and 12 units respectively following AtSPS1. These results suggest that AtSPS1 and AtSPS2 are likely to be most important for SPS enzyme function in sink tissues such as A. thaliana xylem and stem and were therefore chosen for further investigation.     35  Figure 3: Endogenous expression of SPS genes; AtSPS1 (red dot), AtSPS2 (blue dot), AtSPS3 (green dot) and AtSPS4 (brown dot) in select A. thaliana tissues. Units represent relative expression; error bars represent SD. Created with Genevestigator (Hruz et al., 2008). P. trichocarpa  Endogenous expression of P. trichocarpa homologs was examined in source leaf and cambium using RNA sequencing data (Hefer et al., 2015) in order to determine if there is differential contribution of those homologs to SPS enzymatic function in source and sink tissues. PtSPS1 was found to be the most highly expressed gene in cambium at 5.69±1.08*105 fpkm, followed by PtSPS2, 1.61±1.35*105 fpkm, and PtSPS3, 1.64±1.31*105, which have roughly the same level of expression in that tissue (Figure 4). PtSPS3 is most highly expressed in source leaf at 3.27±0.61*105 fpkm followed by PtSPS1 that is expressed at 2.01±0.42*105 fpkm. Interestingly, PtSPS4 shows low expression in both leaf and cambium even though it is orthologous to AtSPS4, which was found to have the second highest expression in A. thaliana leaf. These results   36 suggested that PtSPS1, PtSPS2 and PtSPS3 could be majorly responsible for SPS enzymatic activity in the cambium and were consequently chosen for further study. In addition, there is differential expression of A. thaliana and P. trichocarpa orthologs in source and sink tissues.    Figure 4: Endogenous expression of SPS genes in P. trichocarpa cambium and source leaf. Created with data from Hefer et al. (2015), error bars represent SD.   3.1.3 atsps1-1, atsps1-2 and atsps2 A. thaliana TDNA insertional mutants Relative expression of AtSPS1 and AtSPS2 in TDNA insertional mutants TDNA insertional lines were chosen to study the effects of loss of SPS enzyme protein in A. thaliana. atsps1-1 contains a TDNA insertion in the first exon and atsps1-2 an insertion in the eleventh exon of AtSPS1. atsps2 is a mutant of AtSPS2 and harbors a TDNA insertion in the fifth exon (Figure 5, A). Semi-quantitative PCR was utilized to determine whether atsps1-1, atsps1-2    37 and atsps2 are mutants lacking expression of SPS transcript (Figure 5, B). By amplifying the full length coding sequence (CDS), it was seen that atsps1-1 and atsps1-2 lack complete AtSPS1 transcript. Lack of amplification of AtSPS2 transcript was observed in the atsps2 line, but not AtSPS1 transcript. These results suggested that atsps1-1, atsps1-2 are TDNA insertional mutants for AtSPS1 and atsps2 for AtSPS2. In addition, AtSPS1 is more highly expressed than AtSPS2 in the wild type (Figure 5, B lanes 1 and 2), corroborating AtSPS endogenous expression metadata.     Figure 5: atsps1-1, atsps1-2 and atsps2 are TDNA insertional mutants. A) Exon map of AtSPS1 and AtSPS2, black triangles represent position of TDNA insertions. B) Semi-q PCR of AtSPS1 and AtSPS2, AtEF1a is a reference control.                aatsps1-1 aatsps1-2                 atsps2 A) B)   38 Phenotypic analysis of plant growth To determine if loss of SPS enzyme protein affects the growth of A. thaliana during development, TDNA insertional lines were grown and phenotypically analyzed. At 21 days after germination (DAG), atsps1-1 had an average rosette diameter of 1.65±0.50 cm, significantly smaller than wild type, which averaged at 2.77±0.26 cm (Figure 6, A). atsps1-2 and atsps2 were not affected and grew similar to the wild-type plants (Figure 6, B). At senescence, the stem heights of all three TDNA mutant lines were not significantly different than the wild type, which measured 34.91±2.66 cm (Figure 6, C). In addition, atsps1-1 bolted 9 days earlier than the wild type. These results suggest that loss of AtSPS1 protein has an effect on plant development, and that atsps1-1 is a more severe mutation than atsps1-2, likely representing a complete knockout (KO). The same seed lines were grown twice with reproducibility of the phenotypic results.   A)   39   Figure 6: Rosette diameter and stem height of atsps1-1, atsps1-2 and atsps2. A) Rosette diameter 21 DAG. B) Rosettes pictured at 21 DAG, grey bar = 1cm. C) Stem height at 42 DAG. n = 6, * represents a significant B) C)   40 difference from the wild type at p < 0.05, * represents a significant difference from the wild type at a Bonferroni critical value of p < 0.02, error bars represent SD. Non-structural carbohydrates in leaf and stem Non-structural carbohydrates were determined in A. thaliana TDNA mutants to assess whether loss of SPS protein affects the amount and content of the soluble carbohydrates, and if this effect is contrasting between source and sink organs. Sucrose and glucose contents were found to be significantly increased in both the leaf and stem of all three TDNA mutant lines compared to the wild type plants, with atsps1-1 always having the most pronounced effect (Figure 7, A and B). While the wild type averaged 2.45±0.41 mg/g of sucrose in the leaf and 2.0±0.95 mg/g in the stem, atsps1-1 was found to have 11.38±3.60 mg/g and 8.36±2.37 mg/g respectively in those same tissues. atsps1-2 had 8.84±0.04 mg/g and 4.84±1.30 mg/g, while atsps2 had 6.52±1.56 mg/g and 3.94±1.03 mg/g of sucrose in whole stem and rosette leaf, respectively. As in the wild type, sucrose content in the mutants was higher in the leaf versus the stem.   Glucose content, which appeared to be almost equal in the leaf and the stem of the wild-type plants at 5.05±1.19 mg/g and 4.81±3.01 mg/g, was significantly higher in the stem of atsps1-2 (16.18±8.55 mg/g) and atsps2 (20.47±8.86 mg/g), and was strikingly higher in atsps1-1 (56.74±11.6) mg/g. The glucose content in leaf of atsps1-1, atsps1-2 and atsps3 was also significantly increased compared to the wild type (25.11±12.57 mg/g, 11.88±0.60 mg/g and 18.69±6.36 mg/g, accordingly).     41 Fructose content in wild type plants was 8.68±1.10 mg/g in leaf and 9.73±2.42 mg/g in stem. Fructose was higher in the leaf and stem of atsps1-1 (24.24±9.69 mg/g and 57.24±2.14 mg/g) and atsps2 (24.44±4.40 mg/g and 16.22±2.23 mg/g) compared to the wild type, but not atsps1-2 (Figure 7, C). atsps1-1 had the highest change in non-structural carbohydrate but the trend was similar to wild type, where a higher level of fructose accumulated in the stem. Conversely, atsps1-2 accumulated more fructose in the leaf. These results suggest that loss of AtSPS1 and AtSPS2 protein may affect the amount of non-structural carbohydrate accumulated in the stem and leaf of A. thaliana differentially. Furthermore, there is a difference in the location of accumulation of glucose and fructose that is dependent on the genotype analyzed.   A)   42   Figure 7: Non-structural carbohydrate contents in stem and leaf of atsps1-1, atsps1-2 and atsps2 harvested at midday. A) Sucrose content. B) Glucose content. C) Fructose content. Units in mg/g DW, n = 5, * represents a C) B)   43 significant difference from the wild type at p < 0.05, * represents a significant difference from the wild type at a Bonferroni critical value p < 0.02, error bars represent SD. Starch content in leaf and stem In order to detect whether loss of SPS protein translates to a change in transient carbon storage, percent starch was measured in the stem and leaf. Starch content (Figure 8) was found to be higher in the leaf of atsps1-1 (22.00±3.66%), atsps1-2 (27.77±2.21%), and atsps2 (19.17±3.99%) compared to the wild type plants (9.61±1.35%). Similarly, all mutants had a significant increase of starch in the stems compared to the wild type stem (2.51±0.22%). Based on these experiments, it can be concluded that loss of AtSPS1 or AtSPS2 affects the accumulation of starch in the leaf more strongly than in the stem.     44  Figure 8: Percent starch in the stem and leaf of atsps1-1, atsps1-2 and atsps2 harvested at midday. Percent dry weight (%), n = 5, * represents a significant difference from the wild type at p < 0.05, * represents a significant difference from the wild type at a Bonferroni critical value of p < 0.02, error bars represent SD. Structural carbohydrates and lignin of the stem Analysis of non-structural carbohydrates determined that there is an increase in the amount of sucrose, glucose and fructose in the leaf and stem. To determine if that increase resulted in a change in carbon allocation to the structural components of the stem, the amount of structural carbohydrates in atsps1-1, atsps1-2 and atsps2 stems were measured. No significant changes were observed in most structural sugars, per mass sample of atsps1-1, atsps1-2 and atsps2 main stems (Table 3). Arabinose, however, was found to be slightly higher in atsps1-1 (11.16±1.17 µg/mg) and atsps1-2 (10.39±0.35 µg/mg) compared to the wild type, which had an arabinose content of 9.20±0.90 µg/mg.     45  Table 3: Average quantity of sugar per mass of sample (ug/mg) in atsps1-1, atsps1-2 and atsps2 main stems. Bold values indicate a significant difference from the wild type at p < 0.05, underlined values represent a significant difference from the wild type at a Bonferroni critical value of p < 0.02, n = 5, SD in brackets.   Arabinose Rhamnose Galactose Glucose Xylose Mannose WT 9.20 (0.90) 8.03 (0.45) 16.52 (1.04) 350.84 (16.03) 123.51 (5.29) 21.21 (2.03) atsps1-1 11.16 (1.17) 7.71 (0.60) 16.53 (1.19) 351.38 (20.96) 125.95 (6.13) 20.81 (1.19) atsps1-2 10.39 (0.35) 7.93 (0.20) 16.55 (0.29) 360.26 (5.75) 118.12 (2.38) 19.26 (0.55) atsps2 9.69 (0.33) 7.80 (0.09) 15.93 (0.31) 347.92 (12.25) 120.75 (4.11) 19.14 (0.82)  In order to assess whether changes in the monomeric composition of structural carbohydrates caused an effect on the lignin composition of the primary stem, percent acid soluble and insoluble lignin were measured (Table 4). The soluble fraction of lignin was significantly lower in all three lines examined compared to the wild type based on a Student’s t-test. Particularly, atsps1-1 contained 2.76±0.13%, atsps1-2 was 2.79±0.02% and atsps2 had 2.87±0.02% compared to the wild type at 3.19±0.06%. The insoluble lignin fraction and total lignin were not found to be different in the mutant lines examined compared to the wild type. These results suggest that decrease of SPS enzyme protein potentially affects the composition of lignin content in A. thaliana so that more soluble lignin is released but the total amount of lignin is not affected.     46 Table 4: Average percent insoluble, soluble and total lignin in the main stem of atsps1-1, atsps1-2 and atsps2. Bold values indicate a significant difference from the wild type at p < 0.05, underlined values indicate a significant difference from the wild type at a Bonferroni critical value of p < 0.02, n = 4, SD in brackets.  Insoluble lignin Soluble lignin Total WT 17.42 (1.42) 3.19 (0.06) 20.59 (1.47) atsps1-1 18.08 (0.45) 2.76 (0.13) 20.84 (0.36) atsps1-2 17.30 (1.40) 2.79 (0.02) 20.09 (1.06) atsps2 17.37 (0.24) 2.87 (0.02) 20.18 (0.24)  3.1.4 RNAi hybrid poplar down-regulation Relative expression of PtSPS1 and PtSPS3  Hybrid poplar lines were transformed with vectors targeting the suppression of PtSPS1 and PtSPS3 separately, PtSPS1/3i, or PtSPS1/3i::PtSPS2i combined. While propagating in the greenhouse, these putative RNAi lines were tested for the extent of down-regulation of the respective genes using quantitative PCR of the leaf. Lines found to have a decreased expression in PtSPS1 and PtSPS3 were permitted to continue growing, while the rest were sacrificed. Following the end of the chosen growth period, the remaining lines were harvested and tissue collected for all subsequent analysis. PtSPS1/3i hp RNAi line 10 was found to have a decrease in expression of PtSPS1 by 0.8100 that of the control (1.00) in the developing xylem (Figure 9, B). PtSPS1/3i hp RNAi line 16 was down-regulated for PtSPS3 to 0.0030 compared to the control, at 0.0039, while PtSPS1 expression remained close to the control and PtSPS2 was increased to 0.1398, compared to the control at 0.0934. The hp RNAi vector PtSPS1/3::PtSPS2i lines 1, 2, 7 and 9 were all decreased for the expression of PtSPS1 showing 0.4866, 0.1610, 0.6186 and   47 0.7388 times that of the control. Only line 2 exhibited a decreased expression of PtSPS3 at 0.0017 compared to the control, 0.0039. Therefore, most lines, except line 16, had a decreased expression of PtSPS1 in the cambium compared to the untransformed control. No lines from all three constructs were found to have a decreased expression of PtSPS2 in the cambium and only line 16 of PtSPS1/3i and line 2 of PtSPS1/3::PtSPS2i construct had a decreased expression of PtSPS3. PtSPS1 being the most highly expressed SPS transcript in the developing xylem of P. trichocarpa (Figure 9, A), it can be assumed that some loss of SPS enzyme expression is present in the hybrid poplar lines.    48   A) B)   49  Figure 9: Quantitative PCR on A) untransformed hybrid poplar leaf and cambium, B) pro 35S-driven hairpin RNAi transformed hybrid poplar lines in cambium following four months of growth and C) pro 35S-driven hairpin RNAi transformed hybrid poplar lines in leaf following two months of growth in the greenhouse. Expression in B) and C) was normalized to PtEF1β, relative to the untransformed control (Ctrl), n = 3. Phenotypic analysis of plant growth To establish if a decrease in the amount of SPS transcript affects hybrid poplar growth, stem diameter and height were measured following four months in a greenhouse at harvest (Figure 10). PtSPS1/3i RNAi contruct line 16 had a smaller diameter (16.31±0.77 mm) and line 2 (16.39±1.64 mm) from the PtSPS1/3::PtSPS2i RNAi construct exhibited a decrease in stem diameter compared to the control (19.03±0.75 mm). Conversely, both line 10 (314.14±26.58 cm) and line 16 (329.50±14.97 cm) from the PtSPS1/3i RNAi construct had a significantly smaller stem height compared to the control (351.57±13.35 cm; Figure 10, B). Line 2 (328.28±24.34 cm) C)   50 from PtSPS1/3::PtSPS2i was also decreased in final stem height. The results tentatively suggest that down regulation of PtSPS1 decreases hybrid poplar stem height and diameter.   A)   51  Figure 10: Stem growth parameters of PtSPS1/3i and PtSPS1/3::PtSPS2i RNAi down regulated hybrid poplar following four months of growth in a greenhouse. A) Stem diameter (mm) measured 10 cm above soil. B) Stem height 10 cm above soil to tip. n = 7, * represents significant difference from the control at p < 0.05, * represents a significant difference from the control with a Bonferroni critical value of p < 0.008, error bars represent SD. Non-structural carbohydrates in cambium and source leaf SPS transcript expression is found in both in source and sink tissue. To determine whether change in the regulation of SPS transcript affects the non-structural carbohydrate content in those tissues, sucrose, glucose and fructose levels in source leaf and cambium were measured (Figure 11). Sucrose content of leaf was found to be significantly lower in lines 2 (60.09±0.12 mg/g) and 7 (55.19±5.19 mg/g) of the PtSPS1/3::PtSPS2i RNAi construct compared to the control (71.64±4.14 mg/g), but remained unchanged in lines of PtSPS1/3i (Figure 11, A). In cambium, B)   52 however, sucrose content was significantly increased in all four hybrid poplar lines expressing the PtSPS1/3::PtSPS2i RNAi construct, showing 24.88±2.76 mg/g, 27.17±4.67 mg/g, 28.88±2.76 mg/g and 34.31±4.05 mg/g in lines 1, 2, 7 and 9, respectively. The untransformed control line had a sucrose content of 19.49±1.70 mg/g in the cambium. Line 10, expressing the PtSPS1/3i construct, had an increased sucrose content, 31.68±0.17 mg/g, compared to the control (Figure 11, A).   Glucose content was found to be higher in the cambium of PtSPS1/3i line 10 (27.08±6.53 mg/g) and PtSPS1/3::PtSPS2i lines 7 (10.43±0.57 mg/g) and 9 (31.65±8.10 mg/g) compared to the control (5.94±1.26 mg/g; Figure 11, B). Glucose levels were not found to be significantly different in any of the lines in source leaf compared to the control, which had 21.10±6.74 mg/g of glucose.  Finally, fructose was found to be significantly increased in the cambium (40.85±5.10 mg/g) and source leaf (107.99±2.98 mg/g) of PtSPS1/3i line 10 and solely the cambium of PtSPS1/3::PtSPS2i lines 1 (23.14±3.47 mg/g) and 9 (55.30±1.48 mg/g, Figure 11, C). In comparison, fructose content of the control was 14.39±3.38 mg/g in the cambium and 92.76±5.22 mg/g in the source leaves. These results suggest that a decrease in PtSPS1 transcript changes the amount of non-structural carbohydrate available, particularly in the sink tissue cambium, but also in source leaves.    53   A) B)   54  Figure 11: Non-structural carbohydrates in cambium and source leaf of PtSPS1/3i and PtSPS1/3::PtSPS2i RNAi down-regulated hybrid following four months of growth in a greenhouse and harvested at midday. A) Sucrose content. B) Glucose content. C) Fructose content. Units in mg/g DW, n = 3, * represents significant difference from the control at p < 0.05, * represents a significant difference from the control at a Bonferroni critical value of p < 0.008, error bars represent SD. Starch content in cambium and source leaf Starch is a transient storage form of carbon. To establish if a change in SPS transcript regulation caused variation in this form of carbon storage between plant source and sink tissue, starch content was measured (Figure 12). An increase in starch was detected in the leaf of PtSPS1/3i line 16 (10.44±2.25 mg/g) that was twice the amount found in the control line (4.83±0.72 mg/g). PtSPS1/3i line 10 also had a one-third increase in leaf starch but the change was not significant. Conversely, starch was significantly decreased in the cambium of PtSPS1/3i line 10 RNAi down-C)   55 regulated hybrid poplar (0.37±0.03 mg/g) compared to the control (0.50±0.06 mg/g). Lastly, PtSPS1/3::PtSPS2i lines were not found to have a starch content significantly different from the control in either source leaf or cambium.    Figure 12: Percent starch content in the cambium and source leaf of PtSPS1/3i and PtSPS1/3::PtSPS2i RNAi down regulated hybrid following four months of growth in a greenhouse and harvested at midday. Units in percent DW, n = 3, * represents a significant difference from the control at p < 0.05, * represents a significant difference from the control at a Bonferroni critical value of p < 0.008, error bars represent SD.   3.2 Hexokinase  3.2.1 Phylogenetic analysis of A. thaliana and P. trichocarpa Hexokinase protein families Phylogenetic analysis was undertaken with two objectives: to determine the number of HXK genes in A. thaliana and second, to define any orthologous relationships to P. trichocarpa and    56 other annotated HXKs. It was found that A. thaliana contains protein-coding sequences for six HXKs: AtHXK one through three, and AtHXKL one through three (Figure 13). Black cottonwood also contains six HXK protein-coding sequences named PtHXK one through three and PtHXKL one through three. AtHXK1, AtHXK2 and PtHXK1, PtHXK2 are orthologous and fall within group 6, as previously determined by R. Karve et al. (2010). AtHXK3 and PtHXK3 are orthologous and fall within group 4 as well as AtHXKL3 and PtHXK3, which are orthologous within group 5. Finally, AtHXKL1, AtHXKL2 and PtHXKL1, PtHXKL2 are orthologous and fall within group 3b. Since P. trichocarpa has the same number of known HXK protein-coding sequences as A. thaliana, this implies that there was no gene expansion event in the HXK family of black cottonwood or, any sequences resulting from an expansion were subsequently lost.     57   PGSC0003DMG400000295 | StHXK5 Q6Q8A1 | NtHXK5 Solyc12g008510.1 | SlHXK Q6Q8A3 | NtHXK4a Q6Q8A2 | NtHXK4b Eucgr.C00559 | EgHXK Eucgr.C00983 | EgHXK Eucgr.C00569 | EgHXK Eucgr.C00975 | EgHXK Eucgr.C03728 | EgHXK Potri.018G088300 | PtHXK1 Medtr6g088795 | MtHXK Medtr8g102460 | MtHXK AT2G19860 | AtHXK2 AT4G29130 | AtHXK1 Medtr8g014530 | MtHXK Potri.001G190400 | PtHXK2 PGSC0003DMG400016521 | StHXK Solyc06g066440.2 | SlHXK Q6Q8A4 | NtHXK3 Q6Q8A6 | NtHXK1a Q9SEK2 | NtHXK1 Solyc03g121070.2 | SlHXK PGSC0003DMG400002525 | StHXK7 Q6BDB4 | NtHXK76 GRMZM2G171373 | ZmHXK LOC Os01g52450 | OsHXK9 LOC Os05g45590 | OsHXK2 GRMZM2G051806 | ZmHXK LOC Os05g09500 | OsHXK7 LOC Os07g26540 | OsHXK GRMZM2G104081 | ZmHXK LOC Os01g09460 | OsHXK8 LOC Os05g44760 | OsHXK5 GRMZM2G058745 | ZmHXK GRMZM2G432801 | ZmHXK GRMZM5G856653 | ZmHXK LOC Os01g53930 | OsHXK67 AT4G37840 | AtHXKL3 Potri.007G009300 | PtHXKL3 PGSC0003DMG400030624 | StHXK Solyc02g091830.2 | SlHXK5 AT1G47840 | AtHXK3 LOC Os07g09890 | OsHXK Medtr1g025140 | MtHXK Eucgr.F01647 | EgHXK Potri.005G238600 | PtHXK3 PGSC0003DMG400009861 | StHXK Solyc04g081400.2 | SlHXK4 Phpat.018G040700 | PpHXK Phpat.021G068300 | PpHXK Phpat.019G068600 | PpHXK Phpat.022G035800 | PpHXK Phpat.015G051300 | PpHXK Phpat.008G072000 | PpHXK Phpat.023G002800 | PpHXK Phpat.002G050300 | PpHXK Phpat.010G034000 | PpHXK Phpat.014G021300 | PpHXK Phpat.001G020100 | PpHXK1 GRMZM2G068913 | ZmHXK GRMZM2G467069 | ZmHXK LOC Os01g71320 | OsHXK GRMZM2G046686 | ZmHXK LOC Os05g31110 | OsHXKa Eucgr.B03711 | EgHXK Eucgr.J00734 | EgHXK Potri.001G254800 | PtHXKL2 Potri.009G050000 | PtHXKL1 AT1G50460 | AtHXKL1 AT3G20040 | AtHXKL2 Medtr5g009000 | MtHXK Q6Q8A0 | NtHXK6 PGSC0003DMG400013187 | StHXK6 Solyc11g065220.1 | SlHXKb39 99 96 49 99 79 99 99 39 99 99 95 99 99 95 39 99 99 99 75 26 85 49 29 09 99 99 98 79 37 05 39 99 79 77 09 49 29 37 95 29 99 39 99 99 89 49 59 79 99 99 96 56 66 76 86 4  58 Figure 13: Condensed phylogenetic tree of plant Hexokinase proteins. A Neighbor-Joining tree was constructed using predicted and aligned protein sequences from HXK genes using 2000-boostraps. Black square: P. trichocarpa family, black triangle: A. thaliana family. Group designations based on R. Karve et al.  (2010); a and b are subgroups within 3.   It has been suggested that HXKL proteins are not enzymatically functional and only serve a signaling role due to the presence of an indel sequence in the A. thaliana homologs. A multiple sequence alignment of P. trichocarpa and A. thaliana HXK protein sequences revealed that PtHXKL1 and PtHKXL2 contain the same insertion around amino acid 450 – 460 as AtHXKL1 and AtHXKL2, while PtHXKL3 contains the same deletion around amino acid 460-465 (Figure 14). Consequently, AtHXK one through three and PtHXK one through three were considered for further investigation, as they are likely to be enzymatically functional.      59  Figure 14: Multiple sequence alignment of AtHXK and PtHXK family of proteins using MUSCLE algorithm. PtHXKL1 and PtHXKL2 contain an insertion around amino acids 450-465 similar to AtHXKL1 and AtHXKL2. PtHXKL3 contains a deletion around amino acids 460-465 similar to AtHXKL3.   3.2.2 Endogenous expression A. thaliana Following the finding that A. thaliana has six HXK genes, it was desirable to assess if there is any difference in the spatial distribution of said transcripts. Using meta-data from the Arabidopsis eFP browser (D. Winter et al., 2007) (data not shown) and Genevestigator (Hruz et al., 2008) it was determined that AtHXK1 is the most ubiquitously expressed transcript in all tissues examined, ranging from 11.5 to 12.75 expression units (Figure 15). AtHXK2 was found to have the second highest transcription abundance in adult leaf (11.5 units) and whole rosette (11.6 units). It was also second highest in whole stem at 11.2 expression units. In xylem, AtHXKL1 is slightly higher in expression compared to AtHXK2, 10.5 versus 10.9 expression units. AtHXK3 is   60 lower in expression in adult leaf (7.9 units) and whole rosette (8.1 units) as well as xylem (7.9 units) and whole stem (7.7 units) compared to AtHXKL1 and AtHXKL2. However, AtHXK3 is considered to be enzymatically functional, while AtHXKL1 and AtHXKL2 are not. Therefore, AtHXK1, AtHXK2 and AtHXK3 were chosen for further investigation on account of being highly expressed in the tissues of interest (leaf and stem) or being the highest functionally enzymatic transcript in that tissue.     Figure 15: Endogenous expression of the HXK family of genes in select A. thaliana tissues. AtHXK1 (red dot), AtHXK2 (blue dot), AtHXK3 (green dot), AtHXKL1 (brown dot), AtHXKL2 (purple dot), AtHXKL3 (yellow dot). Units are relative, created with Genevestigator (Hruz et al., 2008). P. trichocarpa  The genome of black cottonwood encodes for six HXK genes. In order to determine which homologs are expressed either in source or sink tissue, endogenous expression data was   61 examined by RNAsequencing of source leaf and cambium (Figure 16). As in A. thaliana, PtHXK1 homolog was most highly expressed in source leaf (1.23±0.01*106 fpkm), with relatively high expression in cambium, (7.16±1.35*105 fpkm). Unlike A. thaliana, the PtHXK2 and PtHXK3 homologs were found to have higher expression in cambium than PtHXK1; 9.48±3.25*105 fpkm and 1.18±0.19*106 fpkm, respectively. PtHXK2 and PtHXK3 also have about half the expression as PtHXK1 in source leaf; 4.37±7.22*105 fpkm and 5.78±0.85*105 fpkm, respectively. Additionally, PtHXKL1 and PtHXKL2 have comparatively low expression in cambium and source leaf. These data suggest a distinct expression pattern of HXK homologs in P. trichocarpa compared to A. thaliana, where PtHXK2 and PtHXK3 are responsible for the bulk of HXK enzyme in the cambium and were, therefore, selected for further study.    Figure 16: Endogenous expression of HXK transcripts in cambium and source leaf of P. trichocarpa. Units in fpkm, data from Hefer et al. (2015), error bars represent SD.     62 3.2.3 athxk1, athxk2 and athxk3 TDNA insertional mutants Relative expression of AtHXK1, AtHXK2 and AtHXK3 TDNA insertional lines athxk1, athxk2 and athxk3 were chosen to study the effects of loss of HXK protein in A. thaliana. athxk1 is predicted to contain a TDNA insertion in the 3! UTR of AtHXK1, athxk2 an insertion in the ninth exon of AtHXK2, and athxk3 in the second exon of AtHXK3 (Figure 17, A). To determine the relative amount of endogenous transcription abundance of the TDNA insertional mutants for AtHXK1, AtHXK2 and AtHXK3, semi-quantitative PCR was employed (Figure 17, B). As seen in lanes 3 and 4, athxk1 was found to retain transcriptional expression of AtHXK1 with primers spanning the CDS, to the same intensity as wild type. athxk2 was found to express transcript for AtHXK1 and AtHXK3 (note faint bands of reference control), but not AtHXK2. Finally, athxk3 was found to express transcript for AtHXK1 and AtHXK2, but not AtHXK3. It is concluded that athxk1 is not a knockout for AtHXK1, but may possess a decrease in the amount of functional AtHXK1 protein produced. athxk2 and athxk3 do not accumulate transcript for their respective genes and, therefore, likely also do not accumulate functional HXK enzyme protein.              athxk1  athxk2             athxk3            A)   63  Figure 17: athxk1, athxk2 and athxk3 are TDNA insertional mutants. A) Exon map of AtHXK1, AtHXK2 and AtHXK3, black triangles represent location of TDNA insertion. B) Semi-qPCR amplifying AtHXK1, AtHXK2 and AtHXK3 in TDNA mutant lines, AtUBQ5 is a reference control.  To ascertain if athxk1 is not a knockout of AtHXK1 and therefore allelic to gin2 (glucose insensitive 2; Moore et al., (2003)), TDNA lines were germinated on high glucose media (Figure 18). All three TDNA lines, athxk1, athxk2 and athxk3 exhibited sensitivity to high glucose media, suggesting that they are not allelic to gin2 and validating evidence from the semi-q PCR experiment.  B)    64  Figure 18: athxk1, athxk2 and athxk3 TDNA seedlings on control media, 2-D-glucose; a non-metabolizable glucose analog, or 6% glucose after 7 DAG. Phenotypic analysis of plant growth To establish whether a decrease or complete loss of HXK protein affects plant growth during development, rosette diameter and stem height where measured. At 21 DAG, athxk1 was found to have a smaller rosette diameter compared to the wild type, 1.4±0.25 cm versus 3.16± (Figure 19, A) and a slightly shorter stem height at 32 DAG (Figure 20, A). athxk2 and athxk3 rosettes were not significantly different compared to the wild-type plants (Figure 19, A and B) at 21    65 DAG, but were found to have shorter stems at 42 DAG (Figure 20, B), which was defined as senescence. athxk2 had a stem height of 28.15±5.44 cm, athxk3 stem height was 30.15±2.95 cm while wild type possessed a stem height of 37.75±6.05 cm.   A)   66  Figure 19: Rosette diameter at 21 DAG of athxk1, athxk2 and athxk3 TDNA insertional mutant lines. A) Rosette diameter of athxk1 was significantly smaller than wild type, n = 6, * represents a significant difference from the wild type at p < 0.05, * represents a significant difference from the wild type at a Bonferroni critical value of p < 0.02, error bars represent SD. B) Rosettes pictured at 21 DAG, grey bar represents 1 cm.  It can be concluded, that a decrease or loss of AtHXK1 protein affects plant growth early in development, whereas a decrease or loss of AtHXK2 and AtHXK3 enzyme protein disturbs plant growth later in development.  B)    67   Figure 20: Stem height of athxk1, athxk2 and athxk3 TDNA mutant lines. A) Stem height pictured at 32 DAG. B) athxk2 and athxk3 have a shorter stem compared to wild type at 42 DAG (senescence), n = 10, * represents a significant difference from the wild type at p < 0.05, * represents a significant difference from the wild type at a Bonferroni critical value of p < 0.02, error bars represent SD.  B) A)   68 Non-structural carbohydrate in leaf and stem  Due to the differential expression of A. thaliana HXK transcript between source and sink tissue, it was desirable to establish whether a decrease or loss of HXK protein would change the amount of non-structural carbohydrate content in the leaf or stem. To this end, sucrose, glucose and fructose contents were measured in the three A. thaliana TDNA insertional lines (Figure 21). Sucrose content was significantly higher in the leaf of all three: athxk1 (8.25±1.70 mg/g), athxk2 (8.36±1.51 mg/g), and athxk3 (11.97±2.10 mg/g), compared to the wild type plants (4.21±0.85 mg/g; Figure 21, A). Sucrose in the stem of the wild type was found to be 5.27±2.82 mg/g, where athxk3 reached 9.94±1.97 mg/g, but athxk1 or athxk2 did not vary compared to the control plants.   Glucose and fructose constituted a significantly higher amount of non-structural carbohydrate in the leaf of athxk3, 38.41±8.28 mg/g and 29.71±7.71 mg/g, respectively. Glucose and fructose were not significantly higher in the stem or leaves of athxk1 and athxk2. Wild type stems were found to have 12.69±3.8 mg/g of glucose and 4.98±1.74 mg/g of fructose. Glucose and fructose levels in the leaf and stem of athxk1 were almost identical to the wild type. Intriguingly, athxk2 exhibited a small decrease in glucose and fructose content in both the leaf and the stem, which was not significant. From these observations, it may be concluded that a decrease in AtHXK1 and AtHXK2 may cause an accumulation of sucrose but not other soluble sugars, and that AtHXK3 an increase in sucrose, glucose and fructose of leaf tissue.      69   A) B)   70  Figure 21: Non-structural carbohydrate in the stem and leaf of athxk1, athxk2 and athxk3 TDNA insertional lines harvested at midday. A) Sucrose content. B) Glucose content. C) Fructose content. Units in mg/g DW, n = 5, * represents a significant difference from the wild type at p < 0.05, * represents a significant difference from the wild type at a Bonferroni critical value of p < 0.02, error bars represent SD. Starch content in leaf and stem  HXK is known to be involved in the breakdown of starch and, to assess if a decrease in HXK protein would change the amount of transient carbon stored in the leaves or stem, starch accumulation was estimated. As with non-structural carbohydrates, athxk3 had a large and significant increase in the percent starch found in the leaf compared to wild type, 88.32±7.97% versus 42.52±16.62% (Figure 22). A significant increase was also noted in the stem of athxk3, which was 7.24±1.05%, while wild type starch stem content was 4.59±0.93%. athxk2 had very similar values to the wild type and athxk1 appeared to have a small but non-significant increase C)   71 in starch in both leaf and stem. Therefore, AtHXK3 may be involved in the regulation of transient carbon storage as starch.    Figure 22: Percent starch in the stem and leaf of athxk1, athxk2 and athxk3 TDNA mutant lines harvested at midday. Units in percent DW, n = 5, * represents a significant difference from the wild type at p < 0.05, * represents a significant difference from the wild type with a Bonferroni critical value of p < 0.02, error bars represent SD. Structural carbohydrates and lignin in the stem   An increase in non-structural carbohydrate may be explained by a redirection of carbon from structural carbohydrate. Structural carbohydrate content of the stem was therefore measured in athxk1, athxk2 and athxk3 A. thaliana TDNA insertional lines. The structural carbohydrate content changed for some sugars in athxk1 and most sugars of athxk2 and athxk3 (Table 5).    72 Arabinose and galactose content was significantly lower in athxk1, athxk2 and athxk3 compared to the wild type. For arabinose, the quantity of sugar per mass sample was 7.78±0.40 µg/mg in athxk1, 6.46±0.30 µg/mg in athxk2 and 7.73±0.64 µg/mg in athxk1, while wild type contained 8.87±0.34 µg/mg arabinose. Galactose content was found to be 13.67±0.33 µg/mg for athxk1 stem, 11.34±0.40 µg/mg for athxk2 stem and 13.11±0.95 µg/mg for athxk3 stem, while wild type galactose levels were 15.48±0.46 µg/mg. athxk2 but not athxk1 or athxk3 had a significant decrease in rhamnose, 6.56±0.31 µg/mg compared to the wild-type; 7.53±0.28 µg/mg.   On the other had, athxk2 and athxk3 had a significant increase in glucose, xylose and mannose in the stems. For athxk2 those values were 387.12±11.09 µg/mg glucose, 142.97±5.54 µg/mg xylose, and 23.07 µg/mg mannose. For athxk3, 388.60±9.67 µg/mg glucose, 142.31±4.74 µg/mg xylose and 24.92±1.86 µg/mg mannose. This data suggests that the main crystalline component of the stem, cellulose, is unaffected by a loss or decrease of HXK2 and HXK3 enzyme protein, but other structural components of the cell wall are affected.         73 Table 5: Average quantity of sugar per mass sample (ug/mg) in athxk1, athxk2 and athxk3 primary stems. Bold values indicate a significant difference from the wild type at p < 0.05, underlined values indicate a significant difference from the wild type at a Bonferroni critical value of p < 0.02, n = 5, SD in brackets.  Arabinose Rhamnose Galactose Glucose Xylose Mannose WT 8.87 (0.34) 7.53 (0.28) 15.48 (0.46) 359.00 (13.44) 125.38 (3.93) 20.52 (1.14) athxk1 7.78 (0.40) 7.24 (0.33) 13.67 (0.33) 363.48 (12.53) 128.01 (5.50) 22.10 (2.55) athxk2 6.46 (0.30) 6.56 (0.31) 11.34 (0.40) 387.12 (11.09) 142.97 (5.54) 23.07 (1.32) athxk3 7.73 (0.64) 7.30 (0.66) 13.11 (0.95) 388.60 (9.67) 142.31 (4.74) 24.92 (1.86)  Following the determination of a change in the monomeric composition of cell wall carbohydrates in athxk1, athxk2 and athxk3, it was of interest as to whether this change would affect other components of the cell wall such as lignin. Consequently, average percent soluble and insoluble lignin in the stem was determined and total lignin represented as the addition of both (Table 6). The soluble lignin fraction of athxk1; 2.69±0.04%, athxk2; 2.17±0.08% and athxk3; 2.29±0.04% was found to be slightly but significantly decreased compared to the wild type; 3.22±0.06%. To compensate, the insoluble lignin fraction of athxk3 stem was higher; 19.01±0.37% compared to the wild type (18.07±0.74%), resulting in an equal total percent lignin composition of the stem. The insoluble portion of athxk1 and athxk2 lignin appeared to be slightly but non-significantly increased as well (18.14±0.98% and 18.91±0.55%, respectively). These results indicate that a decrease in the amount of functional AtSPS1 and AtSPS2 enzyme in the stem of A. thaliana is able to change the composition of lignin in the stem, without affecting total lignin deposition.     74 Table 6: Average percent insoluble, soluble and total lignin in the main stem of athxk1, athxk2 and athxk3. Bold values indicate a significant difference from the wild type at p < 0.05, underlined values indicate a significant difference from the wild type with a Bonferroni critical value of p < 0.02, n = 5, SD in brackets.  Insoluble lignin Soluble lignin Total WT 18.07 (0.74) 3.22 (0.06) 21.29 (0.80) athxk1 18.14 (0.98) 2.69 (0.04) 20.83 (0.97) athxk2 18.91 (0.55) 2.17 (0.08) 21.08 (0.62) athxk3 19.01 (0.37) 2.29 (0.04) 21.30 (0.36)    75 Chapter 4: Discussion  4.1 Effects of loss of SPS in A. thaliana and P. trichocarpa  SPS is a soluble cytosolic enzyme that catalyzes the formation of sucrose in photosynthetic and non-photosynthetic tissues by combining UDP-G and F6P to produce S6P in the following reaction (Huber & Huber, 1996):  UDP-G + F6P !"S6P + UDP Subsequently, S6P is rapidly metabolized to sucrose by the cleavage of the phosphate unit by SPP, recently found to interact in an in vivo complex with SPS (Maloney et al., 2015) to form a metabolic channel. SPS is a low abundance enzyme, even in source leaf where in most species it is present in the highest amounts (Huber and Huber, 1996). Nonetheless, the flux control coefficient for sucrose synthesis by SPS in A. thaliana leaf is 0.2 (Strand et al., 2000) and 0.5 in potato leaf (Geigenberger et al., 1995) showing that SPS plays a significant role in the anabolism of sucrose, but may not be the only enzyme involved (H. Winter & Huber, 2000). This significance is additionally exemplified by the numerous levels at which SPS is regulated, including transcriptional in response to diurnal changes, external cues and developmental changes (Gibon et al., 2004; Okamura et al., 2011; Privat et al., 2008). SPS is regulated post-translationally through phosphorylation by 14-3-3 protein, which inactivates activity during light/dark transitions (Toroser, Athwal, & Huber, 1998) and allosterically by metabolic effectors. G6P is a positive regulator of SPS activity, while inorganic phosphate is a negative regulator (Huber & Huber, 1996). It has also been found, however, that not all enzymes in A. thaliana are regulated in the same way as it has been proposed for spinach and maize SPS: only AtSPS3 was positively regulated by G6P, where its activity was 6.8-fold higher with the regulator, and its   76 activity was reduced 4-fold with the addition of Pi (Volkert et al., 2014).  The importance of SPS in sucrose metabolism has been illustrated numerous times with overexpression studies in poplar (Park et al., 2009), tobacco (Park et al., 2008), tomato (Galtier, Foyer, Huber, Voelker, & Huber, 1993; Worrell, Bruneau, Summerfelt, Boersig, & Voelker, 1991)  and rice (Ishimaru, Ono, & Kashiwagi, 2004) where results show dramatic changes in carbohydrate content, photosynthetic capacity and sometimes, plant phenology. Overexpression of A. thaliana SPS in hybrid poplar resulted in increased intracellular sucrose concentrations in both leaf and stem. These changes were associated with a prolonged onset of senescence and an earlier onset of budflush the following growth season, while no other phenotypic changes were observed compared to the control. In tobacco, only sink sucrose levels were increased, but stem heights were longer due to internode elongation and longer stem fibers (Park et al., 2008). Total dry biomass was also increased compared to the control plants. Maize SPS overexpression in tomato resulted in increased sucrose and a reduction in leaf starch without any detrimental effects to the plant growth (Worrell et al., 1991). Total change in plant biomass was not observed, but SPS activity closely correlated with an increased shoot to root ratio of dry mass. Photosynthetic rate was increased only at saturating CO2 concentrations by 20% with total RUBISCO activity remaining the same (Galtier et al., 1993). And, rice plants overexpressing maize SPS were found to be significantly taller than Nipponbare (control rice variety) during the early growth stage. This increase was suggested to be due to higher SPS enzymatic activity, allowing for the translocation of higher amounts of sucrose from the leaf (Ishimaru et al., 2004). These overexpression studies point to SPS enzyme as being responsible for increased intracellular   77 sucrose concentrations, which may or may not result in an accumulation of biomass likely due to the subsequent utilization of sucrose by the particular species. For example, this has been demonstrated in tomato fruit, where an SPS overexpressing line also exhibited at 25% increase in SuSy activity (N'tchobo et al., 1999).   Far fewer experiments have attempted the down regulation of endogenous SPS transcript in the plant, and most have traditionally focused on the effects of photosynthetic SPS activity. For example, Strand et al. (2000) down regulated the expression of SPS in A. thaliana and examined the effects on photosynthesis. Sucrose accumulation was inhibited in fully expanded source leaf but phosphorylated intermediates did not accumulate and carbon was not redirected towards starch synthesis. Pi was shown to be recycled by end product synthesis, as PGA to Triose-P ratios remained low and ATP to ADP ratios high. Photosynthesis, instead, was inhibited in the regeneration of ribulose-1,5-bisphosphate. The authors suggested that the unexpected response to decrease of SPS was due to a threefold decrease in PPi, an increase in UDP-G to hexose ratio and a lack and increase in AGPase activity. In tobacco, Chen et al. (2005) decreased the transcript levels of NtSPSA and NtSPSC, where the A isoform is ubiquitously expressed and C is specific to source leaves. NtSPSA suppression had no effect on plant photosynthesis. NtSPSC down regulation resulted in increased leaf starch that was due to an impairment of transitory starch reserve remobilization during the dark and the accumulated maltose. NtSPSC transcript was found to be diurnally regulated, with high expression during the night, suggesting it is responsible for sucrose reformation from starch breakdown (Chen et al., 2005). Recently, Volker et al. (2014)  used TDNA insertional lines of AtSPS1 and AtSPS4 (AtSPSA1 and AtSPSC in   78 Volkert et al. (2014) to study decreased SPS protein levels in leaf tissue. Both knockouts had decreased sucrose levels and AtSPS4 knockout had increased starch levels at the end of the light and dark periods. A double mutant of the two knockout lines was strongly inhibited in growth and accumulated significant quantities of starch. It was concluded that a decrease in AtSPS1 and AtSPS4 protein does not cause a redirection of carbon into starch but an impairment to remobilize starch during the light or the dark period as sucrose exudate from the petiole was significantly lower compared to the wild type plants (Volkert et al., 2014). In combination, these findings suggest that different SPS gene family members have diverse functions with respect to sucrose synthesis and that decrease of SPS enzyme activity does not inhibit sucrose synthesis during the light period, possibly due to compensation by other SPS enzymes.   4.1.1 Phylogenetic relationships between plant SPS genes and putative function of A. thaliana and P. trichocarpa enzymes based on endogenous expression. Sucrose synthesis is thought to have arisen in green plants (Viridiplantae) that carry out oxygenic photosynthesis through the endosymbiotic cyanobacteria, the ancestors of modern chloroplasts (Cavalier-Smith, 2000). Lutfiyya et al. (2007) recognize one group of bacterial SPS genes and three groups of plant SPSs that contain sequences from both monocot and dicot species, suggesting that the three plant forms arose before the monocot/dicot split. Phylogenetic analysis undertaken in this study utilized the same approach to evaluate the relationships between A. thaliana, and other previously analyzed SPS genes from maize, rice, tomato, potato, M. truncatula, P. patens and tobacco, as well as the unanalyzed species P. trichocarpa, cotton and Eucalyptus grandis. In congruence with previous phylogenetic studies (Castleden et al., 2004;   79 Langenkämper et al., 2002; Lutfiyya et al., 2007), three groups of plant protein coding genes were identified (Figure 2). The genes analyzed in the study all fall within group 2d (dicotylenous): AtSPS1, AtSPS2, PtSPS1, PtSPS2, PtSPS3. It has been suggested that the different groups represent functional significance and indeed, group 2d contains sequence specific regulatory binding sites for light-dark regulation, 14-3-3 binding (proposed to inactivate SPS by phosphorylation), osmotic regulation, as well as F6P and UDP-G binding sites (suggesting active enzymatic functionality). Conversely, group 2m lacks the conserved 14-3-3 binding and the osmotic regulatory sites (Lutfiyya et al., 2007). Furthermore, group 2d contains NtSPSA, a protein that was found to be ubiquitously expressed in tobacco and likely responsible for sucrose synthesis from Triose-P. Indeed, in leaf and stem tissues analyzed in silico, AtSPS1 transcript was consistently highest in expression (Figure 3). Promoter-GUS analysis found that in 11-day old seedlings, AtSPS1 is ubiquitous (Volkert et al., 2014) and consistently highest in source and sink tissue such as mature leaf, young leaf, flower, root, silique and bud throughout the lifecycle of the plant. In contrast, AtSPS2 GUS signal was found only in root of 11 day old seedlings with no signal in later stages (Volkert et al., 2014). Another study found that AtSPS2 enzyme is highly abundant after cold stress and most abundant of the SPS enzymes in the stem (Lehmann, Wienkoop, Tschoep, & Weckwerth, 2008). PtSPS1, PtSPS2 and PtSPS3 are also highest in expression in P. trichocarpa; however, PtSPS1 and PtSPS2 are higher in cambium than in source leaf and PtSPS3 expression is higher in source leaf.   NtSPSC was shown to fall in group 4, which also contains AtSPS4 and PtSPS4, and was found to be leaf specific, diurnally regulated and functions to synthesize sucrose from starch breakdown (Chen et al., 2005). Volkert et al. (2014) who found that AtSPS4 is highly expressed in leaf and   80 its mutant lacks the ability to mobilize starch during the dark period corroborated this finding. Interestingly, Lehman et al. (2008) found that AtSPS4 levels are higher during the day than at night, which is unexpected if AtSPS4 functions to resynthesize sucrose from starch breakdown products.   AtSPS3, PtSPS5 and PtSPS6 fall within group 3d, sequences that are believed to lack a conserved osmotic regulatory site. In 11-day old seedlings, AtSPS3 promoter expression was shown to be ubiquitous, but expression later on is confined to the reproductive organs (Volkert et al., 2014). Even though AtSPS3 appears to be the only A. thaliana SPS enzyme allosterically regulated by G6P and Pi, protein levels remain low even during cold stress (Lehman et al., 2008). PtSPS5 and PtSPS6 show almost no transcript presence in either source leaf or cambial tissue (Figure 4).   Thus, based on phylogenetic relationships to previously analyzed SPS enzymes, it can be speculated that group 2d enzymes, including AtSPS1, AtSPS2 in A. thaliana and PtSPS1, PtSPS2 and PtSPS3 in P. trichocarpa, are involved in the synthesis of sucrose from photosynthetic derivatives in source and sink organs. Due to the differential expression of the closely related enzymes, it can be speculated that there is functional specialization between source and sink tissues of AtSPS1 and AtSPS2 (PtSPS1, PtSPS3 and PtSPS2). Group 4; AtSPS4 and PtSPS4, likely synthesize sucrose in the dark. And group 3; AtSPS3, PtSPS5 and PtSPS6 remain low in expression but have the ability to be highly stimulated in activity (6-fold in A. thaliana), potentially to respond quickly to external stimuli.    81 4.1.2 Loss of photosynthetic SPS  The most unexpected result of the study was that as opposed to a decrease in sucrose and other soluble sugars, loss of group 2d SPS enzymes resulted in an increase of non-structural carbohydrates in both A. thaliana and P. trichocarpa representative source and sink tissues. Although no groups have previously reported a non-structural carbohydrate increase, A. thaliana SPS down-regulated lines were less inhibited in sucrose production in source leaf than anti-cytosolic fructose-1,6-bisphosphatase (cFBPase), an enzyme upstream of the point where sucrose synthesized from the day and night converge (Strand et al., 2000). When SPS activity is decreased, UDP-G to hexose ratios increased (Strand et al., 2000), and this accumulation is likely due to the inhibition of SuSy to cleave sucrose (Geigenberger & Stitt, 1993; Schaffer & Petreikov, 1997) and reverse the reaction to sucrose synthesis resulting in an accumulation of sucrose in the leaf (Figure 7, A; Figure 11, A). Sucrose is exported and transported to sink organs such as stem, but its utilization is inhibited so that instead of increasing plant growth, lines likely lacking a large percentage of active SPS enzyme are stunted in growth (Figure 6, Figure 10), consistent with previous reports of SPS down regulation in A. thaliana (Strand et al., 2000). Decreases in growth were previously linked to lower rates of photosynthesis in anti-SPS lines, manifesting from a lower total amount of protein synthesis. Carbohydrate metabolism regulates inorganic nitrogen assimilation (Krapp & Stitt, 1995) and a perturbation in protein synthesis, therefore, can be expected in SPS-lacking plants. Cell wall and vacuolar INV likely remain active due to the availability of substrate (sucrose) and continue forming hexoses (Figure 7, B & C; Figure 11, B & C). In cold-girdling manipulations of spinach source leaves, sucrose, glucose and fructose increased two to five-fold, while SPS enzyme levels slightly decreased (Chen et al., 2008; Krapp & Stitt, 1995). These findings suggest that decreases in the amount of active SPS   82 enzyme promotes a stress response. Accumulation of sucrose in leaves has closely been linked to osmotic and cold stress (Chen et al., 2008; Geigenberger, Reimholz, Deiting, Sonnewald, & Stitt, 1999), resulting in a phenotype where non-structural carbohydrates are increased in sink tissue but cannot be employed in growth. Wild type potato tubers induced sucrose synthesis under osmotic stress (Geigenberger et al., 1999) and cold treatment for 34 days at 4° C (Chen et al., 2008). Sucrose was found to accumulate in SPP down-regulated plant lines and suggested that residual SPP is substrate saturated and therefore operates at rates similar to wild type lines. The A. thaliana genome encodes for four SPS enzymes while the P. trchocarpa genome encodes for six. This suggests a possibility of residual functional enzymes not targeted by TDNA and RNAi. In addition, AtSPS3 was shown to be able to be upregulated six fold in planta (Volkert et al., 2014) and could, therefore, be compensating for the low activity of AtSPS1 and AtSPS2 in TDNA lines. This suggests that increased sucrose levels in the leaf could be attributed to up-regulated AtSPS3 enzyme protein. Furthermore, AtSPS4 is also likely active in the leaf tissue to compensate for loss of AtSPS1 and AtSPS2, as it is the second major SPS leaf isoform (Figure 3; Volkert et al. (2014)).   Starch content was found to be higher in leaf tissue of atsps1-1 and atsps2 lines (Figure 8). This increase could be due to increased partitioning of Triose-P to starch or alternatively to lack of remobilization of starch following the dark period. Previously, it has been shown that decreasing SPS1 activity to 44% lead to slight increases in leaf starch at the end of the light and dark period (Volkert et al., 2014). Although in a single knockout mutant of AtSPS1, there was no change in maltose levels at the end of the dark period, in double knockouts of AtSPS1 and AtSPS4 sucrose levels in phloem exudates in the petiole of A. thaliana source leaf reached only 25% of the wild   83 type (Volkert et al., 2014). The lack of accumulation of starch in Volkert et al. (2014) single AtSPS1 knockouts in leaf compared to those found in this research could be attributed to that plants were grown on short day cycles (8 hours) versus the long day cycles used in this study (16 hours), exacerbating the effect of loss of SPS enzyme. In tobacco, NtSPSC, orthologous to AtSPS4, PtSPS5 and PtSPS6, was found to be responsible for resynthesis of sucrose from starch following a dark period, while down regulating NtSPSA, orthologous to AtSPS1, AtSPS2, PtSPS1, PtSPS2 and PtSPS3, had no effect on leaf carbohydrate metabolism (Chen et al., 2005).   Loss of AtSPS1, but not AtSPS2 resulted in increased arabinose content, a structural component of the cell wall (Table 3), while stem cellulose crystallinity and other structural components of the cell wall were similar to wild type (data not shown). The apparent increase in arabinose could not be explained. Nonetheless, these results are consistent with the fact that plants lacking SPS enzyme have increased sucrose levels that cannot be utilized effectively by the sink for growth of the cell wall or fiber production. Plants with increased levels of SPS through overexpression have the ability to maintain high sucrose concentrations and presumably channel this metabolite to the production of fiber as a terminal sink and increase biomass in tobacco (Park et al., 2008). Overexpression of sucrose in hybrid poplar did not lead to a utilization of sink organs to produce more biomass (Park et al., 2009). This may be as result of the difference between crop species, which are generally bred for fast growth versus a tree. Expressing SPS as a fusion construct with SPP promoted plant growth in hybrid poplar (Maloney et al., 2015), implying that sucrose needs to be formed in the appropriate metabolic channel to be utilized efficiently by the tree.     84 4.2 Effects of loss of HXK in A. thaliana  In less than a decade, it will be one hundred years since the discovery of hexokinase (HXK) in an extract from baker’s yeast (Saccharomyces cerevisiae) by Otto Meyerhof, the 1922 Nobel Prize winner in Physiology or Medicine (Claeyssen & Rivoal, 2007). The reaction it catalyzes was described a decade later, where:  hexose + ATP " hexose-6-phosphate + ADP by Von Euler and Adler (1935), and separately by Meyerhof (1935). In the following decade, HXK was established as an integral enzyme in the Embden–Meyerhof–Parnas pathway, now termed glycolysis, an ancient and conserved mechanism in plants and animals to produce ATP and pyruvate (Plaxton, 1996). Despite HXK being integral to many living organisms and its basic function known to scientists, several remaining gaps in knowledge exit. The challenge behind explaining the function of HXK in a whole plant systems-biology approach is that the enzyme not only plays an enzymatic function in the metabolism of sucrose, itself a complicated multi-path system, but a signaling role, a stress-related function, and a hormone cross-talk function (Claeyssen & Rivoal, 2007; Granot et al., 2013; Rolland, Baena-Gonzalez, & Sheen, 2006) .   In plants, HXKs have been found to segregate into HXK enzymes that retain phosphorylating capability, and HXK-likes (HXKL, enzymes that cannot phosphorylate hexoses (A. Karve et al., 2008), and both have been found to exist in basal organisms such as Physcomitrella patens and Selaginella moellendorffii (R. Karve et al., 2010). The purpose for the presence of non-phosphorylating HXKLs in a wide array of organisms, including basal plants has not been fully established. While some are able to bind glucose, like AtHXKL1 and AtHXKL2, AtHXKL3 is   85 thought not to be able to bind the ligand (R. Karve et al., 2010). HXKLs are therefore, predicted to have only a regulatory function with AtHXKL1 believed to be a negative regulator of plant growth under high glucose availability and its downstream effects may be hormone regulated (A. Karve & Moore, 2009). HXKs, on the other hand, have been shown to have the ability to phosphorylate several different hexoses including glucose, fructose, mannose and with some debate, glucosamine and galactose (Claeyssen & Rivoal, 2007; Granot et al., 2013; Saltman, 1953). Therefore, unlike animals, no glucose-specific glucokinase has been found and glucose can only be phosphorylated by HXK unlike fructose, which can be phosphorylated by HXK and fructokinase (FRK) (Dai et al., 2002). Although HXKs can phosphorylate a variety of hexose sugars, the Michaelis-Menten kinetics of the various reactions suggest that in vivo HXK mainly reacts with glucose due to a low Km and a high intracellular concentration particularly in sink organs (Claeyssen & Rivoal, 2007). In potato tuber, cytosolic glucose concentrations may be as high as 30 mM, while those of fructose and mannose are either undetectable or three orders of magnitude lower (Farre et al., 2001).  The choice of substrate may also be determined by the intracellular location of the enzyme (Granot et al., 2013). N-terminal sequences of HXKs have been shown to localize enzymes in the plastid (A group) and the outer mitochondrial membrane (B group). Group A HXKs are characterized by a ~30 amino acid N-terminal transit peptide, while type B contain a 24 amino acid hydrophobic membrane anchor domain. A third type of cytosolic HXK has also been found to occur only in monocots (Y. Cho, Yoo, & Sheen, 2006). In addition, type A HXKs, such as AtHXK3, have been found to localize to the stroma of chloroplasts in A. thaliana, tobacco,   86 tomato, P. patens and rice (J. Cho et al., 2006; Giese, Herbers, Hoffmann, Klösgen, & Sonnewald, 2005; Kandel-Kfir et al., 2006; A. Karve et al., 2008; Olsson, Thelander, & Ronne, 2003). In moss, it was found that plastidic HXK is responsible for 80% of the phosphorylation capacity of glucose and 47% of phosphorylation of fructose, and that growth in the dark is severely impaired in PtHXK1 mutants (Olsson et al., 2003). Thus, it was suggested that plastidic HXK supplies the reactions of that organelle with G6P in the dark and in sink tissues where energy supply might be more limited. With the absence of a HXK in the plastid, glucose would have to be exported, phosphorylated and reimported by a G6P translocator to supply reactions of the plastid such as glycolysis, the oxidative pentose phosphate pathway (OPPP) and starch synthesis (Claeyssen & Rivoal, 2007). Promoter expression analysis of tobacco NtHXK2, the first higher plant plastidic HXK to be examined, indicated that its expression is restricted to the starch sheath (the inner most layer of the cortex, specialized to store starch), xylem parenchyma, guard cells and root tips (Giese et al., 2005). The authors hypothesized that due to its localized expression, NtHXK2 produces G6P that is fed into OPPP to yield erythrose-4-phosphate, the starting material for the shikimate pathway and further phenylpropanoid synthesis (Giese et al., 2005; Hatton et al., 1995). Tomato plastidic HXK, LeHXK4, was found to be expressed in all tissues and to have a high affinity for glucose (Kandel-Kfir et al., 2006). The presence of plastidic HXK in starchless tissue of tomato suggests that it may function to phosphorylate incoming glucose rather than glucose from the breakdown of starch (Kandel-Kfir et al., 2006).  Group B HXKs, famously known for AtHXK1, the most ubiquitously expressed HXK in all A. thaliana organs, and AtHXK2, a homolog of AtHXK1, have been found to localize   87 intracellularly to the mitochondrial membrane through proteomic analysis and GFP localization (Giege et al., 2003; A. Karve et al., 2008). Homologous proteins in P. trichocarpa; PtHXK1, tobacco; NtHXK1, rice; OsHXK2, 3, 5, 6, 9 and spinach, SoHXK1 were also found to associate with the mitrochondrial membrane (J. Cho et al., 2006; Damari-Weissler et al., 2007; R. Karve et al., 2010; M. Kim et al., 2006). The removal of the putative N-terminal membrane anchor domain was shown to target membrane-associated HXKs to the cytosol (Damari-Weissler et al., 2007; M. Kim et al., 2006) and therefore confirmed its requirement for localization to the mitochondrial membrane. There, in non-photosynthetic tissues, HXKs have been thought to perform their primary role to produce G6P from imported sucrose during the day. At night, HXKs are also necessary to phosphorylate glucose from the breakdown of starch (Granot et al., 2013). It was unexpected that HXK was found to be expressed throughout the diurnal cycle in photosynthetic tissue as well as sink tissue (Y. Cho et al., 2006; J. Jang, León, Zhou, & Sheen, 1997; Kandel-Kfir et al., 2006) as presumably there would be no free glucose or fructose produced in the light in photosynthetic organs. Spatio-temporal data of specific cell types and any minor changes in transcript abundance, however, are still unavailable for the HXK gene family (Granot, Kelly, Stein, & David-Schwartz, 2014) .   Perhaps unsurprisingly, due to their ubiquitous expression and convergence of sucrose metabolism, B type HXKs have been implicated in a variety of physiological functions including: hexose phosphorylation, glucose sensing coupled with photosynthetic gene repression at excess glucose levels, programmed cell death, mediation of sugar and hormonal interactions, growth and development including initiation of flowering, transpiration, oxidative stress   88 response, directional root growth and leaf senescence (Aki et al., 2007; Balasubramanian, Karve, & Moore, 2008; J. Cho et al., 2006; Dai et al., 1999; J. Jang et al., 1997; A. Karve et al., 2008; M. Kim et al., 2006; Moore et al., 2003; Pourtau, Jennings, Pelzer, Pallas, & Wingler, 2006; Rolland et al., 2006; Sarowar, Lee, Ahn, & Pai, 2008; Yanagisawa et al., 2003). The pathways, which feed the HXK reaction as well as the products of that reaction, are varied, giving HXKs the ability to have diverse effects. In heterotrophic tissues, sucrose is transported via the phloem and imported into the cell symplastically or apoplastically through sucrose transporters (SUTs). Alternatively, sucrose is broken down in the apoplast by cell wall INV to glucose and fructose and imported through monosaccharide transporters (MSTs) located on the plasma membrane (Lalonde et al., 2004). High turnover rates of sucrose in heterotrophic tissues (Nguyen-Quoc & Foyer, 2001) suggest that sucrose is rapidly metabolized to monomers, likely by SuSy in the cytosol since cytosolic INV activity has been reported to be significantly lower than cell wall or vacuolar INV (J. Y. Kim, Mahe, Brangeon, & Prioul, 2000). UDP-G produced by the breakdown of sucrose can be used for cell wall synthesis, a variety of interconversion reactions, or resynthesis of sucrose-P by the action of SPS (Bar-Peled & O'Neill, 2011; Dennis & Blakeley, 2000). In order for sucrose to be resynthesized, fructose must be phosphorylated to F6P by the action of either FRK or HXK. SuSy is feedback inhibited by its product, fructose at concentrations of 0.5-1 mM (Schaffer & Petreikov, 1997) where at higher concentrations SuSy reverts to synthesis of sucrose rather than cleavage (Claeyssen & Rivoal, 2007). Thus, phosphorylation of fructose by FRK has been suggested to be an important regulatory step in the heterotrophic sucrose cycle. Antisense expression of tomato FRK2 was found to be essential for the development of phloem and xylem, secondary cell wall deposition and normal cambial activity (Damari-Weissler et al., 2009). It is currently unknown what the overall impact of   89 phosphorylation by HXK on fructose is on this pathway, however. In order to be used by the cell, glucose and fructose monomers imported through MSTs on the plasma membrane or the vacuolar membrane must be phosphorylated by HXK. G6P and F6P may directly enter glycolysis for the production of ATP, or G6P may be translocated to the plastid through glucose phosphate transporters (GTPs) to undergo glycolysis and there enter the OPPP or be used for starch synthesis. F6P could be utilized directly to produce sucrose-P with the addition of UDP-G or be isomerized to G6P through phosphoglucose isomerase (PGI), or to mannose-6-phosphate (M6P) through phosphomannose isomerase (PMI). G6P is then converted to G1P by phosphoglucomutase, which may again enter glycolysis or further be transformed to UDP-G by UDP-glucose pyrophosphorylase. At this step, UDP-G is available for resynthesis of glucose by SPS, or entrance into the endoplasmic reticulum along with GDP-mannose (GDP-M) and other nucleotide sugars for the synthesis of cell wall matrix polysaccharides or protein glycosylation (Claeyssen & Rivoal, 2007). HXKs, therefore, activate sugars derived from multiple sources such as phloem import, the vacuole and plastid and allow for the entrance of glucose, fructose and mannose into various metabolic pathways or the resynthesis of sucrose.   This cycle of sucrose breakdown and resynthesis is thought to serve several different metabolic functions. Firstly, sucrose cycling may contribute to buffering variations in metabolite concentrations and modulation of carbon flow (Claeyssen & Rivoal, 2007). Breakdown of sucrose allows for phosphorylation of monosaccharides by HXK and activation of the aforementioned pathways. If more intermediates are required, sucrose can be broken down and utilized. Conversely, if these activated sugars are in excess they can be removed out of their   90 “active” state by sucrose resynthesis from UDP-G and F6P by SPS. This cycle increases the ATP energy demand of the cell (Fernie, Tiessen, Stitt, Willmitzer, & Geigenberger, 2002)  and thus allows for the modulation of carbon flow within the cell via pathways such as glycolysis or the carbon flow to the cell from phloem transport. A further cycle, which may be under (fully or partly) the control of HXK, is the modulation of starch degradation such that at night when maltose is broken down into glucose (S. C. Zeeman, Kossmann, & Smith, 2010) , phosphorylation of glucose by HXK allows for the maintenance of the glucose gradient from the plastid (Claeyssen & Rivoal, 2007). Finally, a variety of plants employ the raffinose family of oligosaccharides (RFOs) for storage and transport of soluble sugars (Turgeon, 2006). A. thaliana has been shown to transport a small amount of RFO’s (Panikulangara, Eggers-Schumacher, Wunderlich, Stransky, & Schoffl, 2004) as has P. trichocarpa, which expresses galactinol synthase (GolS) in a diurnal pattern with a peak in the middle of the light period (Unda et al., 2012). Breakdown of RFOs at the sink will produce glucose and fructose that can subsequently be phosphorylated by HXK, and galactose, which is phosphorylated by galactokinase (GalK) to galactose-1-phosphate (Gal1P) (Caspi et al., 2014). Gal1P can be transformed into UDP-G through UDP-glucose/UDP-galactose-4-epimerase (UGE1) and enter the UDP-xylose biosynthesis pathway or form sucrose through SPS. Alternatively, Gal1P can enter glycolysis by conversion to glucose-1-phosphate (G1P) (Caspi et al., 2014).   Besides its metabolic role, HXKs have been implicated in sugar sensing. The first set of experiments aimed at elucidating this role used maize protoplasts where exogenous sugar was found to repress the expression of the photosynthetic genes RUBISCO and chlorophyll A/B   91 binding protein (CAB) (J. C. Jang & Sheen, 1994). The substrates of HXK, glucose and fructose, as well as the glucose analog, 2-deoxyglucose, which is phosphorylated by HXK but cannot be metabolized further, were found to repress expression. These results were corroborated with the use of A. thaliana seedlings, which were also inhibited in growth and photosynthesis by high glucose and glucose analogs (J. Jang et al., 1997). HXK was decisively dubbed a “moonlighting protein” when its catalytic and signalling activities were uncoupled in an amino acid substituted catalytic domain. Expressing the catalytically inactive protein in an AtHXK1 mutant background, termed glucose insensitive 2 (gin2) was sufficient to inactivate a glucose-mediated response of chlorophyll decrease and repress photosynthesis genes (Moore et al., 2003). The mode of mechanism of AtHXK1 gene repression is not known, however, it has been shown that AtHXK1 protein translocates to the nucleus where it forms a complex with vacuolar H+-ATPase B1 (VHA-B1) and the 19S regulatory particle of the proteasome subunit (RPT5B) (Y. Cho et al., 2006). This complex has the ability to bind to the promoter of CAB2 and mediate the repression of its expression in a glucose dependent manner. In addition, mutant lines of gin2, vha-B1 and rpt5b showed similar adult phenotypes implying a requirement for all three proteins for normal signalling function (Y. Cho et al., 2006). AtHXK2 has also been found to translocate to the nucleus and mediate photosynthetic gene repression (J. Jang et al., 1997; A. Karve et al., 2008), a function that may be due to its close sequence similarity to AtHXK1 (Figure 13). Type A HXKs have been implicated in sugar sensing. For example, A. thaliana AtHXK3 knockout mutant was found to be insensitive to 7% exogenously applied glucose. This is unlike the results presented here, where AtHXK3 knockouts grown on 6% glucose where found to be repressed in growth and cotylendon expansion following 7 days of growth (Figure 18).   92 The discovery that AtHXK1 mediates repression of photosynthetic gene expression in source tissue such as mesophyll cells contradicts the theoretical suggestion of the lack of free glucose produced in the cell during the day. In fact, free glucose may arise from the cleavage of trehalose, itself found to be an important signalling molecule, the degradation of starch, cleavage of sucrose by cytosolic INV and import of glucose monomers from the apoplast, resulting from sucrose cleavage by cell wall INV (Granot et al., 2013). It is probable that free glucose produced in the cell during the day has the ability to induce signalling as its concentration is likely low during active photosynthesis. When the cell is actively photosynthesizing, starch would not be degraded but stored, exported sugars into the apoplast will likely be transported into the phloem for transportation to sinks, and the activity of cytosolic INV is generally low (J. Y. Kim et al., 2000) as is the concentration of trehalose (Paul et al., 2008).  In addition to its direct control of gene expression, AtHXK1 signalling has been found to interact with a variety of hormone signalling pathways. For example, ABA is thought to act synergistically to AtHXK1 signalling as some of the isolated glucose insensitive mutants were, in fact, ABA-synthesis and ABA-insensitive alleles and addition of exogenous glucose promoted the increase of endogenous ABA levels (Cheng et al., 2002). In contrast, the addition of the ethylene precursor aminocyclopropane-1-carboxylic acid (ACC), promotes cotylendon greening even at high glucose concentrations in the wild type (Gibson, Laby, & Kim, 2001)  and AtHXKL1 is required for the ACC response (R. Karve et al., 2010).     93 4.2.1 Phylogenetic relationships between A. thaliana and P. trichocarpa HXKs Phylogenetic analysis of known and putative HXK coding sequences in various monocot and dicot species and the bryophyte P. patens revealed the existence of six phylogenetic clades (Figure 13). These groups were named according to the designation established by R. Karve et al. (2010). The omission of lycophyte sequences in the analysis precluded the addition of a seventh clade of HXKs, specific to the lycophyte family. Group 3 and group 5 contain the HXKL genes, which according to R. Karve et al. (2010) are most basal and contain representatives from all species queried. Here, group 3 was found to have representatives of all species including A. thaliana (AtHXKL1 and AtHXKL2), P. trichocarpa (PtHXKL1 and PtHXKL2), tomato, potato, tobacco, Medicago truncatula, Eucalyptus grandis, rice and corn. The monocot sequences from rice and corn form a subclade, termed 3a and dicots 3b (Figure 13). Group 3 sequences distinguish themselves by the presence of a 6-10 amino acid N-terminal indel in the adenosine-binding domain (R. Karve et al., 2010) and this indel was also present in P. trichocarpa sequences examined (data now shown). Group 5 on the other hand contained sequences from only four species, A. thaliana AtHXKL3, P. trichocapa PtHXKL3 and one sequence from tomato and potato. HXKLs from this group, including AtHXKL3 and PtHXKL3 contain a different 7 amino acid indel in the N-terminal domain (data not shown). Although both groups have been found to lack catalytic activity, group 3 is able to bind glucose while group 5 is thought not to be able to bind glucose (R. Karve et al., 2010). Group 4 contains plastid stromal localized proteins, which are also referred to as type A. This group includes representatives from a wide variety of plant species, including A. thaliana AtHXK3, P. trichocarpa PtHXK3, as well as sequences from E. grandis, tomato LeHXK4 (Kandel-Kfir et al., 2006), potato, M. truncatula and rice exemplifying that stromal HXKs are widely distributed among different species. A more   94 recently diverged clade (group 6 HXKs; R. Karve et al. (2010)), which contains sequences of dicot glucose sensor/transducer proteins such as the closely related AtHXK1 and AtHXK2 proteins. Interestingly, P. trichocarpa PtHXK1 and PtHXK2 are more closely related to protein sequences from other species than they are to each other. This is supported by the phylogenetic analysis of R. Karve et al. (2010) who found that PtHXK1 forms a clade with AtHXK1, AtHXK2, tobacco NtHXK1 and NtHXK2 and radish RcHXK1, while PtHXK2 forms a clade with radish RcHXK2 and tobacco NtHXK4, NtHXK5 and NtHXK6. This phylogenetic divergence may represent functional specialization as PtHXK2, along with PtHXK3, was found to be more highly expressed in the developing cambium of P. trichocarpa, whereas PtHXK1 was higher in source leaf (Figure 16). Group 7 contains only monocot sequences, which may be localized to the outer mitochondrial membrane or cytosol. Finally, groups 1 and 2 are most basal and contain sequences from the moss P. patens, and lycophyte Sellaginella moellendorffii, and are not presented in this phylogenetic analysis.   4.2.2 Effects of loss of AtHXK1, AtHXK2 and AtHXK3 functional enzymes in A. thaliana Due to the presence of AtHXK1 transcript in athxk1 A. thaliana TDNA insertional line (Figure 17, A and B) and its sensitivity to high glucose (Figure 18) one would not expect a dramatic phenotypic change compared to the wild-type plants. athxk1, however, has approximately half the rosette diameter of the wild type, athxk2 and athxk3 at 21 days after germination (DAG) (Figure 19). These results suggest that although AtHXK1 is transcribed, the amount of functional protein synthesized may not be enough to sustain normal metabolic activity. Some functional protein must be remnant as glucose signaling is not inhibited in these plants because athxk1 not allelic to the - AtHXK1 loss of function - gin2 mutation (Moore et al., 2003).  In plant cells, 3!   95 UTRs longer than 300 nucleotides induce mRNA instability and degradation by nonsense mediated decay, a mechanism that eliminates transcripts derived by aberrations in gene structure or expression (Schwartz et al., 2006). Additionally, there have been reports of other 3! UTR disrupted genes in A. thaliana such as histone H2A (RAT5) that exhibit a phenotype. RAT5 is unable to be transformed with Agrobacterium tumefaciens and contains two tandem 3! UTR TDNA insertions upstream of the polyadenylation signal (Mysore, Nam, & Gelvin, 2000). Although the exact position of the TDNA insertion within the athxk1 allele is not known, it can be assumed that less protein is produced, manifesting in a decreased rosette growth phenotype.    During the day, AtHXK1 likely functions as a signalling protein to fine-tune expression of photosynthesis genes according to the “excess” glucose released in the cytoplasm. At night however, AtHXK1 is required for the synthesis of G6P (Granot et al., 2013). In the source cells G6P is required for the maintenance of its own ATP levels though glycolysis, as well as for the synthesis of sucrose from vacuolar-released glucose and fructose. The sucrose translocated to sink cells during the day or night requires phosphorylation by AtHXK1 in order to be activated and enter glycolysis, cell wall synthesis or the OPPP in the plastid (Claeyssen & Rivoal, 2007) . AtHXK2 may be able to compensate for some of the phosphorylation function of AtHXK1 in the athxk1 mutant as the two are closely related (Figure 13) making the phenotype less severe than would be expected for decrease in such a major metabolic enzyme. Furthermore, AtHXK1 glucose signal transduction is required for early seedling growth (Granot et al., 2013) where AtHXK2 may not be able to compensate.  Later during adult development, AtHXK2 could be more successful in offsetting for a decrease in AtHXK1 function, which effectively explains the presence of a shorter stem height in athxk2 at senescence but the lack thereof in athxk1 (Figure   96 20, A & B). As mentioned earlier, however, there is a lack of precise spatio-temporal expression data of the HXK gene family during development making it difficult to assess how much, if any, this proposed compensation may be contributing. AtHXK2 has previously been implicated in stress response, having a 2-3 fold increase in transcript during cold, osmotic, and salt stress (Kreps et al., 2002). Sucrose is a known cryoprotectant in many species and an increase in the leaf of athxk1 (Figure 21, A) may be stimulating the expression of stress-related genes such as AtHXK2.   No increase in non-structural glucose and fructose was found in either source or sink tissue of athxk1 and athxk2 (Figure 21, B & C) and there were no changes in the starch content between these mutants and the wild-type plants. This is contrary to what was previously found in tobacco, where down regulation of NtHXK1, homologous to AtHXK1 and AtHXK2, resulted in a 46-fold increase in glucose, 22-fold increase in fructose and starch in source leaf tissue. Maltose concentration in the transgenics was found to be significantly higher, implicating NtHXK1 in the remobilization of starch for delivery to sink organs during the night (Y. KIM et al., 2013). In A. thaliana, an increase in structural glucose content was found to be present in the stem of athxk2 and athxk3 as well as xylose and mannose (Table 5). Xylose and mannose are both derived from the interconversions of glucose and are incorporated into the cell wall as hemicellulose polymers (Buchanan, Gruissem, & Jones, 2000). Xylose is derived from the oxidation of UDP-glucose to UDP-glucuronate by an unidentified UDP-glucose dehydrogenase in A. thaliana. UDP-glucuronate, which may also be derived from myo-inositol, is decarboxylated to UDP-xylose by an UDP-xylose synthase (UDP-glucuronate decarboxylase). Myo-inositol is itself derived from glucose-6-P by the action of inositol-1-phosphate synthase; found to be subcellularly localized   97 the cytoplasm (Donahue et al., 2010) followed by inositol-1-monophosphatase (Caspi et al., 2014). Finally, as previously mentioned, UDP-glucose is formed in the cell through phosphoglucomutase (PGM) that produces glucose-1-phosphate (G1P) and the phosphate is exchanged for a UDP on the 1! position by UDP-glucose-1-phosphate uridylyltransferase. Additionally, athxk1, athxk2 and athxk3 had a decrease in arabinose and galactose, and athxk2 was found to have lower rhamnose levels in the stem (Table 5). In the cell wall, galactose and arabinose are incorporated into arabinogalactan proteins (AGP), xylose in xylan, xyloglucan or pectic components and rhamnose is exclusively incorporated into the pectic polysaccharides rhamnogalacturonan I (RGA I) and rhamnogalacturonan II (RGA II) although all are, through a variety of pathways, ultimately derived from glucose (Buchanan et al., 2000). Therefore, while soluble glucose and fructose levels have not increased in all three athxk mutant lines, glucose and fructose, in the form of excess leaf sucrose, is incorporated into the cell wall as glucose, xylose and mannose, likely in the form of hemicellulose in athxk2 and athxk3, while pectin content in the cell wall might be lower due to decrease in arabinose and galactose in all three mutants (Table 5). Cellulose is likely unaffected as there was no change in the percent crystalline material of the stem in all three lines compared to the wild type (data not shown) and cellulose typically represents a major component of stem crystallinity (Fujita et al., 2011).  The production of phosphorylated glucose following the degradation of sucrose after import and, possibly further cycles or resynthesis is one of the initial steps of integration of photosynthate into the sink cell. Consequently, it is difficult to establish a clear hypothesis as to how a decrease or absence of different HXK isozymes, with their diverse localization and kinetics, result in a relocation of glucose from one major cell wall polymer (hemicellulose) to another (pectin). To   98 assert the finding that removal of HXK enzyme affects the cell wall lignin levels were found to be different in A. thaliana HXK mutant lines. Acid soluble lignin was lower in athxk1, athxk2 and athxk3, while acid insoluble lignin was increased in athxk3 compared to the wild-type (Table 6). In down regulation of p-Coumaroyl-CoA 3!-hydroxylase (C3!H) in poplar, which catalyzes the 3! hydroxylation of p-coumaroyl shikimate and p-coumaroyl quinate, a rate-limiting step in the synthesis of lignin polymer, it was found that a perturbation in lignin deposition resulted in an increase of glucose and xylose content (Coleman, Park, Nair, Chapple, & Mansfield, 2008) . Perturbations in other cell wall sugars, however, were not present and starch content, similar to athxk1 and athxk2, was not different from the wild type. It was proposed that in perturbing lignin content, sink strength was adjusted so that carbon pooled in the form of free sugars at the source (Coleman, Samuels, Guy, & Mansfield, 2008). Accordingly, sucrose levels were found to increase in the leaf of all three mutant lines as well as the stem of athxk3, suggesting a decrease in sink strength.   From the three manipulated HXK alleles, athxk3 was consistently found to have the most severe chemical phenotype. athxk3 was the only TDNA insertional line to have a sucrose increase in both the leaf and stem, as well as a significant increase in glucose and fructose in the non-structural carbohydrate portion of the leaf (Figure 21). Furthermore, its soluble lignin was decreased, as were the other two lines compared to the wild type, but its insoluble portion of lignin was slightly increased (Table 6). Thus, AtHXK3 seems to be an important enzyme for the maintenance of sink strength in A. thaliana as there was no growth phenotype in that line during vegetative development but stem height was decreased at senescence (Figure 19, Figure 20).   99 Similar, but more mild results were observed for athxk2, which could be attributed to the subcellular localization of the two isozymes; AtHXK3 in the plastid where it directly feeds several metabolic pathways, and AtHXK2 in the cytosol, whose function, in theory could be compensated by the closely related AtHXK1 still present in the athxk2 mutant line (Figure 17). Zhang et al. (2010) proposed a signalling role for AtHXK3, similar to AtHXK1, as an athxk3 mutant line was found insensitive to repression of photosynthesis genes at 7% glucose. This signalling role, even if valid, would likely not be present in sink tissue or would mediate a different type of gene repression other than photosynthetic.      100 Chapter 5: Conclusion  5.1 Proposed function of SPS enzyme  The function of SPS enzyme in source tissue has been speculated to be of two-fold: synthesize sucrose from photosynthetic components following their export into the cytoplasm and the resynthesis of sucrose following starch mobilization in the dark. Corroborating evidence from previous studies (Volkert et al., 2014), AtSPS1 and AtSPS2 indeed likely function to remobilize starch in the leaf due to presence of increased starch in the leaf of atsps1-1, atsps1-2 and atsps2 (Figure 12) as well as to synthesize sucrose from photosynthate, which is illustrated by pooled hexose monomers in leaf tissue (Figure 12). Expression of SPS in sink tissue has, however, raised questions as to the purpose of sucrose resynthesis where it would presumably be broken down for assimilation. In stem, SPS performs the same function as in leaf, i.e. synthesizes sucrose from apoplastic sucrose and unutilized glucose and fructose in the cytoplasm. If any starch is being broken down in sink organs at night, SPS may also function to synthesize sucrose from its remobilization, which could in this case be implied by the evidenced of the increase of starch in the stem of atsps1-2 (Figure 12). Additionally, AtSPS1 and AtSPS2 are required to resynthesize sucrose in the stem sink and the components arise from the breakdown of sucrose in the apoplast from apoplastic transport and cleavage and from the numerous sucrose breakdown cycles. These cycles are required in the cell to control carbon flux from the source so that the two can functionally coordinate and regulate each other’s gene expression through sucrose itself, TPS producing T6P and SnRKs signaling (Li et al., 2002; Rolland et al., 2006; Ruan, 2014). Furthermore, a more immediate consequence is that breakdown and resynthesis of sucrose controls intracellular metabolites like UDP-G, G1P, G6P, Pi and others, which are involved in a   101 variety of enzymatic reactions and also allosterically affect the plant metabolism. Removal of one SPS isoform likely disturbs the balance of those metabolites in a way that another isoform cannot fully compensate, possibly due to functional specialization. For example, AtSPS3 is highly sensitive to G6P and Pi but not AtSPS1, AtSPS2 and AtSPS4 (Volkert et al., 2014). This imbalance can then change the carbon flux for a variety of reactions resulting in compositional changes of cell wall carbohydrates and possibly lignin (Table 3, Table 4).  In order to tease out the particularities of SPS enzymes in the sink, I propose the following future experiments. Utilizing double and triple mutants of SPS isozymes for non-structural and structural carbohydrate determination will enable a better understanding if there is any compensation by SPS isozymes and any functional specialization. Additionally, concomitant down regulation of the various SPS enzymes in hybrid poplar will shed light on the enzymes’ function. The lack of a strong developmental or chemical phenotype in the down regulation of SPS can partially be explained by redundancy in the gene family. It would be useful to determine the enzyme kinetics of P. trichocarpa SPS to understand if they are regulated in a manner similar to A. thaliana, maize, spinach and other enzymes examined. Finally, metabolite profiling of these mutants, single, double or more, would be able to elucidate where carbon flux is redirected to, in order to understand changes in cell wall carbohydrate and lignin. Reciprocal grafting experiments in other model organisms such as tomato using wild-type shoots and down-regulated stems could be investigated to determine if carbohydrate pooling is occurring in the stem when the leaf is fully functional.    102 5.2 Proposed function of HXK enzyme  Hexokinase’s role was previously thought to be required only in the sink organs of plants where sucrose is broken down to glucose and fructose and those hexoses need to be phosphorylated for further metabolism. In the last twenty years a clearer role for the presence of HXK enzyme in source cells has been established where the enzyme takes on signalling roles and phosphorylates free hexoses produced in the cytoplasm. HXK plays a central role in the sink metabolism where it phosphorylates sugars following the breakdown of photosynthate; sucrose, RFOs or other polyols, which are transported in some species. This metabolic centrality, coupled with the possibility of HXK-related glucose signalling in the sink as well as the source allow HXKs to be responsible for changing the landscape for a variety of plant processes. Firstly, HXK likely plays a role in modulating sink strength by removal of glucose to G6P in the system. This would allow for more sucrose to be broken down and therefore allow for bulk flow and is illustrated by the build-up of sucrose in the leaf of HXK mutant lines (Figure 21). This effect however, may not be as strong as other enzymes controlling carbon flux such as SPS, SuSy and INV which are sucrose cleaving even though HXK is considered the rate limiting step in the cycling of sucrose (Nägele & Weckwerth, 2014). Second, HXK activity is an entranceway for glucose into various metabolic pathways in the cytosol and the plastid. This is evidenced by the presence of an early and late growth phenotype, a change in non-structural and structural carbohydrate content and a decrease of insoluble lignin. This broad effect is probably due to a change in metabolite pool since in an in silico HXK model overexpression, it was found that T6P and sucrose synthesis were affected more than twice the wild type when external sucrose was applied to the system (Nägele & Weckwerth, 2014) . The rate of reaction of SPS increased 7.42 fold, FRK 2.92 fold, UDP-G levels increased 5.83 fold and T6P concentration increased 2.14 fold compared to the   103 wild-type Arabidopsis. Therefore, the authors suggest that the main change associated with increasing HXK activity in the T6P/SnRK1/SPS interaction pathway is the concentration of UDP-G (Nägele & Weckwerth, 2014) . UDP-G is central in many carbohydrate interconversion reactions, sucrose resynthesis by SPS and cell wall synthesis, and a change in this metabolite could account for the change in structural carbohydrate composition and soluble lignin of athxk mutants (Table 5, Table 6).   To determine the functional role of HXK enzymes in A. thaliana and P. trichocarpa, I propose the following future experiments. Foremost, in Arabidopsis, double and triple mutants of HXK enzymes specifically expressed in a sink or source tissue followed by extensive carbohydrate and metabolite profiling should further the scientific community’s understanding of 1) how HXKs affect source and sink tissue, 2) what is the communication pathway between major sucrose enzymes like SPS and HXK and cell growth regulators such as SnRK1, and 3) compensatory functions inherent in the HXK proteins. Metabolite profiling is a difficult task; however, given the strong effects associated with changing HXK expression and its centrality to cell metabolism, it will have a high scientific return. Spatio-temporal gene expression, perhaps in the form of promoter-GUS assay throughout plant development and under differential stresses could provide further understanding of HXK isozymes. This could address, if AtHXK2 is indeed a stress related protein as shown in transcriptional profiling (Kreps et al., 2002). Do AtHXK2 and AtHXK3 compensate for the function of AtHXK1 in the sink later in development? It would be interesting to know if all HXKs share the signalling ability of AtHXK1 and AtHXK2 and what is the difference in signalling, if any, between source and sink tissue. This could be achieved with a ChIP-on-chip assay under ambient and exogenous biotic and abiotic stresses. Finally, it would be   104 interesting to know what is the functional reason for an increase in some structural carbohydrates and a decrease in others in the same or similar fashion in all three AtHXK3 and if this change is responsible for a 3-D, structural change of the cell wall matrix, inducing a change in lignin.  5.3 An integrated view of sucrose metabolism The cycle of sucrose degradation begins with SuSy catalyzing its breakdown to UDP-G and fructose (Figure 23). Fructose, also derived from degradation by INV, is phosphorylated by FRK to F6P and may act as the precursor to form S6P with an UDP-G via SPS or may be isomerized by PGI to G6P. G6P can also be derived from phosphorylation by HXK, via glucose derived from the action of INV or the vacuole. G6P is isomerized by PGM to G1P and then has its phosphate exchanged for a UDP nucleotide by UGPase. This marks a second UDP-G input, derived from carbon other than SuSy sucrose breakdown. A small amount of G6P combines with UDP-G to form T6P, which is metabolized to Trehalose by TPP. The signalling aspects of these catabolic and anabolic cycles involve SnRK1, an inhibitor of plant growth. SnRK1 is inhibited by UDP-G and more potently by T6P, G6P and G1P. One of SnRK1’s targets is SPS, therefore, decreasing SnRK1 activity increases sucrose cycling by releasing inhibition of SPS. Additionally, INV is feedback inhibited by its products, glucose and fructose, as well as FRK by F6P and HXK by G6P and F6P (Nägele & Weckwerth, 2014). SuSy reverses its typical catabolic action to synthesis when it is feedback inhibited by its products, UGP-G and fructose.        105  Figure 23: An integrated view of sink cell carbohydrate metabolism. SPS: Sucrose Phosphate Synthase, SPP: Sucrose Phosphate Phosphatase, SuSy: Sucrose Synthase, FRK: Fructokinase, TPS: Trehalose Phosphate Synthase, HXK1/2: Hexokinase 1/2, HXK3: Hexokinase 3, PGI: Phosphoglucoisomerase, PGM: Phosphoglucomutase, UGPase: UDP-Glucose Pyrophosphorylase, SnRK1: SNF1-related kinase, aINV: acid Invertase, cwINV: cell wall Invertase.  Based on these observations, when SPS activity is decreased, there is a build-up of UGP-G and F6K, which would reserve the SuSy reaction and feedback inhibit FRK. Fructose and glucose would build-up as they are underutilized, as would be the accumulating sucrose. The pooling of sucrose will decrease the rate of mass flow from the phloem and some of the excess photosynthate will be transported to the phloem for storage. A higher availability of UDP-glucose will result in the production of more T6P that will inhibit SnRK1 leaving other remaining SPS isoforms, active. HXK may also be repressed due to F6P conversion to G6P, resulting again in more glucose pooling.    106 When HXK is repressed in the same system, a small build-up of glucose may inhibit INV, leading to sucrose pooling. Less carbon will enter the pathway, since UDP-G can now only be produced from the breakdown of sucrose via SuSy. In this case, SnRK1 could be inhibited to a smaller extent because of the lower concentrations of available G6P and G1P, as well as lower T6P production and that will result in decreased SnRK1 activity (Vsps) due to phosphorylation. Therefore, SnRK1 will inhibit plant growth and UDP-G will be used at a lower rate, possibly allowing it to enter other UDP-hexose interconversion pathways and lead to cell wall synthesis.     107 References:   Aki, T., Konishi, M., Kikuchi, T., Fujimori, T., Yoneyama, T., & Yanagisawa, S. (2007). Distinct modulations of the hexokinase1-mediated glucose response and hexokinase1-independent processes by HYS1/CPR5 in arabidopsis. 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