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Modification of cellulose biosynthesis through varied expression of sucrose metabolism genes in tobacco… Coleman, Heather Dawn 2008

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MODIFICATION OF CELLULOSE BIOSYNTHESIS THROUGH VARIED EXPRESSION OF SUCROSE METABOLISM GENES IN TOBACCO AND HYBRID POPLAR by  HEATHER DAWN COLEMAN B. S.F., University of British Columbia, 2002  A THESIS SUBMITTED TN PARTIAL FULFILMENT OF THE REQUIREMENTS FOR THE DEGREE OF  DOCTOR OF PHILOSOPHY  in  THE FACULTY OF GRADUATE STUDIES (Forestry)  THE UNIVERSITY OF BRITISH COLUMBIA June 2008  © Heather Dawn Coleman, 2008  Abstract UDP-glucose, the precursor for cellulose biosynthesis, can be produced via the catalysis of sucrose by sucrose synthase (SuSy) or through the phosphorylation of glucose-I-phosphate by UDP-glucose pyrophosphorylase (UGPase). As such, these genes, together with sucrose phosphate synthase (SPS) which recycles fructose (an inhibitor of SuSy), are interesting targets for altering carbon allocation in plants. In an attempt to alter cell wall biosynthesis in plants, targeted overexpression of SuSy, UGPase and SPS independently and in a pyramiding strategy was assessed in tobacco. All lines displayed enhanced growth and biomass production, and in the case of double and triple transgenics, there was an additive effect. Despite the increased growth rates, there was no consistent change in soluble carbohydrate pools. Furthermore, only the triple transgenics had constant changes in structural carbohydrates: with increased hemicellulose content and slight increases in cellulose. Collectively, these results support the role of SPS, SuSy and UGPase in maintaining sink strength, but suggest that the reallocation of carbon to cellulose production in tobacco may not be possible by overexpressing these genes. In contrast, transgenic poplar overexpressing UGPase produced significantly more cellulose than wild-type trees.  However, this was accompanied by a severe  reduction in growth and the production of a salicylic acid glucoside (SAG) in significant quantities.  The UDP-glucose generated by UGPase overexpression appeared to  participate in both the synthesis of cellulose and SAG, suggesting that cellulose biosynthesis may be limited by the cellulose synthase complex. Poplar transformed with SuSy and with SuSy x UGPase also had increased cellulose production. The trees were phenotypically normal, with only minor reductions in height growth in some lines. It appears that UDP-glucose may be channelled directly to the cellulose synthase complex by SuSy. The increased cellulose content was associated with an increase in cell wall crystallinity, but there was no change in microfibril angle, confirming the re-allocation to cellulose synthesis was not the result of tension wood formation, again supporting the hypothesis that the cellulose synthase complex is the limiting factor.  Clearly, it is possible to alter cellulose deposition in trees by augmenting sucrose metabolism to produce UDP-glucose, the precursor to cellulose biosynthesis.  III  Table of Contents Abstract  .  ii  Table of Contents  iv  List of Tables  vi  List of Figures  viii  Acknowledgements  x  Co-authorship Statement  xi  Introduction  I  Background  I  Cellulose Biosynthesis  1  Sucrose Metabolism  3  Promoters  20  Objectives  21  References  23  Upregulation of Sucrose Synthase and UDP-Glucose Pyrophosphorylase Impacts Plant 31 Growth and Metabolism Introduction  31  Methods  33  Results  38  Discussion  53  Acknowledgements  57  References  58  Altered Sucrose Metabolism Impacts Biomass Production and Flower Development in 61 Tobacco Introduction  61  Methods  63  Results  66  Discussion  80  Acknowledgements  86  References  87  Overexpression of UDP-Glucose Pyrophosphorylase in Hybrid Poplar Affects Carbon Allocation 91 Introduction  91 iv  Methods  .93  Results  99  Discussion  113  Supplemental Data  119  Acknowledgements  119  References  120  Overexpression of Sucrose Synthase and UDP-Glucose Pyrophosphorylase in Hybrid 124 Poplar Affects Cellulose Partitioning and Ultrastructure Introduction  124  Methods  126  Results  132  Discussion  140  Acknowledgements  147  References  148  Conclusions and Recommendations for Future Work Future Work Recommendations Appendix A 3-glucuronidase (GUS) Transformation and Staining -  153 155 156  Methods  156  Results  157  References  160  Appendix B  —  Supplemental Data Chapter 3  161  V  List of Tables Table 1.1. Enzymatic reactions catalyzed by UGPase, SuSy, SPS and SPP  5  Table 2.1. Transcript level and enzyme activity in leaf and stem tissue for single transgene and non-transformed control plants  41  Table 2.2. Concentration of total soluble carbohydrates (jig mg 1 dry weight tissue) in stem tissue of single transgene and non-transformed control tobacco plants  43  Table 2.3. Transcript level and enzyme activity of UGPase and SuSy in leaf and stem issue of double transgene and non-transformed control tobacco plants  48  Table 2.4. Concentration of total soluble carbohydrates (jig mg 1 dry weight tissue) in stem tissue of double transgene and non-transformed control tobacco plants  50  Table 3.1. Transcript level and enzyme activity in leaf and stem of transgenic and wildtype tobacco lines  67  Table 3.2. Soluble carbohydrate and starch content (mg g ) in leaf and stem of 1 transgenic and wild-type tobacco lines Table 3.3. Time to flowering of transgenic and wild-type tobacco lines  71  78  Table 3.4. Chemical composition (% dry weight) of cell wall of stem of UGPase x SuSy x  SPS triple transgenic lines  81  Table 4.1. Transcript level and enzyme activity in leaf tissue and developing xylem for transgenic and wild-type trees  103  Table 4.2. Total soluble carbohydrates (mg g ) in leaves and developing xylem of 1 transgenic and wild-type trees  106  Table 4.3. Starch content (mg g’) in leaves and developing xylem of transgenic and wild-type trees  107  Table 4.4. Chemical composition of stem material of transgenic and wild-type trees.. 108 Table 4.5. Syringyl, guaiacyl and p-hydroxyphenyl monomer contents (%) of transgenic and wild-type trees as determined by thioacidolysis  111  Table 4.6. Metaboiltes identified in the developing xylem of 2x35S::UGPase hybrid poplar relative to levels in wild-type trees  112  Table 5.1. Mean transcript abundance and enzyme activity in leaf and developing xylem tissue for transgenic and wild-type poplar trees  135 vi  Table 5.2. Total soluble carbohydrates and starch in leaf and developing xylem tissue of transgenic and wild-type trees  136  Table 5.3. Chemical composition of stem segments (P1=5 to P1=15) of transgenic and wild-type trees  138  Table 5.4. a-cellulose content, cell wall crystallinity and microfibril angle (MFA) of stem segments (P1=5 to P1=1 5) of transgenic and wild-type trees  139  Table BI. Transcript abundance of the native poplar UDP-glucose pyrophosphorylase genes in leaf and developing xylem tissue for transgenic and wildtype trees  161  Table B2. Transcript abundance of the cell wall biosynthetic genes involved in lignin and cellulose deposition in the developing xylem tissue for transgenic and wild-type trees  162  Table B3. Fold changes in salicylic acid 2-Q-f3-glucoside in the developing xylem of all transgenic 2x35S::UGPase hybrid poplar relative to levels in wild-type trees  163  vii  List of Figures  Figure 1.1. Schematic depicting sucrose metabolism from source to sink Figure 2.1. Plant height (A), internode length (P1  =  5 to P1  =  4  15) (B) and total plant  biomass (C) in single transgene plants  40  Figure 2.2. Starch (A) and cellulose (B) content, and fibre length (C) and coarseness (D) of single transgene lines and non-transformed control line Figure 2.3. Plant height (A), internode length (P1  =  5 to P1  =  45  15) (B) and total plant  biomass (C) in double transgene tobacco lines  46  Figure 2.4. Starch (A) and cellulose (B) content, and fibre length (C) and coarseness (D) of double transgene lines and non-transformed control line  51  Figure 25. Light and UV-fluorescence microscopy of control (A) and transgenic (4CL::UGPasexSuSy 12) (B) tobacco  52  Figure 3.1. Biomass measurements of UGPase x SPS transgenic lines compared to wild-type tobacco plants: height (A), calliper (B), and leaf dry weight (C)  74  Figure 3.2. Biomass measurements of SuSy x SPS transgenic lines compared to wildtype tobacco plants: height (A), calliper (B) and leaf dry weight (C) Figure 3.3.  75  Biomass measurements of UGPase x SuSy x SPS transgenic lines  compared to wild-type tobacco plants: height (A), calliper (B) and leaf dry weight (C)  76  Figure 3.4. Morphological alterations of 4CL::UGPase x SuSy x SPS (A&C&E) plants relative to wild-type tobacco plants (B&D&F)  79  Figure 4.1. Plant height (A), diameter (B), leaf area (C) and internode length (D); for transgenic and wild-type trees  100  Figure 4.2. Image depicts axial shoot elongation and leaf size in a representative 2x35S::UGPase (right) and wild-type (left) poplar  101  Figure 4.3. Schematic representation of the effect of overexpression of 2x35S::UGPase on the transcript abundance of the lignin biosynthetic genes in the developing xylem  104  Figure 4.4. cx-cellulose content of stem material from transgenic and wild-type trees. 110  vi”  Figure 5.1. Biomass measurements of transgenic and wild-type poplar. Tree height (A), calliper (B), leaf dry weight (C) and stem dry weight (D)  133  Figure 5.2. Calcofluor (A, B) and phioroglucinol (C, D) staining of wild-type (A, C) and  2x35S::SuSy transformed (B, D) poplar trees  141  Figure Al. 3-glucuronidase (GUS) staining in tobacco  158  Figure A2. -glucuronidase (GUS) staining in poplar  159  ix  Acknowledgements I would like to start by thanking my supervisor, Shawn Mansfield, for countless hours of work and countless laughs. I am thankful for your support and for having had the opportunity to work with you. I also want to thank my supervisory committee, David Ellis, John Kadla and Lacey Samuels.  Thank you to each of you for your insight and probing questions.  Thank you also for your support and encouragement throughout this process. My thanks goes to present and past post-docs and fellow students in the Mansfield lab for both scientific assistance and emotional support. In particular I would like to thank Thomas Canam for always going above and beyond with any question or request for help. Thanks to the summer students  —  Leigh Beamish, Anya Reid, and Jimmy Yan for  your fantastic help, for being awesome people to work with and for each adding a happy memory to my PhD experience. To the volunteers and exchange students Margaret Kearney, Aaron Fung, and Tony Einfeldt for their assistance with some of the mundane. I would also like to thank Minako Kaneda for help with all things staining and microscopy. Thank you to my family for pretending to read my papers and for teaching me to work hard. Thank you especially to my parents for supporting me in my endeavours. To my friends, thank you for allowing me to vent, celebrating the good, and encouraging me to keep going. Thanks especially to Lisa time, to Jackie  -  -  you arrived in the nick of  we really should do this more often, and to Sharon  -  for always being  ready for a coffee break. And finally thank you to Gabe. Thank you for being a sympathetic ear and an unwavering support.  x  Co-authorship Statement This thesis contains four manuscripts written with the intent of publication in peer-reviewed journals.  Chapters 2 and 4 comprise manuscripts that have been  published. Chapters 3 and 5 consist of manuscripts that will be submitted for publication. Author contributions to the work are as follows: Chapter 2: Heather Coleman helped design the research, performed the research, conducted all data analyses, and prepared the manuscript. Margarita Gilbert assisted in the design of the research. David Ellis helped identify the research opportunity, assisted in design of research and edited the manuscript. Shawn Mansfield contributed to research opportunity identification and research design, edited the manuscript, and supervised the work. Chapter 3: Heather Coleman helped design the research and performed the research, conducted all data analyses, and prepared the manuscript. Leigh Beamish and Anya Reid assisted in performing research. Ji-Young Park assisted in the design of the research. Shawn Mansfield contributed to research opportunity identification and research design, edited the manuscript, and supervised the work. Chapter 4: Heather Coleman helped design the research and performed the research, conducted all data analyses, and prepared the manuscript. Thomas Canam and Kyu-Young Kang assisted in performing the research. David Ellis helped identify the research opportunity and edited the manuscript. Shawn Mansfield contributed to research opportunity identification and research design, edited the manuscript, and supervised the work. Chapter 5: Heather Coleman helped design the research and performed the research and conducted all data analyses, and prepared the manuscript. Jimmy Yan assisted in performing the research.  Shawn Mansfield contributed to research  opportunity identification and research design, edited the manuscript, and supervised the work.  xi  Introduction Background Cellulose is the most abundant organic polymer in the world and is produced by plants at an approximate rate of 1011 tons per year (Preston, 1974). The majority is deposited in the stems of woody plants and is commonly utilized for fuel, timber, forage, fibre and chemical cellulose. It is for this reason that carbon partitioning to cellulose production is a key question for researchers globally (Haigler et al., 2001). The biosynthesis of cellulose utilizes the products of photosynthesis, and competes for these resources with a number of other pathways including starch deposition and the non-cellulosic components of cell walls. Carbon sequestration and the intrinsic value of the forest are becoming more important to the global community resulting in the increased preservation of forested land. Concurrently, the global population is growing and the demand for wood fibre is increasing. As such, there is a trend towards more intensive management and shorter crop rotations. While this shift will result in an increase in the amount of fibre supplied, the rapid growth of these trees will result in decreased wood quality. The value of harvestable trees is directly related to the amount and quality of the cellulose fibres. The availability of carbohydrates from photosynthesis is not a limiting factor in cellulose biosynthesis despite having a crucial role in plant growth and survival. Therefore, by increasing the utilization of carbohydrates in cellulose production, more cellulose should be produced, and consequently fibre yield would increase. Sucrose and other translocatable carbohydrates produced during photosynthesis are transferred to sink tissues where they are converted to starch or cellulose.  The creation of a  carbohydrate sink in cellulose-producing cells should allow a greater portion of the photosynthates to be diverted to these cells, and therefore to cellulose production.  Cellulose Biosynthesis Cellulose is the structural backbone of plant cell walls and plays a key role in cell shape and the morphology of plants.  It is a linear polymer comprised of D-glucose I  residues linked by f3-l,4 glycosidic bonds such that every glucose is rotated by approximately 180. The structural repeating unit is cellobiose, in contrast to many other glucan polymers where the repeating unit is glucose (Brown et a!., 1996). Because of its unique structure, cellulose chains can interact with each other, and assemble into microfibrils containing an estimated 36 strands (Reiter, 2002). These strands are aligned in a parallel fashion in cellulose I, the most prevalent natural cellulose type (Reiter, 2002). or bundles.  Microfibrils are often further associated into macrofibrils  This strand association is so precise that microfibrillar cellulose is  essentially crystalline (Delmer and Amor, 1995). The formation of cellulose into microfibrils is associated with the structure of the cellulose synthase complex, or rosette terminal complex. The rosette is an organized enzyme complex, hexagonal in structure, which produces cellulose microfibrils. It is believed to be composed of 6 units that contain a number of cellulose synthase subunits (possibly 6) that each produces a glucan chain (Delmer, 1999). The resultant chain from each unit associates with adjacent chains of the same rosette to form a microfibril.  The cellulose synthase complex is large, with distinct transmembrane  helices and a cytosolic component that is expected to be the location of the active site involving UDP-glucose (Brown and Saxena, 2000; Delmer, 1999). Although the cellulose synthase complex has been identified visually, identifying the number and association of the cellulose synthase genes has been a slower process. The first cellulose synthase gene was cloned and isolated from Acetobacter xyllnum, a bacterium that secretes large quantities of cellulose in long microfibrils (Saxena et a!., 1990). However, an additional 6 years passed before the first identification and isolation of a cellulose synthase gene in plants. The first identified were 2 genes highly expressed during the phase of cellulose synthesis in cotton fibre development (Pear et a!., 1996). Since then, cellulose synthases have been identified in many plants and all have been shown to share the D,D,D,QxxRW signature (Williamson et a!., 2002). The sequencing of the Arabidopsis genome has identified 10 cellulose synthase (CesA) genes and an additional 6 groups of cellulose synthase like (CSL) genes (Richmond, 2000).  The more recently sequenced poplar genome has  identified 18 predicted CesA genes (Djerbi et al., 2005), which are thought to be 9 types of duplicated genes. 2  Sucrose Metabolism Cellulose is produced from the precursor UDP-glucose which can be produced by a number of pathways (Figure 1.1; Table 1.1).  Sucrose synthase (SuSy; EC  2.4.1.13) catalyzes the reaction of sucrose and UDP to UDP-glucose and fructose. The reaction in plants is reversible, and SuSy has been shown to act in the degradation of sucrose. This reaction provides energy for phloem unloading by providing substrate for respiration (Hanggi and Fleming, 2001), and cleavage by SuSy has also been positively correlated with sink strength in storage organs of potatoes, maize kernels, and pea embryos (Sun eta!., 1992; Zrennereta!., 1995; Dejardin etal., 1999). Two types of SuSy have been proposed, one of which is found in the cytosol and one that is directly associated with the plasmalemma (Amor et a!., 1995; Carlson and Chourey, 1996). Soluble SuSy (S-SuSy) exists in high levels in the cytoplasm of nonphotosynthetic tissues where its products are used in general metabolism and for the synthesis of storage polymers such as starch (Haigler et a!., 2001). Particulate SuSy (P-SuSy) is associated with the plasma membrane or cortical cytoskeleton.  It is  believed to play a major role in providing UDP-glucose to the cellulose synthase complex, in particular during high rate secondary wall cellulose synthesis (Haigler et al., 2001). There is evidence that the synthesis of the secondary cell wall may not rely on the pool of UDP-glucose, but may actually have access to UDP-glucose channelled directly from particulate SuSy in the cell cortex (Amor eta!., 1995). This would allow for the conservation of energy normally required for hydrolysis, as well as ensure the availability of UDP-glucose to the cellulose synthase complex despite the demand from other pathways. The second putative UDP-glucose pathway involves a number of enzymes with the final reaction being catalyzed by UDP-glucose pyrophosphorylase (UGPase; EC 2.7.7.9), which phosphorylates glucose-I-phosphate (Table 1.1).  This reaction is  magnesium dependent and is regulated by pyrophosphate concentrations. It has been suggested that the direction of the reaction is likely regulated by the cytosolic availability of its substrates (Nakano et a!., 1989; Sowokinos et a!., 1993).  In source tissues,  UGPase works in conjunction with SPS in the synthesis of sucrose (Kleczkowski, 1994), while in sink tissues, where it is predominantly found, it is a key enzyme in carbohydrate 3  • Sucrose Phosphate Syrithase V Sucrose Phosphate Phosphatase I Soluble SuSy 3 Phosphoglucornutase  • • • •  Glucose-B-phosphate isomerase Fructose-i ,6-bisphosphalase Phosphofructomutase Fructokinase  4  Cytosolic A Hexokinase + UDP Glucose Pyrophosphorylase • Cellulose Synthase  O  Particulate SuSy Vacuolar Invertase Apoplastic Invertase • ADP-Glucose Pyrophosphorylase  Figure 1.1. Schematic depicting sucrose metabolism from source (a)to sink (b).  4  Table 1.1. Enzymatic reactions catalyzed by UGPase, SuSy, SPS and SPP. Gene  Reaction  UGPase SuSy SPS SPP  UTP + a-D-glucose 1-phosphate -* diphosphate UDP-glucose + D-fructose *-÷ UDP + sucrose  +  UDP-glucose  UDP-glucose + D-fructose 6-phosphate UDP + sucrose 6-phosphate sucrose 6-phosphate + H 0 —k sucrose + phosphate 2 —,  5  biosynthesis.  UGPase is thought to work in coordination with SuSy in the cycling  between the hexose phosphate pool and sucrose. Glucose-I-phosphate, along with glucose-6-phosphate and fructose-6-phosphate comprise the hexose phosphate pool. Phosphoglucomutase (PGM; EC 5.4.2.2), which reversibly converts glucose-I-phosphate to glucose-6-phosphate, and phosphoglucose isomerase (PGI; EC 5.3.1.9), which reversibly catalyses the reaction between glucose6-phosphate and fructose-6-phosphate, maintain the equilibrium of the pool. Fructose 1,6-bisphosphate is an additional hexose phosphate, but is not normally considered to be a part of the hexose phosphate pool since in most organisms its conversion to fructose-6-phosphate by fructose-I,6-bisphosphatase (FPBase; irreversible.  EC 3.1.3.11) is  In plants however, there is an additional enzyme, phosphofructokinase  (PFKase; 2.7.1.11), that catalyzes the pyrophosphate dependant reversible reaction between fructose-I ,6-bisphosphate and fructose-6-phosphate, which is utilized in the production of sucrose. An additional key enzyme in sucrose metabolism is sucrose phosphate synthase (SPS; EC 2.4.1.14). SPS works in coordination with sucrose phosphate phosphatase (SPP; EC 3.1.3.24) to synthesize sucrose from fructose-6-phosphate and UDP-glucose. This reaction is believed to play a large role in the recycling of fructose, which is a known inherent inhibitor of SuSy (Doehlert, 1987) and releases phosphate, which can then be recycled through triose-phosphate transporters into the chloroplast for subsequent photosynthetic reactions. As such, SPP has been implicated as having a substantial role in the regulation of the photosynthetic rate (Stitt et a!., 1987). The role of SPS in the synthesis of sucrose will also affect photosynthesis rate, as sucrose and other soluble sugars have a role in the regulation of photosynthetic gene expression (Koch, 1996). It has been shown that cellulose biosynthesis is reliant on sucrose concentration rather than that of UDP-glucose, its immediate substrate.  As such, invertase (EC  3.2.1.26) is another enzyme that may have a regulatory impact on the synthesis of cellulose. lnvertase catalyzes the hydrolysis of sucrose into glucose and fwctose. It is present in higher concentrations than the other sucrose metabolizing enzymes, and may play an important role in the hydrolysis of sucrose at all developmental stages (Canam et a!., 2008).  Invertase works in coordination with SuSy and SPS in “futile 6  cycles” which allow for more efficient storage of sucrose in the vacuole and intercellular space (Nguyen-Quoc and Foyer, 2001).  Sucrose Synthase Herbaceous SDecies SuSy has been identified as having a key role in the metabolism of sucrose and as such has been employed in transgenic and mutant studies in numerous species including carrot (Tang and Sturm, 1999), maize (Carlson et a!., 2002; Chourey et a!., 1998), potato (Zrenner et a!., 1995; Bologa eta!., 2003), cotton (Ruan eta!., 2003), and tomato (Chengappa eta!., 1999; D’Aoust eta!., 1999). The effect of reductions in SuSy has varying phenotypic effects on plants. In potato, the antisense suppression of SuSy resulted in no obvious phenotypic differences, but the tubers showed a pronounced decrease in dry weight. This was attributed to the enhanced levels of free glucose and fructose causing an increase in water uptake (Zrenner et a!., 1995).  In carrot, both the tap roots and above ground  portion of antisense plants were much smaller than those of the corresponding controls. In most lines the leaf-to-root ratio was unchanged, but in the line with the largest reduction in SuSy expression, there was a shift in the leaf-to-root ratio in favour of the leaves (Tang and Sturm, 1999).  The downregulation of SuSy resulted in the  accumulation of sucrose levels ranging from two- to four-fold in carrot tissue, while levels of fructose and glucose were correspondingly reduced to below 20% that of control levels. Both starch and cellulose were reduced to about 38% and 37% that of the control respectively, and UDP-glucose was reduced by up to 70% (Tang and Sturm, 1999). The maize endosperm mutant, shrunken-I, had shrunken seeds with collapsed endosperms, and showed a decrease in sucrose utilization along with increased glucose and fructose contents (Chourey eta!., 1998). In tomato, the dry weight was not affected, but there was a decrease in the number of flowers that set fruit in plants where SuSy activity was decreased to less than 10% that of the activity in the control plants. This suggests that SuSy, by controlling unloading capacity, determines the capability to set fruit (D’Aoust et a!., 1999).  In cotton, the suppression of SuSy inhibited fibre  initiation and elongation. When SuSy was decreased in the ovule epidermis by 70% or 7  more, the result was a fibreless phenotype. One line was also dwarfed and aborted all fruit by 10 days after anthesis (Ruan et aL, 2003). The transgenic fruit contained just as many seeds as the control fruit, but only 5.7% of these seeds were considered normal sized with approximately 70% reduction of fibre mass.  The remaining seeds were  unviable or stunted (Ruan eta!., 2003). These studies provide insight into the role of SuSy in sink tissues.  In some  species and tissues, such as potato and maize endosperm, SuSy is involved mainly in the synthesis and storage of starch. Tang & Sturm (1999) surmised that SuSy is a major determinant of growth in carrot, and appears to be the major enzyme in sucrose cleavage supplying carbohydrates to metabolism. Plant size was greatly reduced as the downregulation of SuSy caused a decrease in the availability of carbon (Tang and Sturm, 1999). Furthermore, the maize and potato tuber studies suggest a role for SuSy in determining sink strength (Zrenner eta!., 1995). This is supported by the decrease in starch accumulation, which parallels the reduction in SuSy activity. The shrunken-I maize mutant showed a 22% reduction in starch content relative to the control. More recently, this has been shown to be a secondary effect, with the real effect of the mutation being cell degeneration during the cell elongation phase (Chourey et a!., 1998). A supporting study illustrated a 99% reduction in SuSy activity resulted in 46% starch reduction. It was concluded that the two SuSy genes that had been identified in maize at the time of this study have different roles: one provides substrate for cellulose biosynthesis, while the other provides precursors for starch biosynthesis (Chourey eta!., 1998). The study by Ruan et al. (2003) in cotton demonstrated a rate-limiting role for SuSy in the initiation and elongation of cotton fibre cells; SuSy can be used as a determinant for the sink strength of fibre cell development (Ruan et al., 2003). This study also showed that the suppression of SuSy in the seed coat reduced fibre length, but if SuSy was also suppressed in the endosperm and embryo, the seed did not develop at all.  This supports the role of SuSy in controlling plant cell and seed  development (Ruan et a!., 2003). Two concurrent studies employing antisense SuSy in tomato plants report the same findings with regards to sugars, starch and cellulose (Chengappa et a!., 1999; D’Aoust et a!., 1999). D’Aoust et a!. (1999) found that fruit growing from flowers that 8  developed in the first week of flowering were smaller than those of the control from the same time period. Fruit set also decreased in the transgenic lines. The conclusion is that SuSy inhibition leads to a reduction in unloading capacity in the young fruit (D’Aoust et a!., 1999). Vacuolar invertase is the main sucrose metabolic pathway when SuSy is absent or inhibited. When the tomato fruit is young, it has small vacuoles thus when SuSy is reduced the fruit will grow more slowly. As the fruit from the transgenic plant gets larger and the vacuoles expand it will begin to grow as quickly as the fruit from control plants because there is an increase in invertase activity. This suggests that SuSy controls phloem loading into the initial fruits and that this unloading consequently determines the growth rate of the remaining growth phase (D’Aoust eta!., 1999).  Woody Species In comparison to the numerous studies in herbaceous species, relatively little work has been carried out in woody species. A few studies of native gene expression (cited below) have been done, but to our knowledge, only one study has involved the use of transgenics (Konishi eta!., 2004). Egger & Hampp (1993) studied the development of spruce needles throughout the transition from sink to source with respect to SPS, SuSy, and invertase. They found that SuSy was dominant in the very young needles when they are strong sinks, where it acts in coordination with invertase to import carbon in the form of sucrose. As the leaves mature, and gain photoassimilatory competence, the activity of both enzymes decreases and SPS begins to increase. A similar pattern was shown in maize leaves (Nguyen-Quoc et a!., 1990).  In spruce needles, SuSy activity increased steeply in  autumn and winter, along with that of SPS, and both enzymes are at their highest rates from October to February.  This is similar to the trend observed in poplar stems  (Schrader and Sauter, 2002), and is thought to be related to frost hardiness and the accumulation of osmotically active compounds (Guy, 1990). In late winter, the needles are filled with starch and high SPS and SuSy activities could be related to the mobilization of chloroplast starch for bud support, in addition to glycolytic turnover (Egger and Hampp, 1993).  In poplar, SuSy shows particularly high activities in  December and January when most of the starch has been converted in sucrose. The 9  activity of SuSy peaks shortly after SPS and is expected to work in coordination with SPS to maintain a sugar cycle where sucrose is synthesized by SPS and degraded by SuSy (Schrader and Sauter, 2002). This cycle is thought to regulate sucrose levels in response to changing temperatures. Hauch and Magel (1999) examined the sucrose metabolism enzymes in Robinia trunk tissues throughout the year. In April and May, SuSy activity was at its highest in the differentiating xylem. This can be attributed to the high demand for cell wall material in this rapidly growing sink tissue. SuSy activity decreased as the tissue matured and was very low during the winter. The same trend was found in poplar, with SuSy activity being highest in the early summer, but only in the outer wood where cell differentiation takes place (Schrader and Sauter, 2002). In the main storage portion of the Robinia trunk, SuSy rates increased until autumn and then decreased, coinciding with starch accumulation.  The transition zone from sapwood to heartwood also showed an  increase from July to a maximum in September and November, and then decreased. The peak in SuSy activity coincides with increased activities of the enzymes involved in the phenylalanine pathway and with the accumulation of phenolic extractives.  The  conclusion was that a high amount of sucrose cleavage is required to provide the energy and substrates for the synthesis of these extractives (Hauch and Magel, 1999). Magel et a!. (2001) also studied non-structural carbohydrates and sucrose metabolizing enzymes in the trunk tissues of two Juglans species. In contrast to the study in Robinia, no season-dependent changes were seen in the radial distribution of non-structural carbohydrates. In Black Walnut, the content of starch strongly decreased with increasing tissue age. Sucrose pools were diminished in the transition zone, and both starch and sucrose were non-existent in the heartwood.  Glucose started to  accumulate at the transition zone and there were high levels in the heartwood.  In  contrast, in the Juglans hybrid, starch content was consistent throughout all living cells, and pools of sucrose decreased slightly towards the centre of the tree. Glucose and fructose levels were low in all tissues. Starch was deposited in the early sapwood until autumn, and decreased from November to February resulting in a significant increase in sucrose, fructose and glucose. Starch storage is correlated with the enhanced activities of SuSy and UGPase, while decreases in starch content and increases in soluble carbohydrates are correlated to an increase in SPS. An increase in sucrose activates 10  the enzymes of sucrolysis.  In the wood of hybrids this increase is attributed to  invertase, but in black walnut, as in many herbaceous plants, it is SuSy whose activity increases. An additional study investigated the effects of carbohydrates and sucrose metabolism enzymes on the transition from early wood to late wood in Scots Pine (Pinus sylvestris) (Uggla et a!., 2001). Sucrose levels were highest in the phloem, with a steep decrease across the cambial zone to its lowest level in the developing xylem cells. Fructose and glucose showed the opposite pattern to sucrose, being present in low levels in the phloem and increasing towards the xylem. The peak amount was found in the zone of secondary wall formation.  SuSy activity increased during the  earlywood to latewood transition, and became predominant in the area of secondary wall formation. This is in agreement with SuSy having a role in both cellulose synthesis (Amor et a!., 1995) and lignin biosynthesis (Hauch and Magel, 1999). This is supported by the high activity of SuSy at the late stages of cell differentiation when cellulose deposition is decreasing and lignification occurs (Uggla et a!., 2001).  The  overexpression of a mutant mung bean (Vigna radiata) SuSy (ShE) in Acetobacter xylinum has been shown to enhance cellulose production (Nakai eta!., 1999). SuSy upregulation has been studied in Popu!us alba using SuSy (SI I E) and the 35S constitutive promoter (Konishi et al., 2004).  The overall morphology of the  transgenics was unchanged, although a few lines were reported to have grown slightly better. SuSy expression was detected in both the leaves and stems, but was slightly higher in the leaves. It was also detected in both the soluble and microsomal fractions with higher expression in the soluble fraction. The study utilized a dual labelling system and provided evidence that at least some amount of sucrose is channelled directly to glucan formation via UDP-glucose. Overexpression of SuSy caused the glucose moiety of sucrose to be predominantly incorporated into the glucan backbone of xyloglucan in stem tissue.  This allows the high-energy bond to be preserved and then used for  polysaccharide synthesis.  It has also been proposed that xyloglucan synthase may  even form a putative synthase complex with SuSy (Konishi et a!., 2004). It appears that fructose is rapidly recycled to UDP-glucose by way of the hexose phosphate pool, or by SPS.  Fructose inhibits SuSy (Doehlert, 1987) and may also be coupled with the  negative signalling related to photosynthesis (Quereix et a!., 2001).  Despite the 11  apparent channelling to glucan synthases, there was no change observed in cellulose or non-cellulosic polysaccharides. This is attributed to the high level of SuSy in leaf tissue relative to sink tissue, possibly retaining sucrose in the leaves and therefore never allowing it to reach the sink tissues.  UDP-Glucose Pyrophosphorylase Herbaceous Species UDP-glucose pyrophosphorylase (UGPase, EC 2.7.7.9) catalyzes the reversible reaction of glucose-I-phosphate and UTP to UDP-glucose and pyrophosphate. The reaction plays a large role in sugar accumulation (Sowokinos, 1990) and has the potential to restrict the flow of carbon to sucrose formation (Borokov eta!., 1996), due to the negative cooperativity for glucose-I-phosphate and UTP (Sowokinos et a!., 1993). However, it is unlikely that the role of UGPase is rate-limiting as it is generally present in very large quantities compared to other enzymes involved in sucrose metabolism (Jaing et a!., 2003). UGPase has been shown to directly affect cellulose biosynthesis, as in Acetobacter xylinum, cellulose negative mutants were complemented by UGPase and shown to mitigate cellulose accumulation (Valla eta!., 1989). UGPase has been purified from numerous species including potato (Nakano et al., 1989; Sowokinos eta!., 1993), soybean (Vella and Copeland, 1990), rice (Kimura et a!., 1992), and barley (Elling and Kula, 1994). Various isozymes seem to exist within the potato, which has both an acidic and a basic form. Transgenic studies that decreased UGPase activity with very little effect on growth and development support the theory of numerous isozymes (Borokov et a!., 1996; Spychalla et a!., 1994; Zrenner et a!., 1993).  In Dictyostelium discoideum, two UGPase genes have been identified  (Bishop et a!., 2002).  A knockout of udpgpl had no phenotypic effect, while ugpB  mutants were unaffected in early growth but unable to form viable spores. The lack of viable spores has been linked to an inability of the mutants to form cellulose (Bishop et a!., 2002). In barley, 11 cDNA encoding UGPase have been isolated, and differ only in the polyadenation site positioning. It has been suggested that they are likely encoded by the same gene as they are identical at the nucleotide level (Eimert et a!., 1996). 12  UGPase is believed to exist as a monomer, but a study of barley shows it can undergo reversible oligomerization as a form of regulatory mechanism and the monomer is the active form (Martz et a!., 2002). Two UGPase genes have been identified in poplar (Meng et a!., 2007). UGPI was shown to be upregulated by light and sucrose feeding, while both genes were upregulated by cold treatment. Localization of the UGPase activity has been determined in a number of species. In the developing rice endosperm, UGPase is localized mainly in the cytosol (90%) with the remainder being found in the amyloplast and Golgi membranes (Kimura et a!., It has also been found in the microsome fraction of both rice and tobacco  1992).  (Mikami et a!., 2001). In barley, cytosol and membrane fractions suggest that UGPase may bind to membranes (Becker et a!., 1995).  UGPase shows high activity in the  young seeds and vascular tissue of tomato, and is found in the phloem and epidermis hairs on petunia (Sergeeva and Vreugdenhil, 2002). In potato, the enzyme has been identified in roots, tubers, stolons, leaves and stems, but in varying quantities (Sowokinos et a!., 1993). The levels were highest in sink tubers and stolons, and the levels increased during tuber development and with sucrose feeding to detached leaves. It has been shown in barley malt that inhibitory substrates exist in both the synthesis and pyrophosphorylysis reactions (Elling, 1996). In synthesis, UTP surplus has inhibitory effects.  Pyrophosphate is a non-competitive inhibitor for glucose-I-  phosphate and UTP.  While inorganic phosphate is non-competitive inhibitor for  glucose-I-phosphate and UTP, it has a much higher inhibition constant than pyrophosphate. UDP-glucose is a competitive inhibitor for UTP and a non-competitive inhibitor for glucose-I-phosphate. In synthesis, UGPase binds first to UTP and then to glucose-I-phosphate. In pyrophosphorylysis, UTP shows competitive product inhibition with UDP-glucose as a variable substrate. A number of studies have examined the effect of pyrophosphatase on UGPase content. In potato, the overexpression causes a reduction in pyrophosphate content, with little effect on the plant growth (Farre et a!., 2001). The tubers were less dense, which was attributed to a 30-40% decrease in starch content. There was an increase in sucrose and glucose content, but no significant changes in fructose.  UDP-glucose  content increased significantly and there was a 40-60% decrease in glucose-613  phosphate and 30-40% decrease in fructose-6-phosphate. not change appreciably.  Glucose-I-phosphate did  The removal of inorganic pyrophosphatase facilitated the  enhanced conversion of glucose-I -phosphate to UDP-glucose. This is hypothesized to lead to improved sucrose and cell wall biosynthesis which may result in the accelerated sprouting phenotype observed in these plants (Farre eta!., 2001). This study reinforced the finding of a previous study that found the rate of sucrose degradation and starch content  increased  (20-30%)  in  growing  potato  tubers with  pyrophosphatase  overexpressed (Geigenberger et a!., 1998). This seemingly contrary report is related to the differing function of UGPase in source and sink tissues. An investigation in developing potato tubers showed that the UGPase activity does not change with developmental stage (Appeldoorn et a!., 1997).  In developing  tubers UDP-glucose must be rapidly converted to allow the high net flux of sucrose into the tuber.  While fructokinase activity increases, UGPase activity remains constant,  suggesting that the presence of excess enzyme.  Furthermore, it has been  hypothesized that sucrose may regulate UGPase. This is consistent with the study by Spychalla et a!. (1994), who showed an increase in mRNA in potato leaves incubated with high sucrose levels, as well as with the increased activity of the enzyme correlating strongly with sugar amounts accumulated during cold storage. UGPase activity is also regulated strongly by substrate availability (Vella and Copeland, 1990).  It is also  thought to be regulated by the ratio of SuSy to SPS activities (Kleczkowski, 1994). However, in transgenic maize where SuSy was downregulated, UGPase activity did not co-ordinately decrease (Carlson and Chourey, 1996). It has been suggested that the regulation of the gene is translational rather than transcriptional, as significant decreases in activity did not change growth and development (Kleczkowski, 1994; Zrenner et a!., 1993). However, a decrease in levels of inorganic phosphate has been connected to an increase in UGPase activity, which suggests transcriptional or posttranslational control (Ciereszko et a!., 2001). Activities of AGPase, SPS, SuSy and invertase also increase in this situation and the effect may be related to phosphate deficiency causing gene upregulation, or perhaps affecting the catalytic efficiency of the enzyme via protein turnover rate or post-translational  mechanisms such as  phosphorylation. Stress is also known to alter plant inorganic phosphate content and carbohydrate status.  It is possible that the change in UGPase associated with 14  phosphate is related to the maintenance of plant nutritional status (Ciereszko et a!., 2001). There have been a number of studies carried out on potato with decreased UGPase activity. Three significant studies resulted in contradicting results.  The first  showed that the activity of UGPase could be decreased by up to 96% without any detectable changes in sugar content or growth rates (Zrenner et aL, 1993). The other two studies showed that a decrease as low as 30-50% caused a decrease in sugar content (Borokov et aL, 1996; Spychalla et a!., 1994). The difference in findings could be attributed to differences in methods. Zrenner et a!. (1993) utilized the entire coding region of a UGPase gene for the generation of the construct. Spychalla et a!. (1994) used a 0.5 kb fragment from the 5’ end of the cDNA and Borokov et a!. (1996) used a piece of the Ugp genomic clone. The studies also employed different promoters with Zrenner et a!. (1993) and Borokov et a!. (1996) using the 35S cauliflower promoter and Spychalla et a!. (1994) utilizing the patatin promoter. Zrenner et a!. (1993) analyzed tubers that were still growing and acting as sinks, while the other studies looked at tubers stored at varying temperatures and were acting as source tissue. The conclusion from Zrenner et a!. (1993) is that in sink tissues, only 4% of UGPase activity is necessary for normal plant growth.  In mature tubers, it appears that the role of  UGPase is more significant.  Woody Species UGPase activity has been studied in a number of herbaceous species, but only preliminary work has been carried out in woody species, and there has yet to be any research evaluating transgenics. In a study on two species of Juglans, the distribution of UGPase activity was found to be similar to that of SuSy, but ten times higher (Magel et al., 2001), with activity gradually decreased towards the transition zone and non existent in the heartwood. UGPase activity also experienced seasonal fluctuations. In hybrid Jugians, activity increased during the summer and towards autumn, but was low in the winter. Activity in black walnut was similar, but the high autumn activity level was maintained through the winter and declined in the spring. UGPase is thought to play a crucial role in the metabolism of the products of sucrose cleavage, and it has a higher 15  activity than other sucrose metabolism enzymes. Based on the observation of the various sucrose metabolizing enzymes, Magel et aL (2001) presented the hypothesis that the formation of heartwood phenolics is enabled by the enzymes involved in sucrose metabolism as they facilitate the degradation of sucrose. At the same time and in the same location, the activities of enzymes involved in the phenolics pathways are also increased and utilize the products of sucrose degradation in the formation of heartwood extractives.  Sucrose Phosphate Synthase Herbaceous Species SPS is a key enzyme in regulating sucrose synthesis in plants. Protein association studies have shown evidence for a direct interaction between SPS and SPP, creating a metabolic channel from UDP-glucose and fructose-6-phosphate to  sucrose (Echeverria et al., 1997). SPS catalyzes the reaction from UDP-glucose and fructose-6-phosphate into sucrose-6-phosphate and UDP. Sucrose-6-phosphate phosphatase then acts to hydrolyse sucrose-6-phosphate into sucrose and a phosphate group. This specific and highly active phosphatase essentially draws the SPS reaction towards the production of sucrose, by altering the equilibrium of products. It is thought that SPS contributes to the control of flux into sucrose, as it contributes greatly to the biosynthesis of sucrose in both photosynthetic and non-photosynthetic tissues (Geigenberger et a!., 1999; Geigenberger and Stitt, 2000; Strand et a!., 2000). In addition to preventing the accumulation of fructose, a SuSy inhibitor, the reaction also releases phosphate, which has been suggested to have a substantial role in the regulation of photosynthetic rate through its cycling between the chioroplast and the cytosol (Stitt et a!., 1987). SPS has been shown to be under the regulation of a variety of mechanisms including enzyme abundance, allosteric control in which glucose-6phosphate acts as an activator and phosphate as an inhibitor, and phosphorylation. SPS activity is also regulated by developmental, environmental and nutritional signals. Changes in irradiance can affect the transcription level, with an increase in light resulting in an increase in SPS mRNA levels followed by an increase in protein and activity (Klein eta!., 1993; Cheng eta!., 1996). 16  Transgenic plants have been used to examine the effects of SPS. In tomato, the overexpression of maize SPS caused an increase in the synthesis of sucrose, resulting in an increase in the sucrose to starch ratio in leaves, as well as an increase in photosynthetic capacity (Worrell et aL, 1991). This suggests that SPS has some effect on photosynthesis (Galtier et a!., 1993, Galtier et a!., 1995; Micallef et al., 1995). In tobacco, the overexpression of a maize SPS caused the acceleration of flower development and an increase in flower numbers (Baxter et a!., 2003). Similarly, the overexpression of SPS resulted in an increased sucrose to starch ratio in Arabidopsis (Signora et al., 1998) and improved fibre quality and yield in cotton (Haigler et al., 2007). SPS overexpression in tomato resulted in a six-fold increase in SPS activity in leaf tissue, which resulted in slight changes in carbohydrate, but a marked increase in the sucrose to starch ratio (VVorrell eta!., 1991). There have been fewer studies involving the decreased expression of SPS, but a study in potato with decreased expression of SPS demonstrated the inhibition of sucrose synthesis and an increase in starch and amino acids (Krause et al., 1998). Recent studies of SPS overexpression in tobacco have revealed an increase in sink sucrose pools while there was not a concurrent increase in source sucrose pools. The plants were taller with increased biomass, but there were only minor changes in structural carbohydrates (Park et a!., 2008).  Woody Species There is less known about SPS in trees than the body of knowledge in herbaceous plants and much of the research involves conifers. In spruce needles, SPS exhibits activity levels inverse to that of SuSy during the period directly following bud break (Egger and Hampp, 1993).  SPS is negligible in the newest needles, but  increases as the needles mature resulting in a change in the SPS/SuSy ratio as a tissue changes from a sink to a source. In autumn, the patterns of the two enzymes coincide, with both enzymes achieving their highest activity between October and February (Egger and Hampp, 1993). This is attributed to the role of these enzymes in metabolic acclimation to low temperatures. Further studies by the same group showed that SPS activity was low for the first 60 days after bud break; after the needle had achieved a 17  steady state dry weight/fresh weight ratio there was a steady increase in activity (Hampp et a!., 1994).  This increase in the SPS/SuSy ratio is considered to be a  measure of source or sink quality of a given tissue, and shows the transition point from the needle as a sink for carbohydrates to a source. In spruce needles, SPS activity was higher during the period ending with bud break, due to an increase in protein levels and activation under metabolite control, high levels of glucose-6-phosphate and a low inorganic phosphate/glucose-6-phosphate ratio (Egger et a!., 1996).  This period was also characterized by high rates of net  photosynthesis, a large decrease in soluble sugars, and a steep rise in starch content. After bud break, net photosynthesis was greatly reduced (by about 75%), and SPS activity and protein level were decreased.  There was also a reduction in the  concentration  an  of glucose-6-phosphate  and  increased  phosphate/glucose-6-  phosphate ratio. During this period, sucrose synthesis was reduced in older needles and the carbon demand of the developing needles was met in part by the mobilization of starch from older needles (Egger eta!., 1996). SPS activity in spruce can be regulated in an allosteric manner by glucose-6phosphate and inorganic phosphate (Loewe et a!., 1996).  Glucose-6-phosphate  activates SPS by increasing its affinity for fructose-6-phosphate. Inorganic phosphate can inhibit this activation when the fructose-6-phosphate concentration is rate-limiting. The seasonal fluctuations in SPS activity correlate with fluctuations in fructose-6phosphate and glucose-6-phosphate, which are both at higher concentrations in the winter. However, allosteric regulation of SPS seems to be species dependent, because various studies differing results (Loewe et a!., 1996). In spruce, ATP-dependent phosphorylation did not play a major role in seasonal regulation of SPS activity, but the seasonal changes in activity are correlated with protein levels (Loewe et a!., 1996). High SPS activity in the winter indicates a high level of sucrose synthesis, which could be loaded into the phloem for export under mild winter conditions (Loewe et a!., 1996) or used for the formation of raffinose and stachyose, which are synthesized during cold acclimation (Egger et a!., 1996). The recycling of sucrose is necessary to recycle inorganic phosphate to accommodate the considerable rates of photosynthesis during the winter months. The role of SPS in cold 18  acclimated needles in correlation with photosynthetic activity and high levels of sucrose export have been supported by experiments in transgenic tomato (Galtier et a!., 1993). In Robinia, SPS activity was low regardless of season, age, and physiological conditions (Hauch and Magel, 1999). Highest activities were in the middle sapwood (May) and inner sapwood (November), and decreased towards the bark and the sapwood/heartwood transition. Cold-adapted tissues show higher rates of SPS than samples harvested in the summer, which may be regulated allosterically by the seasonally increased pools of glucose-6-phosphate (Hauch and Magel, 1999). Exposure of living wood to decreased temperatures generally results in a decrease in starch content and a related increase in soluble carbohydrates which can act as cryoprotectants (Egger et a!., 1996; Sauter et a!., 1996; Schrader and Sauter, 2002). Increased SPS activities are directly related to the accumulation of sucrose over the winter. Sucrose turnover dominates in the bark of Robinia, which shows a 10-fold higher protein content.  Within the xylem, the radial distribution of sucrose-synthesizing  activities is opposite to that of sucrose-cleaving activities, with SPS located in the mature middle and inner sapwood, versus the outermost wood where sucrose-cleaving activities dominate. In the spring, summer and autumn, the processes within the tree are more spatially defined. In the primary xylem, SPS increases in the spring coinciding with a decrease in starch content, indicating conversion from starch to sucrose (Hauch and Magel, 1999). Once the leaves begin to develop and supply sucrose to the tree, the level of SPS decreased. In the sapwood/heartwood transition zone, SPS levels are highest in autumn, suggesting that this region may be the source of sucrose required for the transition zone (Hauch and Magel, 1999). In Scots pine sampled in the summer months, SPS activity was present in all tissues, but was highest in the phloem (Uggla eta!., 2001). The starch content in living bark decreases steadily from the onset of cambial activity until late summer in both Scots pine and Norway spruce (Egger et a!., 1996). This suggests that sucrose pools in the rays are partly derived from stored starch. In hybrid poplar, SPS activities are high in autumn and winter, which correlates with sucrose content at this time and likely acts as a cryoprotectant mechanism (Schrader and Sauter, 2002).  SPS is thought to play a role in the partitioning of 19  photoassimilates (Galtier et a!., 1993), in relation to the C0 -fixation rate (Galtier et al., 2 1995) and in relation to light/dark-changes (Stitt et a!., 1988). It also plays a role in the ripening of fruit, starch-to-sugar conversion in germination of some seeds, and in stress mechanisms in some plants in response to cold and water stress (Schrader and Sauter, 2002). One study has looked at the overexpression of maize SPS in a poplar hybrid (Populus tremula x P. tremuloides), and the preliminary data showed an increase in total leaf carbohydrates and leaf starch content. There was also a significant increase in photosynthesis and higher growth rates (Mouillon and Hurry, 2001). Diurnal changes in leaf soluble sugar content were also noted, with an increase in the first part of the light period compared to controls.  Promoters Enhanced Tandem Cauliflower Mosaic Virus Promoter The enhanced tandem cauliflower mosaic virus promoter (2x35S) was originally designed to increase the efficiency of transcription obtained by the natural cauliflower mosaic virus 35S promoter. tandem.  The 2x35S promoter contains two 35S promoters in  When compared with the nopaline synthase (NOS) promoter, the 35S  promoter showed a 10-fold increase in neomycin phosphotransferase (NPTII) transcript levels, while the 2x35S promoter showed a 100-fold increase in transgenic tobacco (Kay etal., 1987). Further experimentation on this promoter yielded even greater increases in transcript efficiency. A 40-base leader sequence from alfalfa mosaic virus (AMV leader) was utilized in combination with both the 35S promoter and 2x35S.  Using 3-  glucuronidase (GUS) as a marker gene, the promoters were studied in protoplast suspensions of tobacco and white spruce transformed by electroporation (Datla et a!., 1993)  .  The 2x35S promoter consistently yielded expression levels 4 times that of the  35S promoter with the AMV leader. The promoter is constitutively expressed and yields expression levels of about 100-fold that of the natural cauliflower mosaic virus 35S (Appendix A). 20  4-Coumarate: Coenzyme A Ligase Promoter The 4-coumarate:CoA ligase promoter was examined in a study evaluating the expression of the 4CL enzyme (Hauffe et a!., 1991). In tobacco transformed with GUS under the control of the 4CL promoter, it was found that activity was high in the primary xylem of axillary buds and developing leaf veins, but not observable in other cell types of young leaves or stems. The highest expression was found in xylem tissue during the differentiation of tracheary elements.  In older stems, GUS activity was high in the  secondary xylem, in the ray parenchyma cells.  In the roots, GUS expression was  observed in the vascular tissue, root hairs and subapical cells, while in immature flowers, GUS was expressed in the vascular tissue and developing nectaries. Expression was highest in developing seeds in a single epidermal cell layer and mature stigmas also showed high GUS expression. The 4CL promoter provides an interesting comparison with the 2x35S promoter due to its high expression in vascular tissues and the developing and secondary xylem (Appendix A).  Objectives The objective of this research was to investigate the effect(s) of the constitutive (2x35S) and tissue-specific (4CL) overexpression of SuSy and UGPase in tobacco (Nicotiana tabacum) and hybrid poplar (Populus alba x grandidentata). examined the effects of each gene individually and in combination.  The work  In addition, in  tobacco, these genes were combined with SPS to form double transgenics (SuSy x SPS and UGPase x SPS) as well as triple transgenics. The study focused on the effect of these genes on structural and soluble carbohydrates, as well as molecular characterization of the plants. These analyses permitted the testing of two hypotheses: 1. The tissue specific or constitutive expression of SuSy, UGPase, and SPS individually or in combination can alter the sucrose metabolism pathway, ultimately affecting plant growth rates. 2. The sucrose metabolism pathway can be altered in such a way as to affect the biosynthesis of cellulose, quantitatively and qualitatively. As mentioned, limited transgenic work has been carried out in woody species, with only one study utilizing the 35S promoter and a modified SuSy gene (Konishi et al., 21  2004), and one study examining the effects of the overexpression of SPS (Mouillon and Hurry, 2001). No work has yet been carried out with UGPase transgenic poplar, or with multiple genes from the sucrose metabolism pathway.  22  References  Amor Y., Haigler C., Johnson S., Wainscott M. and Delmer D.P. 1995. A membraneassociated form of sucrose synthase and its potential role in synthesis of cellulose and callose in plants. Proceedings of the National Academy of Sciences of the United States of America 92: 9353-9357. Appeldoorn N.J.G., de Bruijn S.M., Koot-Gronsveld E.A.M., Visser R.G.F., Vreugdenhil D. and van der Plas L.H.W. 1997. Developmental changes of enzymes involved in conversion of sucrose to hexose-phosphate during early tuberisation of potato. Planta 202: 220-226. Baxter C.J., Foyer C.H., Turner J., Rolfe S.A. and Quick W.P. 2003. Elevated sucrosephosphate synthase activity in transgenic tobacco sustains photosynthesis in older leaves and alters development. Journal of Experimental Botany 54: 18131820. Becker M., Vincent C. and Reid J.S.G. 1995. Biosynthesis of (1 ,3)(1 ,4)-beta-glucan and (1,3)-beta glucan in barley (Hordeum vulgare L.): properties of the membranebound glucan synthases. Planta 195: 331-338. Bishop J.D., Moon B.C., Harrow F., Ratner D., Gomer R.H., Dottin R.P. and Brazill D.T. 2002. A second UDP-glucose pyrophosphorylase is required for differentiation and development in Dictyostelium discoideum. The Journal of Biological Chemistiy 277: 32430-32437. Bologa K.L., Fernie A.R., Leisse A., Loureiro M.E. and Geigenberger P. 2003. A bypass of sucrose synthase leads to low internal oxygen and impaired metabolic performance in growing potato tubers. Plant Physiology 132: 2058-2072. Borokov A.Y., McClean P.E., Sowokinos J.R., Ruud S.H. and Secor G.A. 1996. Effect of expression of UDP-glucose pyrophosphorylase ribozyme and antisense RNAs on the enzyme activity and carbohydrate composition of field-grown transgenic potato plants. Journal of Plant Physiology 147: 644-652. Brown R.M.J. and Saxena l.M. 2000. Cellulose biosynthesis: A model for understanding the assembly of biopolymers. Plant Physiology 38: 57-67. Brown R.M.J., Saxena l.M. and Kudlicka K. 1996. Cellulose biosynthesis in higher plants. Trends in Plant Science 1: 149-156. Canam T., Mak S.W.Y. and Mansfield S.D. 2008. Spatial and temporal expression profiling of cell-wall invertase genes during early development in hybrid poplar. Tree Physiology. In press. Carlson S.J. and Chourey P.S. 1996. Evidence for plasma membrane-associated forms of sucrose synthase in maize. Molecular and General Genetics 252: 303-310. 23  Carlson S.J., Chourey P.S., Helentjaris T. and Datta R. 2002. Gene expression studies on developing kernel of maize sucrose synthase (SuSy) mutants show evidence for a third SuSy gene. Plant Molecular Biology 49: 15-29. Cheng W.H., Im K.H. and Chourey P.S. 1996. Sucrose phosphate synthase expression at the cell and tissue level is coordinated with sucrose sink-to-source transitions in maize leaf. Plant Physiology 111: 1021-1029. Chengappa S., Guilleroux M., Phillips W. and Shields R. 1999. Transgenic tomato plants with decreased sucrose synthase are unaltered in starch and sugar accumulation in the fruit. Plant Molecular Biology 40: 213-221. Chourey P.S., Taliercio E.W., Carlson S.J. and Ruan Y.-L. 1998. Genetic evidence that the two isozymes of sucrose synthase present in developing maize endosperm are critical, one for cell wall integrity and the other for starch biosynthesis. Molecular and General Genetics 259: 88-96. Ciereszko I., Johansson H., Hurry V. and Kleczkowski L.A. 2001. Phosphate status affects the gene expression, protein content and enzymatic activity of UDP glucose pyrophosphorylase in wild-type and pho mutants of Arabidopsis. Planta 212: 598-605. Datla R.S.S., Bekkaoui F., Hammerlindl J.K., Pilate G., Dunstan D.l. and Crosby W.L. 1993. Improved high-level constitutive foreign gene translation using an AMV RNA4 untranslated leader sequence. Plant Science 94: 139-149. D’Aoust M.-A., Yelle S. and Nguyen-Quoc B. 1999. Antisense inhibition of tomato fruit sucrose synthase decreases fruit setting and the sucrose unloading capacity of young fruit. The Plant Cell 11: 2407-2418. Dejardin A., Sokolov L.N. and Kleczkowski L.A. 1999. Sugar/osmoticum levels modulate differential abscisic acid-independent expression of two stressresponsive sucrose synthase genes in Arabidopsis. Biochemistry Journal 344: 503-509. Delmer D.P. 1999. Cellulose biosynthesis: Exciting times for a difficult field of study. Annual Review of Plant Physiology and Plant Molecular Biology 50: 245-276. Delmer D.P. and Amor Y. 1995. Cellulose biosynthesis. The Plant Cell 7: 987-1000. Djerbi S., Lindskog M., Arvestad L., Sterky F. and Teen T.T. 2005. The genome sequence of black cottonwood (Populus trichocarpa) reveals 18 conserved cellulose synthase (CesA) genes. Planta 221: 739-746. Doehlert D.C. 1987. Substrate-inhibition of maize endosperm sucrose synthase by fructose and its interaction with glucose inhibition. Plant Science 52: 153-1 57. 24  Echeverria E., Salvucci M.E., Gonzalez P., Paris G. and Salerno G. 1997. Physical and kinetic evidence for an association between sucrose phosphate synthase and sucrose phosphate phosphatase. Plant Physiology 115: 223-227. Egger B., Einig W., Schlereth A., Wallenda T., Magel E., Loewe A. and Hampp R. 1996. Carbohydrate metabolism in one- and two-year-old spruce needles, and stem carbohydrates from three months before until three months after bud break. Physiologia Plantarum 96: 91-100.  Egger B. and Hampp R. 1993. lnvertase, sucrose synthase and sucrose phosphate synthase in lyophilized spruce needles; microplate reader assays. Trees 7: 98103. Eimert K., Villand P., Kilian A. and Kleczkowski L.A. 1996. Cloning and characterization of several cDNAs for UDP-glucose pyrophosphorylase from barley (Hordeum vulgare) tissues. Gene 170: 227-232. Elling L. 1996. Kinetic characterization of UDP-glucose pyrophosphorylase from germinated barley (malt). Phytochemistiy 42: 955-960. Elling L. and Kula M.R. 1994. Purification of UDP-glucose pyrophosphorylase from germinated barley (malt). Journal of Biotechnology 34: 157-163. Farre E.M., Bachmann A., Willmitzer L. and Tretheway R.N. 2001. Acceleration of potato tuber sprouting by the expression of a bacterial pyrophosphorylase. Nature Biotechnology 19: 268-272. Galtier N., Foyer C.H., Huber J., Voelker T.A. and Huber S.C. 1993. Effects of elevated sucrose-phosphate synthase activity on photosynthesis, assimilate partitioning, and growth in tomato (Lycopersicon esculentum var UC82B). Plant Physiology 101: 535-543. Galtier N., Foyer C.H., Murchie E., Aired R., Quick W.P., Voelker T.A., Thepenier C., Lasceve G. and Betsche T. 1995. Effects of light and atmospheric carbon dioxide enrichment on photosynthesis and carbon partitioning in the leaves of tomato (Lycopersicon esculentum L.) plants over-expressing sucrose phosphate synthase. Journal of Experimental Botany 46: 1335-1344. Geigenberger P., Hajirezaei M.R., Geiger M., Deiting U., Sonnewald U. and Stitt M. 1998. Overexpression of pyrophosphatase leads to increased sucrose degradation and starch synthesis, increased activities of enzymes for sucrose starch interconversions, and increased levels of nucleotides in growing potato tubers. Planta 205: 428-437. Geigenberger P., Reimholz R., Deiting U., Sonnewald U. and Stiff M. 1999. Decreased expression of sucrose phosphate synthase strongly inhibits the water stress 25  induced synthesis of sucrose in growing potato tubers. The Plant Journal 19: 119-129. Geigenberger P. and Stitt M. 2000. Diurnal changes in sucrose, nucleotides, starch synthesis, and AGPS transcript in growing potato tubers that are suppressed by decreased expression of sucrose phosphate synthase. The Plant Journal 23: 795-806. Guy C.L. 1990. Cold-acclimation and freezing stress tolerance Role of proteinmetabolism. Annual Review of Plant Physiology and Plant Molecular Biology 41: 187-223. -  Haigler C.H., Singh B., Zhang D., Hwang S., Wu C., Cal W.X., Hozain M., Kang W., Kiedaisch B.M., Strauss R.E., Hequet E.F., Wyatt B.G., Jividen G.M. and Holaday A.S. 2007. Transgenic cotton over-producing spinach sucrose phosphate synthase showed enhanced leaf sucrose synthesis and improved fiber quality under controlled environmental conditions. Plant Molecular Biology 63: 815-832. Haigler C.H., Ivanova-Datcheva M., Hogan P.S., Salnikov V.V., Hwang S., Martin K. and Delmer D.P. 2001. Carbon partitioning to cellulose synthesis. Plant Molecular Biology 47: 29-51. Hampp R., Egger B., Effenberger S. and Einig W. 1994. Carbon allocation in developing spruce needles. Enzymes and intermediates of sucrose metabolism. Physiologia Plantarurn 90: 299-306. Hanggi E. and Fleming A.J. 2001. Sucrose synthase expression pattern in young maize leaves: implications for phloem transport. Planta 214: 326-329. Hauch S. and Magel E. 1999. Extractable activities and protein content of sucrosephosphate synthase, sucrose synthase and neutral invertase in trunk tissues of Robinia pseudoacacia L. are related to cambial wood production and heartwood formation. Planta 207: 266-274. Hauffe K.D., Paszkowski U., Schulze-Lefert P., Hahlbrock K., Dangl J.L. and Douglas C.J. 1991. A parsley 4CL-1 promoter fragment specifies complex expression patterns in trangenic tobacco. The Plant Cell 3: 435-443. Jaing D., Cao W., Dai T. and Jing Q. 2003. Activities of key enzymes for starch synthesis in relation to growth of superior and inferior grains on winter wheat (Triticurn aestivum L.) spike. Plant Growth Regulation 41: 247-257. Kay R., Chan A., Daly M. and McPherson J. 1987. Duplication of CaMV 35S promoter sequences creates a strong enhancer for plant genes. Science 236: 1299-1302.  26  Kimura S., Mitsui T., Matsuoka T. and lgaue I. 1992. Purification, characterization and localization of rice UDP-glucose pyrophosphorylase. Plant Physiology and Biochemistry 30: 683-693. Kleczkowski L.A. 1994. Glucose activation and metabolism through UDP-glucose pyrophosphorylase in plants. Phytochemistiy 37: 1507-1515. Klein R.R., Crafts-Brandner S.J. and Salvucci M.E. 1993. Cloning and developmental expression of the sucrose phosphate gene from spinach. Planta 190: 498-510. Koch K.E. 1996. Carbohydrate-modulated gene expression in plants. Annual Review of Plant Physiology and Plant Molecular Biology 47: 509-540. Konishi T., Ohmiya Y. and Hayashi T. 2004. Evidence that sucrose loaded into the phloem of a poplar leaf is used directly by sucrose synthase associated with various j3-glucan synthases in the stem. Plant Physiology 134: 1146-1152. Krause K.-P., Reimholz R., Hamborg Nielson T., Sonnewald U. and Still M. 1998. Sucrose metabolism in cold-stored potato tubers with decreased expression of sucrose phosphate synthase. Plant, Cell and Environment 21: 285-299. Loewe A., Einig W. and Hampp R. 1996. Coarse and fine control and annual changes of sucrose-phosphate synthase in Norway spruce needles. Plant Physiology 112:641-649. Magel E., Abdel-Latif A. and Hampp R. 2001. Non-structural carbohydrates and catalytic activities of sucrose metabolizing enzymes in twnks of two Juglans species and their role in heartwood formation. Holzforshung 55:135-145. Martz F., Wilczynska M. and Kleczkowski L.A. 2002. Oligomerization status, with the monomer as active species, defines catalytic efficiency of UDP-glucose pyrophosphorylase. Biochemistry Journal 367: 295-300. Meng M., Geisler M., Johansson H., Mellerowicz E., Karpinski S. and Kleczkowski L.A. 2007. Differential tissue/organ-dependent expression of two sucrose- and coldresponsive genes for UDP-glucose pyrophosphorylase in Populus. Gene 389:186-195. Micallef B.J., Haskins K.A., Vanderveer P.J., Roh K.-S., Shewmaker C.K. and Sharkey T.D. 1995. Altered photosynthesis, flowering, and fruiting in transgenic tomato plants that have an increased capacity for sucrose synthesis. Planta 196: 327334. Mikami S., Hon H. and Mitsui T. 2001. Separation of distinct compartments of rice Golgi complex by sucrose density gradient centrifugation. Plant Science 161: 665-675.  27  Mouillon J.M. and Hurry V. 2001. Effects of elevated sucrose-phosphate synthase activity on photosynthesis, carbohydrate partitioning and growth rate in hybrid aspen (Populus tremula x P. tremuloides). Proceedings of the 12th International Conference on Photosynthesis, Brisbane Australia. Nakai T., Tonouchi N., Konishi T., Kojima Y., Tsuchida T., Yoshinaga F., Sakai F. and Hayashi T. 1999. Enhancement of cellulose production by expression of sucrose synthase in Acetobacter xylinum. Proceedings of the National Academy of Sciences of the United States of America 96: 14-18. Nakano K., Omura Y., Tagaya M. and Fukui T. 1989. UDP-glucose pyrophosphorylase from potato tuber: purification and characterization. Journal of Biochemistiy 106: 528-532. Nguyen-Quoc B. and Foyer C.H. 2001. A role for ‘futile cycles’ involving invertase and sucrose synthase in sucrose metabolism of tomato fruit. Journal of Experimental Botany 52: 881-889. Nguyen-Quoc B., Krivitsky M., Huber S.C. and Lecharny A. 1990. Sucrose synthase in developing maize leaves regulation of activity by protein level during the import to export transition. Plant Physiology 94: 516-523. -  Park J.-Y., Canam T., Kang K.Y., Ellis D.D. and Mansfield S.D. 2008. Overexpression of an arabidopsis family A sucrose phosphate synthase (SPS) gene alters plant growth and fibre development. Transgenic Research 17: 181 -1 92. Pear J.R., Kawagoe Y., Schreckengost W.E., Delmer D.P. and Stalker D.M. 1996. Higher plants contain homologs of the bacterial celA genes encoding the catalytic subunit of cellulose synthase. Proceedings of the National Academy of Sciences of the United States of America 93: 12637-12642. Preston R.D. 1974. The physical biology of plant cell walls. London, Chapman and Hall. Quereix A., Dewar R.C., Gaudillere J.-P., Dayau S. and Valancogne C. 2001. Sink feedback regulation of photosynthesis in vines: measurements and a model. Journal of Experimental Botany 52: 2313-2322. Reiter W.D. 2002. Biosynthesis and properties of the plant cell wall. Current Opinion in Plant Biology 6: 536-542. Richmond T.A. 2000. Higher plant cellulose synthases. Genome Biology 1: 3001.13001.6. Ruan Y.L., Llewellyn D.J. and Furbank R.T. 2003. Suppression of sucrose synthase gene expression represses cotton fibre cell initiation, elongation, and seed development. The Plant Cell 15: 952-964. 28  Sauter J.J., Wisniewski M. and Witt W. 1996. Interrelationships between ultrastructure, sugar levels, and frost hardiness of ray parenchyma cells during from acclimation and deacclimation in poplar (Populus canadensis) Moench (robusta). Journal of Plant Physiology 149: 451-461. Saxena f.M., Lin F.C. and Brown R.M.J. 1990. Cloning and sequencing of the cellulose synthase catalytic subunit gene of Acetobacter xylinum. Plant Molecular Biology 15: 673-683. Schrader S. and Sauter J.J. 2002. Seasonal changes of sucrose-phosphate sunthase and sucrose synthase activities in poplar wood (Populus x canadensis Moench <robusta>) and their possible role in carbohydrate metabolism. Journal of Plant Physiology 159: 833-843. Sergeeva L.l. and Vreugdenhil D. 2002. In situ staining of activities of enzymes involved in carbohydrate metabolism in plant tissues. Journal of Experimental Botany 53: 361-370. Signora L., Galtier N., Skot L., Lucas H. and Foyer C.H. 1998. Over-expression of sucrose phosphate synthase in Arabidopsis thaliana results in increased foliar sucrose/starch ratios and favours decreased foliar carbohydrate accumulation in plants after prolonged growth with CO 2 enrichment. Journal of Experimental Botany 49: 669-680. Sowokinos J.R. 1990. Effect of stress and senescence on carbon partitioning in stored potatoes. American Potato Journal 67: 849-857. Sowokinos J.R., Spychalla J.P. and Desborough S.L. 1993. Pyrophosphorylases in Solanum tuberosum: IV. Purification, tissue localization, and physiochemical properties of UDP-glucose pyrophosphorylase. Plant Physiology 101: 10731080. Spychalla J.P., Scheffler B.E., Sowokinos J.R. and Bevan M.W. 1994. Cloning, antisense RNA inhibition, and the coordinated expression of UDP-glucose pyrophosphorylase with starch biosynthetic genes in potato tubers. Journal of Plant Physiology 144: 444-453. Stitt M., Gerhardt R., Wilke I. and Heldt H.W. 1987. The contribution of fructose 2-6bisphosphate to the regulation of sucrose synthesis during photosynthesis. Physiologia Plantarum 69: 377-386. Stitt M., Wilke I., Feil R. and Heldt H.W. 1988. Coarse control of sucrose phosphate synthase in leaves: alterations of the kinetic properties in response to the rate of photosynthesis and the accumulation of sucrose. Planta 174: 217-230. Strand A., Zrenner R., Trevanion S., Stitt M., Gustafsson P. and Gardestrom P. 2000. Decreased expression of two key enzymes in the sucrose biosynthesis pathway, 29  cytosolic fructose-I ,6-bisphosphatase and sucrose phosphate synthase, has remarkably different consequences for photosynthetic carbon metabolism in transgenic Arabidopsis thaliana. The Plant Journal 23: 759-770. Sun J., Loboda T., Sung S.-J.S. and Black C.C.J. 1992. Sucrose synthase in wild tomato, Lycopersicon chmielewskii, and tomato fruit sink strength. Plant Physiology 98: 1163-1169. Tang G.Q. and Sturm A. 1999. Antisense repression of sucrose synthase in carrot (Daucus carota L.) affects growth rather than sucrose partitioning. Plant Molecular Biology 41: 465-479. Uggla C., Magel E., Moritz T. and Sundberg B. 2001. Function and dynamics of auxin and carbohydrates during earlywood/latewood transition in scots pine. Plant Physiology 125: 2029-2039. Valla S., Coucheron D.H., Fjaervik E., Kjosbakken J., Weinhouse H., Ross P., Amikam D. and Benziman M. 1989. Cloning of a gene involved in cellulose biosynthesis in Acetobacter xylinum: Complementation of cellulose-negative mutants by the UDPG pyrophosphorylase structural gene. Molecular and General Genetics 217: 26-30. Vella J. and Copeland L. 1990. UDP-glucose pyrophosphorylase from the plant fraction of nitrogen fixing soybean nodules. Physiologia Plantarum 78:140-146. Williamson R.E., Burn J.E. and Hocart C.H. 2002. Towards the mechanism of cellulose synthesis. Trends in Plant Science 7: 461-467. Worrell A.C., Bruneau J.-M., Summerfelt K., Boersig M. and Voelker T.A. 1991. Expression of a maize sucrose phosphate synthase in tomato alters leaf carbohydrate partitioning. The Plant Cell 3: 1121-1130. Zrenner R., Salanoubat M., Willmitzer L. and Sonnewald U. 1995. Evidence of the crucial role of sucrose synthase for sink strength using transgenic potato plants (Solanum tugerosum L.). The Plant Journal 7: 97-107. Zrenner R., Willmitzer L. and Sonnewald U. 1993. Analysis of the expression of potato uredinediphosphate-glucose pyrophosphorylase and its inhibition by antisense RNA. Planta 190: 247-252.  30  Upregulation of Sucrose Synthase and UDP-Glucose Pyrophosphorylase Impacts Plant Growth and Metabolism 1 Introduction Cellulose, the most abundant organic polymer in the world, is deposited in the stems of plants and is extensively utilized for fuel, timber, forage, fibre and chemical cellulose. As such, how plants control carbon partitioning to cellulose biosynthesis is a key question for researchers globally (Haigler et al., 2001).  The availability of  carbohydrates from photosynthesis is not generally a limiting factor in cellulose synthesis despite carbohydrates having other crucial roles in plant growth and maintenance.  As such, the creation of photoassimilate sinks in cellulose-producing  cells could partition a greater portion of the photosynthate to these cells, and therefore to cellulose production. Cellulose synthesis, in contrast to starch, is essentially an irreversible sink (Haigler et a!., 2001). Cellulose is produced from the precursor UDP-glucose, which can be formed via two potential pathways.  UDP-glucose can be derived from the  cleavage of sucrose in a reaction catalyzed by sucrose synthase (SuSy; EC 2.4.1.13; Figure 1.1) yielding UDP-glucose and fructose.  Alternately, UDP-glucose can be  generated from the phosphorylation of glucose-I-phosphate in a reaction catalyzed by UDP-glucose pyrophosphorylase (UGPase, EC 2.7.7.9; Figure 1.1). SuSy plays an important role in supplying energy for phloem loading by providing a substrate for respiration (Hanggi and Fleming, 2001). The reaction retains the energy of the glycosidic bond in UDP-glucose, conferring an energetic advantage over the hydrolysis of sucrose catalyzed by invertase. SuSy has been characterized as existing both in the cytosol and in association with the plasmalemma (Carlson and Chourey, 1996), with the latter membrane-associated form, hypothesized to provide UDP-glucose  A version of this chapter has been published. Coleman HD, Ellis D, Gilbert M, Mansfield SD (2006). Up-regulation of sucrose synthase and UDP-glucose pyrophosphorylase impacts plant growth and metabolism. Plant Biotechnology Journal. 4: 87-101.  31  directly to the cellulose synthase complex (Amor et a!., 1995) through metabolic channelling. Direct channelling would facilitate the recycling of UDP into UDP-glucose, as well as ensure the availability of UDP-glucose to the cellulose synthase complex despite demands from other pathways. Salnikov et a!. (2001) showed that during secondary wall formation in Zinnea elegans, SuSy is strongly recruited to the plasma membrane and is highly enriched beneath the secondary cell wall thickenings of differentiating tracheary elements. These findings support the putative relationship between SuSy and cellulose biosynthesis. SuSy activity has also been positively correlated with sink strength in storage organs of potatoes, maize kernels, and pea embryos (Dejardin et a!., 1999; Sun eta!., 1992; Zrenner et a!., 1995). UGPase in contrast, utilizes the hexose phosphate pool and, in sink tissues where it is predominantly found, is considered a key enzyme for carbohydrate biosynthesis. UGPase can therefore serve as an extension of SuSy in cycling between sucrose and the hexose phosphate pool (Kleczkowski, 1994). Furthermore, UGPase plays an integral role in sugar accumulation, and has the potential to restrict the flow of carbon to sucrose formation in sink tissues (Borokov, et al., 1996). In source tissues, UGPase works in conjunction with SPS, in the synthesis of sucrose (Kleczkowski, 1994). The apparent dual roles of UGPase in sink and source tissue makes UGPase an interesting target for the production of cellulose in sink tissues. By increasing the generation of sucrose in source tissues, while decreasing sucrose formation in sink tissues, there is the potential to selectively channel sucrose to sink tissue for use in carbohydrate biosynthesis. Despite what is currently known about these two enzymes, few studies have been conducted to elucidate their role in sucrose metabolism and carbon partitioning to cellulose biosynthesis in plants. The purpose of this study was to investigate the effect of overexpressing SuSy and UGPase individually, and in combination, on cellulose synthesis and related pathways to determine the potential role of each enzyme in carbon allocation.  Both genes were expressed under the control of a constitutive promoter, the double cauliflower mosaic virus (2x35S), and a vascular specific promoter, the parsley 4-Coumarate:CoA ligase (4CL; Hauffe et a!., 1991), in tobacco. Resultant analyses included quantifying soluble and structural carbohydrates and biomass accumulation. Although the results showed little increase in cellulose content 32  as a percentage of total mass, the upregulation of these genes caused dramatic variation in height growth and sucrose metabolism, thus affecting the total mass of the plant and therefore the overall amount of cellulose produced.  Methods Cloning of UGPase and Plasmid Construction UGPase was cloned from Acetobacter xylinum ATCC #23768 and inserted into the pBlN cloning vector with one of two promoters: the enhanced tandem CaMV35S constitutive promoter (2x35S) (Kay, et a!., 1987), or the vascular specific 4CL (Petroselinum crispum 4-Coumarate:CoA ligase) promoter (Hauffe, eta!., 1991). SuSy was cloned from Gossypium hirsutum (Perez-Grau, Genbank U73588) and inserted into pBIN under the control of the same two promoters. This plasmid also contained the NPTII gene under the control of the CaMV35S promoter.  Plant Transformation and Maintenance Nicotiana tabacum cv Xanthi (tobacco) was transformed using Agrobacterium tumefaciens EHAIO5 (Hood, et a!., 1993) employing a standard leaf disk inoculation method. Binary plasmids were inserted into EHAIO5 using the freeze-thaw technique, and incubated overnight in liquid Murashige and Skoog media (Murashige and Skoog, 1962) supplemented with 3% sucrose (MS+3%) and I OOpM acetosyringone. Leaf disks were cut and co-cultured with EHAIO5 for one hour at room temperature, blotted dry and plated abaxailly onto MS+3% supplemented with 0.lpM each a-naphthalene acetic acid (NAA) and 6-benzylaminopurine (BA) and solidified with 3% (w/v) agar and 1.1% (w/v) phytagel (MS+NAA/BA). After three days the discs were transferred to MS+NAA/BA supplemented with carbenicillin disodium (500 mg L ) and cefotaxime 1 sodium salt (250 mg L ). Following three additional days of selective growth, the discs 1 were transferred to MS NAA/BA containing carbenicillin, cefotaxime and kanamycin (25 mg L ). After two consecutive five-week periods on this media, shoot tips were isolated 1 to solid MS+3% containing no antibiotics. 33  Plants were confirmed as transgenic by PCR screening of genomic DNA employing gene specific oligonucleotides: specifically, for UGPase UGP-1 (5’atcgaggaattctgcctcgt-3’) and UGP-2 (5’-tcgcaagaccggcaacaggatt-3’) where used, while for SuSy SUS-1 (5’-ctcaacatcacccctcgaat-3’) and S US-2 (5’-accaggggaaacaatgttga-3’) were employed.  Genomic DNA was isolated using the Red Extract and Amp Kit  (Qiagen). All shoot cultures, including transgenic and non-transformed control lines, were maintained on solid MS+3% in GA-7 vessels at 22°C under a 16 hour photoperiod with an average photon flux of 40 pmol m 2 s . 1  Plants were maintained by transferring  apical regions at four-week intervals.  Plant Growth Primary transformed plants and control lines (T ) were grown to maturity and 0 selfed to generate T 1 lines of all the single transformants and the associated controls. The pods from each controlled cross were collected, and the seeds removed and sterilized by washing for 2 minutes in a 10% bleach solution, followed by a 1 minute rinse in sterile water. The seeds were washed in 70% ethanol for two additional minutes and then in sterile water for three consecutive 1 minute washes. Seeds were germinated on solid % MS with 2% sucrose and kanamycin (25 mg L ). The surviving 1 seedlings were then PCR screened using the aforementioned primer sets. Seedlings were grown in GA-7 vessels prior to transfer into 7.5 L pots containing a 50% peat, 25% fine bark, 25% pumice soil mixture in the greenhouse, and covered with l6oz clear plastic cups for one week to aid in acclimation. Each line, transgenic and control, was represented by twelve individual plants (each from an individually selected seed). The greenhouse plants were harvested at the onset of flowering as indicated by the formation of flower buds.  Plant height, from base to tip of the highest bud, was  measured prior to harvest.  Developmental stages of tissues were standardized by employing a plastichron index (P1), where leaf plastichron index P1=0 was defined as the first leaf greater than 5cm in length, and where P1=1 is the leaf immediately below P1=0. A portion of the stem from each plant spanning P1=5 to P1=15 was excised and immediately weighed for total stem fresh weight measurements and leaf biomass. This 34  same section was then dried at 1O5C for 48 hours for dry weight determination, and retained for further analysis. Internode distance represents the average length between each internodes spanning P1 =5 to P1 =15. The lower section of the stem (below P1=15) was dried at room temperature for fibre quality analysis. All data were analysed using single variable ANOVA and Scheffe tests.  Production of Double Transformants 1 tobacco plants transformed with only one construct (single transgene plants) T were grown to maturity in the greenhouse and selectively crossed with transformed lines expressing another construct to produce reciprocal double transgene plants. Double transgene plants are therefore defined as plants derived from the progeny of a controlled cross between a single SuSy transgene plant and a single UGPase transgene plant (under the regulation of the same promoter).  The pods from each  controlled cross were collected, and the seed removed and sterilized as described previously. The seeds were germinated on  1/2  MS with 2% sucrose and kanamycin (25  mg L ). Transformants were confirmed to contain both transgenes via PCR, using the 1 same primers as used for single transformant confirmation.  Transcription Levels Real time PCR was used to determine the transcript level of each transgene. Leaf and stem sections weighing approximately 100 mg were ground in liquid nitrogen, and RNA extracted using Trizol reagent (Gibco BRL, Gaithersburg, MD) according to manufacturers’ instructions. Following extraction, 10 pg of total RNA was treated with 10 units of DNase I (Fermentas, Burlington, ON, Canada) in 6 mM MgCl. The reaction was incubated at 37°C for 30 minutes and then heat inactivated at 80°C for 10 minutes. Following the addition of one volume of phenol:chloroform:isoamyl alcohol (25:24:1), the sample was briefly vortexed, and then centrifuged at 13,000 rpm for 5 minutes at 4°C. The upper phase was transferred to a fresh tube and 10 pL of 3 M sodium acetate and 200 pL of 100% ethanol added. The samples were incubated at  -  80°C for one hour and then centrifuged at 13,000 rpm for 15 minutes at 4°C. The supernatant was removed and the RNA pellet resuspended in 500 pL of 70% ethanol. 35  The sample was centrifuged at 13,000 rpm for 10 minutes at 4°C, the supernatant carefully removed and the pellet resuspended in 40 pL of RNase-free water. Equal quantities of RNA (1 pg) were employed for the synthesis of cDNA using Superscript II Reverse Transcriptase (Invitrogen, Carlsbad, CA) and dT 16 primers according to manufacturer’s instructions. Samples were run in triplicate with Brilliant SYBR Green QPCR Master Mix (Stratagene, La Jolla, CA) on an Mx3000P Real-Time PCR System (Stratagene). The primers for the RT-PCR analysis of UGPase were AU RTF (5’-tggaagcaacccgcgtcatc-3’) and AU-RTR (5’-gccaaggcccagcggttcc-3’) and for SuSy  were  GS-RTF  (5’-ccgtgagcgtttggatgagac-3’)  and  GS-RTR  (5’-  ggccaaaatctcgttcctgtg- 3’). Conditions for the RT-PCR reactions were as follows: 95°C for 10 minutes, followed by 40 cycles of 95°C for 30 seconds, 62°C (64°C) for 1 minute, and 72°C for 30 seconds. Transcript levels (transcript copy number ug 1 total RNA) were based on standard curves derived from known concentrations of plasmid DNA run under the same conditions.  Enzyme Activity Leaf and stem samples (approximately 1 g) were ground in liquid nitrogen with I mg of insoluble PVPP and four volumes of extraction buffer (50 mM HEPES-KOH pH 7.5, 10 mM MgCl, 1 mM EDTA, 2 mM DTT, 1 mM PMSF, 5 mM EAmino-n-caproic acid, 0.1% v/v Triton X-100, 10% v/v glycerol). The samples were centrifuged at 14,000 rpm for 20 minutes at 4°C. The extract was passed through a DG 10 desalting column (BioRad) pre-equilibrated with ice-cold extraction buffer without Triton X-1 00 and PVPP. Extracts were collected into pre-chilled vials and employed immediately. SuSy activity was assayed in the direction of sucrose breakdown, as described in Chourey (1981), using 50 jiL of extract.  The resultant fructose content was determined using a  tetrazolium blue assay (Kennedy & White, 1983). This SuSy assay employs the appropriate controls without the supplementation of UDP to quantify inherent invertase activity, and therefore represents the breakdown of sucrose by SuSy. UGPase activity was determined as per Appeldoorn et a!. (1997) using 100 jiL of extract, and employing UGPase (Sigma) to develop a standard curve.  36  Soluble Carbohydrate and Starch Analysis  Soluble carbohydrates (glucose, fructose and sucrose) were extracted from ground freeze-dried plant material using methanol:chloroform:water (12:5:3) in a -20CC overnight incubation. The sample was centrifuged and the supernatant removed. The remaining pellet was washed twice with fresh methanol:chloroform:water (12:5:3) and all fractions pooled. Five mL of water was added to the combined supernatant and centrifuged to facilitate phase separation.  The aqueous fraction was removed to a  round bottom flask and rotary evaporated to dryness. The sample was resuspended in 3 mL of distilled water and analyzed using anion exchange HPLC (Dionex, Sunnyvale, CA) on a DX-600 equipped with a Carbopac PA2O column and an electrochemical detector. The residual pellet was hydrolyzed using 4% sulfuric acid at 121 C for 4 minutes. The liberation of glucose represented starch content, and was directly quantified by HPLC employing similar conditions.  Determination of Cellulose and Holocellulose Content  Dried plant stem material was ground using a Wiley mill to pass through a 30mesh screen, and then soxhlet-extracted with acetone for 24 hours. The extractive free material was used for all further analyses.  Holocellulose and a-cellulose were  determined using a modified microanalytical method developed by (Yokoyama et a!., 2002). In short, 200 mg of ground sample was weighed into a 25 mL round bottom flask and placed in a 90°C oil bath. The reaction was initiated by the addition of I mL of sodium chlorite solution (400 mg 80% sodium chlorite, 4 mL distilled water, 0.4 mL acetic acid). An additional 1 mL of sodium chlorite solution was added every half hour and the samples removed to a cold water bath after two hours. Samples were then filtered through a coarse crucible, dried overnight, and holocellulose composition determined gravimetrically. Fifty mg of this dried holocellulose sample was weighed into a reaction flask and allowed to equilibrate for 30 minutes.  Four mL of 17.5%  sodium hydroxide was added and allowed to react for 30 minutes, after which 4 mL of distilled water was added. The sample was macerated for 1 minute, allowed to react for an additional 29 minutes and then filtered through a coarse filter.  Following a five 37  minute soak in 1.0 M acetic acid, the sample was washed with 90 mL of distilled water and dried overnight. The a-cellulose content was then determined gravimetrically.  Fibre Quality Analysis  To determine the fibre length and coarseness, a portion of the lower stem of tobacco plants was cut into representative samples of approximate dimensions of 2 mm x  2 mm x 30 mm, and reacted in Franklin solution (1:1 30% peroxide and glacial acetic  acid) with 3.6% sodium hypochlorite for 24 hours at 70°C. The solution was decanted and the remaining fibrous material was reacted in pure Franklin solution for an additional 48 hours at 70°C. The solution was decanted and the remaining material washed under vacuum with deionized water until a neutral pH was achieved.  The  samples were dried overnight at 105°C, and then resuspended in 10 mL of deionized water. Further dilution was used to obtain a count of 25-40 fibres per second on a Fibre Quality Analyzer (FQA). All samples were run in triplicate.  Microscopy  Tobacco stems were hand-sectioned using a double-edged razor blade. The samples were then fixed in FAA (formalin, acetic acid, alcohol), dehydrated through as series of acetone and xylene exchanges, and embedded in paraffin. Sections were cut to 10pm and mounted on glass slides, and images taken on a Leica microscope with a Q-imaging camera.  Results Single Transformants UGPase from Acetobacter xylinum and Gossypium hirsutum SuSy were inserted into the pBIN cloning vector under the regulation of one of two promoters: 2x355 and 4CL. Tobacco was transformed with one of the four resulting constructs (2x35S::SuSy, 2 x35S: : UGPase, 4CL: :SuSy, or 4CL:: UGPase) using an Agrobacterium-mediated transformation technology.  The single transgene plants were PCR-confirmed as 38  transgenic, grown to maturity in a greenhouse, and the flowers bagged to produce 1 seed. T selfed-T 1 seed was then germinated on % MS with kanamycin (25 mg L ) to 1 produce T 1 single transgene plants which were grown in the greenhouse until the onset of flower buds, measured, and a portion of the stem retained for further analysis.  Biomass Accumulation At the onset of flower bud formation, all plants appeared phenotypically normal and displayed no observable pleiotropic effects. Of the lines analyzed, plants from at 1 single trangene lines from each of the four constructs had statistically least two T significant increases in height growth compared to the corresponding control lines (Figure 2.IA). There was also an increase in internode distance for all lines, however, at a a-value of 0.05 this difference was only significant in one line, 4CL::SuSy-3, while three additional lines (4CL::UGPase-4, 4CL::UGPase-5 4CL::SuSy-8) were significant at an a-value of 0.10 (Figure 2.IB). Despite the increased height growth and internode length, only one line, 4CL::UGPase-4, had a significant increase in total dry weight of the stem (Figure 2.1C).  Transcription Levels Although transcript levels within single transgene lines were relatively constant, there was wide variation in transcript levels between lines transformed with either the UGPase or SuSy construct. In general, the transcript levels of genes under the regulation of the 4CL promoter were lower in the leaves but similar in the stems to that of the 2x35S plants, which is consistent with expectations for relative expression of the two promoters. Interestingly, 2x35S::UGPase-8 showed very low transcript levels while the other three 2x35S::UGPase lines displayed the highest transcript levels. Additionally, UGPase transcript levels were up to 10-fold greater than that of SuSy under the regulation of either promoter (Table 2.1). Promoter activity also varied with tissue source (stem vs leaves) in the single transgene lines. In six of the seven lines evaluated, the transcript abundance in leaves of both transgenes under the regulation of the 2x35S promoter was equal to, or greater, than levels in stems. However, in line 2x35S::SuSy-17 transcript levels were reduced in 39  -  o  2.0  o  c  -‘  -  -o  r  -ri  II  Q  °  1•11  -&  -h  ci a’—  —.  II  —  -  •0  D 0 aD  *  ‘  o a —-2-s.  D  0 B  DW CD 0  =  __3  CD C’.)  ••‘  I D 0 0) H—  D D 0  —  CD CD 0—•—-i D 0 D 0 DO  a  ODD  —.  •  o 0 DCC  a  0  .  a  —l  I T I  -i  I IIIIJlIIIIllIItfllI—’  iiiiiiiiiminiiiiiviiiiiiiiijiiiiiw  JIIIPtIIIIIIIIIIIIlIIIIllIl  IIIuIpIIIIIIIlIIIqiiIiiI  //////////p4_,  I  I  \\•.•.•.‘.•.I.I  Dry Weight (g)  a o W  ‘  ] )  lIIIIIIIIIIIllIIIlllIIllIIIIPIIIIIIIIIIIIlliNtljf—  IIIIIIIllIIIlIIIIIIIIPfI-’ a  I  “‘  )IIIIIIIIIIIIIUIIIIIIHIIUIIIIIIIJIIIIIW+wt—’.  ]iiiiiiiiipiiiiiiiiiiiiiit—ii————*  ] 1111111111111 liii rim,_________ 0  *  *  ‘///////7//////////////////, *  Dliiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiitttt—’.  *  flhiiiwiwnwtwiwiiwijwii I  ________________  : ////////////////4—  (1111111111111111114111111  a  —  -  ]iiiiiiiiiiiiiiit+t+t—’  ‘ci  6)  C  9WI  ///h9—  Height (cm)  W/////////////AØZ1  —  -J  4,\  g  j•  )2  t  C  1.I.I...  t  Internode Distance (cm)  Table 2.1. Transcript level and enzyme activity in leaf and stem tissue for single transgene and non-transformed control plants. Mean and SE calculated from 3 plants per line. nd = no transcripts detected  UGPase Transcript Level x10 3 copy number pg 1 total RNA Line  Leaf  UGPase Enzyme Activity Units g 1 fresh weight  Stem  Leaf  Mean  SE  Mean  SE  Mean  2x35S::UGPase 7  1946.5  591.5  1254.0  466.3  0.304  2x35S::UGPase 8  6.0  0.8  2.9  0.9  2x35S::UGPase 10  1729.0  268.2  535.7  2x35S::UGPase 12  1834.6  42.1  4CL::UGPase4  195.8  4CL::UGPase 5 4CL::UGPase 10 Control  Stem SE  Mean  SE  0.227  0.253  0.150  0.175  0.109  0.119  0.087  221.3  0.086  0.059  0.012  0.051  382.5  61.3  0.012  0.010  0.043  0.021  37.1  1116.4  172.5  0.031  0.002  0.232  0.063  44.5  29.6  244.2  21.8  0.034  0.003  0.153  0.087  68.8  53.6  153.5  36.2  0.042  0.001  0.007  0.009  0.081  0.063  0.133  0.090  nd  -  nd  -  SuSy Transcript Level xl copy number Jg 1 total RNA Line  Leaf  SuSy Enzyme Activity ig fructose g 1 fresh weight  Stem  Leaf  Mean  SE  Mean  SE  Mean  2x35S::SuSy9  248.6  46.1  8.7  4.8  0.010  2x35S::SuSy 13 2x35S::SuSy 17  309.1  96.2  315.8  179.4  175.1  40.3  315.4  4CL::SuSy3  nd  4CL::SuSy6  nd  4CL::SuSy 8  4.1  Control  nd  -  -  3.3 -  Stem SE  Mean  SE  0.000  0.010  0.002  0.010  0.002  0.010  0.001  128.3  0.013  0.003  0.011  0.001  65.1  0.0  0.010  0.001  0.010  0.001  312.8  119.6  0.009  0.000  0.009  0.000  83.0  35.7  0.008  0.001  0.009  0,001  0.009  0.001  0.010  0.001  nd  -  41  the leaves relative to the stems, although the difference was not significant. In contrast, the transcript levels of the 4CL promoter were consistently higher in the stems, although the difference between leaf and stem tissue in two of the 4CL::UGPase lines was not significant. Interestingly, no transcripts were detected in the leaves of two 4CL::SuSy lines, while all 4CL::UGPase lines had relatively high UGPase transcripts levels in the leaves.  Enzyme Activity UGPase activity was determined using an indirect assay measuring the production of NADH.  In the 2x35S::UGPase lines, the mean UGPase activity was  generally increased in leaf material (Table 2.1), however, this increase was not significant. Two out of three 4CL::UGPase lines had decreased UGPase activity in the leaves compared to the controls, while one line (4CL::UGPase-1O) showed an increase in UGPase activity in the leaf relative to the stem. UGPase activity in the 4CL::SuSy lines were similar to that of the control plants (data not shown). SuSy activity was determined using a direct assay that quantified the production of fructose from the catalysis of sucrose, and considered (subtracted) any inherent invertase activity in the tissue. Single SuSy transgene plants showed slight increases in SuSy activity compared to controls. In contrast to UGPase activity in UGPase single transformed lines, SuSy activity in the single transgene lines, also did not show a tissue-specific response between the leaves and stems, regardless of the promoter used (Table 2.1).  Soluble Carbohydrates In general, all single transgene lines had increased total soluble sugar content. This increase in total sugar content was not due to increases in sucrose, as no significant differences in sucrose levels between the controls and single transgenic lines were detected.  However, as expected with the overexpression of genes encoding  enzymes that degrade sucrose, significant increases in both glucose and fructose content were evident (Table 2.2).  Further, in the control plants, soluble sucrose  concentrations were higher than either glucose or fructose, while in all transformed lines 42  1 dry weight tissue) in Table 2.2. Concentration of total soluble carbohydrates (big mg stem tissue of single transgene and non-transformed control tobacco plants. Mean and SE calculated from 4 plants per line. Bold indicates significance at a=O.05.  Glucose  Line  Fructose  Sucrose  Total  SE  Mean  SE  Mean  SE  Mean  SE  7.57  2.97  5.52  2.75  6.23  0.74  19.32  6.36  2x35S::UGPase 8  7.19  1.93  6.95  3.03  4.74  0.19  18.88  4.77  2x35S::UGPase 10  6.20  1.42  6.21  1.37  4.24  0.86  16.64  3.59  2x35S::UGPase 12  6.10  2.11  6.38  2.38  3.59  0.60  16.07  4.91  4CL::UGPase4  7.02  1.85  9.01  3.64  3.83  0.78  19.86  6.27  7.99  0.34  10.05  1.14  5.08  0.24  23.12  1.24  4CL::UGPase 10  7.28  1.40  8.01  2.18  4.50  0.72  19.79  4.30  2x35S::SuSy9  6.32  0.55  7.54  0.48  4.43  0.86  18.30  1.88  2x35S::SuSy 13  6.52  0.28  7.93  0.21  4.16  0.34  18.62  0.43  2x35S::SuSy 17  6.91  0.32  8.43  0.58  5.10  0.42  20.43  0.69  4CL::SuSy3  6.33  0.26  6.36  0.78  5.05  1.17  17.74  0.14  4CL::SuSy6  4.47  0.59  5.93  1.17  4.61  0.22  15.00  1.79  4CL::SuSy8  4.20  1.00  4.10  0.70  3.53  0.56  11.83  1.98  Control  3.48  1.01  3.51  0.68  4.49  1.19  11.48  1.99  Mean 2x35S::UGPase 7  4CL::UGPase5  -  43  glucose and fructose levels were higher than sucrose levels. Furthermore, while the concentration of glucose and fructose in control plants were roughly equal, in both the 4CL::UGPase and 2x35S::SuSy lines there were higher fructose levels relative to glucose content.  Polymeric Carbohydrates No significant differences in starch content were detected between control plants and single transgene plants (Figure 2.2A). Similarly, with the exception of the one line, 2x35S::SuSy-9, there were also no significant differences in cellulose content between control plants and single transgene plants (Figure 2.2B).  Fibre Quality Analysis In the UGPase-expressing plants, no significant differences in fibre length and coarseness were detected; however, fibre length was generally decreased while fibre coarseness generally increased (Figures 2.2C & D). Similarly, two SuSy-expressing lines, 2x35S::SuSy-9 and 4CL::SuSy-6, had significantly shorter fibres relative to the controls, but were not different in terms of coarseness.  Double Transgene Plants Biomass Accumulation Single transgene plants were grown in the greenhouse through flowering and reciprocal crosses were made between 2x35S::UGPase and 2x35S::SuSy single transgene plants, and 4CL::UGPase and 4CL::SuSy single transgene plants. The double transgene progeny seed was collected, planted on selective media, and kanamycin resistant offspring were selected and confirmed for the presence of both genes using genomic PCR screening. The double transgene tobacco seedlings were then planted in the greenhouse and allowed to reach reproductive maturity prior to growth analysis and destructive harvesting.  44  26  A  2.4  .‘  B  24  2,0 i.a —  8,1.6 0  0 14 1.2 0 1.0 D.C  HI C1r17  8  10 12 4  22 C C  18 16 0 5  10 9  13 17 36  J in  oo1._ NNNN 000 Clii 7 8 10 12 4 5 10  8  2x35S:UGPase 4CL:UGPase 2x35S:SuSy 4CLSuSy  q 9  Fl 13 17  00 3  8  8  2x35S:UGPas6 4C1:UGPase 2x35S:SuSy 4CL:SuSy  C  D  0.12 0.11  E  6  (0  0.10  0.8  T  T  C  6 U-  N  8,  0.09 0 8)  .0 0.08 U-  0 1 P N N N N N H N H H P 0 0 Hi ci 7  8  10 12 4  5  10  9  13 17  3  6  8  NNNN ONO C1r17 8 10 12 4 5 10  PP 9 13 17  3  6  8  2x35S:UGPe 4CLUGPe 2x35S:SuSy 4CIS,Sy 2x35S:UGPase 4CL:UGPase 2x35S:SuSy 4CL:SuSy  Figure 2.2. Starch (A) and cellulose (B) content, and fibre length (C) and coarseness (D) of single transgene lines and non-transformed control line. Starch and cellulose mean ±SE calculated from 5 plants per line. Fibre quality mean ±SE calculated from 4 plants per transgene line and 12 plants per non-transformed control. *indicates significance at a=O.05. ** indicates significance at a=O.1O.  45  *  A  *  120  E 110  100  c1’’, Clii 1 2 3 4 5 6 9 10 11  .  12 13 14  4CL:UGP8SC/SuSy  2x35S:UGPOSe1SuSy  B 5.0 *  E HflHH Cliii  234569101121314  2x35S:UGPase/SuSy  4CL:UGPase/Susy  C 3.0  2.5 a) 2.0  :: hdh o:oi  Clii  I  2  3  4  5  6  2x35S:UGPase/SuSy  9  10 11  12  13  14  4CL:UGPase/SuSy  Figure 2.3. Plant height (A), internode length (P1 = 5 to P1 = 15) (B) and total plant biomass (C) in double transgene tobacco lines. Mean ±SE were calculated from 12 plants per transgenic line and 40 plants in the non-transformed control line. * indicates significance at a=0.05. ** indicates significance at a=0.10.  46  All of the double transgene lines showed statistically significant increases in height growth, when compared to the corresponding non-transformed control plants (Figure 2.3A).  There was also a significant increase in internode length in many (2x35S::UGPase/SuSy-2 and 4CL::UGPase/SuSy-1 1, 4CL::UGPase/SuSy-1 2, and  4CL::UGPase/SuSy-13) of the double transgene lines evaluated, with all other double transgene lines showing increases in absolute values (Figure 2.3B). In addition, plant biomass was clearly increased (Figure 2.3C) in the double transgene lines relative to the non-transformed control, particularly in the lines under the control of the 4CL promoter, in which half of the lines (4CL::UGPase/SuSy-9, 4CL::UGPase/SuSy-1 1, and 4CL::UGPaseISuSy-12) showed significant increases in biomass. One line (4CL::UGPase/SuSy-1 1) had over 2 times the biomass of the control line.  Transcription Levels Similar to single transgene lines, there was wide variation in both UGPase and SuSy transcript levels in the lines expressing two transgenes.  Transcription levels  were, however, correlated with transcript levels of the parental lines. For example, 2x35S::UGPase/SuSy-1 and 2x355::UGPase/SuSy-2 are reciprocal crosses and had similar relatively low transcript levels for both genes compared to the other 2x35S double transgene lines (Table 2.3). The transcript level detected in the stem of these lines correlates well to the low transcript levels detected in the stem of associated parental lines, 2x35S::SuSy-9 and 2x35S::UGPase-8, both of which had very low transcript levels. In general, the transcript levels of the double transgene lines were slightly lower than their corresponding parental lines. Similar to single transformed lines, transcript levels in 4CL-regulated double transformed lines were significantly higher in the stems than in the leaves and 4CL transcript levels were generally lower than with the 2x35S promoter.  Additionally,  UGPase transcript abundance was shown to be up to 200-fold that of SuSy expression.  Enzyme Activity The 2x35S double transformed lines showed a consistent increase in UGPase activity in the leaf tissue, with one line (2x35S::UGPase/SuSy-6) having a significant 47  Table 2.3. Transcript level and enzyme activity of UGPase and SuSy in leaf and stem tissue of double transgene and non-transformed control tobacco plants. Mean and SE calculated from 3 plants per line. Bold indicates significance at a=O.05. nd = no transcripts detected. UGPase Transcript Level x10 3 copy number lJg 1 total RNA Line  Leaf  UGPase Enzyme Activity Units g 1 fresh weight  Stem  Leaf  Stem  Mean  SE  Mean  SE  Mean  SE  Mean  SE  2x35S::UGPase/SuSy-1  10.8  6.6  8.3  12.6  0.329  0.270  0.142  0.198  2x35S::UGPaseISuSy-2  2.1  2.6  20.2  8.1  0.208  0.201  0.195  0.227  2x35S::UGPase/SuSy-3  4126.5  1818.6  3695.6  1888.9  0.330  0.277  0.170  0.189  2x35S::UGPaseISuSy-4  756.7  402.9  843.5  115.7  0.204  0.202  0.035  0.079  2x35S::UGPase/SuSy-5  506.4  136.4  566.0  328.8  0.338  0.349  0.230  0.079  2x35S::UGPaseISuSy-6  693.1  479.4  969.1  619.8  0.461  0.040  0.346  0.127  4CL::UGPase/SuSy-9  15.7  3.1  402.9  67.9  0.291  0.063  0.186  0.029  4CL::UGPase/SuSy-1O  10.1  11.1  220.5  124.3  0.361  0.015  0.297  0.109  4CL::UGPase/SuSy-11  4.5  1.0  124.9  39.4  0.081  0.084  0.203  0.036  4CL::UGPase/SuSy-12  30.4  25.4  193.5  172.1  0.233  0.042  0.116  0.080  4CL::UGPase/SuSy-13  29.9  19.9  228.5  41.9  0.371  0.021  0.292  0.055  4CL::UGPase/SuSy-14  14.8  9.9  233.5  61.1  0.606  0.246  0.450  0.020  0.081  0.063  0.133  0.090  Control  nd  -  nd  -  SuSy Transcript Level xl copy number pg 1 total RNA Line 2x35S::UGPase/SuSy-1  Leaf  SuSy Enzyme Activity pg fructose g 1 fresh weight  Stem  Mean  SE  Mean  48.2  35.5  nd  Leaf SE -  Stem  Mean  SE  Mean  SE  0.016  0.006  0.010  0.002  2x35S::UGPase/SuSy-2  89.8  39.4  57.1  19.4  0.016  0.002  0.013  0.001  2x35S::UGPase/SuSy-3  529.8  144.1  1210.6  339.0  0.022  0.005  0.009  0.005  2x35S::UGPase/SuSy-4  194.2  94.9  183.4  92.4  0.015  0.001  0.011  0.004  2x355::UGPase/SuSy-5  316.4  38.2  158.4  76.8  0.013  0.002  0.013  0.002  2x35S::UGPase/SuSy-6  326.6  60.6  139.4  100.5  0.010  0.000  0.013  0.004  4CL::UGPase/SuSy-9  21.1  21.1  70.8  5.0  0.009  0.010  0.013  0.010  4CL::UGPase/SuSy-10  nd  46.7  14.4  0.009  0.008  0.009  0.009  4CL::UGPase/SuSy-11  nd  36.2  24.5  0.009  0.009  0.009  0.008  4CL::UGPase/SuSy-12  3.7  23.1  9.0  0.007  0.008  0.007  0.003  4CL::UGPase/SuSy-13  nd  13.4  13.4  0.008  0.009  0.010  0.008  4CL::UGPase/SuSy-14  3.8  32.5  21.5  0.011  0.007  0.007  0.007  Control  nd  0.009  0.001  0.010  0.001  -  -  3.1 -  3.8 -  nd  -  48  increase in UGPase activity relative to the control (Table 2.3). However, no significant differences were seen in UGPase activity in the stem of the 2x35S::UGPase/SuSy lines. Interestingly, despite the desire to have a xylem directed expression pattern with 4CL, four of the six 4CL::UGPase/SuSy lines had significantly elevated UGPase activity in the leaves, while only one of the six lines had significantly higher activity in the stems. In fact, line 4CL::UGPase/SuSy-14 had the highest UGPase activity detected and this activity was in the leaves, not the stems. The UGPase activity of this transformed line was also the highest detected in the stem as well. SuSy activity in double transgene plants was substantially increased in a number of lines relative to the non-transformed control (Table 2.3). The leaves of the 2x35S double transgene lines had a large increase in SuSy activity with half the lines (three out of six lines) showing a significant difference from the controls. In contrast to the leaves, the SuSy activity in the stem tissue remained relatively consistent with that of the control plants with only one line (2x35S::UGPase/SuSy-2) having a significant increase relative to the control. The leaves and stems of the 4CL lines did not show the same increase in SuSy activity as observed in the 2x35S lines.  Soluble Carbohydrates While no significant differences in total soluble sugar content were detected in the 2x35S double transformed plants, five of the six 4CL double transgene lines had significant increases (cc<O.1O) in total soluble sugars (Table 2.4). Similar to the single transgene lines, this increase in the 4CL lines was attributed to elevated glucose and fructose concentrations and not elevated sucrose levels except in one line (4CL::UGPase/SuSy-14) where total sugars as well as glucose and fructose were significantly increased. While only 3/6 and 2/6 2x35S double transformed lines had elevated glucose and fructose levels respectively, 6/6 and 5/6 4CL double transformed lines had elevated glucose and fructose levels, respectively.  Polymeric Carbohydrates No significant differences compared to the controls were detected in either starch or cellulose content (Figure 2.4A & B). Three of the five 2x35S lines and one 4CL line, 49  Table 2.4. Concentration of total soluble carbohydrates (.tg mg 1 dry weight tissue) in stem tissue of double transgene and non-transformed control tobacco plants. Mean and SE calculated from 5 plants per line. Bold indicates significance at a0.1O.  Line  Glucose  Fructose  Sucrose  Total  Mean  SE  Mean  SE  Mean  SE  Mean  SE  2x35S::UGPase/SuSy-1  6.00  1.06  5.48  0.97  3.57  0.46  15.04  2.38  2x35S::UGPase/SuSy-2  5.20  0.56  5.03  0.51  3.83  0.39  14.06  1.34  2x35S::UGPase/SuSy-3  6.60  0.30  5.98  0.25  3.66  0.35  16.24  0.37  2x35S::UGPase/SuSy-4  5.16  1.94  4.45  1.66  4.81  0.26  14.43  3.63  2x35S::UGPasefSuSy-5  2.86  0.76  2.83  0.84  3.15  0.26  8.84  1.64  2x35S::UGPase/SuSy-6  4.44  1.04  4.36  1.00  3.26  0.42  12.07  2.32  4CL::UGPase/SuSy-9  5.42  0.71  4.89  0.68  4.83  0.50  15.14  1.83  4CL::UGPase/SuSy-10  6.67  0.65  5.68  0.70  5.45  0.89  17.80  1.08  4CL::UGPase/SuSy-11  7.83  0.73  6.79  0.81  4.84  0.49  19.46  1.66  4CL::UGPase/SuSy-12  7.36  0.87  6.39  0.81  5.39  0.36  19.13  1.77  4CL::UGPase/SuSy-13  7.45  0.91  6.07  0.68  5.03  0.35  18.55  1.75  4CL::UGPase/SuSy-14  10.35  0.74  7.56  0.84  8.35  0.95  26.26  1.66  Control  3.48  1.01  3.51  0.68  4.49  1.19  11.48  1.99  50  26  H mu i  .2’ 2.6  2 ;1.8  02 C C  1.6 1.4  0.01  .2’ C 2  ‘22  3  2  4  5  6  9  10 II  2x35S1JGPel2o5y  H Ub  20  I1  E o  o w  16  ‘6 0  o  MM1MM 061 1  B  24  16  VA 01  N N N ‘(V/A  ON ON ON ON ON ON  051  12 13 14  1  2  3  4  5  6  Osl 01 01 01 09 09f 10 ii 12 63 14  9  2x35NJGPaseISoSy  4CL:UGPe/SuSy  4CL:UGPase/SuSy  0.9  2 2  2  S 08  :  -J  2  C  9-  0  0  a  2  a  0.0  NI  051  1  2  3  4  5  2x350:UWaseISuSy  6  h\l]l.  9  1011  1213 14  4CL:UGPase/SuSy  0.i I  o.ooMNNNNNNN0N1 Cut  I  2  3  4  5  2x350:UGPaseiSuSy  6  9  10 11  12 13  14  4CLUGPase/SuOy  Figure 2.4. Starch (A) and cellulose (B) content, and fibre length (C) and coarseness (D) of stem tissue of double transgene lines and non-transformed control line. Starch and cellulose mean ±SE calculated from 5 plants per line. Fibre qualities mean ±SE calculated from 4 plants per transgene line and 12 plants per non-transformed control.  51  \  I.  -  Figure 2.5. Light and UV-fluorescence microscopy of control (A) and transgenic (4CL::UGPasexSuSy 12) (B) tobacco. Arrows depict the observed increased cell wall thickening in transgenic lines.  52  4CL::UGPase/SuSy-1 1, showed an increase in cellulose content.  However, the  remaining 4CL lines had reduced cellulose content (Figure 2.4B).  Fibre Quality Analysis As with the single transgene lines, fibres were consistently shorter and had increased cell wall coarseness in the double transformed lines relative to the controls, although these differences were significant in only one line (Figures 2.4C & D).  Discussion This study investigated the effects of employing two promoters, a tissue-specific (4CL) and a ubiquitously expressed (2x35S) promoter, to overexpress exogenous SuSy and UGPase genes in tobacco. As expected, the expression profiles of the genes and enzyme activity varied amongst the four constructs (2x35::UGPase, 4CL::UGPase, 2X35::SuSy, 4CL::SuSy). The highest expression in leaf tissue was observed in single transgene plants under the control of the 2x35S promoter, where expression was equal to or higher than in the stems. In contrast, expression controlled by the 4CL promoter was generally higher in the stems. This is consistent with previous work which showed 4CL to be principally expressed in the xylem and tracheary elements in young tobacco (Hauffe et a!., 1991), while the 2x35S gave a more constitutive expression patterns (Kay et a!., 1987).  The double transgene lines also showed similar patterns of  expression, but expression was generally not as highly as in single transgene plants. Neither the single nor double transgene plants showed any visible phenotypic differences when compared to the non-transformed controls.  However, biomass  measurements of plants suggested that the overexpression of UGPase and SuSy, both individually and in combination, has a significant impact on plant height growth in tobacco.  Most single transgene lines and all of the double transgene lines showed  significant increases in height growth. Furthermore, each of the double transgene lines had at least one parental line that also showed significant increases in height compared to the non-transformed controls. An increase in height growth was also reported with the overexpression of SuSy in poplar under the control of the CaMV35S promoter 53  (Konishi et a!., 2004), and similarly the increased height growth was not evident in all transformed lines.  These findings corroborate SuSy antisense suppression studies  which have shown reduced plant size and leaf number in carrot (Tang and Sturm, 1999), and significantly reduced fruit size in tomatoes (D’Aoust, eta!., 1999). While there is some evidence for the role of SuSy in sink tissue, limited work has been conducted in elucidating a function for UGPase in the same tissue. The current study provides evidence and suggests that UGPase plays a role in the strength of sink tissues as demonstrated by the increased accumulation of biomass in both 2x35S and 4CL lines.  There is evidence, although indirect, that UGPase may possess a similar  role in poplar, as it was recently found to be upregulated during late cell expansion and secondary cell wall formation (Hertzberg et al., 2001).  However, in growing potato  tubers, a reduction in UGPase activity (up to 96%) by antisense suppression caused no change in carbohydrate metabolism (Zrenner et a!., 1993). Similarly, in Arabidopsis, antisense suppression of UGPase (30% reduction) did not confer a phenotypic effect under normal light conditions (Johansson, 2003).  However, Johansson (2003)  attributed these findings to the existence of multiple UGPase isoforms. In addition to a general increase in height due to the overexpression of SuSy and UGPase individually, when the data for single and double transgene line are compared, it is apparent that the double transgene lines were generally taller than the single transgene lines. Lines 2x35S::UGPase/SuSy-2 and 2x355::UGPase/SuSy-4 and lines 4CL::UGPase/SuSy-1 I and 4CL::UGPase/SuSy-12 were all significantly taller than either of their single transgene parental lines (2x35S::UGPase-8 and 2x35S::SuSy-9; 2x35S::UGPase-12 and 2x35S::SuSy-17; 4CL::UGPase-5 and 4CL::SuSy-3; 4CL::UGPase-5 and 4CL::SuSy-3). It is plausible that the combined overexpression of SuSy and UGPase affects the plants’ ability to more effectively utilize photosynthates and produce more biomass. As SuSy has previously been identified as a marker of sink strength (D’Aoust eta!., 1999), it is not surprising that in combination with a second key enzyme in sucrose metabolism that a greater increase in hexose sugar sink strength is prevalent, and consequentially that plant growth is affected. As the direction of UGPase has been hypothesized to be affected by the availability of substrates, UGPase has the potential to utilize the UDP-glucose produced by SuSy and directly incorporate it in the production of hexose phosphates (Kleczkowski et a!., 2004). This 54  could facilitate the observed increased plant growth, and explain the lack of increase in cellulose content. Alternately, if there is a quantifiable supply of glucose-I-phosphate, LJGPase could be effective in producing UDP-glucose. Fructose generated by SuSy mediated catabolism of sucrose could be the indirect supply for glucose-I-phosphate (Kleczkowski et a!., 2004). The single SuSy and UGPase transgene lines, as well as the double transgene lines had longer intemodes.  This suggests that the altered expression of these  genes/enzymes does not affect plant development, but rather invokes either an increase in cell number and/or cell elongation in a given intemodal region. Given there were no statistically significant changes in fibre morphology, the increased internodal distance, and consequently height, is the result of an increase in the number of fibres. Thus, plants displaying increased growth are likely producing fibres at a higher rate than control plants.  This is further supported by the fact that the quantified fibre length  tended to be slightly shorter, while fibre coarseness (wall thickness) was generally slightly greater for the transgenic plants in both single and double transgene lines. The increased height growth in the transgenic plants resulted in the formation of reactiontype fibres, analogous to tension and compression wood fibres in angiosperms and gymnosperms, respectively. Compression wood cells, in particular, are well known to be shorter in length and posses thicker cell walls as a result of growth stresses (Zobel and van Buijtenen, 1989).  These results are further supported by microscopic  sectioning (Figure 2.5). Previous studies involving the antisense suppression of SuSy in potato (Zrenner et a!., 1995), carrot (Tang and Sturm, 1999), and tomato (D’Aoust et a!., 1999) have all demonstrated significant changes in the soluble carbohydrate content in sink tissue. Similarly, shrunken 1, a maize endosperm SuSy mutant, (Chourey and Nelson, 1976) was shown to possess elevated soluble sugar concentration in sink tissue.  The  compromised SuSy activity showed a direct increase in localized sucrose levels (2-4 fold). The altered phenotypes (substantially reduced dry weight) in both potato and maize were hypothesized to be related to the hexose concentration, and an associated increase in water influx (Tang and Sturm, 1999). The upregulated SuSy activity in the current study did not demonstrate an increase in the concentration of sucrose in sink 55  tissue. However, there was a significant increase in both  tose and glucose, with  fructose levels increasing more than that of glucose. This inaicates that the plant was maintaining a basal concentration of sucrose in the stem, and that the upregulation of SuSy may only be associated with the increased degradation of sucrose and the consequential accumulation of glucose and fructose monomers. When SuSy activity is suppressed, there is a build up of sucrose, suggesting that SuSy is a limiting step in sucrose catabolism. In contrast, these findings imply that an increase in SuSy activity facilitates elevated rates of sucrose degradation, and conquentially allows more sucrose to be transported to the stem tissue, and ultimately  eases the metabolizing  cells accessibility to photosynthates. Studies examining the effect of UGPase on soluble sugar content are limited. In potato tubers, the antisense reduction of UGPase (up to 96%) caused no changes in soluble sugars (Zrenner et a!., 1993), while a 30% antisense reduction in arabidopsis resulted in a decrease in sugar content (Johansson, 2003). In the present study, much like the results with SuSy transformed lines, tobacco overexpressing UGPase had increased glucose and fructose content, yet only minor changes in sucrose concentrations. What is particularly interesting in both the UGPase and SuSy transformed lines is that the elevated levels of fructose and glucose were independent of the measured transcript levels or enzyme activities. These data suggest that only a small increase in either of these genes can have a dramatic effect on carbohydrate levels and that there may be other factors limiting the accumulation above the modest, yet significant changes seen in the present study. In plants with suppressed SuSy activity, a consistent decrease in starch content has been shown: 62% in maize (Chourey and Nelson, 1976), 34-63% in potato (Zrenner  eta!., 1995), and 26% in carrot (Tang and Sturm, 1999). In contrast, the upregulation of SuSy in tobacco did not show a significant increase in starch content, however, all 4CL::SuSy lines showed an increasing trend for starch accumulation. While the suppression of SuSy causes a profound effect on the production of starch, the increased activity does not directly alter starch content. A similar trend is found with cellulose content. In SuSy suppressed carrot plants, there was an associated decrease in cellulose content (Tang and Sturm, 1999). The suppression of SuSy in cotton also 56  caused a profound effect on the production of cellulose, resulting in an almost fibreless phenotype (Ruan et a!., 2003). While no consistent trends in cellulose content were obvious among the SuSy transgenic tobacco, one line (2x35S::SuSy-9) did have a significant increase in cellulose content. The results from the antisense work suggest that although SuSy may be critical and necessary for starch biosynthesis, as with carbohydrate levels, other factors become limiting as SuSy is overexpressed. 2x35S::UGPase and 4CL::UGPase transgenic tobacco lines showed no significant change in either starch or cellulose content.  While this suggests that  UGPase is not rate limiting in cellulose production, it does not address the fact that despite an increase in soluble sugars there is no concurrent increase in cellulose or starch. Other studies have shown that UGPase plays a role in providing substrate for starch (Johansson, 2003). UGPase is also known to play a role in the production of cellulose in yeast and bacteria, as demonstrated by the reduction of cellulose in UGPase deficient mutants (Daran et al., 1995; VaIla et a!., 1989). One difference in these bacterial studies and the present study is the availability of SuSy in plants, as this enzyme is not present in either yeast or bacteria (Kleczkowski, et a!., 2004). Despite the common hypothesis that implicates SuSy in regulating sink strength in storage organs, there is clearly a limit to its capability. When suppressed, it has a profound effect not only on the soluble sugar content of the sink organ, but also on the storage and structural carbohydrates cellulose and starch. Conversely, its increased expression can be expected to potentially increase the availability of the hexose sugar pool. Our results show that in tobacco under the control of either the 2x35S or 4CL promoters, the overexpression of SuSy or UGPase significantly influences and regulates the growth of plants, but does not generally increase the partitioning of storage of sugars into starch or cellulose. Clearly, both SuSy and UGPase have the potential to increase overall plant growth, and thus increase total cellulose yield attainable from an individual plant.  Acknowledgements Funding for this project was provided by CelIFor Inc., Natural Sciences and Engineering Research Council of Canada and the Canadian Forest Service. 57  References Amor Y., Haigler C., Johnson S., Wainscott M. and Delmer D.P. 1995. A membraneassociated form of sucrose synthase and its potential role in synthesis of cellulose and callose in plants. Proceedings of the National Academy of Sciences of the United States of America 92: 9353-9357. Appeldoorn N.J.G., de Bruijn S.M., Koot-Gronsveld E.A.M., Visser R.G.F., Vreugdenhil D. and van der Plas L.H.W. 1997. Developmental changes of enzymes involved in conversion of sucrose to hexose-phosphate during early tuberisation of potato. Planta 202: 220-226. Borokov A.Y., McClean P.E., Sowokinos J.R., Ruud S.H. and Secor G.A. 1996. Effect of expression of UDP-glucose pyrophosphorylase ribozyme and antisense RNAs on the enzyme activity and carbohydrate composition of field-grown transgenic potato plants. Journal of Plant Physiology 147: 644-652. Carison S.J. and Chourey P.S. 1996. Evidence for plasma membrane-associated forms of sucrose synthase in maize. Molecular and General Genetics 252: 303-310. Chourey P.S. 1981. Genetic control of sucrose synthetase in maize endosperm. Molecular and General Genetics 184: 372-376. Chourey P.S. and Nelson S.E. 1976. The enzymatic deficiency conditioned by the shrunken-I mutation in maize. Biochemical Genetics 14:1041-1055. D’Aoust M.-A., Yelle S. and Nguyen-Quoc B. 1999. Antisense inhibition of tomato fruit sucrose synthase decreases fruit setting and the sucrose unloading capacity of young fruit. The Plant Cell 11: 2407-241 8. Daran J.M., Dailies N., Thines-Sempoux D. and Francois J. 1995. Genetic and biochemical characterization of the UGPI gene encoding UDP-glucose pyrophosphorylase from Saccharomyces cerevisiae. European Journal of Biochemistiy 233: 520-530. Dejardin A., Sokoiov L.N. and Kleczkowski L.A. 1999. Sugar/osmoticum levels modulate differential abscisic acid-independent expression of two stressresponsive sucrose synthase genes in Arabidopsis. Biochemist,y Journal 344: 503-509. Haigler C.H., Ivanova-Datcheva M., Hogan P.S., Salnikov V.V., Hwang S., Martin K. and Delmer D.P. 2001. Carbon partitioning to cellulose synthesis. Plant Molecular Biology 47: 29-51. Hanggi E. and Fleming A.J. 2001. Sucrose synthase expression pattern in young maize leaves: implications for phloem transport. Planta 214: 326-329. 58  Hauffe K.D., Paszkowski U., Schulze-Lefert P., Hahlbrock K., Dangl J.L. and Douglas C.J. 1991. A parsley 4CL-1 promoter fragment specifies complex expression patterns in trangenic tobacco. The Plant Cell 3: 435-443. Hertzberg M., Aspeborg H., Schrader J., Andersson A., Erlandsson R., Blomqvist K., Bhalerao R., Uhlen M., Teen T.T., Lundeberg J., Sundberg B., Nilsson P. and Sandberg G. 2001. A transcriptional roadmap to wood formation. Proceedings of the National Academy of Sciences of the United States of America 98: 1473214737. Hood E.E., Gelvin S.B., Melcher L.S. and Hoekema A. 1993. New Agrobacterium helper plasmids for gene transfer to plants. Transgenic Research 2: 208-218. Johansson H. 2003. Gene regulation of UDP-glucose synthesis and metabolism in plants. Umea, Sweden, Umea University. Kay R., Chan A., Daly M. and McPherson J. 1987. Duplication of CaMV 35S promoter sequences creates a strong enhancer for plant genes. Science 236: 1299-1302. Kennedy J.F. and White C.A. 1983. Bioactive carbohydrates in chemistry, biochemistry and biology. New York, USA, Haistead Press. Kleczkowski L.A. 1994. Glucose activation and metabolism through UDP-glucose pyrophosphorylase in plants. Phytochemist,y 37:1507-1515. Kleczkowski L.A., Geisler M., Ciereszko I. and Johansson H. 2004. UDP-glucose pyrophosphorylase. An old protein with new tricks. Plant Physiology 134: 912918. Konishi T., Ohmiya Y. and Hayashi T. 2004. Evidence that sucrose loaded into the phloem of a poplar leaf is used directly by sucrose synthase associated with various 3-glucan synthases in the stem. Plant Physiology 134:1146-1152. Murashige T. and Skoog F. 1962. A revised medium for rapid growth and bioassays with tobacco tissue culture. Physiologia Plantarum 15: 473-497. Ruan Y.L., Llewellyn D.J. and Furbank R.T. 2003. Suppression of sucrose synthase gene expression represses cotton fibre cell initiation, elongation, and seed development. The Plant Cell 15: 952-964. Salnikov V.V., Gnimson M.J., Delmer D.P. and Haigler C.H. 2001. Sucrose synthase localizes to cellulose synthesis sites in tracheary elements. Phytochemisti’y 57: 823-833. Sun J., Loboda T., Sung S.-J.S. and Black C.C.J. 1992. Sucrose synthase in wild tomato, Lycopersicon chmielewskii, and tomato fruit sink strength. Plant Physiology 98: 1163-1169. 59  Tang G.Q. and Sturm A. 1999. Antisense repression of sucrose synthase in carrot (Daucus carota L.) affects growth rather than sucrose partitioning. Plant Molecular Biology 41: 465-479. Valla S., Coucheron D.H., Fjaervik E., Kjosbakken J., Weinhouse H., Ross P., Amikam D. and Benziman M. 1989. Cloning of a gene involved in cellulose biosynthesis in Acetobacter xylinum: Complementation of cellulose-negative mutants by the UDPG pyrophosphorylase structural gene. Molecular and General Genetics 217: 26-30. Yokoyama T., Kadla J.F. and Chang H.M. 2002. Microanalytical method for the characterization of fiber components and morphology of woody plants. Journal of Agriculture and Food Chemist,y 50: 1040-1044. Zobel B.J. and Buijtenen v. 1989. Wood variation: its causes and control. Heidelberg, Springer-Verlag. Zrenner R., Salanoubat M., Willmitzer L. and Sonnewald U. 1995. Evidence of the crucial role of sucrose synthase for sink strength using transgenic potato plants (Solanum tugerosum L.). The Plant Journal 7: 97-107. Zrenner R., Willmitzer L. and Sonnewald U. 1993. Analysis of the expression of potato uredinediphosphate-glucose pyrophosphorylase and its inhibition by antisense RNA. Planta 190: 247-252.  60  Altered Sucrose Metabolism Impacts Biomass Production and Flower Development in Tobacco 2 Introduction Plant breeders and biotechnologists are consistently pursuing improved plant productivity (biomass) as a target trait (White et a!., 2006), the impetus for which incorporates several factors, including crop yield, carbon capture and more recently increased biomass for biofuels production. Generally, plant productivity can be viewed as a process governed by resource allocation. Plants have the innate ability to adapt their resource allocation to match resource acquisition, acquire resources more effectively, or avoid deleterious conditions. This plasticity in resource acquisition and allocation can have a profound effect not only on the development and physiology of plants, but on the industrial utility of the plant matter. Generating or breeding plant genotypes where resource allocation is directed to vegetative biomass and/or altered fibre properties, is therefore directly related to altering sink tissue. Increasing the transport of photoassimilate to the sink tissue, and the subsequent catabolism and use of sucrose within this tissue has the potential to disrupt source-sink relationships, and therefore stimulate plants to respond by increasing the mobilization of stored carbohydrates or to alter the photosynthetic machinery to compensate. In many plants, the primary sink exists as secondary cell walls, in the form of cellulose and lignin, the two most abundant polymers on earth. Thus, to create a stronger sink within cellulose producing cells could allow plants to metabolize photoassimilate more rapidly and consequently stimulate plants to alter growth rates.  2  A version of this chapter will be submitted for publication. Coleman HD, Beamish, L, Reid, A, Park, JY, Mansfield SD (2008). Altered sucrose metabolism impacts biomass production and flower development in tobacco. 61  UDP-glucose is the immediate precursor molecule in the synthesis of cellulose and can be formed by two pathways: UDP-glucose pyrophosphorylase (UGPase; EC 2.7.7.9) catalyses the production of UDP-glucose from glucose-I-phosphate and UTP, while sucrose synthase (SuSy; EC 2.4.1.13) cleaves sucrose into fructose and UDP glucose. The latter reaction has an energetic advantage over that of UGPase as it retains the glycosidic bond for use in cellulose formation. There is additional evidence for the direct association of SuSy with the cellulose synthase complex which would permit the recycling of UDP into UDP-glucose (Salnikov et a!., 2001).  Furthermore,  SuSy activity has been positively associated with sink strength in many species, including potato (Zrenner et a!., 1995), tomato (Sun et a!., 1992; D’Aoust et a!., 1999), carrot (Tang & Sturm, 1999) and tobacco (Coleman eta!., 2006). Fructose is a known inhibitor of SuSy activity (Doehlert, 1987). However, plants employ sucrose phosphate synthase (SPS; EC 2.4.1.14) to aid in fructose recycling and limit SuSy inhibition. SPS synthesizes sucrose-6-phosphate from fructose-6-phosphate and UDP-glucose, and concurrently provides additional substrate for SuSy (Delmer, 1999). In non-photosynthetic tissue, SPS has dual-functionality: the re-synthesis of sucrose following cleavage during import, and the involvement in carbohydrate regulatory cycles involving starch degradation and sucrose re-synthesis (Geigenberger eta!., 1997; Stitt eta!., 1988). Furthermore, SPS has been identified as playing a role in diurnal carbohydrate allocation (Huber and Huber, 1996), flower development (Baxter et a!., 2003), fruit development (Laporte et a!., 2001) and cell wall growth and expansion (Haigler et a!., 2001). In source tissues, UGPase can act in concert with SPS in the formation of sucrose (Kleczkowski, 1994), while in sink tissues it has the potential to restrict carbon flow to sucrose formation (Borokov et a!., 1996) as it works co-ordinately with SuSy in the cycling between sucrose and the hexose phosphate pools (Kleczkowski, 1994). The objective of this study was to investigate the effects of misregulating multiple genes involved in sucrose metabolism on plant growth and secondary cell wall cellulose biosynthesis.  62  Methods Crossing (transformation) Transgenic tobacco overexpressing UGPase and SuSy (Coleman et a!., 2006) and SPS (Park et a!., 2008) under the control of either the enhanced tandem CaMV35S constitutive promoter (2x35S) (Kay et a!., 1987; Datla et a!., 1993) or the vascular specific 4CL (Petroselinum crispum 4-Coumarate:CoA ligase) promoter (Hauffe et al., 1991) were grown in the greenhouse until flowering. The transgenic plants were then crossed to produce plants harbouring differing combinations of two exogenous transgenes, including: UGPase x SPS and SuSy x SPS. The resulting seeds were planted on media containing the respective selective antibiotics and double transgenic plants confirmed by PCR screening of genomic DNA using the following primer sets: UG P-F (5’-atcgaggaattctgcctcgt-3’) and UG P-R (5’-tcgcaagaccggcaacaggatt-3’) for UGPase  confirmation,  SUS-1  (5’-ctcaacatcacccctcgaat-3’)  accaggggaaacaatgttga-3’)  for  SuSy  ggctatcgttcaagatgcctctg-3’)  and  SPS-R  confirmation  and and  SUS-2 SPS-F  (5’-aggcctcgcaagggcaagta-3’)  for  (5’(5’SPS  confirmation. The successful crosses were grown and multiplied in tissue culture to produce a minimum of 12 individually double transformed plants per line. UGPase x SuSy plants (Coleman at aL, 2006) were also crossed with SPS tobacco plants, and the resulting plantlets screened for the presence of all three genes. All shoot cultures, including transgenic and non-transformed control lines, were maintained on solid MS+3% in GA-7 vessels at 22CC under a 16 hour photoperiod with an average photon flux of 40 pmol m 2  . Plants were maintained by transferring apical regions at four-week intervals. 1 s All transgenic combinations, along with appropriate non-transformed control  tobacco plants, were transferred to a greenhouse into 7.5 L pots containing a 50% peat, 25% fine bark, 25% pumice soil mixture. Each multi-gene pyramiding combination had a corresponding control group as the growth periods were staggered. UGPase x SPS th plants were planted on the 16 of December, 2005. SuSy x SPS plants were planted th on the 23r of December, 2005 and UGPase x SuSy x SPS plants were planted 4 of July, 2006.  Following the formation of flower buds, the plants were harvested and biomass measurements taken. Plant height was measured from the base (root collar) 63  to the tip of the highest bud. The stage of tissue development was standardized using a leaf plastichron index, where the first leaf larger than 5cm was defined as P1=0 and the leaf immediately below was P1=1. Leaves corresponding to P1=4 and P1=5, and the associated stem section, were harvested for enzyme assays, RNA transcript analysis and soluble sugars content determination. The section of stem spanning P1=5 to P1=15 was retained for cell wall analysis, as well as stem dry weight determination.  The  leaves associated with this section were also used for leaf dry weight determination. All data was analysed using students t-test assuming unequal variance.  Transcription Levels Real time PCR was employed to determine transcript abundance of each transgene. RNA was isolated from 100 mg liquid nitrogen ground samples of stem and leaf tissue of plants using Trizol reagent (Gibco BRL, Gaithersburg, MD) according to manufacturers’ instructions. Ten pg of RNA was then treated with TURBO DNaseTM (Ambion, Austin, TX) to remove residual DNA.  One pg of DNase-treated RNA was  used for the synthesis of cDNA using Superscript II Reverse Transcriptase (Invitrogen, Carlsbad, CA) and dT 16 primers according to the manufacturer’s instructions. Samples were run in triplicate with Brilliant SYBR Green QPCR Master Mix (Stratagene, La Jolla, CA) on an Mx3000P Real-Time PCR System (Stratagene) to determine critical thresholds (Ct). The primers employed for RT-PCR analysis primers pairs: UGPase  -  AU-RTF (5’-tggaagcaacccgcgtcatc-3’) and AU-RTR (5’-gccaaggcccagcggttcc-3’); SuSy -  GS-RTF (5’-ccgtgagcgtttggatgagac-3’) and GS-RTR (5’-ggccaaaatctcgttcctgtg-3’);  SPS  -  AtSPS-F3  (5’-ccacagtggcaaagtgatgatggc-3’)  and  AtSPS-R4  (5’-  tctgacctctccagtgatccc-3’). As a house-keeping control, the transcript abundance of Actin-9 was employed for normalization using primers described previously: NtAct-RTF (5’-ctattctccgctttggacttggca-3’) (Volkov  eta!.,  and  NtAct-RTR  (5’-aggacctcaggacaacggaaacg-3’)  2003). Conditions for the RT-PCR reactions were as follows: 95°C for 10  minutes, followed by 40 cycles of 95°C for 30 seconds, 62°C SuSy (65°C SPS, 60 °C UGPase, 55°C Actin,) for 1 minute, and 72°C for 30 seconds. Relative expression was determined according to Levy  et a!.  (2004) using the following equation: ct  or SuSy or SPS— ctNtActin)  64  Enzyme Activity Leaf and stem samples (Ig f.w.) were ground in liquid nitrogen with 1 mg of insoluble PVPP and four volumes of extraction buffer (50 mM HEPES-KOH pH 7.5, 10 mM MgCl, 1 mM EDTA, 2 mM DTT, 1 mM PMSF, 5 mM EAmino-n-caproic acid, 0.1% v/v Triton X-100, 10% v/v glycerol). The samples were centrifuged at 15,000xg for 20 minutes at 4°C. The extract was passed through a desalting column (DG 10  —  BioRad)  pre-equilibrated with ice-cold extraction buffer without Triton X-1 00 and PVPP. Extracts were collected in pre-chilled vials and used immediately.  UGPase activity was  determined spectrophotometrically at 340nm as per Appeldoorn et a!. (1997) using 100 j.tL of plant extract and a NADH molar extinction coefficient of 6.22 mM cm . SuSy 1 activity was assayed in the direction of sucrose breakdown (Chourey, 1981), using 50 pL of plant extract. The resultant fructose content was determined using a tetrazolium blue assay (Kennedy and White, 1983). This SuSy assay employs the appropriate controls without the supplementation of UDP to quantify inherent invertase activity, and therefore represents only the breakdown of sucrose by SuSy. SPS activity was determined according to Iraqi and Tremblay (2001) and Baxter et a!. (2003). Total protein content of the extracts was determined using a Bio-Rad Protein Assay (Bio-Rad, Hercules, CA).  Soluble Carbohydrate and Starch Analysis Soluble carbohydrates (glucose, fructose and sucrose) were extracted from ground freeze-dried tissue overnight at -20°C using methanol:chloroform:water (12:5:3) as previously described (Coleman et a!., 2006). supernatant  removed,  and  the  remaining  The sample was centrifuged, the pellet  washed  twice  with  fresh  methanol:chloroform:water (12:5:3). All fractions were then pooled. Five mL of water was added to the combined supernatant and centrifuged to facilitate phase separation. The aqueous fraction was rotary evaporated to dryness and re-suspended in 3 mL of distilled water. Soluble carbohydrates were then analyzed using anion exchange HPLC (Dionex, Sunnyvale, CA) on a DX-600 equipped with a Carbopac PAl column and an electrochemical detector. 65  The residual pellet after soluble sugar extraction was then hydrolyzed in 4% sulfuric acid at 121’C for 4 minutes.  The liberation of glucose, representing starch  content, was directly quantified by HPLC under similar conditions.  Cell Wall Composition Oven dried tobacco stem segment spanning P1=5 to P1=15 were ground using a Wiley mill to pass through a 40-mesh screen and soxhlet extracted with acetone for 24 hours.  Lignin and carbohydrate contents were determined using a modified Kiason  method (Huntley et a!., 2003) on 0.2g of extract free tissue. Carbohydrate content was determined using HPLC (Dionex DX-600, Dionex, CA) equipped with an anion exchange PAl column, a pulsed amperometric detector with a gold electrode and postcolumn detection.  Acid insoluble lignin was determined gravimetrically, while acid  insoluble lignin was determined using spectrophotometric analysis at 205 nm according to TAPPI useful method UM-250.  Results Transcript Expression and Enzyme Activity Transcript abundance of all three transgenes was measured in each plant, and expressed relative to p-actin.  All transgenics (doubles and triple lines) clearly  demonstrated substantial expression of the exogenous transgenes, while no detectable transcripts were apparent in the wild-type tobacco plants, as would be expected (Table 3.1). Transgene expression levels were variable among the plants and crosses, which is also expected, as each plant represents the selection of independent transformation events.  Furthermore, the independently selected double transformed lines were not  selfed to select for homozygous lines prior to reciprocal mating, nor were the number of transgene insertion events quantified by Southern analysis. Therefore, the selection of double, and later triple transgenic lines represents the products stemming from allelic variation in heterozygous, segregating selected parents, and as such variability in gene dose effect is not accounted for. This variability likely accounts for the inconsistencies 66  Table 3.1. Transcript level and enzyme activity in leaf and stem of UGPase x SPS, SuSy x SPS and UGPase x SuSy x SPS transgenic and wild-type tobacco lines. Average ± standard error (n = 5). Bold indicates significance at a=O.1O. UGPase Transcript Lev& LCtx 10000  UGPase Enzyme Activity pM0INADH min 1 mg 1 protein  Leaf  Stem  Leaf  Stem  2x35S::UGPase x SPS A  2.43 ± 0.49  7.05 ± 1.62  125.5 ± 6.9  464.9 ± 88.0  2x35S::UGPase x SPS B  131.44 ± 24.45  91.43 ± 11.81  214.0 ± 48.6  267 ± 74.1  2x35S::UGPase x SPS C  90.67 ± 54.00  186.53 ± 58.60  168.2 ± 21.8  418.7 ± 137.1  732.56 ± 173.19  241.3 ± 70.5  439.7 ± 122.1  4CL::UGPase x SPS A  562.30 ±47.51  4CL::UGPase x SPS B  403.94 ± 60.19  520.03 ± 83.71  186.6 ± 33.6  779.0 ± 149.6  4CL::UGPase x SPS C  545.99 ± 74.91  709.20 ± 81.06  144.2 ± 11.4  879.8 ± 83.9  n.d.  n.d.  89.7 ± 7.5  140.4 ± 17.3  Control A-C  SPS Transcript Level tCt x 10000 Leaf  SPS Enzyme Activity pg sucrose min 1 mg 1 protein  Stem  Leaf  Stem  2x35S::UGPase x SPS A  5.11 ± 1.59  5.54 ± 1.66  1.42 ± 0.45  9.16 ± 0.81  2x35S::UGPase x SPS B  7.20 ± 1.05  10.58 ± 2.46  2.73 ± 0.63  11.63 ± 2.27  2x35S::UGPase x SPS C  6.63 ± 2.09  11.86 ± 2.40  1.03 ± 0.24  10.66 ± 1.97  4CL::UGPase x SPS A  69.55 ± 14.33  31.72 ± 10.07  1.90 ± 0.10  12.91 ± 2.33  4CL::UGPase x SPS B  42.84 ± 22.04  16.08 ± 4.78  2.05 ± 0.51  9.71 ± 1.53  4CL::UGPase x SPS C  65.71 ± 22.66  62.34 ± 13.34  2.36 ± 0.69  24.94 ± 5.59  n.d.  n.d.  1.24 ± 0.14  5.73 ± 0.97  Control A-C  67  SuSy Transcript Level Ct x 10000  SuSy Enzyme Activity pg fructose min 1 mg 1 protein  Leaf  Stem  Leaf  Stem  2x35S::SuSyx SPS D  0.20 ± 0.04  10.26 ± 9.53  1.53 ± 0.37  7.19 ± 2.20  2x35S::SuSy x SPS E  0.25 ± 0.05  0.05 ± 0.02  0.94 ± 0.17  8.69 ± 3.42  2x35S::SuSy x SPS F  53.67 ± 49.74  65.71 ± 61.61  1.03 ± 0.13  18.83 ± 10.88  4CL::SuSy x SPS D  24.98 ± 7.75  40.54 ± 10.07  1.09 ± 0.20  9.04 ± 1.54  4CL::SuSyx SPS E  42.09 ± 14.83  44.55 ± 18.33  4.49 ± 1.25  7.22 ± 2.14  4CL::SuSyxSPS F  24.80±6.03  34.42±8.16  0.74±0.06  5.32±2.09  nd.  n.d.  0.33 ± 0.13  6.78 ± 1.40  Control D-F  SPS Transcript Level tCt x 10000  SPS Enzyme Activity 1 mg pg sucrose min 1 protein  Leaf  Stem  Leaf  Stem  2x35S::SuSyx SPS D  0.58 ± 0.16  0.83 ± 0.19  3.96 ± 0.89  25.52 ± 7.87  2x35S::SuSy x SPS E  0.88 ± 0.05  0.52 ± 0.33  4.07 ± 0.71  19.46 ± 5.21  2x35S::SuSy x SPS F  1.01 ± 0.50  1.72 ± 0.50  9.56 ± 3.67  32.84 ± 5.67  4CL::SuSy x SPS 0  22.78 ± 9.71  8.43 ± 2.61  5.42 ± 0.33  20.39 ± 5.43  4CL::SuSy x SPS E  15.91 ± 2.49  8.19 ± 1.32  4.30 ± 0.73  21.26 ± 5.35  4CL::SuSy x SPS F  21.34 ± 7.76  21.29 ± 6.12  7.71 ± 0.39  30.58 ± 9.47  n.d.  n.d.  1.97±0.34  6.36±0.10  ControlD-F  68  UGPase Transcript Level Ct x 10000 Leaf  UGPase Enzyme Activity pMol NADH min 1 mg 1 protein  Stem  Leaf  Stem  2x35S::UGPase  x  SuSy x SPS G  38.39 ± 22.48  0.31 ± 0.09  21.86 ± 5.25  93.40 ± 11.41  2x35S::UGPase  x  SuSy x SPS H  56.73 ± 31.17  73.31 ± 27.90  20.88 ± 4.60  73.58 ± 10.68  2x35S::UGPase  x  SuSy x SPS I  2.01 ± 0.66  1.05 ± 0.12  10.23 ± 1.08  90.09 ± 18.59  4CL::UGPase  x  SuSy x SPS G  98.89 ± 41.30  224.73 ± 179.13  11.04 ± 1.75  112.73± 14.56  4CL::UGPase  x  SuSy x SPS H  169.92 ± 54.57  404.41 ± 9.72  20.84 ± 3.60  80.92 ± 14.53  4CL::UGPase  x  SuSy x SPS I  177.12 ± 5.01  562.58 ± 210.13  11.66 ± 1.53  55.95 ± 4.84  n.d.  n.d.  5.70 ± 0.36  84.70 ± 7.26  Control G-l  SuSy Transcript Level Ct x 10000 2x35S::UGPase  x  SuSy x SPS G  2x35S::UGPase  x  SuSy x SPS H  2x35S::UGPase  x  SuSy x SPS I  4CL::UGPase  x  SuSy x SPS G  4CL::UGPase  x  SuSy x SPS H  4CL::UGPase  x  SuSyx SPS I  Control G-l  SuSy Enzyme Activity 1 mg pg fructose min 1 protein  Leaf  Stem  Leaf  Stem  56.52 ± 41.41  14.73 ± 5.77  1.14 ± 0.61  2.27 ± 0.65  0.75 ± 0.60  0.68 ± 0.27  1.84 ± 1.24  1.33 ± 0.19  10.81 ± 4.12  29.80 ± 1.93  0.43 ± 0.31  2.01 ± 0.20  55.63 ± 14.85  72.51 ± 2.85  0.53 ± 0.09  2.52 ± 0.34  15.11 ± 6.80  122.35 ± 54.89  1.10 ± 0.11  2.78 ± 0.80  8.89 ± 1.46  110.06 ± 66.84  1.77 ± 0.49  6.56 ± 2.59  n.d.  n.d.  0.52 ± 0.09  0.36 ± 0.06  SPS Transcript Level Ct x 10000  SPS Enzyme Activity 1 mg pg sucrose min 1 protein  Leaf  Stem  Leaf  Stem  2x35S::UGPase  x  SuSy x SPS G  69.95 ± 46.06  77.46 ± 25.25  0.75 ± 0.16  3.66 ± 0.50  2x35S::UGPase  x  SuSy x SPS H  63.70 ± 58.61  101.83 ± 37.83  0.57 ± 0.02  2.06 ± 0.30  2x35S::UGPase  x  SuSy x SPS I  40.44 ± 17.55  87.09 ± 20.63  1.14 ± 0.23  7.10 ± 1.56  4CL::UGPase  x  SuSyx SPS G  56.40 ± 24.31  121.33 ± 82.00  0.90 ± 0.08  2.83 ± 0.86  4CL::UGPase  x  SuSy x SPS H  41.95 ± 9.13  38.34 ± 19.79  1.60 ± 0.08  8.56 ± 0.79  4CL::UGPase  x  SuSy x SPS I  40.06 ± 15.43  342.67 ± 88.64  1.24 ± 0.27  4.50 ± 2.09  n.d.  n.d.  0.43 ± 0.05  2.41 ± 0.67  Control G-l  69  in gene expression observed for any given transgene when comparing among lines within a cross, as well as when expression patterns of lines between crosses. Despite the variability in transgene expression, some generalities regarding the influence of promoter and tissues can be made. When the transgenes were under the control of the 2x35S promoter no clear differences were apparent when comparing tissue specificity. 4CL-driven expression appeared to be consistently higher in the stem tissue, when compared to the leaf samples with the exception of the SPS transcript level. This latter finding is consistent with the targeted expression of this promoter (Hauffe et al., 1991). Interestingly, the 4CL promoter appears to drive expression of these genes to a greater extent in both tissues when compared to the 2x35S promoter. In general, there is good agreement between enzyme activity and transcript abundance, as in all cases the activity of the sucrose metabolizing enzymes is substantially greater than the native enzyme activity present in the control tobacco plants (Table 3.1). Furthermore, as with the transcript abundance, activity of all three proteins is generally higher in the stem sections as compared to the leaf tissue, and when the transgenes were under the control of the 4CL promoter. In the UGPase x SPS lines increases of —5- and —4-fold where apparent in UGPase and SPS activity, respectively in the stem segments of the transgenic plants. Similarly, in the SuSy x SPS transgenic lines the SPS activity was shown to be as much as 4.5 times greater than control plants, while one line showed a 12-fold increase in SuSy activity. However, the mean increase in SuSy activity across plants in this cross is approximately 3 times higher than the corresponding SuSy activity native to the control plants.  The triple  transgenic lines also displayed similar increases in enzyme activity.  Soluble Carbohydrates and Starch While variability exists among individual lines and crosses, starch content appears to be influenced by the overexpression of these three transgenes in the tobacco plants compared to the corresponding controls (Table 3.2). It is not possible to draw comparisons between different transgene combinations, as the greenhouse growth trials were conducted at different times during the year. However, comparisons  70  Table 3.2. Soluble carbohydrate and starch content (mg g ) in leaf and stem of 1 UGPase x SPS, SuSy x SPS and UGPase x SuSy x SPS transgenic and wild-type tobacco lines. Average ± standard error (n = 5). Bold indicates significance at a=O.1O.  Leaf  Glucose  Fwctose  Sucrose  Total  Starch  6.30 ± 2.14  5.24 ± 1.52  12.32 ± 2.85  23.86 ± 6.42  38.93 ± 10.71  8.69±2.27  5.60± 1.27  10.86±2.13  25.14±5.48  41.62±6.72  6.34 ± 1.16  5.11 ± 1.09  7.50 ± 1.18  18.95 ± 2.58  36.84 ± 5.53  4CL::UGPase x SPS A  9.08 ± 1.55  5.73 ± 0.66  11.94 ± 1.92  26.75 ± 1.10  54.93 ± 8.93  4CL::UGPase x SPS B  8.49 ± 0.72  4.48 ± 0.23  15.59 ± 2.00  28.57 ± 2.58  40.04 ± 10.40  4CL::UGPase x SPS C  7.04 ± 1.45  4.77 ± 0.83  10.55 ± 1.46  22.35 ± 2.38  41.82 ± 6.70  controlA-c  12.98±1.11  6.23±0.93  12.51 ±2.04  33.21 ±3.63  60.07±7.70  2x35S::SuSy x SPS D 2x35S::SuSyxSPS E 2x35S::SuSy x SPS F  8.52 ± 1.91  7.20 ± 1.05  14.42 ± 2.60  30.61 ± 4.45  43.72 ± 7.96  8.81 ±0.56  6.45± 1.26  14.66±4.19  29.93±3.41  58.05±2.77  7.76 ± 2.47  8.22 ± 0.91  9.81 ± 2.42  25.79 ± 5.69  49.71 ± 5.28  4CL::SuSy x SPS D  10.14 ± 0.56  9.62 ± 0.65  9.16 ± 0.84  28.92 ± 1.27  60.16 ± 9.59  4CL::SuSy x SPS E  5.93 ± 0.20  8.15 ± 0.23  8.66 ± 1.45  22.74 ± 1.63  55.02 ± 12.47  4CL::SuSyx SPS F  9.47 ± 1.27  9.35 ± 1.38  8.16 ± 1.89  26.98 ± 1.40  53.31 ± 10.50  Control D-F  9.23 ± 1.21  11.34 ± 2.70  9.25 ± 1.21  29.83 ± 2.59  55.83 ± 1.72  2x35S::UGPase x SPS A 2x35S::UGPasexSPS B 2x35S::UGPase x SPS C  2x35S::UGPase  x  SuSyx SPS G  41.97 ± 6.23  31.48 ± 1.69  77.64 ± 10.82  151.09 ± 7.66  66.97 ± 10.65  2x35S::UGPase  x  SuSy x SPS H  28.69 ± 3.77  17.38 ± 0.81  59.44 ± 3.91  105.51 ± 8.08  45.85 ± 2.69  2x35S::UGPase  x  SuSy x SPS I  28.61 ± 2.27  24.28 ± 2.67  70.93 ± 4.50  123.81 ± 9.38  54.33 ± 7.89  4CL::UGPase  x  SuSy x SPS G  33.84 ± 7.35  32.41 ± 2.12  63.32 ± 2.39  119.43 ± 13.10  61.63 ± 20.71  4CL::UGPase  x  SuSyxSPSH  26.44±2.29  24.40±0.77  71.29±4.43  118.76±7.99  42.25±10.00  4CL::UGPase  x  SuSy x SPS I  35.95 ± 3.55  26.39 ± 1.63  68.01 ± 7.84  130.35 ± 11.08  28.81 ± 0.12  48.25 ± 5.51  27.45 ± 2.16  78.80 ± 3.34  148.01 ± 5.65  93.63 ± 7.26  Control G-l  71  Stem  Glucose  Fructose  Sucrose  Total  Starch  2x35S::UGPase x SPS A 2x355::UGPase x SPS B  52.30 ± 7.46  46.29 ± 5.89  24.95 ± 3.82  123.55 ± 12.61  18.84 ± 2.75  39.90 ± 4.36  35.19 ± 6.00  24.80 ± 3.93  99.89 ± 11.40  15.34 ± 2.09  2x35S::UGPase x SPS C  47.21 ± 4.68  41.31 ± 5.78  31.57 ± 6.14  120.95 ± 12.48  13.03 ± 0.68  4CL::UGPase x SPS A  49.67 ± 7.75  42.77 ± 4.31  29.02 ± 2.47  121.46 ± 12.94  15.94 ± 1.37  4CL::UGPase x SPS B  50.93 ± 7.13  47.90 ± 4.21  23.53 ± 2.93  122.36 ± 8.64  14.63 ± 1.38  4CL::UGPase x SPS C  38.10 ± 8.40  37.92 ± 4.86  27.87 ± 5.90  103.90 ± 16.39  17.08 ± 1.82  Control A-C  53.78 ± 2.82  52.43 ± 5.09  21.28 ± 4.11  127.49 ± 7.49  16.85 ± 1.78  2x35S::SuSy x SPS D 2x35S::SuSy x SPS E 2x35S::SuSy x SPS F  29.46 ± 4.73  24.96± 4.30  26.41 ± 2.93  80.83 ± 9.27  15.43 ± 1.33  44.16 ± 7.56  30.20 ± 4.87  24.67 ± 5.11  99.03 ± 16.43  18.23 ± 1.47  36.37 ± 3.80  25.62 ± 2.88  22.53 ± 2.58  84.52 ± 7.27  18.43 ± 0.60  4CL::SuSy x SPS 0  28.78 ± 8.63  19.69 ± 5.57  25.23 ± 1.04  73.70 ± 14.11  16.86 ± 1.35  4CL::SuSyxSPS E  25.17±4.57  24.58±2.96  24.95±2.47  74.70±5.93  20.99±1.49  4CL::SuSy x SPS F  24.17 ± 3.71  19.72 ± 3.78  33.08 ± 9.11  76.98 ± 2.27  15.47 ± 2.49  Control D-F  26.00 ± 4.55  21.62 ± 2.75  24.03 ± 4.37  71.65 ± 9.76  14.22 ± 0.41  2x35S::UGPase  x  156.42 ± 7.62  84.94 ± 5.86  381.78 ± 35.66  14.34 ± 1.22  x  SuSy x SPS G SuSy x SPS H  183.84 ± 14.52  2x35S::UGPase  84.29 ± 17.30  122.25 ± 27.29  46.96 ± 16.27  253.51 ± 28.31  13.34 ± 0.53  2x35S::UGPase  x  SuSy x SPS I  158.85 ± 22.64  134.62 ± 9.94  99.55 ± 11.24  362.06 ± 51.04  12.93 ± 1.41  4CL::UGPase  x  SuSy x SPS G  134.44 ± 6.96  107.04 ± 4.39  88.75 ± 6.62  322.54 ± 16.22  16.48 ± 3.30  4CL::UGPase  x  SuSy x SPS H  113.68 ± 34.83  139.79 ± 38.98  88.22 ± 18.36  341.69 ± 92.17  19.33 ± 0.32  4CL::UGPase  x  SuSy x SPS I  79.06 ± 6.74  146.14 ± 1.68  91.36 ± 10.45  316.56 ± 5.38  17.62 ± 1.15  196.53 ± 11.77  140.67 ± 5.65  97.30 ± 4.49  442.80 ± 9.24  16.91 ± 1.23  Control G-I  72  with the independent corresponding controls demonstrate common trends. In particular, it appears that starch accumulation in leaf tissue is reduced in the transgenic lines, while the starch content in the stem segments appears to be unaltered or slightly increased. Similar findings were observed previously in the stem of UGPase x SuSy double transgenics (Coleman et a!., 2006). Furthermore, the selection of promoter does not appear to affect starch metabolism trends. Despite the significant overexpression, as evident by transcript and enzyme activity, of three major sucrose metabolizing enzymes, the overall total soluble carbohydrate levels, and more specifically sucrose contents were generally unchanged or only marginally altered (Table 3.2). In the double transgenics it appears that sucrose levels in leaf tissue are comparable to the control plants, while some lines appear to have elevated sucrose contents in the stem segments.  Plant Growth Total plant biomass of the transgenic lines and corresponding controls plants was assessed by measured height growth, calliper, and total leaf dry weight at harvest. All transgenic lines had significantly increased height growth regardless of transgene combination. These findings are consistent with previously reported transgenic tobacco overexpressing UGPase and SuSy alone and in tandem (Coleman et al., 2006) and SPS alone (Park et a!., 2008).  In the current study, the UGPase x SPS double  transgenic lines showed 23-31% increases in height growth compared to the corresponding control plants (Figure 3.IA), while the SuSy x SPS lines growth enhancement ranged from 18-48% over controls (Figure 3.2A). The triple transgene tobacco, containing upregulated UGPase x SuSy x SPS transgenes demonstrated increases in height growth ranging from 20-57% over controls (Figure 3.3A). These observed gains do not appear to be dominated by either the constitutively expressed or vascular-specific promoter. Consistent with the observed increases in height growth, stem size, as determined by calliper measurements, was generally increased in all transgenic lines at harvest. The combined increase in height and calliper clearly indicated that the overall  73  izu  *  *  *  p9  *  ,,  A  ioofl  77 60 7 9 k\ 6 £60 0  I  40//\\\ 20%/%\  2.0 1.8 1.6 1.4  I  0.8  C)  0.6  0.4 0.2  t  B  i U /  Figure 3.1. Biomass measurements of UGPase x SPS transgenic lines compared to wild-type tobacco plants: height (A), calliper (B), and leaf dry weight (C). *indicates significance at a=O.05.  74  140 120 100  80 60 40 20  1.8 1.6 1.4 1.2 E 2. 1.0 0.8 0.6 0.4 0.2 0.0  .5 2’ 4  -2  Figure 3.2. Biomass measurements of SuSy x SPS transgenic lines compared to wildtype tobacco plants: height (A), calliper (B) and leaf dry weight (C). *indicates significance at a0.05.  75  E  0  a, a,  I  a, a. a,  C)  20  0,  I Figure 3.3. Biomass measurements of UGPase x SuSy x SPS transgenic lines compared to wild-type tobacco plants: height (A), calliper (B) and leaf dry weight (C). *indicates significance at a=O.05.  76  volume of plant biomass accrued during the growth trial by the transgenic lines was significantly greater than that observed in the corresponding control tobacco lines. The most significant increases in calliper were apparent in the UGPase x SPS double transgenics (Figure 3.IB). While increases in calliper were evident in the SuSy x SPS and UGPase x SuSy x SPS transgenics, not all lines evaluated consistently demonstrated improvements in this phenotype (Figures 3.2B & 3.3B). Total stem dry biomass (data not shown) were reflective of the combined growth traits. promoter choice did not offer a selective advantage.  Again,  Total dry leaf biomass was also shown to be increased in the UGPase x SPS double transgenics lines compared to the corresponding control lines. However, similar significant changes were not apparent in the SuSy x SPS and UGPase x SuSy x SPS transgenics, consistent with the calliper measurements. However, most lines showed comparable leaf biomass as compared to the appropriate controls (Figures 3.1C, 3.2C, & 3.3C). Interestingly, the parallel upregulation of multiple sucrose metabolizing genes altered time to set flower buds in many of the transgene combinations (Table 3.3). Initially, the extended time to flower development was thought to be responsible for the increase biomass accrual during the growth period. However, there was no change in timing of flower development in UGPase x SPS transgenics lines compared to the control plants, and these double transgenics displayed the largest and most consistent increases in plant biomass.  Similarly, in a previous study (Coleman et a!., 2006),  substantial increases in plant biomass were attainable by overexpressing a UGPase x SuSy transgene combination, without any change in timing to flower development. In contrast, the SuSy x SPS transgenic lines, and the triple upregulated transgenic lines, UGPase x SuSy x SPS, all showed extended days to floral developmental. In both of the latter two combinations, the length to complete flower development was increased by approximately 21 days, and these differences were independent of timing of the growth trial. In addition to the extended time for developmental completion, many of the 4CL lines showed foliar stipules and morphological alterations in flowers, while the 2x35S driven transgenics only showed similar abnormal morphological flower phenotypes (Figure 3.4). 77  Table 3.3. Time to flowering of UGPase x SPS, SuSy x SPS and UGPase x SuSy x SPS transgenic and wild-type tobacco lines. Average ± standard error (n = 12). Bold indicates significance at a=O.1O. Days to Flowering 2x35S::UGPase x SPS A 2x35S::UGPase x SPS B  50.2 ± 2.32  2x35S::UGPase x SPS C  51.8 ± 2.17  4CL::UGPase x SPS A  47.8 ± 1.19  4CL::UGPase x SPS B  47.7 ± 1.50  4CL::UGPase x SPS C  47.2 ± 0.89  50.0 ± 1.56  Control A-C  51.7 ± 4.63  2x35S::SuSyx SPS D 2x35S::SuSy x SPS E 2x35S::SuSy x SPS F  57.3 ± 0.88  4CL::SuSy x SPS D  61.7 ± 4.18  4CL::SuSy x SPS E  69.4 ± 4.29  4CL::SuSy x SPS F  62.0 ± 7.29  Control D-F  56.2 ± 2.23  78.2 ± 4.98 78.0 ± 3.37  2x35S::UGPase  x  SuSy x SPS G  35.3 ± 1.94  2x35S::UGPase  x  SuSy x SPS H  39.1 ± 2.09  2x35S::UGPase  x  SuSy x SPS I  39.6 ± 1.90  4CL::UGPase  x  4CL::UGPase  x  SuSy x SPS H  51.4 ± 2.70  4CL::UGPase  x  SuSy x SPS I  48.4 ± 1.29  Control G-I  SuSy x SPS G  37.3 ± 2.06  30.4 ± 0.40  78  Figure 3.4. Morphological alterations of 4CL::UGPase x SuSy x SPS (A&C&E) plants relative to wild-type tobacco plants (B&D&F). A&B show foliar stipules on 4CL lines, C&D show altered reproductive bud and flower morphology. 79  Cell Wall Components As a means to quantify carbon skeleton allocation, the stem segment spanning P1=5 to P1=15 was characterized for the changes in cell wall chemistry. The double transgenics did not show any measurable differences in wall chemistry (data not shown), and these findings concur with the previous observations of Coleman et a!. (2006). In contrast, the triple overexpressing UGPase x SuSy x SPS transgenic lines showed statistically increased levels of total hemicellulose-derived sugars, represented by arabinose, rhamnose, galactose, mannose and xylose (Table 3.4).  Additionally,  these same plants appear to be synthesizing increased levels of cellulose, represented by liberated glucose monomers, and reduced levels of lignin. These latter two cell wall components seem to offset one another, showing an approximately 2% change in dry matter shift.  Discussion Tobacco independently transformed with one of three key sucrose metabolizing enzymes, UGPase, SuSy and SPS, were generated under the regulation of two different promoters: a putative constitutive promoter (2x35S) and a vascular specific promoter (4CL). The ensuing single transgenic tobacco lines were then reciprocally crossed to produce double transgenic plants, which were subsequently employed to generate tobacco lines overexpressing all three sucrose metabolizing transgenes. In short, six transgenic combinations were employed for all growth and biochemical analysis, including: 2x35S::UGPase x SPS, 4CL::UGPase x SPS, 2x35S::SuSy x SPS, 4CL::SuSy x SPS, 2x35S::UGPase x SuSy x SPS, and 4CL::UGPase x SuSy x SPS. All six combinations clearly showed evidence of transgene expression, as determined by real time PCR evaluation.  Correspondingly, the associated enzyme activity was  significantly increased in either leaf or stem tissue of all lines relative to their corresponding non-transformed control plants. All lines showed increases in height growth, which is consistent with previous reports of these genes in tobacco (Coleman et a!., 2006; Park et a!., 2008). The triple lines showed slightly higher percent increase in growth over the doubles. This is also 80  Table 3.4. Chemical composition (% dry weight) of cell wall of stem of UGPase x SuSy x SPS triple transgenic lines. Average ± standard error (n = 5). Bold indicates significance at a=O.1O.  2x35S::UGPase  x  2x35S::UGPase  x  2x35S::UGPase  x  Cellulose  Hemicellulose  Lignin  SuSy x SPS G  25.04 ± 0.92  11.15 ± 0.29  21.75 ± 1.32  SuSy x SPS H  25.94±1.47  11.25 ± 0.25  20.70 ± 1.34  SuSy x SPS I  26.43 ± 1.96  11.64 ± 0.57  22.51 ± 0.34 23.57 ± 1.36  4CL::UGPase  x  SuSy x SPS G  26.28 ± 2.25  12.08 ± 0.57  4CL::UGPase  x  SuSy x SPS H  25.83 ± 0.66  12.32 ± 0.39  24.49 ± 1.80  4cL::uGpase  x  SuSy x SPS I  25.27 ± 0.81  11.45 ± 0.47  23.06 ± 0.12  24.07 ± 1.49  10.29 ± 0.21  25.49 ± 2.16  control G-I  81  consistent with what was previously observed with UGPase x SuSy double transgenic tobacco plants (Coleman et al., 2006) which displayed 10-48% increases in height growth, and performed better than plants transformed with only one of the genes (615% height increase). Furthermore, the extent of improved height growth in the current transgenics tobacco plants is consistent with the improvements attainable with SPS alone (Park et al., 2007). The increased growth observed in this study is consistent with other studies examining the misregulation of these three sucrose metabolizing genes individually in numerous other plant species. For example, the overexpression of a modified mung bean SuSy in Populus alba has been associated with slight increased height growth (Konishi et al., 2004), while SuSy antisense studies have clearly shown reduced plant size in carrot (Tang and Sturm, 1999) and reduced fruit size in tomato (D’Aoust et a!., Overexpression of SuSy in tobacco resulted in increased height growth and internode length, despite no significant changes in fibre length or coarseness (Coleman 1999).  et a!., 2006). In poplar overexpressing SuSy under the control of both the 2x35S and 4CL promoters, there was a slight reduction in height, but a significant increase in cellulose production (Coleman et al., unpublished). SuSy has consistently been shown in previous studies, and in the current study, to strongly influence growth whether in height or in increased secondary growth. The majority of studies investigating UDP-glucose pyrophosphorylase have focused primarily on downregulating enzyme activity, as evidence suggests that UGPase is present in ample supply in plants (Appeldoorn et a!., 1997, Magel et a!., However, the findings surrounding UGPase misregulation are inconsistent, ranging from no observed phenotypic effect in potato tubers despite a decrease in UGPase by 96% (Zrenner et al., 1993) to substantial decreases in soluble sugar  2001).  concentrations in potato tubers with a 30-50% reduction in activity (Borokov et a!., 1996; Spychalla et a!., 1994). In Arabidopsis, similar reductions in soluble sugars were observed when UGPase activity was reduced by -50% (Johansson 2003), however, In contrast, when UGPase activity was without observable plant phenotype. upregulated in tobacco, increased plant biomass and changes in carbohydrate metabolism were observed, albeit without altering partitioning to cellulose (Coleman at a!., 2006).  In addition, the overexpressing Acetobacter xylinum UGPase in poplar 82  resulted in severe reductions in height growth and biomass (Coleman et al., 2007). The difference between these latter two studies using the same gene construct in two different species was not directly attributed to the activity of the UGPase gene, but rather to side effects in poplar in which a defence response appeared to be elicited by an increase in UDP-glucose (Coleman et a!., 2007).  In addition, this also points to  differences in sink strength between the two species, as alterations in UGPase activity appear to directly impact secondary growth in poplar, while in tobacco the impact is apparent on height and biomass accumulation. SPS has also been shown to affect biomass production. Tomato overexpressing a maize SPS showed increased shoot growth (Galtier et a!., 1993), while antisense suppression of SPS resulted in a 50% reduction in tomato plant growth (Strand et al., 2001). In rice, SPS has been identified as a targeted gene underlying a quantitative trait locus for plant height (Ishimaru et a!., 2004). These results were confirmed with the overexpression of a maize SPS gene in rice, which resulted in significantly taller plants at the early growth stage.  In tobacco, the overexpression of an Arabidopsis SPS  resulted in increased total dry biomass, attributed to both height growth and diameter growth (Park et a!., 2007). These plants also had increased internode length and fibre length, but only minor changes in soluble and structural carbohydrates. Consistent with the results of the current study, in SPS overexpressing tobacco, there was a slight decrease in lignin content attributed largely to a decrease in acid soluble lignin. However, in the case of the single SPS transgenics, there was also a reduction in cellulose and hemicelluloses which was attributed to a general reduction in secondary cell waIl deposition related to increased growth rate (Park at a!., 2007). In potato, the overexpression of maize SPS resulted in a 20% increase in tuber weight and yield, along with increased tuber sucrose content (lshimaru et a!., 2008). As the tuber is the main sink in potato, this corresponds well with the results seen in this study where the overexpression of SPS in conjunction with other genes results in an increase in sink biomass. With the three sucrose metabolism genes being expressed in combination, it was anticipated that the effects would be additive, as seen with the UGPase  SuSy  transgenic tobacco. However, SPS does appear to have the most significant effect on primary growth, and the addition of the UGPase and SuSy genes do not increase the 83  height gains achieved with SPS alone. However, the crossing of SuSy with the SPS plants added additional phenotypic variations in flower morphology and time to flowering. Interestingly, the time to flowering in the case of tobacco transformed with SPS in combination with SuSy and in combination with both SuSy and UGPase is increased. Also, in the triple transgenic lines, the number of flowers per plant decreased greatly (Figure 3.4) with only one or two morphologically altered flowers per plant. This is inconsistent with previous findings in tobacco in which SPS overexpression resulted in decreased time to flowering and increased the number of flowers per plant (Baxter et al., 2003). The decreased time to flowering was also seen in tomato overexpressing SPS (Micallef et a!., 1995). In both cases, the SPS was from maize. As well, in both cases there was an increase in sucrose to starch ratio in transgenic lines, which concurs with the current findings in triple transgenic lines. Sucrose content has been directly associated with flowering in some species (Lejeune et a!., 1993), however, the changes in intracellular pools manifested by the overexpression of these sucrose metabolizing genes does not appear to driving these substantial abnormalities in floral development.  It is surprising that despite similarities in the sucrose and starch  components in the plants, the opposite effect is seen in time to flowering and in number of flowers. It is possible that this is related to the type of SPS gene being used. Both previous papers (Baxter eta!., 2003; Micallef eta!., 1995) employed the maize Family B SPS gene originally employed by Worrell et a!. (1991), while this study uses an Arabidopsis Family A SPS gene. B-Family genes in maize are found throughout the plant, but appear to be restricted almost exclusively to the anthers in wheat and barley (Castleden et a!., 2004). The effects of this gene appear to be consistent with changes seen as a result of the Arabidopsis Family A genes, with increased sucrose:starch ratios, but differences, include time to flowering, frequency of flowering and biomass production. UGPase and SuSy have also emerged as key genes involved in regulating, directly or indirectly, floral development.  UGPasel suppression has recently been  shown to play a key role in pollen development in rice, where seed set is virtually eliminated (VVoo et al., 2008). SuSy activity has also been recently characterized in tobacco pollen tubes to occur in two distinct pools as soluble SuSy (S-SuSy) and 84  membrane associated SuSy (P-SuSy) forms, and is thought to play a role in extracellular matrix construction (Persia et al., 2008). There did not appear to be changes in cell wall chemistry, despite the substantial growth effects resulting from the misregulation of UGPase x SPS or SuSy x SPS. These results are consistent with what has previously been reported in tobacco expressing these three genes independently (Coleman et a!., 2006; Park et a!., 2008). Similarly, it was shown in UGPase x SuSy double transgenic tobacco, that there was no change seen in carbon allocation (Coleman et a!., 2006). In the present study, there does however appear to be a change, although minor, in the allocation of carbon skeletons to the various cell wall moieties in the triple transgenic lines. The UGPase x SuSy x SPS plants clearly show increased levels of the hemicellulose-derived carbohydrates and a 2% change in the balance of cellulose to lignin composition, with cellulose content increasing and lignin portion decreasing.  Recently, it has been shown that cotton  overexpressing a spinach SPS showed an increase in sucrose:starch ratio, and also had altered fibre qualities, with increased micronaire (wall thickness and diameter), maturity ratio and fibre length, all of which are consistent with increased deposition of fibre cellulose (Haigler et a!., 2007). In studies examining natural variation in wheat, increased SuSy activity was also correlated with an increase in CesA activity and cellulose production.  This was associated with a decrease in water soluble  carbohydrates (Xue et a!., 2007). Contrary to these results, in SuSy x SPS tobacco, there was a trend towards increased total soluble carbohydrates and increased starch in the stem. This could be associated with the regulatory role of the cellulose synthase complex in the production of cellulose.  In tobacco, primary growth could take  precedence over secondary growth, thus increased availability of sugars is stored and utilized for increased height growth, rather than for cellulose production as observed in cotton and wheat (Haigler et al., 2007; Xue et a!., 2007). In triple transgenic tobacco, the addition increased UGPase activity further alters the sucrose metabolism, preventing increased starch storage and potentially increasing the plants ability to utilize sugars that would otherwise be stored for hemicellulose production. Interestingly, these same lines that have SPS and SuSy working in an apparent futile cycle (Nguyen-Quoc and Foyer, 2001) also have altered flower morphology. 85  In summary, the overexpression of UGPase, SuSy and SPS alone or in combination, clearly demonstrate an advantage in plant biomass accumulation in tobacco. As has previously been reported with these transgenes, there is an added advantage in employing a combined pyramiding strategy to alter sucrose metabolism. However, it is apparent from this study, that no single combination of two of these genes has an added advantage over any of the other combinations with respect to growth traits in tobacco.  Pyramiding all three genes again has only slight, if any  improvements, in growth traits, but does appear to challenge the normal cell wall chemical deposition. However, it is unknown whether this is true carbon re-allocation, or a simple a cellular wall response, and as such a chemical response, to alter growth patterns.  Despite the slight advantage in altered cell wall chemistries, favouring  carbohydrate deposition, the triple transgenic lines appear to have severely impaired floral development  —  a phenomena that warrants further investigation. Furthermore, this  study suggests that choice of promoter did not, in these cases, offer an improved advantage in altering plant biomass accumulation.  Acknowledgements The authors thank Thomas Canam, Tony Einfeldt and Jimmy Yan for their technical assistance. Funding for this project was provided by NSERC, Cellfor Inc and CFS.  86  References Appeldoorn N.J.G., de Bruijn S.M., Koot-Gronsveld E.A.M., Visser R.G.F., Vreugdenhil D. and van der Plas L.H.W. 1997. Developmental changes of enzymes involved in conversion of sucrose to hexose-phosphate during early tuberisation of potato. Planta 202: 220-226. Baxter C.J., Foyer C.H., Turner J., Rolfe S.A. and Quick W.P. 2003. Elevated sucrosephosphate synthase activity in transgenic tobacco sustains photosynthesis in older leaves and alters development. Journal of Experimental Botany 54: 18131820. Borokov A.Y., McClean P.E., Sowokinos J.R., Ruud S.H. and Secor G.A. 1996. Effect of expression of UDP-glucose pyrophosphorylase ribozyme and antisense RNAs on the enzyme activity and carbohydrate composition of field-grown transgenic potato plants. Journal of Plant Physiology 147: 644-652. Castleden C.K., Aoki N., Gillespie V.J., MacRae E.A., Quick W.P., Buchner P., Foyer C.H., Furbank R.T. and Lunn J.E. 2004. Evolution and function of the sucrosephosphate synthase gene families in wheat and other grasses. Plant Physiology 135: 1-12. Chourey P.S. 1981. Genetic control of sucrose synthetase in maize endosperm. Molecular and General Genetics 184: 372-376. Coleman H.D., Canam T., Kang K.Y., Ellis D.D. and Mansfield S.D. 2007. Overexpression of UDP-glucose pyrophosphorylase in hybrid poplar affects carbon allocation. Journal of Experimental Botany 58: 4257-4268. Coleman H.D., Ellis D.D., Gilbert M. and Mansfield S.D. 2006. Up-regulation of sucrose synthase and UDP-glucose pyrophosphorylase impacts plant growth and metabolism. Plant Biotechnology Journal 4: 87-101. D’Aoust M.-A., Yelle S. and Nguyen-Quoc B. 1999. Antisense inhibition of tomato fruit sucrose synthase decreases fruit setting and the sucrose unloading capacity of young fruit. The Plant Cell 11: 2407-2418. Datla R.S.S., Bekkaoui F., Hammerlindl J.K., Pilate G., Dunstan D.I. and Crosby W.L. 1993. Improved high-level constitutive foreign gene translation using an AMV RNA4 untranslated leader sequence. Plant Science 94: 139-149. Delmer D.P. 1999. Cellulose biosynthesis: Exciting times for a difficult field of study. Annu. Rev. Plant Physiol. Plant Mol. Biol. 50: 245-276. Doehlert D.C. 1987. Substrate-inhibition of maize endosperm sucrose synthase by fructose and its interaction with glucose inhibition. Plant Science 52: 153-157. 87  Galtier N., Foyer C.H., Huber J., Voelker TA. and Huber S.C. 1993. Effects of elevated sucrose-phosphate synthase activity on photosynthesis, assimilate partitioning, and growth in tomato (Lycopersicon esculentum var UC82B). Plant Physiology 101: 535-543. Haigler C.H., Ivanova-Datcheva M., Hogan P.S., Salnikov V.V., Hwang S., Martin K. and Delmer D.P. 2001. Carbon partitioning to cellulose synthesis. Plant Molecular Biology 47: 29-51. Haigler C.H., Singh B., Zhang D., Hwang S., Wu C., Cai W.X., Hozain M., Kang W., Kiedaisch B.M., Strauss R.E., Hequet E.F., Wyatt B.G., Jividen G.M. and Holaday A.S. 2007. Transgenic cotton over-producing spinach sucrose phosphate synthase showed enhanced leaf sucrose synthesis and improved fiber quality under controlled environmental conditions. Plant Molecular Biology 63: 81 5-832. Hauffe K.D., Paszkowski U., Schulze-Lefert P., Hahlbrock K., Dangi J.L. and Douglas C.J. 1991. A parsley 4CL-1 promoter fragment specifies complex expression patterns in trangenic tobacco. The Plant Cell 3: 435-443. Huber S.C. and Huber J.L. 1996. Role and regulation of sucrose-phosphate synthase in higher plants. Annu. Rev. Plant Physiol. Plant Mol. Biol. 47: 431-444. Huntley S.K., Ellis D., Gilbert M., Chapple C. and Mansfield S.D. 2003. Significant increases in pulping efficiency in C4H-F5H transformed poplars: Improved chemical savings and reduced environmental toxins. Journal of Agriculture and Food Chemistry 51: 6178-6183. Iraqi D. and Tremblay F.M. 2001. Analysis of carbohydrate metabolism enzymes and cellular contents of sugars and proteins during spruce somatic embryogenesis suggests a regulatory role of exogenous sucrose in embryo development. Journal of Experimental Botany 52: 2301-2311. Ishimaru K., Hirotsu N., Kashiwagi T., Madoka Y., Nagasuga K., Ono K. and Ohsugi R. 2008. Overexpression of a maize SPS gene improves yield characters of potato under field conditions. Plant Production Science 11: 104-1 07. Ishimaru K., Ono K. and Kashiwagi T. 2004. Identification of a new gene controlling plant height in rice using the candidate-gene strategy. Planta 218: 388-395. Johansson H. 2003. Gene regulation of UDP-glucose synthesis and metabolism in plants. Umea, Sweden, Umea University. Kay R., Chan A., Daly M. and McPherson J. 1987. Duplication of CaMV 35S promoter sequences creates a strong enhancer for plant genes. Science 236: 1299-1302.  88  Kennedy J.F. and White C.A. 1983. Bioactive carbohydrates in chemistry, biochemistry and biology. New York, USA, Haistead Press. Kleczkowski L.A. 1994. Glucose activation and metabolism through UDP-glucose pyrophosphorylase in plants. Phytochemistiy 37: 1507-1515. Konishi T., Ohmiya Y. and Hayashi T. 2004. Evidence that sucrose loaded into the phloem of a poplar leaf is used directly by sucrose synthase associated with various b-glucan synthases in the stem. Plant Physiology 134: 1146-1152. Laporte M.M., Galagan J.A., Prasch A.L., Vanderveer P.J., Hanson D.T., Shewmaker C.K. and Sharkey T.D. 2001. Promoter strength and tissue specificity effects on growth of tomato plants transformed with maize sucrose-phosphate synthase. Planta 212: 817-822. Lejeune P., Bernier G., Requier M.C. and Kinet J.M. 1993. Sucrose increase during floral induction in the phloem sap collected at the apical part of the shoot of the long-day plant Sinapis alba L. Planta 190: 71-74. Levy M., Edelbaum 0. and Sela I. 2004. Tobacco mosaic virus regulates the expression . Plant Physiology 135: 2392-2397. 1 of its own resistance gene N Magel E., Abdel-Latif A. and Hampp R. 2001. Non-structural carbohydrates and catalytic activities of sucrose metabolizing enzymes in trunks of two Juglans species and their role in heartwood formation. Holzforshung 55: 135-145. Micallef B.J., Haskins K.A., Vanderveer P.J., Roh K.-S., Shewmaker C.K. and Sharkey T.D. 1995. Altered photosynthesis, flowering, and fruiting in transgenic tomato plants that have an increased capacity for sucrose synthesis. Planta 196: 327334. Nguyen-Quoc B. and Foyer C.H. 2001. A role for ‘futile cycles’ involving invertase and sucrose synthase in sucrose metabolism of tomato fruit. Journal of Experimental Botany 52: 881-889. Park J.-Y., Canam T., Kang K.Y., Ellis D.D. and Mansfield S.D. 2008. Overexpression of an arabidopsis family A sucrose phosphate synthase (SPS) gene alters plant growth and fibre development. Transgenic Research. Persia D., Cai G., Del Casino C., Faleri C., Willemse M.T.M. and Cresti M. 2008. Sucrose synthase is associated with the cell wall of tobacco pollen tubes. Plant Physiology doi: 10.1 104/pp.108.1 15956. Salnikov V.V., Grimson M.J., Delmer D.P. and Haigler C.H. 2001. Sucrose synthase localizes to cellulose synthesis sites in tracheary elements. Phytochemistty 57: 823-833. 89  Spychalla J.P., Scheffler B.E., Sowokinos J.R. and Bevan M.W. 1994. Cloning, antisense RNA inhibition, and the coordinated expression of UDP-glucose pyrophosphorylase with starch biosynthetic genes in potato tubers. Journal of Plant Physiology 144: 444-453. Stitt M., Wilke I., Feil R. and Heldt H.W. 1988. Coarse control of sucrose phosphate synthase in leaves: alterations of the kinetic properties in response to the rate of photosynthesis and the accumulation of sucrose. Planta 174: 2 17-230. Sun J., Loboda T., Sung S.-J.S. and Black C.C.J. 1992. Sucrose synthase in wild tomato, Lycopersicon chmielewskii, and tomato fruit sink strength. Plant Physiology 98: 1163-1169. Tang G.Q. and Sturm A. 1999. Antisense repression of sucrose synthase in carrot (Daucus carota L.) affects growth rather than sucrose partitioning. Plant Molecular Biology 41: 465-479. Volkov R.A., Panchuk 1.1. and Schoffi F. 2003. Heat-stress-dependency and developmental modulation of gene expression: the potential of house-keeping genes as internal standards in mRNA expression profiling using real-time RT PCR. Journal of Experimental Botany 54: 2343-2349. White T.L., Adams W.T., and Neale D.B. 2006 Forest Genetics. Cabi International, Oxford, UK. Woo M.O., Ham T.H., Ji H.S., Choi M.S., Jiang W., Chu S.H., Piao R., Chin J.H., Kim J.A., Park B.S., Seo H.S., Jwa N.S., McCouch S. and Koh H.E. 2008. Inactivation of the UGPasel gene causes genic male sterility and endosperm chalkiness in rice (Oiyza sativa L.). The Plant Journal doi: 10.1111/j.136531 3X.2008.03405.x. Worrell A.C., Bruneau J.-M., Summerfelt K., Boersig M. and Voelker T.A. 1991. Expression of a maize sucrose phosphate synthase in tomato alters leaf carbohydrate partitioning. The Plant Cell 3: 1121-1130. Xue G.-P., Mcintyre C.L., Jenkins C.L.D., Glassop D., van Herwaarden A.F. and Shorter R. 2007. Molecular dissection of variation in carbohydrate metabolism related to water soluble carbohydrate accumulation in stems of wheat (Triticum aestivum L.). Plant Physiology 146: 441-454. Zrenner R., Salanoubat M., Willmitzer L. and Sonnewald U. 1995. Evidence of the crucial role of sucrose synthase for sink strength using transgenic potato plants (Solanum tugerosum L.). The Plant Journal 7: 97-107. Zrenner R., Willmitzer L. and Sonnewald U. 1993. Analysis of the expression of potato uredinediphosphate-glucose pyrophosphorylase and its inhibition by antisense RNA. Planta 190: 247-252. 90  Overexpression of UDP-Glucose Pyrophosphorylase in Hybrid Poplar Affects Carbon Allocation 3 Introduction To maximize resource acquisition and to minimize exposure to deleterious phenomena, trees constantly monitor endogenous and environmental cues, and use this information to regulate resource allocation.  Ultimately, these responses are  controlled by gene expression patterns, resulting in the synthesis of a variety of metabolites, including carbohydrates, which can accumulate, participate in metabolite channelling and/or be polymerized into more complex macromolecules. Studies have shown that altered carbon partitioning can manifest changes in the chemical composition of plants by the altered regulation of genes involved in the synthesis of lignin or cellulose (Li et a!. 2003, Canam et a!. 2006).  Despite these findings, how  carbon partitioning occurs and the factors that regulate plants to produce more or less cellulose still remain largely unanswered. It may be possible to alter and/or regulate carbohydrate utilization by increasing the availability of precursor substrates such as UDP-glucose in the production of cellulose, or ADP-glucose in starch synthesis. The availability of soluble carbohydrates would therefore be a key component to altering and/or regulating carbohydrate utilization,  thus by increasing  available sugar  metabolites in the sink cells (i.e. UDP-glucose), there may be the potential to augment cellulose production and content. Two pathways have been identified that lead to the direct production of UDP glucose. The first is based on the cleavage of sucrose by sucrose synthase (SuSy; EC 2.4.1.13) liberating fructose and UDP-glucose, while the second relies on the hexose phosphate pool and the active phosphorylation of glucose 1-phosphate by UDP-glucose pyrophosphorylase (UGPase, EC 2.7.7.9).  UGPase is considered a key enzyme in  A version of this chapter has been published. Coleman HD, Canam T, Kang, K.Y., Ellis D, Mansfield SD (2007). Over-expression of UDP-glucose pyrophosphorylase in hybrid poplar affects carbon allocation. Journal of Experimental Botany. 58: 4257-4268.  91  carbohydrate biosynthesis and plays an important role in sucrose metabolism (Coleman et a!., 2006). In sink tissues, UGPase works in coordination with the sucrolytic enzymes (i.e. SuSy, fructokinase and SPS/SPP) in the metabolism of sucrose and hexose phosphates, while in source tissues, UGPase works closely with SPS in the synthesis of sucrose (Kleczkowski 1994). As such, the dual functionality of UGPase makes it an interesting target for altering cellulose production; it has the potential to increase the amount of sucrose in source tissues, and concurrently decrease sucrose in sink tissues, thereby modulating the solute potential gradient and facilitating sucrose channelling to sink tissues. Previous investigations have focused primarily on downregulating UGPase enzyme activity, as evidence suggests that UGPase is present in ample supply in plants (Appeldoom et a!. 1997, Magel et a!. 2001). However, the results of such studies are mixed, ranging from no observed phenotypic effect in potato tubers despite a decrease in UGPase by 96% (Zrenner at a!. 1993) to substantial decreases in soluble sugar concentrations in potato tubers with a 30-50% reduction in activity (Borokov et a!. 1996; Spychalla et a!. 1994).  In Arabidopsis, similar reductions in soluble sugars were  observed when UGPase activity was reduced by -P50% (Johansson 2003). In contrast, when UGPase activity was upregulated in tobacco, increased plant biomass and changes in carbohydrate metabolism were observed, albeit without altering partitioning to cellulose (Coleman at a!. 2006). Studies evaluating native UGPase gene expression patterns in poplar clearly demonstrate the onset of UGPase upregulation during late cell expansion and secondary cell wall formation, which is consistent with the theory that UGPase contributes in providing the immediate substrate for cellulose synthesis, and is co-ordinately up regulated with the cellulose synthase complex (Hertzberg et a!. 2001). The current study attempts to further elucidate the role of UGPase in secondary cell wall biosynthesis in hybrid poplar, and its effects on carbon partitioning.  Poplar  trees were transformed with UGPase from Acetobacter xylinum under the control of the constitutive (2x35S) promoter and were employed to evaluate changes in tree growth characteristics, biochemistry and cell wall chemistry. The Acetobacter-derived UGPase gene was chosen for these studies since it shows a high specificity for UDP-glucose (Brede et a!. 1991), in contrast to the broader substrate specificity of plant-derived non specific UDP-sugar pyrophosphorylases (Kotake et a!. 2004). 92  Methods Cloning of UGPase and Plasmid Construction UGPase (M76548) was cloned from Acetobacter xylinum ATCC #23768 and inserted into the pBIN cloning vector under the control of the enhanced tandem CaMV35S (2x35S) constitutive promoter (Datla eta!. 1993; Kay eta!. 1987). Sequence analysis was used to confirm the proper insertion of the promoter and gene into the binary vector.  Plant Transformation and Maintenance Hybrid  poplar  (Popu!us a!ba  x  grandidentata) was transformed  using  Agrobacterium tumefaciens EHA1O5 (Hood et a!. 1993) employing a standard leaf disk inoculation.  Binary plasmids were inserted into EHA1O5 using the freeze-thaw  technique, and incubated overnight in liquid Woody Plant Media (WPM: McCown and Lloyd 1981) with 100 iiM acetosyringone.  Leaf disks were cut and co-cultured with  EHA1O5 for one hour at room temperature, blotted dry and plated abaxailly onto WPM supplemented with 0.1 pM each a-naphthalene acetic acid (NAA), 6-benzylaminopurine (BA), and thiadiazuron (TDZ) and solidified with 3% (w/v) agar and 1.1 % (wlv) phytagel (WPM 0.1/0.1/0.1).  After three days the discs were transferred to WPM 0.1/0.1/0.1  supplemented with carbenicillin disodium (500 mg L ) and cefotaxime sodium salt (250 1 mg L ). Following three additional days, the discs were transferred to WPM 0.1/0.1/0.1 1 containing carbenicillin, cefotaxime and kanamycin (25 mg L ). 1  After five weeks,  shoots and callus material were transferred to WPM with agar and phytagel, 0.01 pM BA, carbenicillin, cefotaxime and kanamycin.  Once individual shoots were visible,  plantlets were transferred to solidified WPM with 0.OlpM NAA and carbenicillin, cefotaxime and kanamycin to induce rooting. After two consecutive five-week periods  on this media, shoot tips were isolated to solidified antibiotic-free WPM with 0.01 pM NAA. Plants were confirmed as transgenic by PCR screening of genomic DNA employing gene specific oligonucleotides: specifically, UGP-F (5’-atcgaggaattctgcctcgt3’) and UGP-R (5’-tcgcaagaccggcaacaggatt-3’). 93  All shoot cultures, including transgenic and non-transformed control lines, were maintained on solid WPM with 0.01 pM NAA in GA-7 vessels at 22CC under a 16-hour photoperiod with an average photon flux of 40 pmol m 2  Plant Growth and Biomass Tissue culture plantlets were transferred into 7.5 L pots containing a 50% peat, 25% fine bark, 25% pumice soil mixture in the greenhouse, and covered with 16 oz clear plastic cups for one week to aid in acclimation. Each poplar line, transgenic and wild type, was represented by 12 clonally-propagated trees.  The greenhouse trees  were harvested after four months growth, at which time tree height, from base to tip,  and stem diameter (10 cm above root collar) were measured. Developmental stages of tissues were standardized by employing a plastichron index, where leaf plastichron index P1=0 was defined as the first leaf greater than 3cm in length, and where P1=1 was the leaf immediately below P1=0. Portions of the stem from each plant spanning P1=5 to P1=15, and from P1=15 to P1=25 were excised and dried at 105CC for 48h for dry weight determination, and retained for further analysis. Leaves were also collected in ten node groups (P1=6-I 5, P1=16-25) and analysed using an Area Meter (Li-Cor Environmental, Lincoln NE), and then dried at 105C for 48 hours for dry weight determination. Developing xylem was scraped and flash frozen in liquid nitrogen for future analysis of enzyme activity, RNA transcript abundance, and soluble sugar analysis.  Transcription Levels Real time PCR was used to determine transcript abundance of the transgene. Leaf and developing xylem samples weighing approximately 1 g (f.w.) were ground in liquid nitrogen, and RNA was extracted according to the method of Kolosova et al. (2004).  One pg of RNA was used for the synthesis of cDNA using Superscript II  Reverse Transcriptase (Invitrogen, Carlsbad, CA) and dT 16 primers according to the manufacturer’s instructions. Samples were run in triplicate with Brilliant SYBR Green QPCR Master Mix (Stratagene, La Jolla, CA) on an Mx3000P Real-Time PCR System (Stratagene). The primers for the RT-PCR analysis of the Acetobacter xylinum UGPase 94  were AU-RTF (5’-tggaagcaacccgcgtcatc-3’) and AU-RTR (5’-gccaaggcccagcggttcc-3’). Conditions for the RT-PCR reactions were as follows: 95°C for 10 minutes, followed by 40 cycles of 95°C for 30 seconds, 62°C for 1 minute, and 72°C for 30 seconds. Transcript levels were based on standard curves derived from known concentrations of plasmid DNA run under the same conditions. Transcript abundance of the two native poplar UGPase genes, as per Meng et a!. (2007) was concurrently evaluated by RT PCR using the following primers: PtUGP1 F (5’-ggcttcttcagatttgcttctg-3’), PtUGP1 R (5’ccagtttcacaccagatttcac-3’) and PtUGP2F (5’-gcaacttcagatctgcttcttg-3’), PtUGP2R (5’tccaatttcacaccagattttg-3’). Additionally, the transcription abundance of key genes involved in lignin and cellulose production  in the developing xylem were also quantified,  including:  phenylalanine ammonia-lyase, PAL-RT-FW (5’-aaaggtgccgaaattgccatgg-3’), PAL-RT RV (5’-tgcagaaatcaagcccaaggag-3’); gtggggaattgctgagcttgt-3’),  C4H-RT-RV  C4H-RT-FW (5’-  (5’-cgcaacttcttctggatttca-3’);  coumarate  3(5’-  C3H-RT-RV  (5’-atggcttcgttggatgtttc-3’),  C3H-RT-FW  hydroxylase,  cinnamate 4-hydroxylase,  atccataatagctctagtga-3’); caffeoyl CoA 3-0-methyltransferase, CCOMT-RT-FW (5’tttgcatgcttcctgttggtga-3’), CoA  reductase,  CCOMT-RT-RV  CCR-RT-FW  (5’-aatgcagcccctcacttgatcc-3’);  ci nnamoyl  (5’-atggtttactctatgtgcttctct-3’),  CCR-RT-RV  (5’-  dehydrogenase,  CAD-RT-FW  (5’-  gctcctccttcacaaaccttaa-3’);  cinnamyl  atgaagtggttggtgaggttgt-3’),  CAD-RT-RV (5’-acaccgacaacatctccaactt-3’); ferulate 5-  hydroxylase,  F5H-RT-FW  alcohol  (5’-agctcgcagacgtggtgggtttag-3’),  F5H-RT-RV  (5’-  gaaataaccagcaacctcagcatct-3’); caffeic acid 3-0-methyltransferase, COMT-RT-FW (5’gccagtgcttcagttctaccaa-3’); COMT-RT-RV (5’-ggtcgagttcaatggctgtttt-3’); 4C L-RT-FW (5’gcacctaaaactcaccatctctcc-3’),  4CL-RT-RV  (5’-aaggtttttcgggatgtagatgtc-3’);  sucrose  SuSyPt-RT-RV  (5’-  CESA-RT-FW  (5’-  agagctgtgatcattatgcgactg-3’), CESA-RT-RV (5’-acccaagaaaatgcaaaccagatc-3’).  The  synthase,  SuSyPt-RT-FW  gcaacacgcaaatcctcaacaa-3’);  (5’-ccatggattgctcttgctctgc-3’), and  cellulose  synthase,  genes selected and employed for transcript analysis of the lignin branch of the phenylpropanoid pathway and cellulose biosynthesis were based on previously reported high levels (highest of each isoform) during EST expression profiling of the cambial zone and tension wood formation in poplar (Sterky et a!. 2004). Critical threshold (Ct) 95  values for all genes were quantified in triplicate and normalized to -actin transcript levels.  Enzyme Activity Leaf and developing xylem samples (approximately I g f.w.) were ground in liquid nitrogen with I mg of insoluble PVPP and four volumes of extraction buffer (50 mM HEPES-KOH pH 7.5, 10 mM MgCl, 1 mM EDTA, 2 mM DTT, 1 mM PMSF, 5 mM EAmino-n-caproic acid, 0.1% v/v Triton X-100, 10% v/v glycerol). The samples were centrifuged at 14,000 rpm for 20 minutes at 4°C. The extract was passed through a desalting column (DG 10 BioRad) and pre-equilibrated with ice-cold extraction buffer —  without Triton X-100 and PVPP. Extracts were collected into pre-chilled vials and used immediately. UGPase activity was determined spectrophotometrically at 340 nm as per Appeldoorn et al. (1997) using 100 iL of plant extract and a molar extinction coefficient . Total protein content of the extracts was determined using a Bio-Rad 1 of 6.22 mM cm Protein Assay (Bio-Rad, Hercules, CA).  Soluble Carbohydrate and Starch Analysis  Soluble carbohydrates (glucose, fructose and sucrose) were extracted overnight at -20°C from ground freeze-dried plant material using methanol:chloroform:water (12:5:3). The sample was centrifuged, the supernatant removed, and the remaining pellet washed twice with fresh methanol:chloroform:water (12:5:3) and all fractions pooled.  Five mL of water was added to the combined pooled supernatant and  centrifuged to facilitate phase separation.  The aqueous fraction was removed to a  round bottom flask and rotary evaporated to dryness. The sample was resuspended in 3 mL of distilled water and analyzed using anion exchange HPLC (Dionex, Sunnyvale, CA) on a DX-600 equipped with a Carbopac PAl column and an electrochemical detector, as per Coleman et a!. (2006). The residual pellet was hydrolyzed using 4% sulfuric acid at 121CC for 4 minutes. The liberation of glucose represented starch content, and was directly quantified by HPLC employing similar conditions. 96  Cell Wall Compositional Analysis  Greenhouse grown plant stem material was ground using a Wiley mill to pass through a 40-mesh screen, and then soxhlet extracted with acetone for 24 hours. The extractive free material was used for all further analyses. Lignin content was determined using a modified Kiason, where extracted ground stem tissue (0.2 g) was treated with 3 0 as per Coleman et al. (2006). Carbohydrate concentrations in the 4 S 2 mL of 72% H hydrolyzate were determined using high-performance liquid chromatography (HPLC) (Dionex DX-500, Dionex, CA) equipped with an ion exchange PAl (Dionex) column, a  pulsed amperometric detector with a gold electrode, and a Spectra AS 3500 autoinjector (Spectra-Physics, Los Angeles, CA). 20 tiL of hydrolyzate was loaded on the column equilibrated with 250 mM NaOH and eluted with deionized water at a flow rate , followed by a post column addition of 200 mM NaOH at a flow rate of 1 of 1.0 mL min . Each experiment was run in duplicate. 1 0.5 mL min  Determination of x-Cellulose Content  a-cellulose was determined from the extract free wood using a modified microanalytical method developed by (Yokoyama et al., 2002).  In short, 200 mg of  ground extract-free wood was weighed into a 25 mL round bottom flask and placed in a 90°C oil bath. The reaction was initiated by the addition of I mL of sodium chlorite solution (400 mg 80% sodium chlorite, 4 mL distilled water, 0.4 mL acetic acid). An additional I mL of sodium chlorite solution was added every half hour and the samples removed to a cold waterbath after two hours. Samples were then filtered through a coarse crucible, dried overnight, and holocellulose composition determined gravimetrically. Fifty mg of this dried holocellulose sample was weighed into a reaction flask and allowed to equilibrate for 30 minutes. Four mL of 17.5% sodium hydroxide was added and allowed to react for 30 minutes, after which 4 mL of distilled water was added. The sample was macerated for 1 minute, allowed to react for an additional 29 minutes and then filtered through a coarse filter. Following a five-minute soak in 1.0 M acetic acid, the sample was washed with 90 mL of distilled water and dried overnight. The a-cellulose content was then determined gravimetrically. 97  Monolignol Analysis Thioacidolysis (Rolando et a!. 1992) was employed to determine the lignin monomer ratios (syringyl:guaiacyl; S:G), using 10 mg of oven-dried extractive free wood 1 in dichloromethane). The and tetracosane as an internal standard (2 mL of 25 mg mL siylation reaction proceeded for a minimum of 2 hours, and gas chromatography was carried out on a ThermoFinnigan Trace GC-PolarisQ ion trap system with an AS2000 autosampler and a splitlsplitless injector. The GC was equipped with a 30 m, 0.25 mm internal diameter J&W DB-5 column. The GC conditions were as follows: initial injector temperature of 250°C, detector temperature of 270°C, and initial oven temperature of 130°C. Following a 2 pL injection, the oven remained at 130°C for 3 minutes and then 1 to 260°C and held for 5 minutes. ramped at a rate of 3°C min  Soluble Metabolite Analysis Liquid nitrogen frozen developing xylem tissue was ground using a dental amalgam mixer, and extracted using a two-phase methanol:chloroform extraction as per Robinson et a!. (2005). In short, 600 j.iL of methanol was added to 20 mg of frozen tissue to stop biological activity. The samples were vortexed and 40 pL of water, 10 pL i of lipophilic internal standard 1 ribitol in water) and 10 1 of internal standard (10 mg mL 1 nonadecanoic acid methyl ester in methanol) were added. The samples (10 mg mL were then incubated for 15 minutes at 70°C with constant agitation, and then centrifuged for 5 minutes at 13000 rpm. The supernatant was removed, and 800 pL of methanol was added to the pellet, which was vortexed to resuspend the pellet, and then incubated at 35°C for 5 minutes with constant agitation. The sample was centrifuged for 5 minutes at 13000 rpm and then the supernatant removed and combined with the previous methanol supernatant. 600 pL of water was added to the pooled methanol extract, vortexed and centrifuged at 4000 rpm for 15 minutes. A 900 pL sample was removed from the upper potion (water/methanol) and dried at 40°C using a speedvac. The pellet was resuspended in 50 iiL of methoxyamine hydrochloride solution 1 in chloroform) and incubated for 2 hours at 60°C with constant agitation. (20 mg mL Following the addition of 200 pL of N-methyl-N-trimethylsilyltnfluoroacetamide (MSTFA), the sample was incubated at 60°C for 30 minutes with constant agitation.  Samples 98  were left at room temperature overnight to allow the reaction to progress to completion, and then filtered through tissue paper to remove particulates. GC/MS analysis was carried out using a 30m, 0.25mm internal diameter Restek Rtx-5MS column using the following conditions: inlet temperature 250°C, injector split ratio 10:1, resting oven temperature 70°C, and GC-MS transfer line temperature of 300°C. Following a I pL injection, the oven remained at 70°C for 2 minutes and then ramped at a rate of 8°C 1 to 325°C and held for 6 minutes. min  Results Transformed hybrid poplar trees regenerated from the AxUGPase-agrobacterium treated leaf explants were propagated from single shoots from individual unique explants, and represented independent transformed lines which were confirmed by PCR screening of genomic DNA to amplify a diagnostic fragment specific to the AxUGPase gene.  From  independent transgenic  the  lines 6  2x35S::UGPase  transformants (based on high real time qPCR expression levels) and corresponding control (wild type) shoots were propagated in vitro as shoot cultures, and a minimum of 12 individual trees of each line were transferred into the greenhouse and grown for four months under 18 hour days supplemented with overhead lighting with a radiant flux . At harvest, all of the transgenic lines showed substantially altered 2 density of 300 W m growth characteristics, demonstrating significantly impaired height and diameter growth (Figure 4.1A & B). Furthermore, the trees displayed significantly reduced internodal length (Figure 4.1 D), as well as decreased stem dry weight (data not shown). Morphologically, the 2x35S::UGPase transgenic lines also displayed a greater abundance of significantly smaller leaves per stem, and consistently had elongated axial shoots at each leaf node (Figure 4.2). A quantitative evaluation of leaf characteristics confirmed that the 2x35S::UGPase leaves were 75% smaller, had reduced total leaf area per stem and reduced total leaf dry weight (Figure 4.1C). In addition, the trichomes appeared to be more highly concentrated in the 2x35S::UGPase leaves  when  compared  to  the  wild-type  poplar  (data  not  shown).  99  180  12  A  160 140 120 100 0  80  I  60 40 20 0  iiiilii  200 180 160  140 120  100 0  80 60 40 20 0  C  Iiii, I—i WT  A  B  C  D  E  F  10 8 B 6 B ‘a 4 2  3.5 3.0 2.5 2.0  a, a’  -J  1.5  B a) 1.0 0.5 0.0  Figure 4.1. Plant height (A), diameter (B), leaf area (C) and internode length (D); for transgenic and wild-type trees. Mean (±SE) were calculated from 10 plants per line. All transgenic lines are statistically significant different from the control trees at a=0.05.  100  Figure 4.2. Image depicts axial shoot elongation and leaf size in a representative 2x35S::UGPase (right) and wild-type (left) poplar (inset is a close up of elongated axial shoots of transgenic poplar).  101  Transcription Levels  Real time quantitative PCR was used to evaluate transcript abundance of the exogenous UGPase gene in the transgenic lines (Table 4.1). All the 2x35S::UGPase transgenic lines had significantly higher transcript levels in the leaf material compared to 1 the stem (developing xylem). Transcript abundance ranged from 68 to 173 copies Jg 1 total RNA in the developing xylem, total RNA in leaf tissue and 2.1 to 27.9 copies pg among the transgenic poplar trees. The expression of the two native poplar UGPase genes was also shown to be affected in most lines (Table BI), with smaller increases in PtUGPI when compared to the observed increases in PtUGP2. Transcript abundance of genes involved directly in cellulose and lignin biosynthesis in the developing xylem were also compared using real time quantitative PCR.  PAL, 4CL, CST, CQT and COMT showed no change in the transgenic lines  relative to wild type levels, while all other genes in the lignin biosynthetic pathway were up regulated (Figure 4.3).  SAD, CAD, C3H, and F5H were increased between 2-3 fold  relative to wild type levels, and C4H was increased to 4-fold the wild type level. CCR and CCoAMT showed the highest relative increases at over 6 times the abundance seen in wild-type trees (Table B2). Similarly, the CesA gene family was also shown to be up regulated approximately 4-fold over the corresponding wild type CesA transcript levels, while SuSy expression levels remained comparable to the wild-type trees.  Enzyme Activity  UGPase activity was determined using an indirect assay measuring the production of the reduced form of nicotinamideadenine dinucleotide (NADH). All of the 2x35S::UGPase transgenic trees showed consistent increases in UGPase activity in the leaf tissue compared to the wild-type trees (Table 4.1). However, in contrast, only one line (2x35S::UGPase line C) demonstrated a statistically significant increase in enzyme activity in the developing xylem, despite the slight general upward trend in activity in the other transgenic lines.  102  Table 4.1. Transcript level and enzyme activity in leaf tissue and developing xylem for transgenic and wild-type trees. Mean (±SE) were calculated from 3 plants per line. Bold denotes significant difference from control values at a=O.1O. UGPase Transcript Level copy number pg 1 total RNA Transgenic line  Leaf  Developing Xylem  Control  UGPase Enzyme Activity 1 mg protein 1 iJmolNADH min Leaf  Developing Xylem  0.18 ± 0.05  0.71 ± 0.05  2x35S::UGPase A  99.47 ± 14.94  13.13 ± 12.10  0.50 ± 0.10  0.75 ± 0.01  2x35S::UGPase B  172.99 ± 15.17  20.79 ± 11.65  0.34 ± 0.05  0.80 ± 0.05  2x35S::UGPaseC  89.81 ±44.64  2.07± 1.23  0.54±0.12  1.69±0.40  2x35S::UGPase D  89.61 ± 48.89  19.24 ± 17.07  0.47 ± 0.02  0.68 ± 0.06  2x35S::UGPase E  67.51 ± 25.22  12.66± 11.13  0.39±0.08  0.86±0.18  2x35S::UGPase F  76.60 ± 42.27  27.92 ± 25.99  0.84 ± 0.16  0.82 ± 0.05  103  NC  X2-3  X4.-5  X6+  Expression relative to control trees  J I)-’ (l  Cellulose  Lignin  Guaiacyl Lignin  Lignin  Figure 4.3. Schematic representation of the effect of overexpression of 2x35S::UGPase on the transcript abundance of the lignin biosynthetic genes in the developing xylem. All mRNA levels were calculated by qPCR from threshold cycle values and are relative to controls and normalized with respect to f3-actin transcript abundance.  104  Soluble Carbohydrates In general, all transgenic lines showed elevated levels of total soluble sugars in the leaf tissue, which was a result of a commensurate increase in all three soluble sugars quantified  -  sucrose, fructose and glucose (Table 4.2).  The total soluble  carbohydrate concentrations, as well as concentrations of each individual sugar, were significantly increased in the leaves of five out of the six 2x35S::UGPase transgenic lines evaluated relative to the controls. The exception was Line F, which despite having higher glucose and fructose levels did not have statistically significant higher levels of sucrose, and thus total carbohydrates. In contrast, significant increases in total soluble carbohydrates, as well as significant increases in glucose and sucrose, were only found in two 2x35S::UGPase lines (B and C) in the developing xylem.  Starch Content Three of the six 2x35S::UGPase transgenic lines (A, B, and F) had significant increases in starch content in the leaves compared to starch levels in control leaves (Table 4.3). As well, five of the six 2x35S::UGPase transgenic lines (A, B, C, D and F) had significantly elevated starch in the developing xylem relative to control trees, with two lines (C and F) displaying three times the concentration of wild-type trees. And, although not statistically significant the sixth transgenic line (line E) did have higher levels of starch accumulation in the developing xylem, similar to the other transgenics.  Cell Wall Chemistry The total cell wall carbohydrate content, as a measure of percent total dry weight, of the 2x35S::UGPase lines increased dramatically compared to the wild-type trees, where all six lines showed statistically significant changes in polymeric cell wall moieties. Wild-type hybrid poplar trees were shown to be composed of -64.4% total carbohydrates, while the 2x35S::UGPase transgenic trees ranged from 69.2 to 73.1%. Although all cell wall carbohydrates (arabinose, galactose, glucose, xylose, mannose, and rhamnose) were shown to be elevated in the 2x35S::UGPase transgenic lines, the 105  Table 4.2. Total soluble carbohydrates (mg g ) in leaves and developing xylem of 1 transgenic and wild-type trees. Mean (±SE) were calculated from 3 plants per line. Bold denotes significant difference from control values at a=0.10. Tissue (collected between P1=3-5) was collected from greenhouse plants between 10:00 and 12:00 am.  Leaf  Glucose  Fructose  Sucrose  Total  Control  7.49 ± 2.07  2.61 ± 0.58  49.90 ± 3.19  60.00 ± 2.57  2x35S::UGPase A  18.79 ± 1.10  6.00 ± 0.75  68.96 ± 4.25  93.75 ± 5.60  2x35S::UGPase B  14.57 ± 6.29  4.93 ± 1.22  61.93 ± 3.95  81.42 ± 8.22  2x35S::UGPase C  17.54 ± 3.76  5.90 ± 1.61  66.33 ± 4.76  89.77 ± 6.79  2x35S::UGPase 0  13.60 ± 3.11  6.59 ± 1.68  65.60 ± 6.63  85.79 ± 11.08  2x35S::UGPase E  24.35 ± 3.35  6.06 ± 1.83  64.45 ± 3.74  94.86 ± 2.56  2x35S::UGPase F  19.71 ±4.45  6.89±0.56  51.30±4.67  77.91 ± 9.12  Developing Xylem  Glucose  Fructose  Sucrose  Total  Control  3.61 ± 0.87  0.00 ±0.00  23.71 ± 5.47  27.31 ± 6.12  2x35S::LJGPase A  8.65± 3.12  0.00 ± 0.00  35.05 ± 7.90  43.71 ± 10.64  2x35S::UGPase B  5.82 ± 0.97  0.00 ± 0.00  43.33 ± 1.93  49.15 ± 1.05  2x35S::UGPase C  13.19 ± 4.76  0.00 ± 0.00  61.95 ± 4.51  75.14 ±9.15  2x35S::UGPase D  3.37 ± 0.51  0.00 ± 0.00  27.83 ± 6.70  31.20 ± 6.55  2x35S::UGPase E  4.38 ± 1.41  0.00 ± 0.00  40.52 ± 10.72  50.06 ± 15.77  2x35S::UGPase F  3.38 ± 0.53  0.00 ± 0.00  35.06 ± 6.36  38.44 ± 6.71  106  Table 4.3. Starch content (mg g ) in leaves and developing xylem of transgenic and 1 wild-type trees. Mean (±SE) were calculated from 3 plants per line. Bold denotes significant difference from control values at a=0.05. Tissue (collected between P1= 3-5) was collected from greenhouse plants between 10:00 and 12:00 am.  Leaf  Developing Xylem  Control  2.6±0.1  8.3±0.7  2x35S::UGPase A  4.3 ± 0.5  18.3 ± 4.2  2x35S::UGPase B  4.2 ± 0.3  16.0 ± 4.0  2x35S::UGPase C  1.3 ± 0.0  27.6 ± 6.3  2x35S::UGPase D  2.9 ± 0.4  11.2 ± 0.6  2x35S::UGPase E  2.4 ± 0.0  14.0 ± 6.4  2x35S::UGPase F  3.7 ± 0.1  31.2 ±7.9  107  C  -  o  16.9±0.7 3.2±0.1 0.5±0.1  1.8±0.1  17.4±0.4  48.8±0.7  1.2±0.1  0.5±0.0  2x35S::UGPaseF  17.6±0.3 3.2±0.1 0.6±0.0  1.9±0.2  18.8±0.6  46.3±0.8  1.1 ±0.1  0.5±0.0  2x35S::UGPaseE  17.5±0.7 3.4±0.1  0.7±0.0  1.8±0.2  19.6±0.9  46.6±2.1  1.3±0.2  0.6±0.0  2x35S::UGPaseD  16.4±0.9 3.5±0.2  0.6±0.0  1.8±0.1  18.9±1.2  47.7±2.9  1.3±0.3  0.6±0.0  2x35S::UGPaseC  15.9±0.5 3.4±0.1  0.5±0.0  1.8±0.1  18.0±0.3  50.6±0.9  1.7±0.2  0.5±0.0  2x35S::UGPa5eB  15.4±0.5  3.3±0.1  0.6±0.0  2.0±0.1  18.8±0.7  49.0±1.4  1.4±0.1  0.6±0.0  2x35S::UGPaseA  20.7±0.1  3.0±0.0  0.6±0.1  1.5±0.1  17.7±0.7  43.2±0.2  1.0±0.1  0.4±0.0  Control  Insoluble  % Lignin Soluble  Rhamnose  Mannose  Xylose  Glucose  Galactose  Arabinose  % Carbohydrates  Table 4.4. Chemical composition of stem material of transgenic and wild-type trees. Mean (±SE) were calculated from 3 plants per line. Bold denotes significant difference from control values at a=O.1O.  most significant changes were observed in glucose content (Table 4.4). The observed increases in glucose content is reflective of the overall increase in cellulose composition in all transgenic lines compared to the corresponding wild-type trees (Figure 4.4), as determined by cL-cellulose quantification. The total cell wall lignin content of the 2x35S::UGPase transgenic lines was concurrently shown to be significantly decreased. As measured by percentage weight of total dry mass, the total lignin was reduced from 23.7% of the dry weight in wild-type trees to a range of 18.7 to 20.1% in the 2x35S::UGPase transgenic lines.  This  represents a decrease of 12% to 21% across the lines. Furthermore, the drop in total lignin content appears to be directly related to a reduction in the acid-insoluble lignin moieties (Table 4.4). Significant increases in the syringyl:guaicyl (S:G) ratio in the lignin was also observed in the stems of these 2x35S::UGPase transgenic lines (Table 4.5), with the % mol S concentration increasing from an average of 69.2% in wild-type poplar, to between 76.6% to 78.2% in the six 2x35S::UGPase lines with decreased lignin content. The increase in syringyl-based lignin monomers is proportionately related to a decrease in the guaiacyl monomers. Although significant, the change in the S:G ratio had a minor effect on the level of the p-hydroxyphenyl monomer (H-lignin) composition. However, H-lignin is only minor component in poplar trees relative to total lignin content.  Soluble Metabolite Analysis Total soluble metabolites, extracted independently from leaf tissue and developing xylem, were evaluated by metaboilte profiling with GCIMS analysis. Table 4.6 lists 18 (of -280 compounds investigated) whose levels are changed two-times or greater in 2x35S::UGPase trees compared to wild type. In general, many of the identified compounds are associated with plant/tree defence or stress, including myo inositol, galactinol, galactitol, and pinitol.  As well, compounds related to carbon  allocation, such as maltose and other carbohydrates were identified. However, a single compound was shown to dominant the pooling metabolites in the 2x35S::UGPase transgenic lines; salicylic acid 2-O-3-D-gIucoside was dramatically increased in all 109  *  *  40 0)  V/////A  .30 ID  20 o 0 ci) °  10 ci 0 WT  A  B  C  D  E  F  Figure 4.4. a-cellulose content of stem material from transgenic and wild-type trees. Mean (±SE) were calculated from 3 plants per line. Symbol denotes significant difference from control values at a=0.10.  110  Table 4.5. Syringyl, guaiacyl and p-hydroxyphenyl monomer contents (%) of transgenic and wild-type trees as determined by thioacidolysis. Mean (±SE) were calculated from 3 plants per line. Bold denotes significant difference from control values at a=O.05.  p-Hydroxyphenyl  Guaiacyl  Syringyl  Mean ± SE  Mean ± SE  Mean ± SE  Control  0.02 ± 0.01  30.78 ± 0.25  69.20 ± 0.26  2x35S::UGPase A  0.06 ± 0.02  21.68 ± 1.25  78.26 ± 1.27  2x35S::UGPase B  0.05 ± 0.01  21.87 ± 0.65  78.08 ± 0.66  2x35S::UGPase C  0.07 ± 0.02  23.20 ± 0.23  76.73 ± 0.20  2x35S::UGPase D  0.08 ± 0.00  23.29 ± 0.39  76.63 ± 0.39  2x35S::UGPase E  0.11 ± 0.00  22.63 ± 1.08  77.26 ± 1.08  2x35S::UGPase F  0.11 ± 0.01  23.28 ± 1.08  76.61 ± 1.07  111  Table 4.6. Metabolites identified in the developing xylem of 2x35S::UGPase hybrid poplar relative to levels in wild-type trees.  Fold change relative to wild-type poplar  Compound  232.6 11.4  Salicylic acid 2-O-f3-D-glucoside Maltose  5.8  Unknown  5.3  Myo-lnositol  5.1  Maltotriose methoxyamine  4.4  Galactinol  4.1  D-pinitol  3.0  Unknown  2.7  Galactitol  2.5  Unknown  2.5  Glucose  2.5  Galactose  2.3  Sucrose  2.2  Unknown  2.2  Sorbose  2.1  4-amino butyric acid  2.0  Fructose  2.0  Picein  112  transgenic lines with increases relative to wild-type poplar ranging from 175 to 270-fold (Table 4.6 and Table B3).  Discussion This study investigated the effects of overexpressing the Acetobacter xylinum UGPase gene in hybrid poplar under the control of a constitutive (2x35S) promoter on tree growth and cell wall biochemistry. Transcript quantification revealed a much higher level of transcript abundance in the leaves of the 2x35S::UGPase lines relative to stem (developing xylem). Consistent with the observed elevated levels of transcription, the total UGPase enzyme activity followed a similar pattern in the leaves. The effect of the introduction of the exogenous UGPase transgene from Acetobacter xyiinum on the transcript levels of the two native poplar UGPase gene was also examined, and an increase in both PtUGPI and PtUGP2 was shown, in both leaf and developing xylem (Table Bi). However, it was apparent that altered levels of expression of the two genes responded differently, with increases ranging from —2-4-fold in developing xylem while a maximum 2-fold change was apparent in the leaf tissue. Meng et a!, (2007), however, have shown tight post-transcriptional! translational control for these two genes, and therefore, increases in transcript abundance, while expected with an increase in sucrose content, may not necessarily correlate to an increase in protein/enzyme activity. The 2x35S::UGPase transgenic lines were shown to be substantially smaller than wild-type trees, with decreased height growth, diameter and stem dry weight (biomass). Morphologically, the 2x35S::UGPase trees, despite having a significantly greater number of leaves, had a reduced total leaf area and biomass compared to the corresponding wild-type trees. These same lines exhibited elongation of the axial shoots, which is consistent with weakening or loss of apical dominance resulting in changes in the auxin/cytokinin ratios. For example, in pea plants it has been shown that the removal of the apex resulted in the elongation of lateral shoots, with the exogenous application of NAA resulting in the retention of unaltered phenotype (Li et a!. 1995). The same result was seen when the apex was removed from chick pea plants, or when exogenous cytokinins were applied (Turnbull et a!. 2004). 113  To date, most of the work elucidating the function of UGPase is mixed and inconclusive, and has evolved from a limited number of studies downregulating UGPase activity.  For example, reductions from 30 to 96% in UGPase activity by antisense suppression showed no change in plant biomass in either Arabidopsis or potato, respectively (Johansson 2003; Zrenner et a!. 1993). In the case of Arabidopsis, Johansson (2003) speculated that the 30% reduction in Arabidopsis UGPase expression did not confer a phenotype as multiple UGPase isoforms exist permitting a compensatory control of pathway regulation (Johansson 2003). These studies imply that UGPase activity is abundant in the plant, and misregulation will likely have little or no effect on plant phenotype. Contrary to this, the overexpression of transgenic UGPase in tobacco resulted in significantly improved growth rates, both under ubiquitous and tissue-specific promoters (Coleman et a!. 2006).  The improved growth characteristics were attributed to the potential role of UGPase in altering sink strength as it participates in the catabolism of sucrose into hexose phosphates. Although similar results were not observed in the current study investigating hybrid poplar, the differences may be attributed to the innate differences in mechanisms of carbon translocation, with tobacco employing an active mechanism (Burkle et al. 1998) and poplar using a passive system moving assimilate along a sucrose concentration gradient (Turgeon and Medville 1998). UGPase in source tissues tends to act in the formation of sucrose, providing the substrate to sucrose transporters for active loading from the leaves (Kleczkowski 1994). In poplar, photoassimilate accumulating in sink cells must be lower than the solute concentration in source cells, and this occurs primarily by temporary storage in starch, or permanent storage in structural carbohydrates such as cellulose. UGPase in sink tissue tends to function in the breakdown of sucrose, while in source cells it is thought to act in the synthesis of sucrose (Kleczkowski et al. 2004), thus creating the concentration gradient necessary for the passive flow of carbon skeletons required for active metabolism. Altered UGPase activity in other plant species has also been shown to manifest changes in carbohydrate biochemistry, however, these changes are inconsistent and  conflicting.  In potato, Zrenner et a!. (1993) showed no change in carbohydrate metabolism with a 96% reduction in UGPase enzyme activity, while other studies with  less extreme reductions in UGPase activity showed significant decreases in stored 114  tuber carbohydrate concentrations (Spychalla et al. 1994; Borokov et al. 1996). Similarly, a 30% reduction in UGPase activity resulted in a decrease in soluble sugar and starch content in Arabidopsis (Kleczkowski et a!. 2004).  Consistent with these  findings, tobacco plants overexpressing UGPase showed increases in stem glucose and fructose levels compared to non-transgenic control plants (Coleman et a!. 2006). In the current study, significant increases in all soluble carbohydrates were observed in the leaf tissue of 2x35S::UGPase hybrid poplar.  In the developing xylem, the  2x35S::UGPase transgenic lines showed significant, yet smaller, increases in only primarily glucose and sucrose. These findings support the theory that UGPase activity can contribute to photoassimilate generation in source tissue and, in the case of symplastic loading plants like hybrid poplar, that this elevated activity in sink tissue can augment carbon allocation. In all of the 2x35S::UGPase transgenic lines, there was a significant increase in transcript, enzyme activity and consequently total soluble carbohydrate. Additionally, many of the transgenic lines showed increased levels of starch accumulation pointing to accumulation of storage carbohydrates resulting from the pooling of available soluble sugars. It appears that the expression of the exogenous AxUGPase in the leaf tissue is resulting in biosynthesis of carbohydrates in source tissue and is likely working co ordinately with SPS in the synthesis of sucrose, as has been proposed by Kleczkowski (1994).  Interestingly,  the  introduction  of  the  exogenous  UDP-glucose  pyrophosphorylase from Acetobacter appears to have manifested an increase in transcript abundance of both endogenous poplar UGPase genes, and therefore it is difficult to ascertain if the observed effects are independent of the native gene/protein. Despite not having measured metabolic flux, it appears that the increased biosynthesis of sucrose in the source tissue has resulted in an increase in the transport of photoassimilate to the sink tissue, as in all transgenic lines an accumulation of both storage  and  structural  carbohydrates  was  apparent.  Although  changes  in  hemicellulose-derived carbohydrates were observed, the most dramatic increase in cell wall carbohydrate chemistry was observed in glucose concentrations, which is derived from cellulose. In these lines, increases in cellulose content ranged from —2.8 to 6.5% (Figure 4.4). These findings are consistent with the theory that an increase in cellulose content would coincide with an increased supply of precursor to the cellulose synthase 115  complex (CSC) from elevated levels of soluble metabolites synthesized and likely transported to the sink (wall developing tissue). Furthermore, this is supported by the 4fold increase in the secondary wall specific CSC gene expression levels in the transgenic lines. Confirmation of this theory and our results are also consistent with the microarray findings of Hertzberg et a!. (2001) who showed that UGPase is upregulated during the period of late expansion and secondary cell wall formation. Similar changes in structural carbohydrates were not observed in tobacco plants transformed with the same constructs (Coleman et a!. 2006), which can be attributed to a lack of any measurable change in stem sucrose content compared to the stem of the poplar. In addition to the observed increases in carbohydrate composition in the 2x35S::UGPase transgenic trees, there was also an associated decrease in lignin content.  This is not thought to be the result of a decrease in the rate of lignin  deposition, but rather the result of a change in the ratio of cellulose to lignin. This was confirmed using real time PCR analysis of the lignin-branch of the phenylpropanoid biosynthetic genes toolbox, which showed that there was no decrease in the transcripts of any of the key lignin biosynthetic gene isoforms surveyed, rather in many of the genes there was an increase (Figure 4.3, Table B2). Correlated to the change in the relative amount of lignin, was a change in lignin monomer composition in favour of syringyl units.  These biochemical findings were  supported by real time PCR evaluations, which demonstrated significant augmentation of ferulate 5-hydroxylase (F5H) consistent with the change in monomer composition of the cell wall lignin. These findings suggest that the elevated transcript abundance of lignin biosynthetic genes may be related to changes in sugar concentrations rather than a change in cellulose deposition.  Sucrose has been shown to serve as a signal  molecule regulating gene expression (Wiese et a!. 2004), and consequently influences associated metabolic pathways and morphological development (Lunn and McRae 2003; Gibson 2005).  A compelling body of evidence indicates that carbohydrates,  particularly the hexose glucose, are essential sources of carbon skeletons and function as important signalling molecules (reviewed in Smeekens 2000; Rolland et al. 2002; Gibson 2005). In Arabidopsis, a relationship between carbon availability and lignin accumulation has been established (Rogers et a!. 2005), clearly demonstrating that metabolizable 116  carbohydrates positively influence the abundance of lignin.  Further, concurrent  transcriptome analysis lends support to the hypothesis that carbohydrates are not merely a source of carbon skeletons for lignification, but also function as a signal to enhance the capacity to synthesize these key cell wall macromolecules. Additionally, diurnal fluxes in lignin biosynthetic capacity were suggested to be modulated at the transcriptional level by at least three different stimuli: light, the circadian clock, and available hexose carbohydrates.  The absolute abundance of these transcripts is  shaped by the amount of available carbohydrates. The link between sugar signalling and lignification is particularly interesting, as carbohydrate-mediated changes in vegetative development have been well documented in dark-grown seedlings (Roldan  et a!. 1999; Baier et a!. 2004). Metabolic profiling was employed to investigate changes in “global” cell wall metabolism as a result of this single gene misregulation.  In addition to the altered  levels of soluble carbohydrates identified by HPLC, the 2x35S::UGPase transgenic poplar trees differentially accumulated several metabolites, many of which have been identified as products of sugar metabolism, such as galactose, maltose, and sorbose, or as metabolites commonly associated with abiotic or biotic stress (Table 4.5). Aside from being an integral component of cell wall hemicellulose, the monosaccharide galactose is also known to be a major carbohydrate participating in the formation of the raffinose family of oligosaccharides known to be elevated in response to stress, particularly in Populus. Maltose on the other hand is the immediate by-product of starch degradation, while sorbose is a precursor ketose monosaccharide involved in the biosynthesis of ascorbic acid, which has been shown to influence leaf development, growth and size (Chen and Gallie 2006), and may be responsible directly or indirectly for the alteration of leaf morphology in the current study.  In addition, other stress-  related compounds were identified including myo-inositol, galactinol, and pinitol. Myo iriositol and galactinol are the immediate precursors to the formation of raffinose family of oligosaccharides, while pinitol is a methylated cyclitol, derived from myo-inositol, thought to be an important osmolyte in plants responding to drought stress. Most interesting was the extremely high quantities of a single compound, the glycoside of salicylic acid (salicylic acid 2-O-3-D-glucoside [SAG]), which was found at significantly elevated levels (up to a 230-fold increase compared to the wild-type trees) 117  in the developing xylem of all 2x35S::UGPase transgenics trees (Table 4.6 and Table B3).  Salicylic acid 2-O-J3-D-glucoside was also identified in the leaf tissue of all  transgenic lines, where it was absent in the wild-type trees. The formation SAG is has been shown in tobacco to be catalyzed by a UDP-glucose:SA glucosyltransferase, which employs UDP-glucose as the sole glucose donor (Lee and Raskin 1999). Furthermore, it has been shown to be induced by the presence of salicylic acid, which has been shown to accumulate in tobacco leaves following TMV infections, resulting in an accumulation of SAG as a major product and glucosyl salicylate (GS) as a minor, less stable metabolite (Lee and Raskin 1998). Our results suggest that the accelerated generation of UDP-glucose, manifested by the overexpression of 2x35S::UGPase in hybrid poplar, resulted in the substantial accumulation of SAG.  It is tempting to  speculate that the allocation of UDP-glucose to salicylic acid may be a direct response of the inability of the cellulose synthase complex to effectively utilize the intracellular UDP-glucose channelled to the formation of cellulose, which was increased by -6%. However, the accumulation of SAG, among other compounds, has also recently been observed in transgenic aspen overexpressing sucrose phosphate synthase (Hjältén et a!. 2006), which should have provided another sink for UDP-glucose. Salicylic acid has been shown to act as a signalling molecule in local defence reactions and also in the induction of systemic resistance (Dumer et a!. 1997), and as such the increases in salicylic acid, and hence SAG and/or GS, may be the catalyst for the increases in other plant defence metabolites and may improve herbivore-plant interactions, as shown by Hjältén et a!. (2006). In summary, the overexpression of iJGPase in hybrid poplar resulted in significant increases in soluble sugars, more-so in the leaves than the developing xylem.  These increases in sugar appear to provide increased substrate to both cellulose and starch synthesis, resulting in changes in the chemical composition of the  stem, with as much as a 6.6% increase in cell wall cellulose content being observed. Contrary to the results observed in tobacco, in hybrid poplar results suggest that the overexpression of UGPase under the control of a ubiquitous promoter can alter carbon partitioning to starch and cellulose. However, the alterations in sucrose metabolism, particularly in the symplastically loading plant Populus, appears to cause other down stream repercussions within the plant, particularly with respect to carbohydrate 118  signalling and sensing, which can augment cell wall biosynthesis, as is evident in the altered lignin biosynthesis.  In addition to changes in wood chemistry, the trees also  produced more defence-related metabolites, which may explain the decreased energy directed to growth.  Supplemental Data The data reported in this manuscript are supported by supplemental data, which is available in appendix B. Included in the supplemental data is transcript abundance of the native poplar UDP-glucose pyrophosphorylase genes in leaf and developing xylem tissue (Table BI), as well as transcript abundance of the cell wall biosynthetic genes involved in lignin and cellulose deposition for transgenic and wild-type trees (Table B2). Additionally, quantification of the fold change in salicylic acid 2-O-3-glucoside in the developing xylem of all transgenic 2x35S::UGPase hybrid poplar relative to levels in wild-type trees is available (Table B3).  Acknowledgements The authors would like to thank Lisa McDonnell, Ji-Young Park, Ahn Nguyen, Andrew Robinson, and Victoria Maloney for their technical assistance. Funding for this project to SDM is acknowledged from the Natural Sciences and Engineering Research Council of Canada, CelIFor Inc., and the Canadian Forest Service.  119  References Appeldoorn N.J.G., de Bruijn S.M., Koot-Gronsveld E.A.M., Visser R.G.F., Vreugdenhil D. and van der Plas L.H.W. 1997. Developmental changes of enzymes involved in conversion of sucrose to hexose-phosphate during early tuberisation of potato. Planta 202: 220-226. Baler M., Hemmann G., Holman R., Corke F., Card R., Smith C., Rook F. and Bevan M.W. 2004. Characterization of mutants in Arabidopsis showing increased sugarspecific gene expression, growth, and developmental responses. Plant Physiology 134: 81-91. Borokov A.Y., McClean P.E., Sowokinos J.R., Ruud S.H. and Secor GA. 1996. Effect of expression of UDP-glucose pyrophosphorylase ribozyme and antisense RNAs on the enzyme activity and carbohydrate composition of field-grown transgenic potato plants. Journal of Plant Physiology 147: 644-652. Brede G., Fjaervik E. and Valla S. 1991. Nucleotide sequence and expression analysis of the Acetobacter xylinum uridine diphosphoglucose pyrophosphorylase gene. Journal of Bacteriology 173: 7042-7045. Burkie L., Hibberd J.M., Quick W.P., Kuhn C., Hirner B. and Frommer W.B. 1998. The W-sucrose cotransporter NtSUT1 is essential for sugar export from tobacco leaves. Plant Physiology 118: 59-68. Canam T., Park J.-Y., Yu K.Y., Campbell M.M., Ellis D.D. and Mansfield S.D. 2006. Varied growth, biomass and cellulose content in tobacco expressing yeastderived invertases. Planta 224: 1315-1327. Chen Z. and Gallie D.R. 2006. Dehydroascorbate reductase affects leaf growth, development, and function. Plant Physiology 142: 775-787. Coleman H.D., Ellis D.D., Gilbert M. and Mansfield S.D. 2006. Up-regulation of sucrose synthase and UDP-glucose pyrophosphorylase impacts plant growth and metabolism. Plant Biotechnology Journal 4: 87-101. Datla R.S.S., Bekkaoui F., Hammerlindi J.K., Pilate G., Dunstan D.l. and Crosby W.L. 1993. Improved high-level constitutive foreign gene translation using an AMV RNA4 untranslated leader sequence. Plant Science 94: 139-149. Durner J., Shah J. and Klessig D.F. 1997. Salicylic acid and disease resistance in plants. Trends in Plant Science 2: 266-274. Gibson S.I. 2005. Control of plant development and gene expression by sugar signaling. Current Opinion in Plant Biology8: 93-102.  120  Hertzberg M., Aspeborg H., Schrader J., Andersson A., Erlandsson R., Blomqvist K., Bhalerao R., Uhien M., Teen T.T., Lundeberg J., Sundberg B., Nilsson P. and Sandberg G. 2001. A transcriptional roadmap to wood formation. Proceedings of the National Academy of Sciences of the United States of America 98: 1473214737. Hjalten J., Lindau A., Wennstrom A., Blomberg P., Witzell J., Hurry V. and Ericson L. 2006. Unintentional changes of defence traits in GM trees can influence plantherbivore interactions. Basic and Applied Ecology 8: 434-443. Hood E.E., Gelvin S.B., Melcher L.S. and Hoekema A. 1993. New Agrobacterium helper plasmids for gene transfer to plants. Transgenic Research 2: 208-218. Johansson H. 2003. Gene regulation of UDP-glucose synthesis and metabolism in plants. Umea, Sweden, Umea University. Kay R., Chan A., Daly M. and McPherson J. 1987. Duplication of CaMV 35S promoter sequences creates a strong enhancer for plant genes. Science 236: 1299-1302. Kleczkowski L.A. 1994. Glucose activation and metabolism through UDP-glucose pyrophosphorylase in plants. Phytochemistiy 37: 1507-1 515. Kleczkowski L.A., Geisler M., Ciereszko I. and Johansson H. 2004. UDP-glucose pyrophosphorylase. An old protein with new tricks. Plant Physiology 134: 912918. Kolosova N., Miller B., Ralph S., Ellis B.E., Douglas C., Ritland K. and Bohlmann J. 2004. Isolation of high-quality RNA from gymnosperm and angiosperm trees. BioTechniques 36: 821-824. Kotake T., Yamaguchi D., Ohzono H., Hojo S., Kaneko S., Ishida H.-K. and Tsumuraya Y. 2004. UDP-sugar pyrophosphorylase with broad substrate specificity toward various monosaccharide 1-phosphates from pea sprouts. Journal of Biological Chemistiy 279: 45728-45736. Lee H. and Raskin I. 1998. Glucosylation of salicylic acid in Nicotiana tabacum Cv. Xanthi-nc. Phytopathology 88: 692-697. Lee H. and Raskin I. 1999. Purification, cloning, and expression of a pathogen inducible UDP-glucose: salicylic acid glucosyltransferase from tobacco. The Journal of Biological Chemistiy 274: 36637-36642. Li C.-J., Guevara E., Herrera J. and Bangerth F. 1995. Effect of apex excision and replacement by 1-naphthylacetic acid on cytokinin concentration and apical dominance in pea plants. Physiologia Plantarum 94: 465-469.  121  Li L., Zhou Y., Cheng X., Sun J., Marita J.M., Ralph J. and Chiang V.L. 2003. Combinatorial modification of multiple lignin traits in trees through multigene cotransformation. Proceedings of the National Academy of Sciences of the United States of America 100: 4939-4944. Lunn J.E. and MacRae E.A. 2003. New complexities in the synthesis of sucrose. Current Opinion in Plant Biology 6: 208-2 14. Magel E., Abdel-Latif A. and Hampp R. 2001. Non-structural carbohydrates and catalytic activities of sucrose metabolizing enzymes in twnks of two Juglans species and their role in heartwood formation. Holzforshung 55: 135-145. McCown B.H. and Lloyd G. 1981. Woody Plant Medium (WPM) a mineral nutrient formulation for microculture for woody plant species. Horticultural Science 16: 453. -  Meng M., Geisler M., Johansson H., Mellerowicz E., Karpinski S. and Kleczkowski L.A. 2007. Differential tissue/organ-dependent expression of two sucrose- and coldresponsive genes for UDP-glucose pyrophosphorylase in Populus. Gene 389: 186-1 95. Robinson A.R., Gheneim R., Kozak R.A., Ellis D.D. and Mansfield S.D. 2005. The potential of metabolite profiling as a selection tool for genotype discrimination in Populus. Journal of Experimental Botany 56: 2807-2819. Rogers L.A., Dubos C., Cullis l.F., Surman C., Poole M., Willment J., Mansfield S.D. and Campbell M.M. 2005. Light, the circadian clock, and sugar perception in the control of lignin biosynthesis. Journal of Experimental Botany 56: 1651-1663. Rolando C., Monties B. and Lapierre C. 1992. Thioacidolysis. Methods in Lignin Chemistry. S. Lin and C. Dence. Berlin, Germany, Springer-Verlag: 334-349. Roldan M., Gomez-Mena C., Ruiz-Garcia L., Salinas J. and Martinez-Zapater J.M. 1999. Sucrose availability on the aerial part of the plant promotes morphogenesis and flowering of Arabidopsis in the dark. Plant Journal 20: 581-590. Rolland F., Moore B. and Sheen J. 2002. Sugar sensing and signaling in plants. Plant Cell 14: S185-S205. Smeekens S. 2000. Sugar-induced signal transduction in plants. Annual Review of Plant Physiology and Plant Molecular Biology 51: 49-81. Spychalla J.P., Scheffler B.E., Sowokinos J.R. and Bevan M.W. 1994. Cloning, antisense RNA inhibition, and the coordinated expression of UDP-glucose pyrophosphorylase with starch biosynthetic genes in potato tubers. Journal of Plant Physiology 144: 444-453. 122  Sterky F., Bhalerao R.R., Unneberg P., Segerman B., Nilsson P., Brunner A.M., Charboneel-Campaa L., Lindvall J.J., Tandre K., Strauss S.H., Sundberg B., Gustafsson P., Uhlen M., Bhalerao R.P., Nilsson 0., Sandberg G., Karisson J., Lundeberg J. and Jansson S. 2004. A Populus EST resource for plant functional genomics. Proceedings of the National Academy of Sciences of the United States of America 101: 13951-13956. Turgeon R. and Medville R. 1998. The absence of phloem loading in willow leaves. Proceedings of the National Academy of Sciences of the United States of America 95: 12055-1 2060. Turnbull C.G.N., Raymond M.A.A., Dodd I.C. and Morris S.E. 2004. Rapid increases in cytokinin concentration in lateral buds of chickpea (Cicer arietinum L.) during release of apical dominance. Planta 202: 271-276. Wiese A., Elzinga N., Wobbes B. and Smeekens S. 2004. A conserved upstream open reading frame mediates sucrose-induced repression of translation. Plant Cell 16: 1717-1729. Yokoyama T., Kadla J.F. and Chang H.M. 2002. Microanalytical method for the characterization of fiber components and morphology of woody plants. Journal of Agriculture and Food Chemistry 50: 1040-1044. Zrenner R., Willmitzer L. and Sonnewald U. 1993. Analysis of the expression of potato uredinediphosphate-glucose pyrophosphorylase and its inhibition by antisense RNA. Planta 190: 247-252.  123  Overexpression of Sucrose Synthase and UDP-Glucose Pyrophosphorylase in Hybrid Poplar Affects Cellulose Partitioning and Ultrastructure 4 Introduction Sucrose is the primary translocatable carbohydrate in the majority of plants. As such its metabolism is vital to the regulation of photoassimilate in sink tissues (Asano et al., 2002). Sucrose hydrolysis can be catalyzed by two enzymes:  invertase, which  cleaves sucrose into glucose and fructose, and sucrose synthase (SuSy), which catalyzes the formation of UDP-glucose and fructose from sucrose.  Both SuSy and  invertase have been shown to be tightly associated with the processes of phloem unloading, and SuSy has been repeatedly identified as playing a central role in modulating sink strength (Sun et a!., 1992; Zrenner et a!., 1995; Dejardin et al., 1999). Furthermore, SuSy can be found in very high levels in companion cells (Nolte and Koch, 1993).  SuSy has also been linked with the synthesis of both storage and  structural carbohydrates, acting as the catalyst in the metabolism of sucrose which results in the liberation of the precursor for the generation of callose (Subbaiah and Sachs, 2001), cellulose (Amor,  1995), and mixed linkage 3(1-3),(1-4)-glucans  (Buckeridge et al., 1999). Finally, SuSy expression has also been correlated with the development and deposition of secondary xylem in trees from the vascular cambium (Hauch and Magel, 1998; Hertzberg eta!., 2001). SuSy has dual functionality, in that it provides the immediate precursor for cellulose biosynthesis and concomitantly recycles UDP which has been identified as an inhibitor of cellulose biosynthesis (Benziman et a!., 1983). It has been suggested that SuSy exists in two forms, a soluble (S-SuSy) and particulate (P-SuSy) form, with the  A version of this chapter will be submitted for publication. Coleman HD, Yan J, Mansfield SD (2008) Overexpression of sucrose synthase and UDP-glucose pyrophosphorylase in hybrid poplar affects cellulose partitioning and ultrastructure.  124  latter being membrane bound and directly supplying UDP-glucose to the cellulose synthase complex for cellulose biosynthesis (Amor et a!., 1995). As such, the high energy bond is retained for use in the synthesis of polysacoharides.  In maize,  phosphorylation of SuSy has been shown to alter the enzyme from a membrane bound to a soluble form (Winter et a!., 1997, Komina et al., 2002), but this phosphorylation is reversible (Winter and Huber, 2000). A mutant form (replacement of Ser-Il by Glu-Il) of the mung bean SuSy (SI I E), which has a higher catalytic efficiency with respect to sucrose, was overexpressed in transgenic poplar (Populus alba) and investigated using a dual labelling system (Konishi et al., 2004). Using this system, it was shown that only a fraction of the sucrose loaded into the phloem is directly used by the CesA complex associated with SuSy, thus conserving the high energy bond for use in polysaccharide synthesis. Altering the expression of SuSy has also been shown to cause changes in structural and storage carbohydrates in other species.  For example, maize SuSy  mutants displayed increased sucrose levels (Chourey & Nelson, 1976), while increased glucose and fructose levels were apparent in tobacco with SuSy overexpression (Coleman et a!., 2006). In wheat, natural variations in SuSy levels demonstrated a clear association between SuSy activity and increased cell wall polysaccharide levels (Xue et al., 2007). Similar results have been observed in wheat roots, with high SuSy activity (caused by hypoxia) being associated with increased cellulose content (Albrecht & Mustroph, 2003). Studies investigating gene expression patterns in poplar have also identified SuSy as being associated with cellulose synthesis and with tension wood formation where increased cellulose deposition occurs (Hertzberg et a!., 2001; Andersson-Gunnerás et a!., 2006).  As such, increasing the expression of SuSy  appears to be a key target to accelerate or improve the production of cellulose within forest trees. In addition to SuSy, UDP-glucose pyrophosphorylase (UGPase) acts as a source for the synthesis of UDP-glucose, the precursor to cellulose biosynthesis.  UGPase  phosphorylates glucose I-phosphate producing UDP-glucose, while SuSy cleaves sucrose supplied to the cell.  UGPase can be involved in the recycling of fructose  produced by SuSy, which can act as an inhibitor of SuSy activity. In source tissues, UGPase works in conjunction with sucrose phosphate synthase in the synthesis of 125  sucrose (Kleczkowski, 1994), while in sink tissues, UGPase works with SuSy in the cycling of sugars between sucrose and the hexose phosphate pools (Borokov et a!., 1996). Overexpression of UGPase in poplar under the control of the 2x35S promoter resulted in significant increases in cellulose, but decreased biomass production (Coleman et a!., 2007) confirming the potential for altering cellulose synthesis by manipulating UDP-glucose pools. This paper investigates the effect of misregulating SuSy and UGPase in hybrid poplar. SuSy was upregulated individually and in combination with UGPase, and the ensuing trees assessed for changes in transcript abundance, enzyme activity, biomass production, storage and structural polysaccharides, as well as cell wall crystallinity and microfibril angle.  The results clearly show that SuSy can indeed modulate the  production of cellulose, and its ultrastructural characteristics.  Methods Plasmid Construction SuSy was cloned from Gossypium hirsutum (Perez-Grau, GENBANK U73588) as previously described (Coleman et a!., 2006) and inserted into pBlN under the  regulation of one of two promoters: the enhanced tandem cauliflower mosaic virus 35S constitutive promoter (2x35S) (Datla et a!., 1993; Kay et a!., 1987) or the vascularspecific 4CL (Petroseilnum crispum 4-coumarate:CoA ligase) promoter (Hauffe et a!., 1991). UGPase was cloned from Acetobacterxylinum ATCC #23768 and inserted into the pBIN cloning vector under the control of the same two promoters. Sequence analysis was used to confirm insertion of the promoter and gene into the binary vector (Coleman et a!., 2006).  Plant Transformation and Maintenance Hybrid  poplar (Populus alba  x  grandidentata)  Agrobacterium tumefaciens EHAIO5 (Hood et a!., 1993).  was transformed  using  In short, wild-type and  UGPase transgenic poplar (Coleman et a!., 2007) leaf disks were cut and co-cultured with EHAIO5 harbouring the 2x35S::SuSy or 4CL::SuSy transgenes.  The explants 126  were plated on Woody Plant Media (WPM) (McCown and Lloyd 1981) supplemented with 0.1 pM each of a-naphthalene acetic acid (NAA), 6-benzylaminopurine (BA), and thiadiazuron (TDZ) and solidified with 3% (w/v) agar and 1.1% (w/v) phytagel (WPM 0.1/0.1/0.1).  After three days the disks were transferred to WPM 0.1/0.1/0.1  supplemented with carbenicillin disodium (500 mg L’) and cefotaxime sodium salt (250 mg L ). Following three additional days, the disks were transferred to WPM 0.1/0.1/0.1 1 ). 1 containing carbenicillin, cefotaxime and kanamycin (25 mg L  After five weeks,  shoots and callus material were transferred to WPM containing agar and phytagel, supplemented with 0.01 pM BA, carbenicillin, cefotaxime and kanamycin.  Once  individual shoots were visible, plantlets were transferred to solidified WPM with 0.01 pM NAA and carbenicillin, cefotaxime and kanamycin to induce rooting.  After two  consecutive five-week periods on this media, shoot tips were isolated to solidified antibiotic-free WPM with 0.01 pM NAA. Plants were confirmed as transgenic using PCR screening of genomic DNA employing gene specific oligonucleotides: specifically, UGP-F (5’-atcgaggaattctgcctcgt3’) and UGP-R (5’-tcgcaagaccggcaacaggatt-3’) were used for UGPase identification, and SUS-1 (5’-ctcaacatcacccctcgaat-3’) and SUS-2 (5’-accaggggaaacaatgttga-3’) were employed for SuSy confirmation. All shoot cultures, including transgenic and non-transformed wild-type lines, were maintained on solid WPM with 0.01 pM NAA in GA-7 vessels at 22CC under a 16-hour 2 s until out-planting to the photoperiod with an average photon flux of 50 pmol m greenhouse. Growth Conditions and Biomass Measurements Wild-type control trees and each transgenic line, represented by a minimum of 12 individual trees, were transferred into 7.5 L pots containing 50% peat, 25% fine bark, and 25% pumice soil mixture in the greenhouse, and allowed to acclimate under 16 oz clear plastic cups for one week. The trees were grown under 16 hour days . Following 2 supplemented with overhead lighting with a radiant flux density of 300 W m four months growth in the greenhouse, the trees were harvested and total height and stem diameter (10 cm above root collar) determined for each tree. 127  Tissue developmental stage was standardized using a plastichron index, where P1=0 was defined as the first leaf greater than 5 cm in length, and P1=1 is the leaf immediately below P1=0.  Stem segments spanning P1=5 to P1=15 were retained for  wood cell wall and chemical analysis. Leaves from P1=3 to P1=5 were frozen in liquid nitrogen and retained for RNA, enzyme and soluble carbohydrate analysis. Developing xylem was scraped and flash frozen in liquid nitrogen for analysis of enzyme activity, RNA transcript abundance, and soluble carbohydrate content.  Transcription Levels Real time PCR was employed to determine transcript abundance of each transgene.  Leaf and developing xylem samples (1 g f.w.) were ground in liquid  nitrogen, and RNA extracted according to the method of Kolosova et al. (2004). Ten pg TM (Ambion, Austin, TX) to remove of RNA was then treated with TURBO DNase residual DNA.  One pg of DNase-treated RNA was used for the synthesis of cDNA  using Superscript II Reverse Transcriptase (Invitrogen, Carlsbad, CA) and dT 16 primers according to the manufacturer’s instructions. Samples were run in triplicate with Brilliant SYBR Green QPCR Master Mix (Stratagene, La Jolla, CA) on an Mx3000P Real-Time PCR System (Stratagene) to determine critical thresholds (Ct). The primers employed for RT-PCR analysis of UGPase were AU-RTF (5’-tggaagcaacccgcgtcatc-3’) and AU RTR (5’-gccaaggcccagcggttcc-3’), while the primers for SuSy were GS-RTF (5’ccgtgagcgtttggatgagac-3’) and GS-RTR (5’-ggccaaaatctcgttcctgtg-3’).  As a house  keeping control, the transcript abundance of transcript initiation factor 5A (TIF5A) was employed for normalization (Ralph et a!., 2006).  The primers for TIF5A transcript  quantification were TI F5A-RTF (5’-gacggtattttagctatggaattg -3’) and TI F5A-RTR (5’ctgataacacaagttccctgc -3’). Conditions for the RT-PCR reactions were as follows: 95°C for 10 minutes, followed by 40 cycles of 95°C for 30 seconds, 62°C (64°C for SuSy, 55°C for TIF5A) for 1 minute, and 72°C for 30 seconds.  Relative expression was  determined according to Levy et a!. (2004) using the following equation: tct or SuSy— ctTIF5A)  128  Enzyme Activity  Leaf and developing xylem samples (1 g f.w.) were ground in liquid nitrogen with 1 mg of insoluble PVPP and four volumes of extraction buffer (50 mM HEPES-KOH pH  7.5, 10 mM MgCl, 1 mM EDTA, 2 mM DTT, 1 mM PMSF, 5 mM EAmino-n-caproic acid, 0.1% v/v Triton X-100, 10% v/v glycerol). The samples were centrifuged at 15,000xg for 20 minutes at 4°C. The extract was passed through a desalting column (DG 10  —  BioRad) pre-equilibrated with ice-cold extraction buffer without Triton X-100 and PVPP. Extracts were collected in pre-chilled vials and used immediately. UGPase activity was determined spectrophotometrically at 340 nm as per Appeldoorn et a!. (1997) using 100 jiL of plant extract and a NADH molar extinction coefficient of 6.22 mM cm . SuSy 1 activity was assayed in the direction of sucrose breakdown (Chourey, 1981), using 50 pL of plant extract. The resultant fructose content was determined using a tetrazolium blue assay (Kennedy and White, 1983). This SuSy assay employs the appropriate controls without the supplementation of UDP to quantify inherent invertase activity, and therefore represents only the breakdown of sucrose by SuSy. Total protein content of the extracts was determined using a Bio-Rad Protein Assay (Bio-Rad, Hercules, CA).  Soluble Carbohydrate and Starch Analysis  Soluble carbohydrates (glucose, fructose and sucrose) were extracted from ground freeze-dried tissue overnight at -20°C using methanol:chloroform:water (12:5:3) as previously described (Coleman et a!., 2006). supernatant  removed,  and  the  remaining  The sample was centrifuged, the pellet  washed  twice  with  fresh  methanol:chloroform:water (12:5:3). All fractions were then pooled. Five mL of water was added to the combined supernatant and centrifuged to facilitate phase separation. The aqueous fraction was rotary evaporated to dryness and resuspended in 3 mL of distilled water. Soluble carbohydrates were then analyzed using anion exchange HPLC (Dionex, Sunnyvale, CA) on a DX-600 equipped with a Carbopac PAl column and an electrochemical detector. The residual pellet was then hydrolyzed in 4% sulfuric acid at 121t for 4 minutes. The liberation of glucose, representing starch content, was directly quantified by HPLC under similar conditions. 129  Cell Wall Chemistry Oven dried stem material was ground using a Wiley mill to pass through a 40mesh screen and soxhlet extracted with acetone for 24 hours. Lignin and carbohydrate contents were determined using a modified Kiason method (Huntley et a!., 2003) and 0.2 g of extract free tissue. Carbohydrate content was determined using HPLC (Dionex DX-600, Dionex, CA) equipped with an anion exchange PAl column, a pulsed amperometric detector with a gold electrode and post-column detection. Acid insoluble lignin was determined gravimetrically, while acid insoluble lignin was determined using spectrophotometric analysis at 205 nm according to TAPPI Useful Method UM-250.  Determination of a-Cellulose and Holocellulose Content Extract-free ground wood was used to determine holocellulose and a-cellulose contents according to the method of Yokoyama et a!. (2002).  In short, 100 mg of  ground tissue was weighed into a 20 mL flask and placed in a 90°C heat block. The reaction was initiated by the addition of 0.5 mL of sodium chlorite solution (200 mg 80% sodium chlorite, 2 mL distilled water, 0.4 mL acetic acid). Additional 0.5 mL aliquots of solution were added every 30 minutes to a total volume of 2 mL. After 2 hours, the samples were removed to a cold water bath and filtered through a coarse crucible. Following overnight drying at 105°C, the holocellulose content was determined gravimetrically. 50 mg of dried holocellulose was then weighed into a reaction flask and allowed to equilibrate for 30 minutes. Four mL of 17.5% sodium hydroxide was added and permitted to react for 30 minutes, after which 4 mL of distilled water was added and the sample macerated for 1 minute. Following 29 minutes of reaction, the sample was filtered through a coarse crucible and placed in 1.0 M acetic acid for 5 minutes before washing with distilled water. a-cellulose content was determined gravimetrically following overnight drying at 105°C.  130  Crystallinity and Microfibril Angle Microfibril angle and cell wall crystallinity were determined by X-ray diffraction using a Bruker D8 Discover X-ray diffraction unit equipped with an area array detector (GADDS) on the radial face of the wood section precision cut to 1.69 mm from the growing stem. Wide-angle diffraction was used in the transmission mode, and the measurements were performed with CuKal radiation (A = 1.54 A), the X-ray source fit with a 0.5 mm collimator and the scattered photon collected by the GADDS detector. Both the X-ray source and detector were set to theta  =  0° for microfibril angle  determination, while 2 theta (source) was set to 17° for wood crystallinity determination. The average T-value of the two 002 diffraction arc peaks was used for microfibril angle calculations, as per the method of Megraw et a!. (1998), while crystallinity determined by mathematically fitting the data using the method of Vonk (1973). Two radii were taken from samples isolated 5 cm above ground on each tree, and these values were averaged for each tree.  Microscopy Cross sections of stems (40 pm) from 5 cm above the ground were cut using a microtome. Sections were mounted on glass slides and visualized using a Leica microscope under UV fluorescence. Histochemical examination of cellulose in the poplar stems was carried out using calcofluor staining. The samples were mounted in 10% KOH (wlv) and 0.1% calcofluor white (w/v) and then viewed after 5 minutes under bright-field illumination with a Leica DMR microscope equipped with a QICAM CCD camera (Q-imaging). Histochemical examination of lignin was analysed using phloroglucinol staining achieved by mounting stem sections in a saturated solution of phloroglucinol in 20% HCI. Samples were viewed under dark-field illumination on the Leica DMR microscope.  131  Results Transformed hybrid poplar trees regenerated from leaf tissue were propagated from single shoots from individual explants, and as such each line represents an individual transformation event.  From the transformed lines produced, three  2x35S::SuSy lines and four 4CL::SuSy lines were selected for greenhouse growth trials and in-depth cell wall characterization, based on enzyme activity. Together with corresponding wild-type controls, 12 individual trees per line were transferred to the greenhouse. At harvest, two of three 2x35S::SuSy lines (lines 1 and 2) and one of four 4CL::SuSy (line 2) lines were slightly shorter than the corresponding wild-type poplar (Figure 5.1), while the other lines exhibited similar height growth. Only one line, 2x35S::SuSy-1, had a decreased calliper and leaf dry weight, while one 4CL::SuSy -1 had an increased calliper (Figure 5.1). All other lines displayed similar phenotypes to the wild-type trees. There was no change in stem weight relative to controls in any of the transformed lines. Existing transgenic hybrid poplar trees expressing 2x35S::UGPase and 4CL::UGPase (Coleman et a!., 2007) were concurrently transformed with SuSy under control  of  the  4CL::UGPasexSuSy).  corresponding  promoter  (2x35S::UGPasexSuSy  and  Trees regenerated from leaf explants were again propagated  from single shoots from individual explants, and each line represents an individual SuSy transformation event. Despite several attempts, no 2x35S double transformed lines were recovered, which  is likely due to the severe phenotype observed in  2x35S::UGPase overexpressing transgenic lines (Coleman et al., 2007).  However,  four of 12 successfully transformed 4CL::UGPasexSuSy lines were selected, based on enzyme activity, and propagated for in-depth analysis. The double transformants were grown simultaneously with single gene transformed lines and controls trees in the greenhouse for four months. At harvest, two 4CL::UGPasexSuSy lines (lines 3 and 4) were  significantly  shorter  than  the  corresponding  wild-type  poplar,  and  4CL::UGPasexSuSy-3 also had decreased calliper. There were no changes in total dry weight (biomass) of leaf or stem (Figure 5.1).  132  g  E  x  26 0  a  24 22 20 18 16  12  0  10  0  E Co  Figure 5.1. Biomass measurements of SuSy and SuSy x UGPase transgenic and wildtype poplar. Tree height (A), calliper (B), leaf dry weight (C) and stem dry weight (D). Mean (±SE) were calculated from 10 plants per line. * Indicates a significant difference between the transgenic and wild-type trees at a=0.10.  133  Transcript Level and Enzyme Activity  Quantitative real time PCR was used to measure transcript abundance of the SuSy and UGPase transgenes relative to actin (Table 5.1). All lines clearly showed the existence of the exogenous gene and its regulation. In general, transcript abundance was higher in leaf tissue than in the developing xylem of single gene transgenic lines. However, in the double transgenic lines, transcript abundance of SuSy was higher in leaf tissue than in the developing xylem, while transcript abundance of UGPase was higher in the developing xylem than in the leaf tissue. Correspondingly, SuSy activity, as determined by the breakdown of sucrose into UDP-glucose and fructose, was increased in developing xylem of all of the transgenic lines (Table 5.1). In some lines, as much as a 2.5-fold increase was apparent.  Furthermore, two 2x35S::SuSy lines  (lines 1 and 2) and one 4CL::SuSy line (line 1) also had increased SuSy activity in the leaf tissue. All of the double transgenic lines showed increased SuSy activity in developing xylem, but not in the leaf tissue (Table 5.1).  Enzyme activity ranged from 13.26 to  17.42 pg fructose mg 1 protein min 1 compared to 4.99 pg fructose mg 1 protein min 1 in the control trees, representing a 2.5- to 3.5-fold increase in transgene enzyme activity. UGPase activity was determined using an indirect assay measuring the production of the reduced form of nicotinamideadenine dinucleotide (NADH).  All lines had  significantly increased UGPase activity in the developing xylem; however, the changes were not as substantial as those observed in the SuSy activity levels (Table 5.1). Activity ranged from 0.52 to 0.65 pmol NADH min 1 mg protein’ compared to 0.42 pmol NADH min’ mg protein 1 in the control trees, representing a 1.25- to 1.5-fold increase in activity.  Soluble Carbohydrates and Starch Content  Changes in soluble sugar content were variable (Table 5.2).  While there  appeared to be an increasing trend in total soluble sugars (glucose, fructose and sucrose) in the transgenic lines, few of the changes were significant. In leaf tissue, 4CL::SuSy-3 and 4CL::SuSy x UGPase-2 had increased total soluble sugars. In the 134  Table 5.1. Mean transcript abundance and enzyme activity in leaf and developing xylem tissue for SuSy and SuSy x UGPase transgenic and wild-type poplar trees. Mean (±SE) were calculated from 3 plants per line. Bold denotes significant difference from control values at a=0.10. SuSy Transcript Level Ct  SuSy Enzyme Activity 1 mg 1 pg fructose min  Transgenic line  Leaf  Developing Xylem  Leaf  Control 2x35S::SuSy 1 2x35S ::SuSy 2 2x35S ::SuSy 3  n.d. 2.50 ± 0.29  n.d. 0.18 ± 0.05  506.71 ± 45.66 1126.43 ± 178.08  2.34 ± 0.71  0.49 ± 0.23 0.14 ± 0.13  845.45 ± 59.84  4.99 ± 0.54 8.18 ± 0.50 9.22 ± 1.17  455.90 ± 63.48  11.95 ± 2.29  0.82 ± 0.53  788.67 ± 101.54  9.10 ± 0.34  0.12 ± 0.01  541.56 ± 24.32 680.56 ± 74.44 581.24 ± 105.83 679.99 ± 215.98  8.06 ± 0.84 12.77 ± 1.34 7.61 ± 0.87  4CL::SuSy 1  1.32 ± 0.06 0.59 ± 0.05  Developing Xylem  4CL::SuSy 2 4CL::SuSy 3 4CL::SuSy 4 4CL::SuSy x UGPase 1  1.17 ± 0.57 0.99 ± 0.19 0.00 ± 0.00 1.10 ± 0.20  0.34 ± 0.15 0.02 ± 0.01 0.27 ± 0.09  4CL::SuSy x UGPase 2 4CL::SuSy x UGPase 3  0.37 ± 0.10 0.30 ± 0.10  0.20 ± 0.08 0.24 ± 0.08  1424.04 ± 121.50 609.55 ± 202.16  13.26 ± 2.79 17.42 ± 0.39  4CL::SuSy x UGPase 4  1.14 ± 0.63  0.28 ± 0.09  514.18 ± 90.41  13.38 ± 2.29  UGPase Transcript Level Ct  UGPase Enzyme Activity pmoINADH min 1 mg 1 total protein  Leaf  Developing Xylem  Leaf  Control 4CL::SuSy x UGPase 1  n.d. 0.13 ± 0.01  n.d. 0.26 ± 0.15  0.15 ± 0.04 0.12 ± 0.01  4CL::SuSy x UGPase 2 4CL::SuSy x UGPase 3  0.09 ± 0.02 0.42 ± 0.04  0.24 ± 0.07 3.46 ± 0.41  4CL::SuSy x UGPase 4  0.61 ± 0.09  2.35 ± 1.54  0.12 ± 0.03 0.13 ± 0.03 0.12 ± 0.07  .  .  Transgenic line  16.70 ± 2.91  Developing Xylem 0.42 ± 0.03 0.65 ± 0.03 0.58 ± 0.07 0.52 ± 0.04 0.54 ± 0.02  135  Table 5.2. Total soluble carbohydrates and starch in leaf and developing xylem tissue of SuSy and SuSy x UGPase transgenic and wild-type trees. Mean (±SE) were calculated from 3 plants per line. Bold denotes significant difference from control values at a0.10. Leaf mg g 1 .  Soluble Carbohydrates  .  Transgenic Line Control 2x35S ::SuSy 1 2x35S ::SuSy 2 2x35S ::SuSy 3 4CL::SuSy 1 4CL::SuSy 2 4CL::SuSy 3 4CL::SuSy 4 4CL::SuSy x UGPase 4CL::SuSy x UGPase 4CL::SuSy x UGPase 4CL::SuSy x UGPase  1 2 3 4  159.66 ± 12.68 166.59± 11.75 191.24 ± 16.60 168.11 ± 16.87 185.51 ± 13.52 175.79 ± 14.73 193.50 ± 8.74 169.83 ± 13.48 185.73 ± 17.24 203.97 ± 20.78 181.16 ± 22.26 146.13 ± 16.62  Developing Xylem mg g Starch  Soluble Carbohydrates  0.76 ±0.15 0.83 ± 0.26 0.88 ± 0.31 0.68 ± 0.11 0.61 ± 0.13 1.28 ± 0.13 0.94 ± 0.35 0.90 ± 0.43 0.73 ± 0.13 1.12 ± 0.55 0,92 ± 0.23 0.44 ± 0.13  54.04 ± 7.36 61.81 ± 3.36 105.22 ± 30.85 134.27 ± 8.71 136.73 ± 21.27 91.52 ± 9.43 121.37 ± 9.48 87.91 ± 20.98 128.29 ± 7.48 93.98 ± 10.99 95.69 ± 38.44 71.09 ± 7.89  Starch 0.30 0.28 0.22 0.26 0.24 0.39 0.30 0.31 0.23 0.29 0.33 0.24  0.05 ± 0.01 ± 0.05 ± 0.08 ± 0.00 ± 0.07 ± 0.04 ± 0.06 ± 0.03 ± 0.02 ± 0.04 ± 0.05 ±  136  developing xylem, 2x35S::SuSy -3, 4CL::SuSy -1, 2, and 3 and 4CL::SuSy x UGPase-1 and 2 had significant increases in total soluble sugars. Starch content was unchanged in the leaves of all but one of the single transgenic plants (Table 5.2). 4CL::SuSy-4 had increased leaf starch content (1.28 mg ) relative to the control at 0.76 mg g 1 g . There were no changes in the starch content 1 of the developing xylem in any of the single transgenic lines. Furthermore, there were no significant changes in the starch content of leaves or developing xylem in any of the double transgenic lines.  Cell Wall Chemistry Cell wall carbohydrate content was substantially altered in the transgenic lines. All lines showed increased glucose content, with all 2x35S::SuSy lines, 4CL::SuSy-1 and 4 and 4CL::SuSy x UGPase-1, 3, and 4 lines displaying statistically significant increases in glucose content. 2x35S::SuSy-3, 4CL::SuSy-1 and 4CL::SuSy x UGPase 1, 2, and 4 also had increased mannose levels.  All lines showed decreases in  arabinose content, while in general galactose, rhamnose, and xylose remained relatively unchanged when compared to the wild-type trees (Table 5.3).  While there  were no changes in overall lignin content, a number of lines showed significant changes in acid soluble lignin content, and all lines showed a trend towards decreased acid soluble lignin.  Phloroglucinol staining confirmed the consistent level of lignin in the  transgenics relative to wild-type trees. The observed increases in glucose content, as determined by Klason analysis, were confirmed to be the result of an increase in cellulose content and not glucoseassociated with hemicellulose biosynthesis, by a-cellulose quantification (Table 5.4). All lines were confirmed to have increased cellulose content, ranging from 2 to 6% by weight more than wild-type trees. As a mechanism to investigate if the changes in cellulose were related to a change in cellulose production and not to a formation of tension wood, crystallinity and microfibril angle of all transgenic and wild-type stems were determined.  Crystallinity was increased in all but two transgenic lines ranging  from I to 12%, while none of the transgenic lines showed any changes in microfibril  137  c3 Co  14.36 ± 0.42  15.95±0.48  39.82±0.86 41.27 ± 2.49  42.33 ± 1.85 38.45 ± 0.37 40.80 ± 0.39  40.37±0.63  1.03±0.01 1.04±0.05 1.03 ± 0.11 0.99±0.04 1.03±0.07 1.18 ± 0.24 0.95 ± 0.02 0.91 ± 0.08  0.41 ±0.01 0.47±0.01 0.37 ± 0.01 0.42±0.05 0.40±0.04 0.36 ± 0.03 0.36 ± 0.06 0.43 ± 0.08  0.39±0.02  2x35S::SuSy3  4CL::SuSyl  4CL::SuSy 2  4CL::SuSy3  4CL::SuSy4  4CL::SuSy x UGPase 1  4CL::SuSy x UGPase 2  4CL::SuSy x UGPase 3  4CL::SuSyxUGPase4  0.96±0.02  0.99 ± 0.02  39.75±0.55  39.32±0.98  41.01 ±1.09  40.71 ± 0.87  15.74 ± 0.26  14.82 ± 0.79  1.98±0.10  0.51 ±0.01  0.48 ± 0.03  0.51 ± 0.04  2.41 ± 0.26 1.75 ± 0.18  0.54 ± 0.06  0.51 ±0.06  2.25 ± 0.19  1.91 ±0.12  15.78±0.43  0.47±0.03  0.49 ± 0.00  2.25 ± 0.42 2.22±0.33  0.54±0.02  0.51 ±0.01  0.51 ± 0.03  0.55 ± 0.02  0.50±0.01  Rhamnose  2.13±0.12  2.41 ±0.25  2.38 ± 0.38  2.34 ± 0.36  1.65±0.05  Mannose  15.86±0.22  15.16 ± 0.03  16.32±0.89  16.03±0.95  15.82 ± 0.74  15.33 ± 0.34  0.43 ± 0.03  40.97 ± 1.06  2x35S::SuSy 2  0.93 ± 0.03  0.36 ± 0.02  2x35S::SuSy 1  15.49±0.22  36.56±1.05  1.00±0.08  0.53±0.03  Control  Xylose  Glucose  Galactose  Arabinose  % Carbohydrates  2.68±0.12  3.05 ± 0.25  3.17 ± 0.17  3.14 ± 0.16  3.36±0.13  3.48±0.10  3.23 ± 0.30  3.26±0.16  3.56±0.43  3.16 ± 0.32  2.72 ± 0.02  3.94±0.26  Acid Soluble  20.69±0.48  20.28 ± 0.63  20.83 ± 1.04  19.57 ± 1.50  20.74±0.47  20.41 ±1.20  19.55 ± 0.51  19.65±0.33  22.50±1.78  20.92 ± 0.12  20.68 ± 0.55  20.15±0.46  Acid Insoluble  % Lignin  Table 5.3. Chemical composition of stem segments (P1=5 to P1=1 5) of SuSy and SuSy x UGPase transgenic and wild-type trees. Mean (±SE) were calculated from 3 plants per line. Bold denotes significant difference from control values at a=O.1O.  Table 5.4. a-cellulose content, cell wall crystallinity and microfibril angle (MFA) of stem  segments (P15 to P1=15) of SuSy and SuSy x UGPase transgenic and wild-type trees. Mean (±SE) were calculated from 3 plants per line. Bold denotes significant difference from control values at a=0. 10.  a-Cellulose (% DW)  Cell Wall Crystallinity  MFA  (%)  (°)  Control  31.96 ± 0.07  51.40 ± 0.51  16.70 ± 0.35  2x35S::SuSy 1  38.29 ± 1.22  56.60 ± 1.75  15.66 ± 0.81  2x35S::SuSy 2  36.61 ± 0.54  54.00 ± 1.00  16.63 ± 0.87  2x35S::SuSy 3  35.76 ± 1.20  56.40 ± 1.44  16.63 ± 0.55  4CL::SuSy 1  37.13 ± 0.04  53.40 ± 0.75  16.43 ± 1.31  4CL::SuSy 2  37.35 ± 0.87  53.25 ± 0.48  19.08 ± 1.22  4CL::SuSy3  35.90±0.47  52.00± 1.00  18.21 ± 0.88  4CL::SuSy 4  35.08 ± 0.46  54.00 ± 1.00  17.30 ± 1.04  4CL::SuSy x UGPase 1  37.43 ± 0.76  63.25 ± 2.17  16.38 ± 1.60  4CL::SuSy x UGPase 2  37.41 ± 0.87  56.80 ± 1.69  17.37 ± 0.53  4CL::SuSy x UGPase 3  33.84 ± 0.51  55.80 ± 1.83  17.45 ± 0.93  4CL::SuSy x UGPase 4  36.41 ± 0.81  60.00 ± 2.61  17.52 ± 0.53  139  angle (Table 5.4). Microscopy and staining of stem cross-sections also demonstrate the lack of a G-layer commonly associated with tension wood formation, as well as an increase in cellulose content, as implied by calcofluor staining, in all transgenic lines (Figure 5.2).  Discussion This study investigated the effects of overexpressing two exogenous genes, SuSy and UGPase (individually and in combination), on tree growth and cell wall chemistry and ultrastructure in hybrid poplar. Quantification of transcript abundance of the transgenes showed variable expression among lines and tissues. In general, the 2x35S::SuSy transgenic lines had higher expression in leaf tissue when compared to developing xylem, and similarly, higher transcript abundance than the 4CL::SuSy transgenic lines examined.  Despite the tissue specificity of the 4CL promoter, there  was no discernable difference in transgene expression in the developing xylem when comparing transcript abundance of 4CL::SuSy and 2x35S::SuSy transgenic lines.  In  the 4CL::UGPasexSuSy double transgenic lines, UGPase transcript level was higher in the developing xylem, while SuSy transcript abundance was higher in the leaf tissue. The difference in exogenous transcript abundance between leaf and xylem tissue was also reflected in the enzyme activity, where SuSy was much more active in the leaf tissue, when compared to the developing xylem. These observations are consistent with the wild-type trees which inherently have an approximate 100-fold difference in SuSy activity between leaf and developing xylem tissue (Table 5.1).  Furthermore,  these findings concur with Le Hir et al., (2005) who observed high levels of SuSy in leaf tissue of Querus robur, with undetectable levels in the stem. An obvious trend towards increased SuSy enzyme activity was apparent in all transgenic lines, with greater than two-fold increases being observed in the leaf tissue in some lines. Similar increases were observed in developing xylem, with greater than a 3-fold increase in some lines. In contrast, UGPase activity is inherently higher in the developing xylem, where as much as three times the activity per gram total protein is apparent. The observed 140  Figure 5.2. Calcofluor (A, B) and phioroglucinol (C, D) staining of wild-type (A, C) and 2x35S::SuSy transformed (B, D) poplar trees (Scale bars: 90 pm). Arrows depict the observed increased cell wall thickening in transgenic lines.  141  increase in UGPase activity was confined to the developing xylem, which may be reflective of tissue-specific targeting of the 4CL promoter. An  assessment of plant growth  and  architecture  suggested that the  overexpression of these targeted transgenes had no pleiotropic effect on the trees, as they appeared phenotypically normal. However, some lines appeared slightly shorter or had minor changes in calliper relative to corresponding wild-type trees. This is contrary to Konishi et a!., (2004) who reported an increase in height growth in some lines of Populus alba transformed with a modified mung bean SuSy under the control of the 35S promoter. In tobacco, the overexpression of the G. hirsutum SuSy under the control of  two different promoters resulted in increased height growth (Coleman et al., 2006). Similarly, in other plants the downregulation of SuSy has been shown to influence height growth.  For example, using antisense suppression, Tang & Sturm (1999)  generated carrot plants that were smaller and had fewer leaves, while D’Aoust et al., (1999) created tomatoes with reduced fruit size and compromised sucrose unloading capacity. As SuSy is generally thought to exert control over sink strength, increased plant growth is consistent with its overexpression, while suppression of SuSy can be associated decreased growth. Despite the apparent abundance of UGPase activity within plants, the overexpression of foreign Acetobacter xylinum UGPase in poplar and tobacco has yielded changes in plant growth and carbon allocation. In tobacco, plants were taller and some lines had increased internode lengths and stem dry weight (Coleman et a!., 2006), while in poplar, UGPase overexpression resulted in stunted growth and significant increases in cellulose content. The latter example implies that the generation of altered levels of UDP-glucose can facilitate the re-allocation of carbon skeletons derived from photosynthate towards cellulose deposition, as UDP-glucose is the immediate precursor to cellulose polymer biosynthesis (Coleman et a!., 2007). Unlike tobacco, the double transgenics in the current study did not show any additive effect as a result of the expression of UGPase along with SuSy, nor were any changes seen in 4CL::UGPase individual lines (Coleman, unpublished). Previous studies evaluating SuSy and  UGPase transgenic plants have  demonstrated significant changes in soluble carbohydrate contents manifested by the misregulation of these genes. For example, a maize endosperm SuSy mutant had a 2142  to 4-fold increase in localized sucrose levels (Chourey & Nelson, 1976). Consistently, the overexpression of SuSy in tobacco resulted in an increase in stem glucose and fructose concentrations (Coleman et a!., 2006). This suggested that plants generally attempt to maintain a basal concentration of sucrose in the sink tissue (stem) despite the substantially increased metabolism of sucrose into glucose and fructose, which consequently pool. With increased SuSy activity in the sink tissue, an associated increase in sucrose catabolism exists, and therefore more sucrose can be translocated to the sink tissue. A comparison of wheat variants (Triticum aestivum) with differing water soluble carbohydrate concentrations, showed that SuSy transcript abundance (and enzyme activity) was inversely correlated with water soluble carbohydrate accumulation (Xue et al., 2007). The same pattern was apparent with the transcript abundance of three cellulose biosynthesis subunits (TaCesAl, TaCesA4-like and TaCesAlO). It was concluded that these gene families along with others, when present in lower abundance play a role in the accumulation of water soluble carbohydrates, which  is  associated with a decrease in  sucrose hydrolysis  and  decreased  polysaccharide accumulation. No associated change in height growth or stem strength was observed with a lower level of cell wall polysaccharides (Xue et a!., 2007). UGPase has also been shown to affect sink strength in plants.  Most studies  have focussed on the downregulation of UGPase as it is thought to exist in abundance in plants (Appeldoorn et a!., 1997; Magel et a!., 2001). UGPase antisense reduction in potato tubers has resulted in conflicting results which range from no change in soluble sugars, despite a 96% reduction in activity (Zrenner et a!., 1993), to significant reductions in soluble carbohydrates (Borokov et a!. 1996; Spychalla et a!. 1994). In Arabidopsis, plants with decreased UGPase activity had lower soluble carbohydrate and starch contents (Kleczkowski et a!., 2004). Tobacco overexpressing UGPase on the other hand showed increased glucose and fructose contents, but only small changes in sucrose concentration and no change in cellulose production (Coleman et a!., 2006).  Poplar overexpressing UGPase under the control of the 2x35S promoter  had significant increases in all soluble carbohydrates in leaf tissue. In the developing xylem, glucose and sucrose were increased although less so than in leaf tissue (Coleman et a!., 2007). Native UGPase expression studies in poplar clearly show the coordinate upregulation of UGPase with late cell expansion and secondary cell wall 143  formation (Hertzberg et al., 2001), as well as dunng the formation of tension wood (Andersson-Gunnerás et a!., 2006), which is known to be closely linked to cellulose production. Although the SuSy and double transgenic trees did not generally show an increase in starch content, 2x35S::UGPase overexpressed poplar demonstrated increased starch accumulation in the developing xylem and leaf tissue of some lines (Coleman, et a!., 2007). These data concur with findings in Arabidopsis mutants with increased UGPase activity that were shown to have higher starch content when grown in both light and dark (Ciereszko et a!., 2005). However, when UGPase is expressed under the control of the 4CL promoter this effect is not duplicated (Coleman, unpublished), nor is it replicated when UGPase is expressed in combination with SuSy under the control of the 4CL promoter, as double transgenics did not show significant additive effects over single transgenics. Contrary to the current results, Konishi et a!. (2004) did not observe altered carbon allocation in poplar, as the xylem cellulose content of trees expressing the modified mung bean SuSy was unchanged. This discrepancy is likely a consequence of the origin of the exogenous gene and resulting differences in expression. Konishi et a!. (2004) attributed the lack of changes in structural carbohydrates to the higher relative expression of SuSy in leaf tissue as compared to stem tissue. Furthermore, the expression of the mung bean SuSy was relatively higher in the soluble fraction than in the microsomal membrane fraction, and as such influenced the recycling of fructose. Subsequently, in feeder studies with these transgenic lines, labelled (H ) fructose was 3 shown to be incorporated into both cellulosic and non-cellulosic polymers at a greater rate than in control plants. In the current study, despite the inherently higher levels of SuSy activity in leaves, there did not appear to be an increase in SuSy activity in the leaves of the transgenic overexpressing poplar when compared to the wild-type control trees. In contrast, SuSy activity in the developing xylem was significantly increased, and this undoubtedly accounts for the increased cellulose production. Furthermore, the SuSy transgene employed in the current study has previously been shown to be strongly associated with cellulose formation in cotton and may associate more closely with the activity of the cellulose synthase complex in the cell wall (Amor et a!., 1995). This hypothesis is supported by evidence in pea, which has clearly shown that different 144  isoforms of SuSy are associated with the different metabolic fates of sucrose (Barratt, 2001). In the hybrid poplar system employed herein, the elevated levels of cellulose deposition in the secondary xylem resulting from the overexpression of cotton SuSy was also mirrored with a significant change in the ultrastructural characteristic of the cellulose, as it had increased crystallinity. The higher crystalline nature of the cell wall was not associated with a commensurate increase in microfibril angle, as would be expected in the formation of tension wood (Joshi 2003). Intuitively, an increase in cellulose deposition would likely come at the expense or be mirrored by a change in ultrastructure, in this case crystallinity. Calcofluor staining also clearly shows that despite increased fluorescence associated with increased cellulose, there is no evidence of the G-layer that would appear in tension wood (Figure 5.2). An evaluation of the gene expression patterns surrounding tension wood formation identified many genes associated with its formation and the inherent higher degree of cellulose production. Both SuSy and UGPase were identified as genes that are significantly upregulated during tension wood formation (Andersson-Gunnerás et a!., 2006). SuSy was identified as one of the most highly expressed genes in tension wood, with ratios of 1.57 (PttSuSl) and 1.39 (PttSuS2) relative to their expression in normal wood. In an independent study, UDP-glucose pyrophosphorylase (PHUGP2) was shown to be present at a ratio of 2.31 relative to normal wood (Andersson-Gunneràs et a!., 2006). The inconsistencies in biomass (i.e. height growth) results among investigations evaluating SuSy expression may be directly related to the observed differences in cell wall constituents (cellulose content). Increasing cellulose deposition, augmented by the misregulation of genes influencing key pathway steps, could manifest in altered tree growth as more energy and carbon skeletons are being directed or committed to cell wall deposition, as opposed to cell initiation or elongation. To this effect, trees with increased cellulose content have shown varying results.  For example, we have  previously shown large increases in cellulose content (2.8 to 6.5%) in UGPase transformed poplar, which were at the expense of biomass accumulation (Coleman et a!., 2007). These plants also had decreased lignin content. Furthermore, these trees were severely stunted, which was attributed in part to synthesis and accumulation of a salicylic acid glucoside, which has been shown in tobacco to be synthesized by a 145  salicylic acid glucosyltransferase that employs UDP-glucose as the sole source of glucose (Lee & Raskin, 1998). The slight reduction in height growth observed in the current transgenic SuSy trees is not as dramatic as that observed with the UGPase transgenics, and is perhaps due to the closer connection between SuSy (over UGPase) and the cellulose synthase complex proteins (CesA subunits) facilitating a direct metabolic channel for UDP-glucose to cellulose biosynthesis. Other studies, however, have shown marginal increases in cellulose content associated with an increase in height (Park et al., 2004; Shani eta!., 2004; Hu eta!., 1999) Similarly, in other plant species, the level of SuSy expression has been shown to be strongly associated with cellulose synthesis. Expression of the modified mung bean SuSy (Si 1 E) in Acetobacter xy!inum caused enhanced cellulose production by preventing the accumulation of UDP, which is known to inhibit cellulose formation in A. xy!inum (Nakai et a!., 1999; Benziman et a!., 1983).  In addition, carrot plants with  suppressed SuSy showed decreased cellulose content (Tang & Sturm, 1999), while suppression of SuSy in cotton resulted in an almost fibreless phenotype (Ruan et a!., 2003).  On the other hand, tobacco plants overexpressing UGPase showed no  significant changes in starch or cellulose (Coleman et al., 2006), despite an increase in soluble sugars. Given that both SuSy and LJGPase independently  increased cellulose  accumulation in poplar, an attempt to create double transgenics harbouring both genes, by pyramiding genes under the regulation of the vascular specific promoter, was attempted. To this end, gene stacking did not offer an advantage in diverting carbon skeletons from photosynthate to cellulose deposition, as additive effects were not apparent. The lack of additive influence is not consistent with our previous findings in tobacco which showed clear effects in height growth, but not carbon re-allocation to cellulose deposition (Coleman et a!., 2006).  In the current case, it appears that in  poplar, SuSy alone provides the largest effect in maintaining height growth and improving the level of cellulose deposition, while the additional effects of UGPase are not evident. This is not particularly surprising, as 4CL::UGPase on its own causes only minor changes in soluble sugars and does not appear to influence growth or cellulose content in poplar (Coleman, unpublished).  Furthermore, under the regulation of the 146  2x35S promoter it was extremely difficult to obtain double transformants, as UGPase alone severely affected plant growth (Coleman et a!., 2007). In summary, the overexpression of SuSy individually and in combination with UGPase in hybrid poplar resulted in increased cellulose production. While no apparent trend in soluble sugars is evident, the plants do indeed synthesize cellulose to a greater extent, thus providing evidence for a direct connection between sucrose supply (sink strength), its breakdown and cellulose deposition through the increased activity of SuSy. The overexpression of UGPase in combination with SuSy appears to manifest similar results, suggesting that SuSy is more closely related to cellulose synthesis, and consequently xylem deposition. These results together with the work of others clearly implicate SuSy as a strong component in the production of cellulose and in sink strength in poplar.  Acknowledgements The authors would like to thank Tony Einfeldt, Lisa McDonnell and Tom Canam for their insight and technical assistance. Funding for this project is acknowledged from the Natural Science and Engineering Research Council of Canada.  147  References Albrecht G. and Mustroph A. 2003. Localization of sucrose synthase in wheat roots: increased in situ activity of sucrose synthase correlates with cell wall thickening by cellulose deposition under hypoxia. Planta 217: 252-260. Amor Y., Haigler C., Johnson S., Wainscott M. and Delmer D.P. 1995. A membraneassociated form of sucrose synthase and its potential role in synthesis of cellulose and callose in plants. Proceedings of the National Academy of Sciences of the United States of America 92: 9353-9357. Andersson-Gunneràs S., Mellerowicz E., Love J., Segerman B., Ohmiya Y., Coutinho P.M., Nilsson P., Henrissat B., Moritz T. and Sundberg B. 2006. Biosynthesis of cellulose-enriched tension wood in Populus: global analysis of transcripts and metabolites identifies biochemical and developmental regulators in secondary wall biosynthesis. The Plant Journal 45: 144-165. Appeldoorn N.J.G., de Bruijn S.M., Koot-Gronsveld E.A.M., Visser R.G.F., Vreugdenhil D. and van der Plas L.H.W. 1997. Developmental changes of enzymes involved in conversion of sucrose to hexose-phosphate during early tuberisation of potato. Planta 202: 220-226. Asano T., Kunieda N., Omura Y., lbe H., Kawasaki T., Takano M., Sato M., Furuhashi H., Mujin T., Takaiwa F., Wu C.-y., Tada Y., Satozawa T., Sakamoto M. and Shimada H. 2002. Rice SPK, a calmodulin-like domain protein kinase, is required for storage product accumulation during seed development: phosphorylation of sucrose synthase is a possible factor. Plant Cell 14: 619-628. Barratt D.H.P., Barber L., Kruger N.J., Smith A.M., Wang T.L. and Martin C. 2001. Multiple, distinct isoforms of sucrose synthase in pea. Plant Physiology 127: 655664. Benziman M., Aloni Y. and Delmer D.P. 1983. Unique regulatory properties of the UDP glucose: b-I ,4-glucan synthetase of Acetobacter xylinum. Journal of Applied Polymer Science 131-143. Borokov A.Y., McClean P.E., Sowokinos J.R., Ruud S.H. and Secor G.A. 1996. Effect of expression of UDP-glucose pyrophosphorylase ribozyme and antisense RNAs on the enzyme activity and carbohydrate composition of field-grown transgenic potato plants. Journal of Plant Physiology 147: 644-652. Buckeridge M.S., Vergara C.E. and Carpita N.C. 1999. The mechanism of synthesis of a mixed-linkage (1-3), (1-4) b-D-glucan in maize. Evidence for multiple sites of glucosyl transfer in the synthase complex. Plant Physiology 120: 1105-1116. Chourey P.S. 1981. Genetic control of sucrose synthetase in maize endosperm. Molecular and General Genetics 184: 372-376. 148  Chourey P.S. and Nelson S.E. 1976. The enzymatic deficiency conditioned by the shrunken-I mutation in maize. Biochemical Genetics 14:1041-1055. Ciereszko I., Johansson H. and Kleczkowski L.A. 2005. Interactive effects of phosphate deficiency, sucrose and light/dark conditions on gene expression of UDP-glucose pyrophosphorylase in Arabidopsis. Journal of Plant Physiology 162: 343-353. Coleman H.D., Canam T., Kang K.Y., Ellis D.D. and Mansfield S.D. 2007. Overexpression of UDP-glucose pyrophosphorylase in hybrid poplar affects carbon allocation. Journal of Experimental Botany 58: 4257-4268. Coleman H.D., Ellis D.D., Gilbert M. and Mansfield S.D. 2006. Up-regulation of sucrose synthase and UDP-glucose pyrophosphorylase impacts plant growth and metabolism. Plant Biotechnology Journal 4: 87-101. D’Aoust M.-A., Yelle S. and Nguyen-Quoc B. 1999. Antisense inhibition of tomato fruit sucrose synthase decreases fruit setting and the sucrose unloading capacity of young fruit. The Plant Cellll: 2407-2418. Datla R.S.S., Bekkaoui F., Hammerlindl J.K., Pilate G., Dunstan D.l. and Crosby W.L. 1993. Improved high-level constitutive foreign gene translation using an AMV RNA4 untranslated leader sequence. Plant Science 94: 139-149. Dejardin A., Sokolov L.N. and Kleczkowski L.A. 1999. Sugar/osmoticum levels modulate differential abscisic acid-independent expression of two stressresponsive sucrose synthase genes in Arabidopsis. Biochemistiy Journal 344: 503-509. Hauch S. and Magel E. 1998. Extractable activities and protein content of sucrosephosphate synthase, sucrose synthase and neutral invertase in trunk tissues of Robinia pseudoacacia L. are related to cambial wood production and heartwood formation. Planta 207: 266-274. Hauffe K.D., Paszkowski U., Schuize-Lefert P., Hahlbrock K., Dangl J.L. and Douglas C.J. 1991. A parsley 4CL-1 promoter fragment specifies complex expression patterns in trangenic tobacco. The Plant Cell 3: 435-443. Hertzberg M., Aspeborg H., Schrader J., Andersson A., Erlandsson R., Blomqvist K., Bhalerao R., Uhlen M., Teen T.T., Lundeberg J., Sundberg B., Nilsson P. and Sandberg G. 2001. A transcriptional roadmap to wood formation. Proceedings of the National Academy of Sciences of the United States of America 98: 1473214737. Hood E.E., Gelvin S.B., Melcher L.S. and Hoekema A. 1993. New Agrobacterium helper plasmids for gene transfer to plants. Transgenic Research 2: 208-218. 149  Hu W.-J., Harding S.A., Lung J., Popko J.L., Ralph J., Stokke D.D., Tsai C.-J. and Chiang V.L. 1999. Repression of lignin biosynthesis promotes cellulose accumulation and growth in transgenic trees. Nature 17: 808-81 2. Huntley S.K., Ellis D., Gilbert M., Chapple C. and Mansfield S.D. 2003. Significant increases in pulping efficiency in C4H-F5H transformed poplars: Improved chemical savings and reduced environmental toxins. Journal of Agriculture and Food Chemistiy5i: 6178-6183. Joshi C.P. 2003. Xylem-specific and tension stress-responsive expression of cellulose synthase genes from aspen trees. Applied Biochemistry and Biotechnology 105108: 17-25. Kay R., Chan A., Daly M. and McPherson J. 1987. Duplication of CaMV 35S promoter sequences creates a strong enhancer for plant genes. Science 236: 1299-1302. Kennedy J.F. and White C.A. 1983. Bioactive carbohydrates in chemistry, biochemistry and biology. New York, USA, Halstead Press. Kleczkowski L.A. 1994. Glucose activation and metabolism through UDP-glucose pyrophosphorylase in plants. Phytochemistiy 37: 1507-1515. Kleczkowski L.A., Geisler M., Ciereszko I. and Johansson H. 2004. UDP-glucose pyrophosphorylase. An old protein with new tricks. Plant Physiology 134: 912918. Kolosova N., Miller B., Ralph S., Ellis B.E., Douglas C., Ritland K. and Bohlmann J. 2004. Isolation of high-quality RNA from gymnosperm and angiosperm trees. BioTechniques 36: 821 -824. Komina 0., Zhou Y., Sarath G. and Chollet R. 2002. In vivo and in vitro phosphorylation of membrane and soluble forms of soybean nodule sucrose synthase. Plant Physiology 129: 1664-1 673. Konishi T., Ohmiya Y. and Hayashi T. 2004. Evidence that sucrose loaded into the phloem of a poplar leaf is used directly by sucrose synthase associated with various 3-glucan synthases in the stem. Plant Physiology 134: 1146-1152. Le Hir R., Pelleschi-Travier S., Viemont J. and Leduc N. 2005. Sucrose synthase expression pattern in the rhythmically growing shoot of common oak (Quercus robur L.). Annals of Forest Science 62: 585-591. Lee H. and Raskin I. 1998. Glucosylation of salicylic acid in Nicotiana tabacum Cv. Xanthi-nc. Phytopathology 88: 692-697. Levy M., Edelbaum 0. and Sela I. 2004. Tobacco mosaic virus regulates the expression of its own resistance gene N . Plant Physiology 135: 2392-2397. 1 150  Magel E., Abdel-Latif A. and Ham pp R. 2001. Non-structural carbohydrates and catalytic activities of sucrose metabolizing enzymes in trunks of two Juglans species and their role in heartwood formation. Holzforshung 55: 135-145. McCown B.H. and Lloyd G. 1981. Woody Plant Medium (WPM) a mineral nutrient formulation for microculture for woody plant species. Horticultural Science 16: 453. -  Megraw R.A., Leaf G. and Bremmer D. 1998. Longitudinal shrinkage and microfibril angle in loblolly pine. Microfibril angle in wood. B. A. Butterfield. Christchurch, New Zealand, University of Canterbury Press: 27-61. Nakai T., Tonouchi N., Konishi T., Kojima Y., Tsuchida T., Yoshinaga F., Sakai F. and Hayashi T. 1999. Enhancement of cellulose production by expression of sucrose synthase in Acetobacter xylinum. Proceedings of the National Academy of Sciences of the United States of America 96: 14-18. Nolte K.D. and Koch K.E. 1993. Companion-cell specific localization of sucrose synthase in zones of phloem loading and unloading. Plant Physiology 101: 899905. Park Y.W., Baba K., Furuta Y., lida I., Sameshima K., Arai M. and Hayashi T. 2004. Enhancement of growth and cellulose accumulation by overexpression of xyloglucanase in poplar. FEBS Letters 564: 183-1 87. Ralph S., Oddy C., Cooper D., Yueh H., Jancsik S., Kolosova N., Philippe RN., Aeschliman D., White R., Huber D., Ritland C.E., Benoit F., Rigby T., Nantel A., Butterfield Y.S.N., Kirkpatrick R., Chun E., Liu J., Palmquist D., Wynhoven B., Stott J., Yang G., Barber S., Holt R.A., Siddiqui A., Jones S.J.M., Marra M.A., Ellis B.E., Douglas C.J., Ritland K. and Bohlmann J. 2006. Genomics of hybrid poplar (Populus trichocarpa x deltoides) interacting with forest tent caterpillars (Malacosoma disstria): Normalized and full-length cDNA libraries, expressed sequence tags (ESTs), and a cDNA microarray for the study of insect-induced defenses in poplar. Molecular Ecology 15: 1275-1 297. Ruan Y.L., Llewellyn D.J. and Furbank R.T. 2003. Suppression of sucrose synthase gene expression represses cotton fibre cell initiation, elongation, and seed development. The Plant Cell 15: 952-964. Shani Z., Dekel M., Tsabary G., Goren R. and Shoseyov 0. 2004. Growth enhancement of transgenic poplar plants by overexpression of Arabidopsis thaliana endo-1 ,4b-glucanase (cell). Molecular Breeding 14: 321-330. Spychalla J.P., Scheffler B.E., Sowokinos J.R. and Bevan M.W. 1994. Cloning, antisense RNA inhibition, and the coordinated expression of UDP-glucose 151  pyrophosphorylase with starch biosynthetic genes in potato tubers. Journal of Plant Physiology 144: 444-453. Subbaiah C.C. and Sachs M.M. 2001. Altered patterns of sucrose synthase phosphorylation and localization precede caliose induction and root tip death in anoxic maize seedling. Plant Physiology 125: 585-594. Sun J., Loboda T., Sung S.-J.S. and Black C.C.J. 1992. Sucrose synthase in wild tomato, Lycopersicon chmielewskii, and tomato fruit sink strength. Plant Physiology 98: 1163-1169. Tang G.Q. and Sturm A. 1999. Antisense repression of sucrose synthase in carrot (Daucus carota L.) affects growth rather than sucrose partitioning. Plant Molecular Biology 41: 465-479. Vonk C.C. 1973. Investigation of non-ideal two-phase polymer structures by small-angle x-ray scattering. Journal of Applied Ctyst. 6: 81-86. Winter H., Huber J.L. and Huber S.C. 1997. Membrane association of sucrose synthase: changes during the graviresponse and possible control by protein phosphorylation. Federation of European Biochemical Societies Letters 420: 151-1 55. Winter H. and Huber S.C. 2000. Regulation of sucrose metabolism in higher plants: localization and regulation of activity of key enzymes. Critical Reviews in Biochemistiy and Molecular Biology 35: 253-289. Xue G.-P., Mcintyre C.L., Jenkins C.L.D., Glassop D., van Herwaarden A.F. and Shorter R. 2007. Molecular dissection of variation in carbohydrate metabolism related to water soluble carbohydrate accumulation in stems of wheat (Triticum aestivum L.). Plant Physiology 146: 441-454. Yokoyama T., Kadla J.F. and Chang H.M. 2002. Microanalytical method for the characterization of fiber components and morphology of woody plants. Journal of Agriculture and Food Chemistiy 50: 1040-1044. Zrenner R., Salanoubat M., Willmitzer L. and Sonnewald U. 1995. Evidence of the crucial role of sucrose synthase for sink strength using transgenic potato plants (Solanum tugerosum L.). The Plant Journal 7: 97-107. Zrenner R., Wilimitzer L. and Sonnewald U. 1993. Analysis of the expression of potato uredinediphosphate-glucose pyrophosphorylase and its inhibition by antisense RNA. Planta 190: 247-252.  152  Conclusions and Recommendations for Future Work The molecular understanding of the biosynthesis of cellulose in plants and forest trees is still a relatively new area of study. The identification of UDP-glucose as the precursor to cellulose synthesis and the key substrate for the cellulose synthase complex has permitted the scientific community to make significant advances in understanding this vital pathway in recent years. For example, the knowledge of the pathway leading to the formation of UDP-glucose within plants has permitted the study of cellulose biosynthesis via the variation in expression of genes involved directly and indirectly in the formation of this precursor molecule. Testing the hypothesis that altering carbohydrate metabolism to generate increased cellular pools of UDP-glucose should result in altered cellulose content by shifting carbon allocation towards cellulose biosynthesis was tested in two model plant species. In tobacco, the overexpression of SPS, UGPase and SuSy individually or in combination resulted in substantial increases in height growth, and consequently plant biomass. Furthermore, it was apparent that pyramiding genes to enhance the catalysis of sucrose to UDP-glucose offers a mechanism to increase growth additively over the ability of a single gene misregulation. However, consistent changes in either soluble or structural carbohydrates were not evident. Therefore, in tobacco, it appears that the reallocation of carbon skeletons from photosynthate to cellulose deposition may not be possible by targeting these sucrose metabolism genes and enzyme products. In contrast, in poplar, the overexpression of UGPase and SuSy individually and in combination resulted in a significant increase in cellulose.  However, in UGPase  transgenics trees this was associated with reductions in growth, as well as a defence response which resulted in the formation of salicylic acid 2-O-f3-salicylic acid-glucoside (SAG). The formation of this glucoside has been shown to be catalyzed by a UDP glucose:salicylic acid glucosyltransferase in tobacco, where UDP-glucose acts as the sole donor. The results described in this thesis suggest that using the current system to increase cellular pools of UDP-glucose in poplar trees does indeed manifest in the formation of elevated levels of cell wall cellulose polymer.  However, it appears that  UDP-glucose formed in these targeted transgenics is excessive and the cellulose synthase complex becomes limiting. Ultimately, to maintain the homeostatic levels of 153  UDP-glucose near the plasma membrane, the excess UDP-glucose is mobilized from cell wall biosynthesis in the generation of the salicylic acid glucoside. Similarly, in transgenic poplar overexpressing SuSy and SuSy x UGPase there is a significant increase in cellulose content. In these lines, while there was a slight reduction in height and biomass in some lines, the decrease was much less severe than in the UGPase transgenic lines. It appears that the close association of SuSy with the cellulose synthase complex (CSC) offers an avenue for the direct channelling of UDP glucose to the CSC, and limits the availability of UDP-glucose to other cellular pathways. As such, the defence response induced in the UGPase transgenics was not apparent. Furthermore, there does not appear to be an additive gain in carbon partitioning to cellulose production by transforming poplar trees with both genes. The elevated cellulose content in the SuSy and SuSy x UGPase transgenic poplar was also associated with significant increase in crystallinity. This increase was not associated with a change in microfibril angle which would be expected in the case of tension wood formation, suggesting that the apparent change in cellulose ultrastructure is indeed a change in the mechanism of cellulose deposition as opposed to the induction of tension wood formation by the poplar trees. The differences  observed  between tobacco  and  poplar,  despite  being  transformed with the identical constructs has been attributed to two major factors. The first is the difference in phloem loading and unloading between the two species, with poplar being a symplastic loader and tobacco an apoplastic loader, and the second is the difference in sinks and in sink strength. In poplar, the stem is the major sink and as such is responsible for regulating much of the gradient in photoassimilate translocation, while in tobacco there are more numerous sinks including leaf and stem tissue, as well as reproductive bud formation. The findings of this research cleaily demonstrate that it is possible to alter cellulose metabolism in forest trees through the misregulation of key sucrose metabolism genes. It also suggests that the formation of UDP-glucose may not be the overall limiting factor in manipulating carbon allocation to cellulose biosynthesis. The formation of cellulose may also be highly limited by the CesA complex, thus encouraging the focus of cellulose research on these genes. 154  Future Work Recommendations There are a number of interesting projects that should be developed and extended based on the findings of the work presented in this thesis. First of all, the gene encoding for UDP-glucose:saticylic acid glucosyltransferase has not yet been identified in poplar. This putative gene could be cloned and gene function confirmed in terms of glucose donor sensitivity. Once identified and cloned, transgenics verifying its catalytic function should be made. In addition, downregulating this gene in the overexpressing UDP-glucose pyrophosphorylase transgenics may offer an opportunity to increase the partitioning to cellulose without eliciting the defence response apparent in the existing transgenic UGPase poplar trees. Within the array of successful poplar SuSy transformants, key lines (those that are not affected in growth and biomass accumulation) should be selected, and combined with misregulation efforts targeting key lignin biosynthetic genes (i.e. downregulation). For example, substantial industrial gains have been made in altering the lignin monomer composition and content in poplar using F5H or RNAi C3’H. By pyramiding SuSy transgenics with one (or both) of these two lignin genes, there is potential for substantial additive gains that could hypothetically manifest in an increased cellulose producing line with either reduced lignin production or altered lignin monomer composition in favour of syringyl lignin sub-units. Ideally, as research progresses with the modulation of the CesA genes, there is the possibility that increased CSC activity will be attainable, and as such UDP-glucose could become the limiting factor in cellulose biosynthesis. Thus, pyramiding the SuSy or UDP-glucose pyrophosphorylase genes with a CSC modified tree with the ability to rapidly synthesize cellulose could provide increased substrate to an already improved system for further advancement. Finally, in poplar, it would also be interesting to pursue the overexpression of SuSy under the control of a promoter that is more specific to the timing and location of cellulose biosynthesis.  In particular, one of the secondary cell wall specific CesA  promoters may facilitate the most useful and opportune expression of the SuSy gene in coordination with the CSC genes.  155  Appendix A  -  f3-glucuronidase (GUS) Transformation and Staining  Methods Construct Design pBI-121 containing the 35S::GUS construct was modified by the replacement of the 35S promoter with either the 2x35S promoter (from pBl-426) or the 4CL promoter (from 99-G1-501). The resulting cassettes (2x35S::GUS::NOS and 4CL::GUS::NOS) were excised from pBI 121 as Hindlil-EcoRl fragments and inserted into the pCAMBIA 1300 binary vector for use in plant transformation.  Transformation of Hybrid Poplar and Tobacco Poplar and tobacco were transformed using Agrobacterium as previously described (Coleman et a!, 2006 and 2007). In short, poplar leaf disks were cut and co cultured with EHA1O5 harbouring the 2x35S::GUS or 4CL::GUS transgenes.  The  explants were plated on Woody Plant Media (WPM) (McCown and Lloyd 1981) supplemented  with  0.1  pM  each  of a-naphthalene  acetic  acid  (NAA),  6-  benzylaminopurine (BA), and thiadiazuron (TDZ) and solidified with 3% (wlv) agar and 1.1% (w/v) phytagel (WPM 0.1/0.1/0.1). After three days the disks were transferred to WPM 0.1/0.1/0.1 supplemented with carbenicillin disodium (500 mg L ) and cefotaxime 1 sodium salt (250 mg L ). Following three additional days, the disks were transferred to 1 WPM 0.1/0.1/0.1 containing carbenicillin, cefotaxime and kanamycin (25 mg L ). After 1 five weeks, shoots and callus material were transferred to WPM containing agar and phytagel, supplemented with 0.01 pM BA, carbenicillin, cefotaxime and kanamycin. Once individual shoots were visible, plantlets were transferred to solidified WPM with 0.01 pM NAA and carbenicillin, cefotaxime and kanamycin to induce rooting. After two consecutive five-week periods on this media, shoot tips were isolated to solidified antibiotic-free WPM with 0.01 pM NAA. Procedures for tobacco were the same with the exception of the use of MS media (Murashige and Skoog, 1962) and the absence of TDZ in the media.  156  -glucuronidase (GUS) Assays GUS staining assays were performed as described by Jefferson (1987) using leaves, shoots, roots, and node sections from transgenic and control tobacco and hybrid poplar. Stems and nodes were hand sectioned, and leaves and roots were used directly. Plant sections were vacuum infiltrated in acetone.  Following the removal of  acetone, the plants were incubated in an x-gluc solution (0.1% Triton X-100, 5% IM 4 pH 7.0, 0.5% 100 mM Fe(CN), 0.05% x-gluc) overnight at 37°C. NaPO  Prior to  imaging, the samples were washed with 70% ethanol. Sections were mounted on glass slides and visualized using a Leica DMR microscope equipped with a QICAM CCD camera (Q-imaging).  Results Expression of the 2x35S and 4CL promoters was assessed using the GUS reporter gene. The resulting staining patterns in tobacco revealed very little difference between the two promoters (Figure Al). Both promoters were shown to be expressed in all tissues examined. In leaf tissue, 4CL expression was somewhat more restricted to the vasculature, but expression of the promoter was seen throughout the leaf tissue. Poplar results were similar to tobacco in that the promoters were expressed in all tissues examined (Figure A2).  While 4CL again appeared to be somewhat more  associated with vascular tissue, it was still apparent throughout the samples. Despite the significant differences seen by others (Hauffe et al., 1991; Kay et al., 1987) in the expression of 2x35S and 4CL, the results presented herein clearly demonstrate that both promoters effectively express throughout the plant, with no real significant differences between promoters.  157  Figure Al. 3-gIucuronidase (GUS) staining in tobacco stem cross sections (A, B, C), roots (D, E, F), longitudinal stem sections (G, H, I) and leaves (J, K, L) of control (A, F, G, J), 2x35S (C, E, H, L), and 4CL (B, D, I, K). Scale bar: 1 mm.  158  r  ——  .•  j  & Pc,  .  Figure A2. 3-glucuronidase (GUS) staining in poplar stem cross sections (A, C, E), leaves (B, D, F), longitudinal stem sections (G, H, I) and petioles (J, K, L) of control (C, D, H, J), 2x35S (E, F, I, L) and 4CL (A, B, G, K). Scale bar: 1 mm.  159  References Hauffe K.D., Paszkowski U., Schulze-Lefert P., Hahlbrock K., Dangl J.L. and Douglas C.J. 1991. A parsley 4CL-1 promoter fragment specifies complex expression patterns in trangenic tobacco. The Plant Cell 3: 435-443. Jefferson R.A., Kavanagh T.A. and Bevan M.W. 1987. GUS fusions: 3-gIucuronidase as a sensitive and versatile fusion marker in higher plants. European Molecular Biology Organization Journal 6: 3901-3907. Kay R., Chan A., Daly M. and McPherson J. 1987. Duplication of CaMV 35S promoter sequences creates a strong enhancer for plant genes. Science 236: 1299-1 302. McCown B.H. and Lloyd G. 1981. Woody Plant Medium (WPM) a mineral nutrient formulation for microculture for woody plant species. Horticultural Science 16: 453. -  Murashige T. and Skoog F. 1962. A revised medium for rapid growth and bioassays with tobacco tissue culture. Physiologia Plantarum 15: 473-497.  160  Appendix B  —  Supplemental Data Chapter 3  Table BI. Transcript abundance of the native poplar UDP-glucose pyrophosphorylase genes in leaf and developing xylem tissue for transgenic and wildtype trees. Mean (±SE) were calculated from 3 plants per line. Bold denotes significant difference from control values at a=O.1O.  Poplar UGPasel  Poplar UGPase2  Ct  Ct  Control  9.08 ± 0.40  0.45 ± 0.01  2x35S::UGPase A 2x35S::UGPase B  9.59 ± 0.27  1.47 ± 0.06  9.38 ± 0.47  1.19 ± 0.01  2x35S::UGPase C 2x35S::UGPase D  12.66 ± 2.76  1.74 ± 0.14  14.62 ± 1.03  2.15 ± 0.12  2x35S::UGPase E  12.19 ± 0.37  0.99 ± 0.47  2x35S::UGPase F  6.82 ± 0.44  0.62 ± 0.01  Developing Xylem  Mean  Mean  Control  3.85 ± 0.43  0.48 ± 0.02  2x35S::UGPase A  5.52 ± 1.25  1.99 ± 0.16  2x35S::UGPase B  5.49 ± 0.10  1.84 ± 0.12  2x35S::UGPase C  4.28 ± 0.12  1.06 ± 0.08  2x35S::UGPase D  6.61 ± 0.37  1.77 ± 0.17  2x35S::UGPase E 2x35S::UGPase F  5.14 ± 0.17  1.80 ± 0.04  2.70 ± 0.08  0.67 ± 0.01  Leaf  161  Table B2. Transcript abundance of the cell wall biosynthetic genes involved in lignin and cellulose deposition in the developing xylem tissue for transgenic and wild-type trees. Mean (±SE) were calculated from 3 plants per line.  Gene  2x35S::UGPase Ct ± Mean  Wild-type Ct ± Mean  PAL  1.51 ± 0.51  1.00±0.06  C4H  3.94±1.10  1.08±0.28  SuSy  1.19±0.09  1.03±0.17  C3H  2.20±0.42  1.01 ± 0.12  COMT  1.69 ± 0.38  1.07 ± 0.26  SAD  1.96 ± 0.22  0.86 ± 0.37  CCoAMT  8.85 ± 1.73  1.02 ± 0.17  CAD  2.18 ± 0.42  0.80 ± 0.03  CESA  4.11 ± 1.33  1.13 ± 0.40  F5H  1.61 ± 0.25  1.03±0.16  4CL  1.18±0.16  1.00±0.00  CCR  6.14±0.93  1.00±0.00  UGPI  0.80±0.19  1.05±0.22  UGP2  2.34±0.67  1.01 ± 0.10  162  Table B3. Fold changes in salicylic acid 2-O-3-.glucoside in the developing xylem of all transgenic 2x35S::UGPase hybrid poplar relative to levels in wild-type trees. Salicylic acid 2-O43-D-glucoside Fold Change Control  1.0  2x35S::UGPase A  221.4  2x35S::UGPase B  177.7  2x35S::UGPase C  239.6  2x35S::UGPase D  232.3  2x35S::UGPase E  270.2  2x35S::UGPase F  268.9  163  

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