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A biochemical and molecular study of lignin biosynthesis Dharmawardhana, Dilsiri Palitha 1996

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A B I O C H E M I C A L A N D M O L E C U L A R STUDY OF LIGNIN BIOSYNTHESIS by DILSIRI P A L I T H A D H A R M A W A R D H A N A B . S c , The University of Colombo, 1983 M . S c , The Australian National University, 1988 A THESIS SUBMITTED IN P A R T I A L F U L F I L L M E N T OF T H E REQUIREMENTS FOR T H E D E G R E E OF DOCTOR OF PHILOSOPHY in T H E F A C U L T Y OF G R A D U A T E STUDIES Biotechnology Laboratory and Department of Botany  We accept this thesis as confirming to the required standard  T H E UNIVERSITY OF BRITISH C O L U M B I A June 1996 © Dilsiri Palitha Dharmawardhana, 1996  In  presenting  degree  this  at the  thesis  in  University of  freely available for reference copying  of  department  this or  publication of  partial  British Columbia, and study.  of  by this  his  or  her  the  The University of British Columbia Vancouver, Canada  that the  I further agree  representatives.  may be It  thesis for financial gain shall not  Department  requirements  I agree  thesis for scholarly purposes  permission.  DE-6 (2/88)  fulfilment  that  advanced  Library shall make it  by the  understood be  an  permission for extensive  granted  is  for  allowed  that  without  head  of  my  copying  or  my written  11  ABSTRACT The objective of the first half of this thesis was to develop an efficient and sensitive method to detect lignin in intact tissues using confocal laser scanning microscopy and to characterize the timing and quality of lignin deposited during Arabidopsis thaliana development. Lignin could first be detected 48h post-imbibition. During subsequent development, lignification of vasculature proceeded in a temporally coordinated manner, with lignin deposition closely following secondary wall thickening in vascular elements, in a cell-by-cell sequence. A spatially and temporally distinct pattern of syringyl lignin deposition was observed along the developing stem. Syringyl type lignin appeared only once fiber differentiation was initiated at day 21. The second half of the thesis is dedicated to identification, purification and characterization of a coniferin specific P-glucosidase from Pinus contorta xylem, and cloning of the corresponding cDNA.  Coniferin accumulates to high levels during  cambial reactivation, and a cinnamyl alcohol glucoside/ p-glucosidase system is thought to play a key role in lignification in gymnosperm tissues. However, evidence for the existence of P-glucosidases specific for cinnamyl alcohol glucosides in lignifying xylem has not been conclusive. Using a range of synthetic aromatic P-glucosides and coniferin as substrates, two major P-glucosidases present in total extractable xylem protein were identified. The enzyme that demonstrated a high specificity/ activity for coniferin was purified to homogeneity using anion exchange, hydrophobic interaction and gel filtration chromatography.  Based on the N-terminal amino acid sequence of this  Ill  protein, the corresponding cDNA was isolated from a library constructed from differentiating pine xylem. A 1909 bp full length cDNA sequence was confirmed to be that of coniferin Pglucosidase on the basis of it's homology to other plant P-glucosidases and by the demonstration of coniferin hydrolytic activity and similar substrate specificity when expressed in E. coll The deduced 513 amino acid protein contains a 23 amino acid N terminal signal peptide that is characteristic of eukaryotic secretory proteins.  The  isolation of the coniferin |3-glucosidase cDNA offers novel opportunities to clarify the ultimate steps of lignin biosynthesis. It also opens a route to the possible modification of lignin in trees by down-regulation of its activity through genetic engineering.  T A B L E OF CONTENTS Abstract Table of Contents List of Abbreviations List of Tables List of Figures Acknowledgement Foreword INTRODUCTION A N D O V E R V I E W  ii iv vi vii viii x xi 1  C H A P T E R 1 Vascular lignification in Arabidopsis thaliana Introduction Materials and methods Plant material Preparation for fluorescence microscopy Flowering stem development and pattern of lignin deposition Peroxidase localization Chemical analysis Results Microscopic detection Vascular development and lignification Flower stem development Pattern of lignin deposition Composition of lignin Discussion  8 9 9 9 11 11 11 12 12 12 17 24 30 33  C H A P T E R 2 Identification, purification and properties of coniferin P-glucosidase Introduction Materials and methods Plant material Biochemicals and buffers Enzyme extraction Assay for P-glucosidases Determination of the profile of P-glucosidases Enzyme purification SDS-PAGE Native P A G E and activity staining N-terminal sequencing Antibodies against the synthetic peptide  35 37 37 37 38 38 39 40 41 41 42 43  V  Western blot analysis and glycoprotein detection Results p-glucosidases in differentiating xylem Purification of coniferin p-glucosidase Properties of coniferin P-glucosidase Catalytic properties N-terminal amino acid sequence Immunological analysis Discussion  44 44 44 49 51 51 53 54 59  C H A P T E R 3 c D N A cloning and heterologous expression of P. contorta coniferin P-glucosidase Introduction Materials and methods R N A isolation Construction of cDNA library PCR amplification of CBG-cDNA Cloning and sequencing Cloning of the 5' end of the C B G c D N A Sequence comparison and analysis Expression of C B G in E. coli Results Isolation of mRNA and construction of c D N A library PCR amplification of C B G - c D N A Nucleotide sequence of C B G Primary structure of C B G Expression of C B G in E. coli Discussion  63 64 64 65 65 68 68 69 69 70 73 77 81 92 97  C H A P T E R 4 In situ localization of lignin biosynthetic enzymes Introduction Materials and methods Plant material Detection of lignin In situ enzyme activity staining Immunolocalization Results Discussion  103 104 104 105 105 106 107 114  G E N E R A L DISCUSSION A N D PERSPECTIVES  117  L I T E R A T U R E CITED  131  LIST OF ABBREVIATIONS  FPLC PVDF SDS EDC c-BSA DEPC 2-NPG 4-NPG 4CL C3H C4H CAD CBG CCo3H CCoMT CCR F5H MUG OMT,COMT PAGE PAL PO FITC PCR VRA-G  Fast Protein Liquid Chromatograph polyvinylidenefluoride sodium dodecyl sulphate 1 ,ethyl-3-(3-dimethylaminopropyl)carbodiimide hydrochloride cationized bovine serum albumin diethyl pyrocarbonate 2-nitrophenyl p-glucoside 4-nitrophenyl P-glucoside hydroxycinnamate: Co Aligase coumarate-3 -hydroxylase cinnamate-4-hydroxylase cinnamyl alcohol dehydrogenase coniferin p-glucosidase coumaroyl-Co A-3 -hydroxylase caffeoyl-CoA 3-O-methyltransferase cinnamyl: Co A reductase ferulate-5 -hydroxylase 4- methylumbelliferyl P-glucoside caffeic acid O-methyl transferase polyacrylamide gel-electrophoresis phenylalanine ammonia lyase peroxidase fluorescein isothiocyanate polymerase chain reaction 5[4-(P-D-glucopyranosyloxy)-3-methoxyphenylmethylene]-2thioxothiazolidine-4-one-3 -ethanoic acid  vii  LIST OF T A B L E S  1.1  Chemical analysis of lignin from Arabidopsis thaliana tissues  32  2.1  Purification of C B G from Pinus contorta xylem  50  2.2  Substrate specificity of purified C B G and partially purified M U G p-  52  glucosidase 3.1  Deduced amino acid composition of C B G  90  3.2  Substrate specificity of C B G and E. coli expressed C B G - c D N A  96  3.3  Deduced codon usage for C B G - c D N A  102  Vlll  LIST OF FIGURES  1.1  Lignin biosynthetic pathway  4  1.1-1.3  Phase contrast (P.C.) and fluorescence (FL.) confocal laser-scanning  15  micrographs (CLSM) of A. thaliana seedlings 1.4-1.10  Merged fluorescence (red) and phase contrast (green) C L S M images of A.  18  thaliana seedling tissue. 1.11-1.14  Phase contrast (P.C.) and fluorescence (FL.) C L S M images of A. thaliana  20  seedlings 1.15-1.18  Fluorescence C L S M images of A. thaliana seedlings during late vascular  22  development 1.19-1.21  Line diagram of flowering stem at several developmental stages and cross  25  sections of stem to show differentiation of vascular bundles and fibers 1.22  Spatial distribution of syringyl and guaiacyl lignin (based on Maule  27  reaction) along the stem of Arabidopsis during development. 1.23-1.30  Histochemical staining for guaiacyl/ syringyl lignin (Maule reaction) and  28  peroxidase localization 2.1  Distribution of P-glucosidases in total extractable Pinus contorta xylem  46  protein, following anion exchange chromatography 2.2  Native P A G E and activity staining of the three major P-glucosidases  47  2.3  Documentation of sequential purification of coniferin P-glucosidase.  47  2.4  Coomassie blue stained preparative SDS-PAGE blot of purified C B G  47  2.5  Native P A G E analysis of purified coniferin P-glucosidase from P. contorta.  Al  2.6  N-terminal amino acid alignment of C B G with other plant P-glucosidases  56  2.7  Detection of peptide/carrier protein coupling  57  2.8  Western blot analysis of coniferin p-glucosidase using polyclonal antibodies  57  raised against the 15- amino acid synthetic peptide 3.1.  P C R amplification strategy used for isolating a complete C B G - c D N A from 67  ix  a lodgepole pine xylem A.ZAP library. 3.2  Total R N A from 3 preparations of lodgepole pine xylem following gel  71  electrophoresis. 3.3  Autoradiograph of first (lanel) and second strand (lane2) cDNA synthesis  71  products 3.4  Autoradiograph of fractions of cDNA following size fractionation and gel  71  electrophoresis 3.5  BamHI digestion products of D N A from 10 randomly picked plaques from  71  the Pine xylem /JLAP-cDNA library 3.6- 3.7  C B G - c D N A amplification from the XZAP-cDNA library  74  3.8  Complete nucleotide sequence (1909 nucleotide) of C B G c D N A and it's  79  translation. 3.9  Deduced amino acid sequence of C B G . letters in bold - signal peptide,  80  arrow - predicted cleavage sites, * - predicted N-glycosylation sites 3.10  Hydropathy index of C B G amino acid sequence computed using an interval  82  of 11 amino acids, arrow head - predicted signal peptide cleavage sites. 3.11  Deduced C B G amino acid sequence aligned with P-glucosidase sequences  84  showing high similarity. 3.12-3.13  Expression of C B G in E.coli and Native-PAGE  94  4.1.- 4.3  In situ enzyme activity localization and lignin staining of Pinus contorta  108  stem sections 4.4-4.5  Maule reaction stained actively growing poplar stem sections from apex to  112  the base of the stem. 4.7- 4.9  Localization of P-glucosidase, peroxidase activity and immunolocalization of O M T in actively growing poplar stems.  112  ACKNOWLEDGEMENT  I thank the members of the supervisory committee: Drs. John E. Carlson, Brian E. Ellis, Carl J. Douglas and G.H.Neil Towers for their guidance.  I express my  gratitude to Dr. Carlson for the support, opportunity and providing an atmosphere for independent research in his lab. M y gratitude also extends to Dr. Ellis for his encouragement, vision and providing ready access to equipment in the Department of Plant Science. M y thanks extend to Dr. Edith Camm for providing the Bio-Rad Econo system, Dr. Yousry El-Kassaby for Pinus contorta trees, Wade Stock for assistance in obtaining xylem scrapings and coniferin extraction, Leroy Scrubb for assistance in lignin chemical analysis and Arthur Yee for useful hints on experimental procedures.  xi  FOREWORD The contents in chapter 2 and parts of chapter 1 and 4 have been published in Plant Physiology (Dharmawardhana et al. 1995) and Canadian Journal of Botany (Dharmawardhana et al. 1992). The author of the thesis conducted all the research and writing under the guidance of Dr. J.E. Carlson and Dr. B.E. Ellis.  1  INTRODUCTION A N D O V E R V I E W  Lignin is a complex polymer of aromatic subunits derived from the phenylpropanoid biosynthetic pathway. The deposition of lignin in the wall of plant cells that have secondary wall thickenings (xylem elements and sclerenchyma) creates a three-dimensional matrix that surrounds the polysaccharide components of the cell wall. This matrix helps provide rigidity and hydrophobicity to the cells. Because of it's importance in providing structural support and in water transport functions, it is thought that the appearance of lignin biosynthesis played a major role in the evolution of terrestrial vascular plants.  In addition, lignin resists microbial attack and is  involved in plant defense and defense responses by sealing off wounding and pathogen infection sites (Dixon and Paiva 1995). Lignin makes up 15-36% of the dry weight of wood (Sarkanen and Hergert 1971), and it is one of the most abundant natural polymers in the biosphere. While it is biologically important to plants, the composition, quantity and distribution of lignin in plants has some negative consequences for the use of plants by humans.  The  quantity and composition of lignin in forage crops affect their digestibility and energy yield (Jung and Vogel 1986). The industrial removal of lignin during pulp and papermaking process involves significant energy and environmental costs. The ease of lignin extraction, however, depends, in part, on its composition and quantity in the wood pulp. Primarily for these economic reasons, there is considerable interest in trying to produce plants with modified lignin. The three monolignols, para-hydroxycoumaryl, coniferyl and sinapyl alcohol  2  are derived from amino acid, phenylalanine, through the phenylpropanoid pathway (Fig 1.1). The general phenylpropanoid pathway gives rise to several other classes of compounds in addition to lignin, such as flavanoids, coumarins, stilbenes and benzoic acid derivatives (Holton and Cornish 1995). The conversion of L-phenylalanine to fr-tms-cinnamic acid by the enzyme phenylalanine ammonia lyase (PAL) is the entry point  and  the  first  committed  phenylpropanoid pathway.  step that channels  phenylalanine  into  the  Hence, P A L is one of the most well-characterized  enzymes in the pathway. The /ra«.s-cinnamic acid is hydroxylated by cytochrome P450 dependent cinnamic acid 4-hydroxylase (C4H) to form para-coumaric acid (a 4coumaric acid). Cell fractionation studies have shown that C 4 H and a fraction of P A L reside in the microsomal fraction.  Radiolabeled feeding experiments have  shown that C 4 H utilizes cinnamic acids produced in situ by P A L more readily than exogenous cinnamic acids. These observations suggest involvement of metabolic channeling in the pathway (Hrazdina and Wagner 1985, Whetten and Sederoff 1995). Such substrate channeling has been suggested to be a means of controlling the accumulation of toxic concentrations of metabolic intermediates within cells without reducing the flux through the path (Stafford 1981, Hrazdina and Wagner 1985). /?ara-Coumaric acid undergoes a series of hydroxylation and methylation steps that are catalyzed by hydroxylases and O-methyl transferases (OMT).  The various  hydroxy cinnamic acids thus formed are activated to their C o A thioesters by 4coumarate:CoA ligase (4CL). Some recent evidence suggests that the O-methylation at 3 and 5 positions of the ring could occur at this activated Co A-ester level as well as  3  at the free cinnamic acids level (Ye et al. 1994). The C o A thioesters of cinnamic acids then undergo sequential reduction to the corresponding aldehydes and to cinnamyl alcohol in reactions catalyzed by cinnamyl-CoA reductase (CCR) and cinnamyl alcohol dehydrogenase (CAD), respectively. A comprehensive review of the enzymes involved in lignin biosynthesis is available in Whetten and Sederoff (1995).  The exact mechanism of polymerization of the differently substituted  monolignols to form lignin within the cell wall still needs clarification. Peroxidase, laccase, and possibly a specific coniferyl alcohol oxidase, have all been implicated and each has been located in lignifying cell walls (Lewis and Yamamoto 1990, O'Malley et al. 1993, Dean and Eriksson 1994, Savidge et al. 1994). Although polymerization of lignin is free radical mediated there is increasing evidence to show that the reaction process is not random; i.e. there is an order to lignin structure in situ (Atalla and Agarwal 1985, Agarwal and Atalla 1986). Most aromatic rings within the polymer appear to be oriented parallel to the cell wall surface. A s in organization of honey comb-like cellulose-hemicellulose structure, hemicellulose is suggested to play a role in the organization of lignin via lignin-hemicellulose complex formation (Terashima et al. 1993). The proportion of the different monolignols incorporated into the lignin polymer varies depending on plant species, developmental stage, tissue/cell type and environmental stresses. In general, lignin in gymnosperms such as conifers is predominantly derived from coniferyl alcohol, giving rise to a guaiacyltype lignin whereas angiosperms incorporate significant proportions of sinapyl  °-c-°  H  C4H  C3H OCH,, OH  Cinnamic acid  OH  para-Coumaric acid  • 1 4CL  Caffeic acid  1 4CL  4CL  . 4CL  O  S-CoA  S-CoA  F5H?  CC03H  CCoOMT  OCH  OH OH para-Coumaroyl-CoA |  r4CL?  3  H C 3  OH 5-Hydroxyferuloyl-CoA  Caffeoyl-CoA  J CCR J CAD  CCR  5-Hydroxyferulyl alcohol Ct^)H  GT G-O para-hydroxycinnamyl alcohol glucoside  'OCH OH  para-Coumaryl alcohol  Q  OH  OH  Coniferyl alcohol  Sinapyl alcohol  PO/Laccase  Lignin  Fig. 1.1. Lignin biosynthetic pathway (modified with permission from Douglas C. 1996). PAL, phenylalanine ammonia-lyase; 4CL, hydroxycinnamate.CoAligase; C3H, coumarate-3hydroxylate; C4H, cinnamate-4-hydroxylase; C A D , cinnamyl alcohol dehydrogenase; PG, coniferin P-glucosidase; GT, glucosyl transferase; CCo3H, coumaroyl-CoA-3-hydroxylase; CCoMT, caffeoyl-CoA 3-O-methyltransferase; CCR, cinnamyl.CoA reductase; F5H, ferulate-5-hydroxylase; PO, peroxidase; COMT, caffeic acid O-methyltransferase.  5  alcohol units into their lignin, giving rise to a mixed guaiacyl-syringyl type lignin. The composition of angiosperm lignin makes it less strongly cross-linked and thus more readily extracted during the pulping process. Differences in substrate affinities, and in the presence of different isozymes of enzymes (such as F5H, C O M T , 4 C L , CAD) involved in monolignol synthesis have been suggested to account for this difference in lignin quality between angiosperms and gymnosperms as well as between cell types at different developmental stages (Whetten and Sederoff 1995). The monolignols are relatively toxic and unstable compounds and do not accumulate in plant cells. The 4-O-glycosylation of the monolignols to form their respective cinnamyl glucosides ( para-hydroxycinnamyl alcohol glucoside, coniferin and syringin) makes them more water soluble and less cytotoxic (Whetten and Sederoff 1995). The glucosides have therefore been viewed as less reactive storage and transport forms of the monolignols. These glucosides have been identified from lignifying stems of all the gymnosperms analyzed and from some angiosperms, notably species belonging to the families Oleaceae and Magnoleaceae (Terazawa et al. 1984, Terashima et al. 1986, Fukushima and Terashima 1990). The glucoside of coniferyl alcohol (coniferin) is reported to accumulate to very high levels in lignifying cells of conifer xylem during spring cambial reactivation (Savidge  1989).  Radiolabeling studies also showed that coniferin is a very effective precursor for lignin synthesis (Terashima et al. 1986, Terashima and Fukushima 1988).  These  observations suggest that a glucosyl transferase/ p-glucosidase system may operate in lignifying tissues for the inter-conversion of cinnamyl glucosides and monolignols. A  6  UDP-glucose: coniferyl alcohol glucosyl transferase has been reported from spruce cambium (Schmid and Grisbach 1982) and other tissues (Ibrahim 1977), while enzymes capable of hydrolyzing coniferin have been reported from spruce seedlings (Marcinowski and Grisbach 1978) and cell suspension culture systems (Hosel et al. 1982, Campbell and Ellis  1992).  However, the evidence linking such a  glucosyltransferase/ glucosidase system to the process of lignin formation in lignifying cells is only fragmentary. Although most of the steps involved in monolignol biosynthesis have been fairly well characterized, the steps following this, i.e. glycosylation, storage, transport into the cell wall and in situ polymerization to form lignin remain to be established. The work reported in the first chapter of this thesis was carried out to characterize lignification in Arabidopsis thaliana which could be then used as a model system (advantages listed in Meyerowitz 1989, Meyerowitz and Pruitt 1985) to assist in identifying lignification mutants.  Chapter 2 describes the identification and  characterization of a coniferin specific p-glucosidase from Pinus contorta var. latifolia xylem. Pinus is known to accumulate high levels of coniferin during spring cambial reactivation  (Savidge 1989) and radiolabelling studies have shown that  coniferin can be effectively incorporated into lignin (Terashima et al. 1986, Terashima and Fukushima 1988). Xylem, differentiating during spring growth is an excellent resource for biochemical and enzymological work on cell wall biosynthesis. Plant cell culture systems have also been extensively used in secondary metabolism related studies, and a Pinus banksiana suspension culture system has been reported to  7  show coniferin hydro lytic activity upon elicitor induction (Campbell and Ellis 1992). However, differentiating pine xylem was considered to be a superior system for this work due to its particular involvement in cell wall biosynthesis which makes it less complex and an enriched source for enzyme purification. One of the main driving forces to obtain a deeper understanding of the lignin biosynthetic pathway and it's regulation is the interest in genetically engineering plants for modified lignin contents and qualities.  This has been attempted using antisense and sense suppression  technologies for a number of genes involved in phenylpropanoid metabolism. These include P A L (Elkind et al. 1990), C O M T (Dwivedi et al 1994, N i et al 1994, Atanassova et al. 1995), 4 C L (Lee and Douglas 1994), C A D (Halpin et al. 1994, Schuch 1993) and peroxidase (Lagrimini et al. 1993); reviewed in Campbell and Sederoff (1996) and Dixon et al. (1994). The use of genes involved in the ultimate steps of lignin biosynthesis would cause the least perturbation of biosynthetic and physiological functions of the plant that are not lignin related.  Coniferin P-  glucosidase is a very appropriate step for down regulation of lignin biosynthesis and isolation of it's c D N A would be a major step towards this goal. Chapter 3 describes the isolation of coniferin P-glucosidase c D N A from a lodgepole pine xylem library and confirmation of it's identity.  In chapter 4 the expression pattern of p-  glucosidases and other lignin biosynthetic enzymes in lignifying pine and poplar stems is examined to establish whether this is correlated in space and time with lignin deposition in the same tissue.  8  CHAPTER 1 Vascular lignification in Arabidopsis thaliana  Introduction: Lignin, one of the most abundant compounds in nature, is of critical importance in normal growth and development of plants. Lignin in angiosperms is mainly composed of syringyl and guaiacyl residues. Different morphological areas in wood have been shown to have different quantities and proportions of these residues by labeled precursor studies, UV-microscopy and bromination Transmission Electron Microscopy - Energy Dispersive X-ray Analysis ( T E M - E D X A ; Fukushima and Terashima 1991, Terashima et al. 1986, Saka and Thomas 1982). Its presence in tree species has special industrial significance, since it is a major determinant of wood and paper quality. The need to extract lignin chemically during pulp and paper production has become a major concern because of the environmental impact associated with this process.  Development of the capability to alter forest species genetically with  reference to lignin content and/or composition therefore is of considerable interest. One of the most useful methods for genetically defining a metabolic trait is the isolation and characterization of mutants affected at various points in the biosynthetic pathway. To achieve this, it is advantageous to make use of a simple model plant system like Arabidopsis that possesses a short generation time and a small genome, and for which a detailed genetic map is available (Meyerowitz 1989). The aim of the present investigation was to develop an efficient and sensitive  9  method for detection of lignin microscopically, to determine the temporal and spatial pattern of lignification during development, histochemically monitor the composition of lignin deposited during development and to chemically characterize A. thaliana lignin.  Confocal scanning microscopy, which has the capability of optical serial  sectioning and imaging thick intact tissues was employed for microscopic detection. Concomitantly, lignin preparations were subjected to alkaline nitrobenzene oxidation. The resulting data establishes a profile of wild type A. thaliana lignin and its pattern of deposition. Since lignification in A. thaliana has not been characterized previously this information would be valuable in evaluating mutant phenotypes.  Materials and methods:  Plant material: Seeds of Arabidopsis thaliana ecotype Columbia were surface sterilized in 30% Chlorox, spread on 0.7% agar minimal nutrient medium (Haughn and Somerville 1986) in six-well culture plates and grown aseptically under 16 h light of 100-150 umol"2s"l illumination at 25°C in a growth chamber. For studies involving the flowering (bolting) stem, the seeds were surface sterilized planted in soil pots and grown under the above conditions.  10  Preparation for fluorescence microscopy: Seedlings were sampled at regular intervals from time zero to 30 days post imbibition, cleared in methanol (4 hr at 25° C) and 10% NaOH (8 hr at 60°C). The following methods were used to detect lignin in the cleared intact (unsectioned) seedlings. (a) Cleared seedlings were mounted in 50% glycerol.  Epi-fluorescence under U V  (365nm) excitation was observed using a 420 nm barrier filter on a Zeiss Axioplan microscope. (b) Cleared seedlings were stained with 0.01% basic fuchsin for 5 min., destained in 70% ethanol, rehydrated and mounted in 50% glycerol. Fluorescence under green (546 nm) excitation was observed using a 590 nm barrier filter on a Zeiss Axiophot photomicroscope or imaged using a Bio-Rad M R C 500 confocal laser scanning microscope (CLSM) with an argon ion laser (lOmW at 514 run). Objectives used were Zeiss Plan-apochromat 100X (1.3 N.A.), Plan-neofluar 40X (0.75 N.A.) or Planneofluar 20X (0.5 N.A.). Digital image processing of C L S M images consisted of filtering for noise reduction through averaging (Kalman filtering) during image acquisition and scaling. Five to ten optical sections (0.2-0.4 u apart) spanning the whole specimen were projected together to obtain a composite image of the entire vascular system. Black and white photographs were taken from a flat screen, high-resolution video display with Technical-pan film at 50 A S A . Colour images were photographed off a standard colour monitor with Kodak Ektar 125 A S A film.  11  Flowering stem development and pattern of lignin deposition: Flowering stems were collected at two day intervals from bolting until maturity.  1-2 mm thick sections were cut from each flowering stem at regular  intervals from the apex to base and fixed in 4 % paraformaldehyde in p H 7 phosphate buffer. Thin hand sections from these stem segments were subjected to the Maule reaction by treating with: 1% KMnC>4 (5 min) and 12% HC1 (10 min) then mounted in saturated N H 4 O H .  The sections were observed under a Zeiss Axiophot photo  microscope and photographed using Kodak Ecktachrome 160 A S A tungsten film or Techpan (at 50 A S A ) black & white film.  Peroxidase localization: Histochemical localization of peroxidase activity was carried out using syringaldazine, diaminobenzidine and guaiacol as substrates, according to Goldberg era/. (1983).  Chemical analysis: Seedlings were extracted in boiling methanol for 30 minutes and subsequently hydrolyzed with 2 M sodium hydroxide for two hours at room temperature.  After  removal of the base, samples of the hydrolyzed tissue were either (a) processed for alkaline nitrobenzene oxidation (Galletti et al. 1989), with the analysis of oxidation products by reverse phase H P L C and U V detection, or (b) processed for  12  determination of the level of thioglycolic acid extractable lignins (Whitmore 1978).  Results:  Microscopic detection: Vascular bundles of mature A. thaliana seedlings exhibited a blue autofluorescence under U V excitation, a response that has been used as a marker for lignification (O'Brien and McCully 1981). Although the absolute specificity of basic fuchsin as a stain for lignin has yet to be determined, it is a very effective stain for the vasculature in cleared plant tissues (Fuchs 1963). The pattern of red fluorescence that I observed in basic fuchsin-stained tissue under green light excitation was identical to the U V autofluorescence pattern, confirming that this staining method was a valid indicator for microscopic detection of lignin. The red fluorescence emitted by basic fuchsin-stained vascular elements is especially useful due to its brightness, sensitivity and the possibility of employing the optical sectioning capability of the C L S M (using the 514 nm line of argon ion laser for excitation) for imaging intact tissue.  Vascular development and lignification: Up to 36 hr post imbibition, there was no evidence of vascular tissue or lignification in the A. thaliana embryo. Patchy fluorescence appeared only in the cytoplasm of cells, and this diminished with maturity.  Emergence of the radicle  13  through the seed coat occurred at around 36 hr and at this stage the first tracheary elements appeared in the radicle and cotyledons.  In the radicle these could be  detected as two rows of elongated cells with thickened walls (Fig. 1.1). Occasionally, two rows of elongated but narrower phloem cells were also discernible, near the proximal end of the radicle. Phloem is reported to begin differentiation before the occurrence of xylogenesis in the embryo (Esau 1965). At this early stage of vascular differentiation, while the secondary wall was visible microscopically along more than half of the axis of the radicle, there was no cytochemical evidence of lignification. In most seedlings, the first sign of lignification was seen at approximately 48h, when fluorescence appeared in mid and distal parts of the radicle tracheary elements (Fig.l.2B). At this stage, the tracheary elements in the radicle were clearly visible, often with closely spaced spiral or annular thickenings. Root hairs were also prominent (Fig.l.2A). The cotyledons showed a median vascular strand and a late developing peripheral loop on both, or occasionally on one side. Those seedlings that germinated earliest and grew most quickly displayed well defined spiral cell wall thickenings by 48 hr, both in the radicle and the cotyledons, and the primary axis had visibly differentiated into hypocotyl and primary root (Fig. 1.3 A). In these precocious seedlings, xylem in both the hypocotyl and the root were well lignified (Fig.l.3B). In addition, cells external to the vascular area of the root also fluoresced, perhaps reflecting the presence of suberin as well as lignin, deposited on endodermal cells (Fig.l.3B). This pattern became more prominent as the root matured (Fig. 1.12). Lignification within the cotyledons was limited to the  14  midvein at this stage and later spread acropetally (Fig.l.3B). By the third day post-imbibition, the cotyledonary leaves had freed themselves from the seed coats and were fully expanded in most seedlings. Lignification of the peripheral xylem strands had also been initiated by this time (Fig. 1.6).  In the  hypocotyl, two new rows of tracheary elements appeared inner (exarch) to the first two strands.  The differentiation and lignification of these elements proceeded  basipetally (Fig. 1.4). Fig. 1.11 shows a similar trend at day 5. Xylem elements at the root apex showed acropetal lignification with non-lignified elements (but with secondary thickenings) closest to the apex (Fig. 1.5). It is evident from Fig. 1.4 and 1.5 that lignification of the tracheary elements proceeds in a well coordinated pattern, subsequent to the laying down of annular and spiral secondary wall thickenings. By day 5, the vascular tissue in the hypocotyl was well developed and contained several rows of strongly lignified elements (Fig. 1.7). After 7 to 8 days, the first true leaves appeared.  The vasculature within these leaves differentiated as a  single strand of tracheary elements that extended along the midvein and showed acropetal lignification, again following behind the formation of spiral wall thickenings (Fig. 1.8). elements  proceeded  (Fig.1.9,1.10).  The differentiation and lignification of peripheral xylem acropetally in some  leaves and basipetally in others  15  Fig 1.1-1.3. Phase contrast (P.C.) and fluorescence (FL.) confocal laser-scanning micrographs (CLSM) of A. thaliana seedlings during early vascular development. Fig. 1.1 P.C. image of a 36 h seedling with first signs of tracheary elements (arrowhead),. Fig. 1.2A,L2B. P.C. and FL. images, respectively, of a 2 day old seedling showing first signs of lignification (arrow-head),. Fig. 1.3A, 1.3B. P.C. and FL. images, respectively, of an early germinating 2 day old seedling. Note lignified mid vein of cotyledon (black arrow-head) and fluorescence in endodermal cells (white arrow-head), Scale bar= 60 u.  17  Flowering stem development: The flowering stem emerges about 15 days after germination and  4-5  vascular strands differentiate within the stem. Fig. 1.13 shows a 18-day old seedling just prior to flowering, with two well-lignified xylem strands in the bolting stem. By day 21 (after flowering) the bolting stem showed 5 vascular bundles (Fig. 1.14,1.15). Each xylem bundle consisted of earlier developed elements with spiral thickenings (these appear stretched due to stem elongation) and recently formed pitted xylem elements (Fig. 1.16).  The hypocotyl at this stage had a thick vasculature with  numerous well lignified pitted xylem elements (Fig. 1.17,1.18). About 6 days after emergence of flowering stem, cells between the vascular bundles at the base of the stem start to differentiate into fibers with thick secondary walls (Fig. 1.20).  This  differentiation of fibers gradually proceeds towards the apex with stem maturation. Fig. 1.22 shows that by day 11 the differentiation of fibers has proceeded halfway up the stem. Cross sections close to the zone of fiber differentiation(Fig.l.20B) show that the population of fibers are separate from the vascular bundles and later fuse to the vascular bundles (Fig.l.20C). At day 22 when the elongation of the flowering stem has ceased (i.e. after pod maturation), the differentiation of fibers has progressed to the apex of the stem (Fig. 1.21). These fibers have the typical characteristics of angiosperm sclerenchymatous fibers with elongated spindle shape and occasional small simple pits.  They are not involved in conduction and probably provide  resistance to compression and bending of the bolting stem.  18  Fig. 1.4-1.10. Merged fluorescence (red) and phase contrast (green) C L S M images of A. thaliana seedling tissue (white arrow-head^ non-lignified tracheary elements). Fig. 1.4. Hypocotyl of a 3 day old seedling showing progressive lignification of tracheary elements, Scale bar= 10 u. Fig . 1.5. Distal part of a 3 day old root. Note non-lignified tracheary elements and fluorescence of endodermal cells (*), Scale bar= 40 u. Fig. 1.6. Three day old cotyledon with lignifying peripheral tracheary elements, Scale bar= 40 u. Fig. 1.7. Five day old hypocotyl with several strands of well lignified tracheary elements, Scale bar= 40 u. Fig. 1.8.  Eight day old hypocotyl  with lignifying tracheary elements of primary leaf mid vein, Scale bar= 40 u. Fig. 1.9. Twelve day old hypocotyl with lignified and non-lignified peripheral tracheary elements of primary leaves, Scale bar= 400 u. Fig. 1.10. Distal part of primary leaf (12 day) with basipetally lignifying peripheral tracheary elements, Scale bar= 10 u.  20  Fig. 1.11-1.14. Phase contrast (P.C.) and fluorescence (FL.) C L S M images of A. thaliana seedlings. Fig. 1.11A, 1.11B. P.C. and FL. images, respectively, of 5 day old hypocotyl with non-lignified inner elements (arrow-head), Scale bar= 20 u. Fig. 1.12. FL. images of root of same seedling as Fig. 11 showing outer tracheary elements with stretched secondary thickenings (arrow-head). *= endodermis, Scale bar= 20 u. Fig. 1.13. Eighteen day old bolting seedling. Note the two lignified xylem strands in the stem (arrow-head). Fig. 1.14. Twenty one day bolting seedling showing 5 lignified vascular bundles in the bolting stem, Scale bar= 400 u.  22  Fig. 1.15-1.18. Fluorescence C L S M images of A. thaliana seedlings during late vascular development Fig. 1.15. Vascular bundles of 21 day bolting stem, Scale bar= 50 u. Fig. 1.16A, 1.16B. Vascular bundles of Fig. 15 at a higher magnification. Note stretched spiral thickenings of early formed elements (arrow-head), Scale bar= 10 u. Fig. 1.17. Xylem elements in the upper hypocotyl of 21 day seedling, Scale bar= 25 u. Fig. 1.18. Higher magnification of Fig. 17 showing well lignified pitted xylem elements, Scale bar= 5 u.  2,3  24  Pattern of lignin deposition: Lignin deposited in vessel elements of vascular bundles (in both proto and metaxylem) is of the guaiacyl type, as shown by their golden-brown color with Maule reagent. This remains true throughout stem development (Fig, 1.23 - 1.26). Five to six days after appearance of the bolting stem, the first signs of syringyl type lignin (Maule reaction- pink) appeared in differentiating fibers between vascular bundles at the base of the stem (Fig. 1.24). With the differentiation of fibers towards the apex during stem development, the syringyl-guaiacyl type lignin deposition would closely follow cell wall thickening in the newly formed fibers (Fig. 1.25 and 1.26). The differentiation and lignification of this population of fibers proceed centripetally as opposed to centrifugal differentiation of xylem in vascular bundles. This pattern is analogous to differentiation of phloem fibers in primary phloem of most dicot angiosperms, suggesting that the fiber bundles of Arabidopsis bolting stem might have a phloem origin. Syringyl-type lignin deposition is apparently switched on at the onset of fiber lignification although the predominantly guaiacyl type lignin may be synthesized simultaneously in other cell types.  By day 22, both fiber  differentiation and syringyl-guaiacyl lignin deposition have nearly reached the apex (Fig. 1.22). A t this stage the base of the stem has a well-lignified cylinder of vascular tissue several cell layers thick and the heterogeneity of predominantly guaiacyl and syringyl-guaiacyl lignin types in vessels and fibers is still maintained (Fig. 1.26). This heterogeneity is present even in hypocotyl (Fig. 1.27) and in root tissue (data not shown).  25  Fig, 1.19-1.21. Line diagram of flowering stem at several developmental stages and cross sections of stem to show differentiation of vascular bundles and fibers. Fig. 1.19. Two day old stem (2 days after bolting) showing developing vascular bundles. Fig. 1.20,1.21. 11 and 22 day old stems, respectively, with the cross sections stained with toluidine blue to show developing vascular bundles (arrow) and fibers (*).  200  150  100  50  8  9  10  11  12  13  14  15  16  17  18  19  20  21  22  Days after bolting  Fig. 1.22. Spatial distribution of syringyl and guaiacyl lignin along the Arabidopsis stem during development (based on staining of sections taken at regular intervals along the stem with Maule reaction)  28  Fig. 1.23-1.26. Histochemical staining for guaiacyl lignin (brown colour) and syringyl lignin (pink colour) using Maule reaction at different developmental stages of the stem.  Fig. 1.27. Cross section of a 30 day old (15 days after bolting) hypocotyl following Maule reaction. Note absence of syringyl lignin in exarch xylem elements . The root displays a similar pattern.  Fig. 1.28A,B- Locali2ation of peroxidase activity using syringaldazine (mauve colour) in a section of a 15 day old stem.  Fig.1.29-1.30. Comparison of wild type (right) and fahl mutant (left) lacking syringyl type lignin using Maule reaction. Fig. 1.29. Intact stem Fig. 1.30. Cross sections of stems.  30  Fig. 1.28 shows peroxidase enzyme activity localization (indicative of ongoing lignification) using syringaldazine as a substrate during two developmental stages of the stem. The other substrates tested showed background activity in non-lignifying parenchyma cells. It is apparent in Fig. 1.23 and 1.24 that the vascular bundle vessel elements are the first to lignify during development.  Only guaiacyl lignin is  deposited in these xylem elements and one could assume that they have ceased lignifying by the fiber differentiation stage. However, Fig. 1.28 demonstrates that at this stem developmental stage, when active fiber lignification (including syringyl lignin deposition) is occurring, the walls of the metaxylem vessels, which contain only guaiacyl lignin, are still actively lignifying. This spatial separation of guaiacyl and syringyl-guaiacyl type in vessels and fibers of A. thaliana. suggests that the syringyl path of lignin biosynthesis is not active during lignin deposition in xylem vessel walls, although the physically adjacent fiber cells have the capability of syringyl lignin synthesis.  Composition of lignin: Analysis of the levels and monomer composition of lignin from A. thaliana showed that young seedlings contained too little lignin for accurate assessment. The bolting stems, however, yielded substantial amounts of thioglycolic acid lignins (Table 1.1). The monomer composition of the bolting stem lignin is typical of that found in many herbaceous angiosperms, with a predominance of guaiacyl units (G:S 77:23) accompanied by a significant contribution of syringyl moieties.  31  The Arabidopsis phenylpropanoid metabolism in mutant fahl is affected in ferulate 5-hydroxylase and has a modified lignin composition that is dominated by guaiacyl lignin (Chappie et al. 1994, Chappie et al. 1992). Fig. 1.29 - 1.30 clearly demonstrates the absence of syringyl lignin in fibers of the fahl mutant, and confirms that the histochemical analysis of wild type lignin distribution accurately reported the relative distribution of guaiacyl and syringyl-type lignin in Arabidopsis thaliana.  Table 1.1.  Chemical analysis of lignin from Arabidopsis thaliana tissue  Sample  Thioglycolic acid lignins*  Nitrobenzene  (A o-g"'fresh wt. 10" )  oxidation products  2  28  4-Day old seedlings  0.11  60-Day old aerial tissue  1.46  N.D.  4-hydroxybenzaldehyde(tr.) vanillin (77%) syringaldehyde (23%)  A l l values represent the average of duplicate analysis. N.D. = not detectable  33  Discussion:  Although Arabidopsis thaliana has a very short life cycle (flowering within 3 weeks) and does not undergo secondary growth, the data presented here show that it still displays extensive lignification throughout the plant, starting as early as 48 h after germination. As the radicle, hypocotyl and cotyledons expanded, lignification proceeded cell-by-cell as a temporally coordinated event closely following the appearance of spiral and annular thickenings in the developing vasculature. Interestingly, developmental regulation in the type of lignin deposited in the flowering stem was observed. Lignin deposited up to 20 - 21 days after germination (5 days after bolting) is predominantly of the guaiacyl type. Syringyl-type lignin appeared only after differentiation of fibers between the vascular bundles in the flowering stem.  Such heterogeneity in the type of lignin deposited in fibers and  vessels is also reported in studies using different labeled lignin precursors (Fukushima and Terashima 1990, Terashima et al. 1986). Since the developmental pattern and the biochemical nature of this lignin formation appear to be typical of other higher plants, A. thaliana should make an effective model in which to study the biochemistry and molecular biology of lignin biosynthesis. The spatial and temporal separation of syringyl-type lignin along the developing stem (Fig. 1.22) also makes it a unique system to study developmental regulation in the syringyl path of lignin biosynthesis, a trait which affects the quality of lignin deposited in wood in trees. The methods developed in this study should also  34  be appropriate for studies of vascular development in other plant tissues and organs. These results also provide a sound basis on which to develop a screening program for possible lignification mutants.  However, an initial screening of 4000 E M S  mutagenized seedlings for reduced lignification using the basic fuchsin induced lignin fluorescence did not yield any viable lignin mutants.  35  CHAPTER 2 Identification, purification and properties of coniferin p-glucosidase  Introduction Lignin is a complex polymer of three cinnamyl alcohols, p-coumaryl, coniferyl and sinapyl, all products of phenylpropanoid metabolism. Depending on the plant species or tissue, the relative proportion of the different monomers in lignin can vary significantly. In gymnosperms, lignin is predominantly composed of coniferyl alcohol monomer units, whereas angiosperms have significant proportions of syringyl moieties. The 4-O-glucoside of coniferyl alcohol, coniferin, is known to accumulate in conifer xylem during cambium reactivation, consistent with a role as the dominant lignin precursor in these species (Freudenberg and Harkin 1963, Savidge 1989). This model is supported by data showing that radioactive monolignol glucosides are readily incorporated into the lignin polymer (Matsui et al. 1994, Terashima and Fukushima 1988). Before being polymerized to lignin, however, coniferin must be hydrolyzed to free the phenolic hydroxyl. While the cinnamyl alcohol glucoside/ p-glucosidase system has been implicated in lignification, the evidence has been primarily correlative.  p-  Glucosidases capable of hydrolyzing coniferin have been detected in cell suspension culture systems (Hosel et al. 1982,  and Todenhagen  1980) and seedlings  (Marcinowski and Grisebach 1978), but, until now, a coniferin glucosidase has not been identified that derives from differentiating xylem of mature trees, the most  36  active site for lignin biosynthesis in woody plants. Modification of quality and quantity of lignin in plants for increasing the strength of wood, or for enhancing pulpability of trees, has been a long-standing interest among tree breeders and, more recently, among molecular biologists. Alteration of lignin levels by antisense and sense suppression of gene expression has already been attempted for several enzymes in the phenylpropanoid pathway including P A L (Elkind et al. 1990), O M T (Ni et al. 1994) C A D (Halpin et al. 1994, Schuch 1993) and 4CL (Lee and Douglas 1994).  It is clear, however, that  interference with early steps in the phenylpropanoid pathway can have undesirable pleiotropic effects (Elkind et al. 1990).  Coniferin hydrolysis to release coniferyl  alcohol is considered to be one of the final steps in the reaction sequence of conifer lignin biosynthesis, which makes coniferin p-glucosidase a particularly suitable target for biotechnological modification of lignin in trees. This possibility gains support from the observation that the only lignification-related phenylpropanoid metabolite accumulating to a significant degree in lodgepole pine xylem tissues is coniferin (Savidge 1988 and 1989). I therefore undertook to characterize such an enzyme in lignifying conifer tissue as an initial step towards cloning the corresponding gene. The following  chapter  describes the  major  P-glucosidases found in  differentiating lodgepole pine xylem, and the properties of a coniferin-specific enzyme that has been purified to apparent homogeneity.  37  Materials and methods  Plant material: Ten to fifteen year-old Pinus contorta var. latifolia Engelm. trees from the U.B.C. Research Forest (Haney, British Columbia) and from Pacific Forest Products Ltd. (Saanich, Vancouver Island) were felled at the beginning of the growing season (late May to early June). The logs were quickly debarked, the differentiating xylem layer scraped off with razor blades, and the tissue flash-frozen in liquid nitrogen for storage at -70°C.  Biochemicals and buffers: The glucosides used for detecting P-glucosidase activity in chromatographic fractions were coniferin, 4-nitrophenyl P-glucoside (4-NPG), 2-nitrpphenyl Pglucoside (2-NPG), 4- methylumbelliferyl p-glucoside (MUG) and 5[4-(p-Dglucopyranosyloxy)-3-methoxyphenylmethylene]-2-thioxothiazolidine-4-one-3ethanoic acid (VRA-G). V R A - G is a substrate analogue of coniferin synthesized by Biosynth International Inc., Skoke, Illinois.  Coniferin was extracted and purified  from pine xylem scrapings as described in van-Uden (1991).  The purity of  recrystallized coniferin was compared with authentic coniferin by H P L C , T L C and U V analysis. Syringin was provided by Prof. Neil Towers (Dept. of Botany, U . B . C ) . Buffers: A = 50mM M E S , pH 6.0; B = 20mM M E S , pH 5.5; C = 20mM M E S , pH 5.5, 0.5M NaCl; and D = 50mM MES, pH 5.8, 2.0M ammonium sulfate.  38  A l l buffers contained 3mM DTT and 10% glycerol.  Enzyme extraction: Frozen xylem tissue was immersed in liquid nitrogen and ground in a prechilled stainless steel blender (Waring). The resulting powder was further ground with liquid nitrogen using a mortar and pestle, and extracted twice ( l g tissue: 2ml buffer)  by  stirring for 30 min. with cold buffer  polyvinylpolypyrrolidone (O.lg/ g fresh weight).  A containing insoluble  The crude extract was filtered  through two layers of fine nylon mesh and clarified by centrifugation at 10,000g for 30 min. To obtain the salt-extractable P-glucosidases, the crude cell wall residue remaining from the buffer extraction was further extracted with two volumes of buffer A containing 0.5M NaCl for 3h at 4°C, followed by filtration and centrifugal clarification as above.  Assay for P-glucosidases: The standard assay containing the enzyme preparation (10-50uT) and substrate (2mM final concentration) in 0.2M MES, pH 5.5 buffer (final volume: 150ul) was incubated at 30°C for 30 min. The reaction was stopped by adding an equal volume of 0.5M C A P S , pH 10.5 and the activity was measured by determining the absorbance of the released aglycone in the alkaline mixture. calculations, the following analysis wavelengths and  For quantitative  s values ( m M ' l x cm~l) were  used: coniferyl alcohol, 325nm, s= 7.0; sinapyl alcohol, 315nm, e= 11.2; 2-  39  nitrophenol, 420nm, s= 4.55, 4-nitrophenol, 400nm, 8=19.3; 4-methyl umbelliferone, 360 nm, s= 18.25; V R A , 490nm, e= 38.6; salicyl alcohol, 295nm, e= 3.3. p-glucosidase activity in all fractions generated chromatographically during enzyme purification was monitored using both I m M coniferin and 4-NPG as substrates. The ability of purified coniferin P-glucosidase and partially purified M U G Pglucosidase to hydrolyze various substrates was determined at 2mM concentration. K m values for these substrates were calculated using both Lineweaver-Burk and Hanes plots. Inhibitor studies with the P-glucosidase inhibitors, bromoconduritol mixed isomers  (6-bromo-4-cyclohexene-1,2,3,-trio 1), conduritol  B  epoxide  (DL-1,2-  anhydro-wyoinositol) and glucono-1,5-lactone, were carried out with 0.1-5.0 m M inhibitor, and with coniferin or M U G as substrates (2mM).  Determination of the profile of P-glucosidases: The extract was brought to 95% saturation with ammonium sulfate, stirred for 2h at 4°C and centrifuged at 12,000g for 30 min. The pellet was dissolved in buffer B and desalted using a column of BioGel P6DG resin in the same buffer.  The  desalted extract was subjected to anion exchange chromatography on a Q M A MemSep 1010 cartridge (Millipore). Proteins were eluted in a shallow gradient of buffer C.  A l l chromatographic fractions were assayed for P-glucosidase activity  using the range of glucosides listed under Biochemicals and Buffers.  40  Enzyme purification: The buffer extract was brought to 70% saturation with solid ammonium sulfate, stirred gently at 4°C for 1 h, and the protein precipitate was collected by centrifugation at 12,000g for 30 min. The pellet was resuspended in buffer B and desalted using a BioGel P6DG column. The desalted extract was applied to a Q Sepharose column (2.5x30cm) equilibrated with buffer B, washed with the same buffer to remove unbound protein and then eluted with a linear gradient of 0.05-0.4M NaCl, using buffer C.  Chromatographic fractions having coniferin glucosidase  activity were pooled and solid ammonium sulfate was added to a final concentration of 2M. After adjusting the pH to 5.8, the sample was fractionated by hydrophobic interaction chromatography on a 1.5x20cm tertiary-butyl Macroprep (Bio-Rad) column. The elution was carried out with a decreasing salt gradient of ammonium sulfate (2.0-0.0M) and a simultaneously increasing pH gradient (5.8 to 6.6).  The  active fractions were desalted into buffer B (pH 5.7) by ultrafiltration on 5K M W cutoff Filtron centrifugal filters or stirred cell device, and subjected to a second anionexchange purification using a Q M A MemSep 1010 (Millipore) cartridge equilibrated with buffer B (pH 5.7) and a 0.02M - 0.12M NaCl gradient. Active fractions were concentrated by ultrafiltration as above and chromatographed on a Superose 12 H R 10/30 (Pharmacia) gel filtration column, equilibrated with buffer B containing 0.25M NaCl.  The most active fractions were used for enzyme characterization and  electrophoretic analysis. A l l the above chromatographic separations were performed  41  using a Bio-Rad Econo low-pressure chromatographic system or an (Pharmacia) system.  FPLC  Protein profiles during chromatography were monitored by  absorbance at 280 nm. Quantification of protein was carried out using a Bio-Rad protein assay kit, based on the Bradford method (Bradford, 1976). For molecular mass determination of the native enzyme, the Superose 12 H R 10/30 (Pharmacia) gel filtration column was calibrated using ferritin (Mr=440,000), catalase '(Mr= 232,000), aldolase (Mr=l 58,000), bovine serum albumin (Mr=67,000) and chymotrypsinogen (Mr=25,000). The isoelectric point of the enzyme was estimated by chromatofocussing of purified enzyme on a Mono-P H R 5/10 (Pharmacia) column equilibrated with 20mM Bis-Tris pH 5.5, using a pH gradient of 5.5 to 4.0, in lOX-diluted Polybuffer 74 (Pharmacia).  SDS-PAGE: Denaturing gel electrophoresis was conducted according to the standard Laemmli (1970) procedure, in 12% polyacrylamide gels.  The molecular weight  standards used were Rainbow prestained markers (Amersham) or Promega mid-range molecular weight markers.  For protein visualization, gels were stained with  Coomassie Brilliant Blue, or silver (Sambrook et al. 1989).  Native P A G E and activity staining: Electrophoresis under non-denaturing conditions was performed at 4°C  42  according to the  Laemmli procedure  but with omission of SDS and P-  mercaptoethanol, and a reduction in separating gel buffer pH to 8.4 to enhance the stability of the enzyme.  For P-glucosidase activity staining, the gels were  equilibrated in 20 m M M E S , pH 5.5 for 20 min., followed by incubation in 2mM V R A - G , or 0.2 m M M U G for l h . The gels incubated with V R A - G were basified with 0.25M CAPS, pH 9 to intensify the orange color of the aglycone. The gels incubated with M U G were washed in three changes of water and the aglycone was detected by its fluorescence under U V light. For SDS-PAGE analysis of native-gel protein bands that had stained positive for P-glucosidase activity, the active bands were excised into 2xSDS loading buffer, equilibrated for 30 min. at room temperature (or boiled for 4 min.), transferred to sample wells of SDS-PAGE gels (with stacking gel at least three times the height of the gel slices) and electrophoresed.  N-terminal sequencing: The purified enzyme was run on preparative native-PAGE gels and stained for P-glucosidase activity using V R A - G . The active band was excised and subjected to SDS-PAGE as described above.  After electroblotting onto Immobilon P 1-PVDF SC  membrane (Millipore), the excised 28 kD and 24 kD protein bands were subjected to N-terminal sequence analysis by automated Edman degradation on an Applied Biosystems 470A gas phase sequencer with on-line PTH-HA analysis (Protein Microchemistry Center, University of Victoria).  43  Database searches were performed with B L A S T P version 1.3.1 IMP (Altschul et al. 1990) and F A S T A version 1.4c (Pearson and Lipman 1988) on SWISSPROT release 28 and PIR release 41.0.  Antibodies against the synthetic peptide: The sequence of amino acid residues 2-16 from the 24 kD protein was used to direct synthesis of a 15-residue long peptide using an Applied Biosystems 430A automated peptide synthesizer at the Nucleic Acid and Protein Services Unit, Biotechnology Laboratory, University of British Columbia. was coupled to cationized B S A (Imject  The synthetic peptide  Super CarrienPierce) by l-ethyl-3-  (dimethylaminopropyl) carbodiimide (EDC)-mediated amide formation, following the manufacturer's recommendations with some modifications.  Briefly, the carrier  (cationized B S A ) and the peptide were dissolved in conjugation buffer (pH 4.3), followed by the addition of 0.5mg/ ml EDC.  After incubation at 25°C for 1.5h,  unconjugated peptide and residual E D C were removed by gel filtration on a BioGel P6DG column. The conjugation was confirmed by electrophoretic analysis on a SDSP A G E gel. Polyclonal antibodies to this conjugate were elicited in a female New Zealand white rabbit by injecting (subcutaneous) 300 \xg of conjugate and complete Freund's adjuvant followed by boosters (intramuscular) every six weeks. Test bleeds were performed 10 days after each booster injection. Affinity removal of anti-BSA antibodies from the serum was carried out by incubating diluted serum (1: 50 dilution with PBS) overnight at 4°C with B S A -  44  loaded nitrocellulose membrane.  Western blot analysis and glycoprotein detection: After SDS-PAGE or native P A G E , proteins were electroblotted onto nitrocellulose membranes and total protein was detected with 0.2% (w/v) Ponceau S in 3% acetic acid. After destaining in PBS, the blots were blocked with 3% gelatin from cold-water fish skin (Sigma) in PBS. The western blots were developed using a Promega western blot kit with alkaline-phosphatase conjugated secondary antibody, following the manufacturer's instructions.  Primary antibody (rabbit anti-peptide  antiserum) was used at a dilution of 1:700. A n SDS-PAGE blot of the purified enzyme was probed for glycoproteins, using a Boerhinger Mannheim digoxigenin glycan detection kit, following manufacturer's recommendations.  Results:  P-Glucosidases in differentiating xylem: Initial experiments revealed that the pH optimum for pine xylem pglucosidases, tested against a variety of substrates, was in the range of pH 5-6. Following anion exchange chromatography, the total extractable xylem protein could be resolved into three major populations of p-glucosidases (Fig.2.1). Only peak 3  45  had coniferin hydrolyzing activity, and it was also active against the coniferin analog, V R A - G . The p-glucosidases in peaks 1 and 2 were able to hydrolyze all the synthetic glucosides used, including V R A - G , but not coniferin. Activity-stained native P A G E gels of proteins from the three p-glucosidase activity peaks revealed a different pattern in each peak (Fig.2.2). The M U G activity of peak 1 migrated only slightly into the separating gel and appeared diffuse (Fig. 2.2, lane 1). On lower percentage (7%) polyacrylamide gels, this activity produced a broad smear without discrete bands on the upper half of the separating gel. The relative activity of peak 1 varied substantially from extraction to extraction, even in tissues of the same tree. This material might be an aggregated or complexed form of the p-glucosidase(s) found in peak 2, given that the substrate preferences were also similar.  In contrast, activity peaks 2 and 3 each yielded single major bands on  activity-stained gels (Fig.2.2, lanes 2 and 3), with the coniferin-active peak (peak 3) showing a band of V R A - G hydrolyzing activity that migrated slightly faster than the MUG-hydrolyzing activity observed in peak 2 . Based on these activity staining responses, the P-glucosidases in the peaks 2 and 3 were designated M U G Pglucosidase and coniferin P-glucosidase, respectively.  1  5  9  13  17  21  25  29  Fraction number  Figure 2.1. Distribution o f P-glucosidases in total extractable Pinus contorta xylem protein, following anion exchange chromatography (0-0.17 M N a C l gradient). Each fraction was assayed for P-glucosidase activity using 4 - N P G , 2 - N P G , V R A - G , M U G and coniferin as substrates.  47  Figure 2.2. Native P A G E and activity staining of the three major p-glucosidase populations (peak 1, 2 and 3 in Fig. 1) resolved by anion exchange chromatography. The gels were stained for P glucosidase activity using M U G or V R A - G as substrates (B) and for protein with Coomassie Brilliant Blue (A). Lanes 1, 2 and 3 are protein profiles and P glucosidase activity of proteins in peak 1,2 and 3 respectively. BSAd, bovine serum albumin dimer (MW 132,000); BSAm, bovine serum albumin monomer (MW 66,000); C E A , chicken egg albumin (MW 45,000).  Figure 2.3. Documentation of sequential purification of coniferin P-glucosidase. Pooled active fractions from each purification stage were subjected to SDS-PAGE on 12% gels and visualized by silver staining ( 1-5): lane 1, desalted 70% ( N F L ^ S C ^ precipitate; lane 2, Q Sepharose pool; lane 3, t-butyl pool; lane 4, Q M A MemSeplOlO pool; lane 5, most active Superose 12 fraction.  Figure 2.4. Coomassie blue stained preparative SDS-PAGE blot of purified enzyme. The 28 kD and 24 kD bands were excised and used for N-terminal amino acid sequencing.  Figure 2.5. Native P A G E analysis of purified coniferin p-glucosidase from P. contorta. Following native P A G E , one lane was sliced into 0.5 cm segments, equilibrated for 20 min. with 0.5M MES pH 5.5, each segment pulverized with 200 ul of 2mM coniferin (pH 5.5), incubated for 2 h and following basification the absorbance of the released aglycone (coniferyl alcohol) was measured at 325 nm. A duplicate lane was stained in situ with V R A - G (B) and subsequently silver stained to detect proteins (A).  49  Further extraction of  buffer-extracted xylem cell walls with 0.5M NaCl  released less than 5% additional coniferin hydrolyzing activity, but it did release another 20%-30% of the P-glucosidases active against M U G , which included other minor species in addition to the major species observed in peaks 1 and 2.  Purification of coniferin P-glucosidase: The optimized purification protocol resulted in > 2700 fold enzyme purification, in four chromatographic steps (Table 2.1 and Fig. 2.3). The resultant preparation produced a single band on silver-stained native P A G E gels(Fig. 2.5A), which coincided with P-glucosidase activity staining on gels using the chromogenic coniferin analog, V R A - G , and with coniferin hydrolytic activity on gel slices from duplicate lanes (Fig.2.5 ). SDS-PAGE of the single band from native gels produced two protein bands of molecular mass 28 kD and 24 kD (Fig. 2.4), the 24 kD band being of lower intensity. A smaller molecular weight band close to the dye front was also discernible on some gels, especially on Coomassie stained gels with high protein loading, but this probably represents a degradation artifact (Fig.2.4). The omission of P-mercaptoethanol from the SDS-PAGE loading buffer did not change the banding pattern or the observed molecular mass.  50  Table 2.1. Purification of coniferin p-glucosidase from Pinus contorta xylem.  Purification stage  0  total activity  specific activity  purification  yield  mg  nKat  nKat/mg  -fold  % 100  crude extract  2010  80.1  0.04  a(NH4)2S04  1100  42.2  0.04  1  52.6  Q Sepharose  40.00  16.9  0.42  11  21.1  Hydrophobic interaction  6.00  8.3  1.38  35  10.4  MemSep Q M A  0.20  2.9  14.50  364  3.6  bo.Ol  1.1  2760  1.3  Superose 12  a  protein  measurement after buffer exchange. estimated values from silver stained native gels  110.0  51  Properties of coniferin P-glucosidase: The molecular mass of the native enzyme, by calibrated Superose-12 gel filtration chromatography, was 60,000 D. This value is consistent with the relative mobility of the coniferin P-glucosidase and of the native molecular weight standards on native P A G E gels.  The molecular mass estimated from the mobility of the  purified protein on SDS-PAGE was 28 kD (and/or 24 kD), implying a dimeric structure. The optimum pH for coniferin hydrolytic activity at 30°C was between 5.4-5.9 and the enzyme was most stable within that pH range.  The isoelectric point,  estimated by chromatofocussing of the pure enzyme, was pH 4.5. The protein of interest reacted as a glycoprotein when blots were probed for the presence of sugars by periodate oxidation and labeling with digoxigenin via a hydrazide conjugate, followed by immunodetection.  Catalytic properties: The activity of both coniferin P-glucosidase (purified) and M U G Pglucosidase (partially purified) on a range of glycoside substrates is summarized in Table 2.2. M U G p-glucosidase could hydrolyze a range of synthetic glucosides, and showed particularly high affinity for the two synthetic substrates, 4-NPG and M U G . The hydrolysis of coniferin and syringin by this preparation was slow and yielded high K m values. Interestingly, the galactoside of 2-nitrophenol was also hydrolyzed to some degree by the M U G p-glucosidase.  52  Table 2.2. Substrate specificity of purified coniferin p-glucosidase and partially purified M U G p-glucosidase from Pinus contorta:  Relative activity substrate  coniferin syringin 4-methyl umbelliferyl-Pglucoside 2-nitrophenyl-p-glucoside 4-nitrophenyl-P-glucoside VRA-G salicin 4-methyl umbelliferyl-aglucoside 2-nitrophenyl-P-galactoside 4-nitrophenyl-P-cellobioside  coniferin P-glucosidase (coniferin =100)'  M U G P-glucosidase (4-methylumbelliferyl-p-glucoside)  100 (0.18mM) 51 (0.29mM) 16(2.3mM)  21(>5mM) 23 (> 5mM) 100 (0.18mM)  57 26(1.9mM) 45 10 1  96 75 (0.17 mM) 62 60 4  12 ND  37 2  N D : not detectable numbers in parentheses are K m values of the respective substrates 12  ' 100% activity represents 13pKat for coniferin and 8pKat for 4-methylumbelliferyl P-glucoside, respectively  2  53  In contrast, coniferin P-glucosidase was very active against the native substrate, coniferin, and the related syringin, for which low K m values were obtained. The only other substrates that were hydrolyzed to a substantial degree by the coniferin P-glucosidase were 2-NPG and the coniferin analog, V R A - G (Table 2.2). At lower substrate concentrations (<0.5mM), V R A - G actually exhibited a higher activity than coniferin.  This could be due to non-linearity of the V R A - G assay at higher  concentrations and/or due to the saturation of enzyme with V R A - G at lower concentrations. Neither enzyme was capable of hydrolyzing a disaccharide-containing glycoside, 4-nitrophenyl P-cellobioside or a-linked M U G . Out of the three p-glucosidase inhibitors bromoconduritol (mixed isomers), conduritol B epoxide and glucono 1,5-lactone, only glucono 1,5-lactone was effective in inhibiting coniferin P-glucosidase. The M U G p-glucosidase (partially pure) was effectively inhibited by both bromoconduritol (mixed isomers) and glucono 1,5lactone.  N-terminal amino acid sequence: The 28 kD protein appeared to be N-terminally blocked since it did not give a signal during automated sequencing. The 24 kD protein, on the other hand, yielded a 17 amino acid sequence, which was found to have high homology to known plant Pglucosidases. The optimal sequence alignment shows that the 24 kD coniferin Pglucosidase N-terminal sequence has a 60%-70% similarity to the N-terminal region  54  of P-glucosidases from Zea mays, Brassica, Arabidopsis, cassava and barley (Fig. 2.6).  Immunological analysis: A synthetic 15 amino acid N-terminal peptide was made immunogenic by coupling it to the carrier protein, cationized-BSA (c-BSA). In addition to eliciting an enhanced immune response compared to unmodified B S A (Muckerheide et al. 1987), use of cBSA enabled me to couple the peptide predominantly through its C-terminal carboxyl group, since the majority of available carboxyl groups on c-BSA have been substituted with aminoethylamide groups. Coupling via carboxyl groups of the two aspartate residues of the peptide was suppressed by acidifying the coupling reaction to pH 4.3. SDS-PAGE analysis of c-BSA before (Fig. 2.7, lane 2) and after (Fig. 2.7, lane 1) peptide coupling confirmed the efficiency of the reaction. Following the removal of anti-BSA antibodies, the induced anti-peptide-cBSA serum reacted with a single protein band in partially purified (after one anion exchange step) coniferin P-glucosidase fractions, as well as with the completely purified protein (Fig. 2.8B) on Western blots of native P A G E gels. On SDS-PAGE blots, it reacted strongly with the 28 kD protein and more weakly with the 24 kD protein from which the antigenic peptide sequence had been derived. However, the intensity of reaction on these blots also reflects the amount of protein present in the 28 kD and 24 kD bands. (Fig. 2.8A). The abundance of the 24 kD band relative to the 28 kD band was low and variable between different enzyme preparations.  These  55  results suggest that the 28 kD and 24 kD proteins are very similar, at least in their N terminal sequence. One possible explanation is that the 24 kD protein is a processing or degradation product of the 28 kD protein subunit (in vivo or post-extraction).  CBG A48860 S52771 S57621 x94986 S56656 S41869 S23940  01 72 32 34 39 39 14 28  KLDRNNFPSDFTFGTAS IPQRDWFPSDFTFGAAT TLSRA'SFPEGFMFGTAT KLSRASFPEGFLFGTAT NFSRSYFP DDFIFGTAT RLNSKGFPKDFIFGVSS GLSRQGFPAGFVFGTAA D FSRKYFP DDFIFGTAT  Figure 2.6. Optimal N-terminal amino acid sequence alignment of the coniferin P-glucosidase 24 kD band and other plant p-glucosidases. Amino acids with letter underlined denote positions of questionable identity. Sequences have been aligned to maximize similarity, and identical residues are in bold. The number to the left of the sequence represent the first residue in each line. C B G , Pinus contorta p-glucosidase; A48860, Zea mays P-glucosidase; S52771, Brassica napus p-glucosidase; S57621, Arabidopsis thaliana thioglucosidase; X94986, Manihot esculenta P-glucosidase; S56656, Brassica napus thioglucosidase; L41869, Hordium vulgare p-glucosidase; S23940, Manihot esculanta p-glucosidase.  57  F i g u r e 2.7. Detection of peptide/carrier protein coupling. The synthetic 15- amino acid long N-terminal peptide following coupling to cationized-BSA was analyzed by SDS-PAGE. Lane 1, peptide-BSA conjugate; lane 2, C-BSA before coupling.  F i g u r e 2.8. Western blot analysis of coniferin P-glucosidase using polyclonal antibodies raised against the 15- amino acid synthetic peptide to the N-terminus of the 24 kD protein. Blots of SDS-PAGE (A) and native P A G E (B) gels of, partially purified ( after Q- Sepharose step, Table 1) or purified coniferin p-glucosidase, were probed with antibody following staining for proteins. Lane 1 and 2 SDS-PAGE protein profile of partially purified and purified enzyme respectively; lane 3 and 4, subsequent immunodetection of proteins in lanes 1 and 2; lane 5 and 6, native P A G E protein profile of purified enzyme and immunodetection respectively. Arrowheads, 28 kD and 24 kD bands.  BSAd, bovine serum albumin dimer ( M W 132,000);  BSAm, bovine serum albumin monomer (MW 66,000); C E A , chicken egg albumin (MW 45,000).  1  2  A  B  59  Discussion  This study shows that lignifying lodgepole pine xylem possesses two major types of aromatic P-glucosidases. One ( M U G p-glucosidase) efficiently hydrolyzed a variety of synthetic aromatic glucosides, while the second enzyme, coniferin Pglucosidase, was far more specific for the native substrate, coniferin. The coniferin-specific p-glucosidase was purified to apparent homogeneity and appears to have a native molecular mass of 60,000 D. SDS-PAGE analysis of the purified enzyme revealed two major bands of 28 kD and 24 kD, but the immunological data, together with the asymmetric quantity of protein in the 2 bands, suggest that the native structure could be a 28 kD homodimer. A P-glucosidase has been reported from barley endosperm that also shows two immunologically similar proteins differing in apparent molecular weight by 3-4 kD on SDS-PAGE (Simos et al., 1994). These proteins also change in their relative abundance during different stages of germination. The N-terminal sequences of the two barley protein bands are identical and it was suggested that the lower molecular weight form is the result of limited in vivo proteolysis of the higher molecular weight form. If this scenario is also applicable to the coniferin P-glucosidase (which shows 60% similarity in N terminal amino acid sequence to the barley protein), then the processed 24 kD protein, of relatively low abundance, may be co-migrating with the dominant 28 kD protein as a homo- or hetero-dimer in native P A G E gels, with the apparent 4 kD subunit  60  difference of this minor species showing up only upon separation of subunits during SDS-PAGE. Although the origin of the 28 kD and 24 kD proteins cannot be specified definitively, their N-terminal amino acid sequences are immunologically crossreactive. The high homology of this sequence to other cloned plant p-glucosidases, together with the demonstration of coniferin-hydrolyzing specificity and V R A - G staining of a single protein band in native gels, strongly support the identification of these subunits as coniferin P-glucosidase. Unlike this coniferin p-glucosidase from P. contorta, the coniferin-hydrolyzing enzyme detected in Norway spruce seedlings (Marcinowski and Grisebach 1978), is an apparently monomeric, 59 kD protein. Most known glucosidases are monomeric, but several plant P-glucosidases are reported to be oligomeric, including a P-glucosidase from cassava (Yeoh and Woo 1992), dhurrinase from sorghum ( Hosel et al. 1987), prunasin hydrolase from black cherry (Li et al. 1992), and linamarases from Trifolium repens (Hughes and Dunn 1982) and flax seeds (Fan and Conn 1985). As with most other glycosidases, the coniferin P-glucosidase from pine is also a glycoprotein. The P. contorta coniferin p-glucosidase displays a clear substrate preference for coniferin and closely-related molecules.  Syringin, the glucoside of sinapyl  alcohol, differs from coniferin only by a single O-methyl group and is also hydrolyzed efficiently. Other coniferin-hydrolyzing glucosidases from Glycine max suspension cultures ( Hosel and Todenhagen 1980) and spruce seedlings were also capable of using both of these hydroxy cinnamyl alcohol glucosides as substrates.  61  Although lodgepole pine tissue appears to accumulate only coniferin during the period of rapid lignification, Freudenberg and Harkin (1963) reported that spruce cambial sap did contain a small amount of syringin in addition to large amounts of coniferin. Coniferin P-glucosidase might therefore be responsible for the hydrolysis of both of these glucosides to release monolignols for lignin biosynthesis in conifers, but the predominance of guaiacyl units in conifer lignin would reflect the composition of the substrate supply stream, rather than the specificity of the p-glucosidase. While the spruce seedling p-glucosidase described by Marcinowski and Grisebach (1978) hydrolyzed coniferin/syringin, its reported  physico-chemical properties  differ  markedly from those of the coniferin P-glucosidase reported here, most notably in the basic nature of the spruce seedling enzyme (pl= 10), as opposed to the acidic nature of the P. contorta enzyme (pl= 4.5). Following the conclusion of this study and submission for publication, a report appeared on two coniferin-hydrolyzing enzymes from Pinus banksiana (Leinhos et al. 1994). Although these two enzymes could hydrolyze coniferin, their molecular weights (100 and 90 kD), pi (pH 3.8) and N-terminal amino acid sequences were very different from the P. contorta coniferin P-glucosidase described in the present study. The purity of the Pinus banksiana enzymes were not convincing, hence their characteristcs should be accepted only with some reservation.  It would also be  interesting to have comparative kinetic data on the substrate specificities of these Pinus banksiana p-glucosidases. Freudenberg and Harkin (1963) proposed that, during lignin formation, the  62  monolignol precursors are transported to the cell wall as glucosides, and that specific P-glucosidases within the cell wall would regenerate the monolignol aglycones for in situ polymerization. The P. contorta coniferin P-glucosidase has the high activity and affinity expected of such an enzyme but the localization of the activity in the soluble fraction of the tissue extracts suggests that it is not strongly associated with the cell wall. Nevertheless, its properties suggest that it performs a crucial role in pine stem lignification. The information and tools generated by this part of the study were next employed to isolate the corresponding cDNA clone for the P. contorta coniferin Pglucosidase.  63  CHAPTER 3  cDNA cloning and heterologous expression of P. contorta coniferin P-glucosidase  Introduction: p-glucosidases have a wide occurrence and diverse functions in different organisms and many of the corresponding genes have been cloned from bacterial and animal species. During the past few years several P-glucosidase cDNAs and genes from plant species have also been identified (Leah et al. 1995, Xue et al. 1995, Lenman et al. 1993, Hughes et al. 1992, Oxtoby et al. 1991). From the information and tools gathered during the study of the purification and characterization of the P. contorta coniferin p-glucosidase, the following avenues were open for cloning its gene: screen an expression library with the anti-peptide antibody, screen an expression library for p-glucosidase active clones using the chromogenic  substrate  VRA-G,  screen  a  library with  labeled  degenerate  oligonucleotide pools based on N-terminal amino acid sequence, and P C R amplify target D N A using N-terminal amino acid sequence-based primers.  Since most of  these possibilities required a c D N A library, a c D N A library was constructed in the XZA? vector using mRNA isolated from lignifying xylem tissue, a source which should be enriched for C B G transcripts. The following account describes the construction of a c D N A library from  64  lodgepole pine xylem, cloning and characterization of full length coniferin (3glucosidase c D N A from the library, and expression of the c D N A in E. coli.  Materials and methods:  R N A isolation: A l l precautions for handling R N A were taken as described in Sambrook et a/. (1989). Total R N A from xylem scrapings was isolated according to Lewinsohn et al. (1994). In brief, the tissue was ground in liquid nitrogen, then extracted with buffer (200 m M Tris pH 8.5, 10 m M EDTA, 300 m M L i C l , 5 m M thiourea, 1 m M aurintricarboxylic acid, 10 m M dithiothreitol and 1% polyvinylpolypyrolidone), centrifuged to remove particulates, and the polysaccharides were precipitated with 10% ethanol. Nucleic acids were precipitated with 33% isopropanol (2 h, -20°C). To isolate R N A the total nucleic acids were subjected to discontinuous CsCl gradient ultracentrifugation. The R N A was dissolved in DEPC treated water, then quantified by absorbance measurements at 260nm.  Electrophoresis in 1.5% T E agarose or  denaturing formaldehyde gels (Sambrook et al. 1989) was carried out to determine the integrity of the R N A based on the intensity of 28S and 18S r R N A bands. PolyA R N A was isolated from 1-2 mg of total R N A using Oligotex mRNA isolation kit (Qiagen) following the manufacturer's instructions.  65  Construction of c D N A library: c D N A synthesis from 5ug xylem polyA R N A was carried out using a Z A P c D N A synthesis kit (Stratagene) according to the manufacturer's instructions. The efficiency and size range of first strand and second strand synthesis were evaluated by 32  incorporating [a- P]dATP during synthesis, followed by alkaline agarose gel electrophoresis of aliquots and autoradiography.  After blunt ending of the c D N A  with Pfu D N A polymerase, EcoRl adapters were ligated.  The modified c D N A  population was then Xhol-digested and size fractionated using a Pharmacia cDNA fractionation column.  The size of cDNA in each fraction was evaluated by  electrophoresis and autoradiography. The fraction containing cDNAs > 300bp was quantified by agarose ethidium bromide plate assay and ligated to E c o R l / X h o l digested A.ZAP-XR vector, which was then packaged using a Stratagene Gigapackll Gold packaging kit, according to manufacturer's instructions. The library was titered on E.coli strain X L 1-Blue M R F ' using L B plates with 1% top agar containing IPTG and X-gal.  Amplification of the library was carried out according to Stratagene  instructions.  PCR amplification of CBG-cDNA: XZAP-cDNA from the amplified library was prepared according to the plate lysate method in Current Protocols in Molecular Biology (Ausubel et al. 1995). Initial P C R reactions contained 200-300 ng XZAP-cDNA as template, 200 n M degenerate  gene-specific  primer  N7A  [GCTCTAGAGCGAC(T)A(C)GI  66  AAC(T)AAC(T)TTTCC] or N7B [GCTCTAGAGCGAC(T)A(C)GIAAC(T)AAC (T)TTCCC], 50 n M vector primer M13F or T7 (BRL), 200uM dNTP, and I X reaction buffer (lOmM TrisHCl pH 8.3, 1.5mM M g C l , 50mM KC1). Prior to adding 2  3 units Taq polymerase (Boehringer), the reaction mixture was heated to 94°C for 2 min.  The thermal cycling regime was as follows: 1-2 cycles (94°C/lmin., 48°-  52°C/2min.,72°/2min.); 30 cycles (94°C/45sec.,55°/lmin., 72°C/2min.) : 72°C/10min. extension. To increase specificity and identify the desired amplification product (Fig. 3.1 step 1), 20ng reaction products from the initial P C R was reamplified using a partially  nested  gene  specific-primer  TTCCCIT(A)C(G)IGA(T)TT  and  N10  vector  [GAC(T)A(C)GIAAC(T)AAC(T) primer  T7  (30  94°C/45sec.,55°/l min., 72°C/2min.and 72°C/10min. final extension).  cycles  of  The reaction  products were analyzed by agarose gel electrophoresis. Following identification of the 1.7kb band as the desired amplification product, the initial PCR reaction was repeated with less (0.9mM) M g C l  2  in the  reaction buffer . The reaction products were separated on a 1% agarose gel, the 1.7kb band excised, purified using a QIAquick gel purification kit (Qiagen) and used for cloning.  c7  N7 - gene specific primer N10 - gene specific primer  C B G 1 7 2 - gene specific primer (based on seq. info, from step 1)  Flg.3.1. PCR amplification strategy used for isolating a complete CBG-cDNA from a lodgepole pine xylem AZAP library.  68  Cloning and sequencing: The PCR products were cloned into EcoRV-digested T-tailed Bluescript II K S vector according to the T/A cloning protocol of Holton and Graham (1991). Fifteen white colonies were picked for plasmid minipreps (Zhou et al. 1990) and digested with appropriate enzymes to confirm the presence of the insert, or for restriction analysis of the insert. Plasmid D N A from clones selected for sequencing (1A6, 1A7, 1A2, 1A2, 1A3 and 1A5) was prepared according to a mini alkaline lysis/PEG precipitation procedure (Ausubel et al. 1995). Primer-directed sequencing of both strands of the insert was carried out using A B I AmpliTaq dye termination cycle sequencing  chemistry  according to the manufacturer's protocol and analyzed on a A B I 373 D N A sequencer (Nucleic Acid and Protein Services unit, Biotechnology Lab., U.B.C.)  Cloning of the 5' end of the C B G cDNA: XZAP-cDNA from the library was again used as template to amplify the 5' end of the C B G using T3 vector primer and a gene-specific primer CBG172 (CACATATCTGTGATATTGGTCG)  based on sequence of C B G clone 1A6  determined previously (Fig.3.1 step2 ).  A second nested gene-specific primer  CBG75(CCATCTTCTCGGACTGCTC) was used to reamplify the former reaction products to confirm the authenticity of the reaction product. sequencing of the PCR product was conducted as described earlier.  The cloning and  69  Sequence comparison and analysis: Nucleotide and amino acid sequence homology searches and comparisons were carried out using B L A S T (Altschul et al. 1990) on Genbank, E M B L , P D B , SWISS-PROT and PIR databases. Nucleotide and amino acid sequence analysis of C B G was performed using PC/Gene (Intelligenetics Inc.) software.  Expression of C B G in E.coli: To express C B G protein in E. coli, the full length coding region for the mature protein (i.e. excluding the signal peptide) was amplified using a N-terminal primer NT1 ( T A G C T A G C A G G C T G G A C A G G A A C A A C T T C ) containing a 5' Nhel site, a C-terminal primer C T l ( C T C G A G A C A A G C A G T C T A A A T G C T ) containing a X h o l site, and clone 1A6 as the template. The resulting 1.5kb D N A fragment was ligated into Bluescript II K S by T / A cloning as described earlier. construct were sequenced to ensure validity.  The two ends of this  The insert was then excised with  Nhel/Xhol and ligated into similarly digested and phosphatase-treated expression vector pET21a (Novagen).  Ligated plasmids were transformed into E.coli strain  DH5a and the presence of the insert was verified by restriction digestion of plasmid minipreps. The plasmid was then introduced into the expression host BL21(DE3) and transformants selected with 100u,g/ml ampicillin. To express the C B G , the bacteria were grown to log phase (A =0.6-0.9) followed by an additional 2-3 h incubation at 600  29-37°C in the presence or absence of 0.4-ImM IPTG. Following incubation, the  70  cells were harvested and washed with buffer (50mM Tris pH 8, 2mM EDTA) and frozen at -70°C until analysis. The soluble and insoluble proteins in both induced and uninduced bacterial cells were isolated according to procedures in Current Protocols in Molecular Biology (Ausubel et al. 1995). The expressed C B G was purified by preparative Q-Sepharose chromatography followed by QMA-Memsep (Millipore) chromatography.  Enzyme assays to detect C B G activity, electrophoresis of the  proteins and detection of glucosidase activity in gels were conducted as described in chapter 2.  Results:  Isolation of mRNA and construction of cDNA library: Initial attempts to isolate total R N A from xylem tissue following the protocol recommended for plant tissues in Current Protocols in Molecular Biology (Ausubel et al. 1995) failed to give adequate yields. However, a protocol optimized for isolating R N A from woody stems of gymnosperms (Lewinsohn et al. 1994) gave good yields ' of total R N A (100 ug/g.fwf).  Based on the relative intensity of 28S and 18S  ribosomal R N A bands following denaturing agarose gel electrophoresis (Fig.3.2) the R N A appeared undegraded and was free of D N A contamination. Isolation of polyA R N A using the Oligotex m R N A isolation kit gave mRNA yields of 15-20u,g/mg of total R N A used.  71  Fig.3.2. Total R N A from 3 preparations of lodgepole pine xylem following gel electrophoresis.  Fig.3.3. Autoradiograph of first (lanel) and second strand (lane2) cDNA synthesis products following alkaline agarose gel electrophoresis.  Fig.3.4. Autoradiograph of fractions of cDNA following size fractionation (using a Pharmacia c D N A fractionation column) and gel electrophoresis.  Fig.3.5. BamHI digestion products of D N A from 10 randomly picked plaques from the Pine xylem AZAP-cDNA library.  kb  M  1  2  3  4  5  M 6 7 8 9 10 M  kb  73  The efficiency of c D N A synthesis was monitored by determining the size range of first and second strand synthesis products by autoradiography (Fig.3.3). The c D N A synthesized had a size range of 0.1-10 kb and was used for ligation of EcoRl adaptors.  Following size fractionation of the cDNA, fraction number 1 (Fig.3.4)  which had a c D N A size range of 0.4 - 10 kb was used for construction of the library by ligating to A.ZAP vector. 350,000 pfu.  The primary library produced contained more than  Fig.3.5 demonstrates the insert size range of 10 randomly picked  plaques. The average insert size was 1.4 kb with a range of 0.5 - 4.0 kb (0.9 kb should be subtracted from the apparent insert size measured in the gels to account for vector sequences included).  PCR amplification of C B G - c D N A : Degenerate gene-specific primers were designed based on the N-terminal amino acid sequence of the purified enzyme determined earlier.  To decrease the  complexity of the oligonucleotide pool, deoxyinosine was incorporated at codon positions with complete degeneracy. The N 7 A and N7B primers based on the first 7 amino acids contained 1 inosine, and a Xbal restriction site was added at the 5' end as another alternative for cloning i f T/A cloning failed. The N 7 A and N7B primers were identical except that the degeneracy at the third base from the 3' end was split between the two primers to carry out separate P C R reactions. Fig.3.6 lane 2 shows the amplification products using N7B and vector primer M l 3 , following optimization  74  Fig.3.6. C B G - c D N A amplification from the A Z A P - c D N A library. Lane 1 - using primers N 7 A and M 1 3 F , lane 2 - using primers N 7 B and M13F.  Fig.3.7. P C R amplification of 5' end of C B G - c D N A from the  A.ZAP-CDNA  library.  Lane 1 - using primers C B G 172 and T 3 , lane 2 - reamplification of product in lanel using nested primer  CBG75  and  T3.  kb  2.0-  M  1 2  1 2  M  kb  •2.0  76  of amplification conditions. Primer N 7 A and vector primer M l 3 needed a lower annealing temperature to produce amplification products and gave numerous bands. Reamplification of the N7B/M13 reaction products with partially nested gene-specific primer N10 and vector primer T7 identified the 1.7 kb band as the most likely C B G cDNA candidate. significant product.  Reamplification of N7A/M13 reaction products did not yield a Together with poor amplification in the initial reaction, this  indicated that the degenerate third 3' base of N7A does not match the C B G template. To reduce chances of P C R errors, the 1.7 kb band was isolated from a fresh P C R reaction (i.e. without reamplification) with a reduced M g C l  2  concentration.  Following gel purification, the 1.7 kb band was cloned into Bluescript II K S and sequenced. The 5' end sequence of the cDNA matched the N-terminal amino acid sequence of the enzyme determined previously, except at the 12th position which was reported as Thr in the protein but appears as Met in the D N A sequence. To amplify the 5' end of C B G cDNA, a gene specific primer (CBG172), 200 nt internal to the N-terminus of the mature protein was designed based on the sequence of the 1.7 kb band from previous step. The amplification of X-ZAP-cDNA with C B G 172 and vector primer T3 produced a single band at 0.5 kb (Fig.3.7 lane 1). Reamplification of a smaller 0.4 kb band with the nested primer CBG75 and vector primer T3 confirmed the authenticity of the amplification product (Fig.3.7 lane 2). The amplification of only one band in the initial reaction, with no background smear demonstrates that almost all of the longer CBG-cDNAs in the library are full length. To clone the 5' end fragment of the CBG-cDNA, the P C R was repeated  on the  77  library using CBG75 and the T3 vector primer, but with reduced M g C l levels. Since 2  the reaction products contained only one band, it was directly used for cloning into Bluescript II KS.. Fifteen clones containing the expected size of insert when digested with an enzyme (BamHl) having a cut site within the insert all demonstrated identical restriction fragments. This and an exact sequence match of the 79 bp overlapping region between 3' end clones (1A6 and 1A7) and these 5' end clones ( F l , F4 and F14) confirmed that this fragment is the matching 5' end of the C B G sequence. Although the traditional approach of screening a library for isolating a full length c D N A was not used in this study, the above discussed facts confirm the authenticity of the obtained full length C B G c D N A sequence.  Nucleotide sequence of C B G : The c D N A of two clones (1A6 and 1A7) representing the nucleotide sequence from the N-terminus of the mature protein to the 3' end of the mRNA (3' end clones) were sequenced in both directions. The sequences of the two clones were identical except at nucleotide 438 where 1A6 had the base pair A / T instead of C / G in 1A7. Since this could be a P C R error during the initial isolation of the cDNA, 3 more clones (1A2, 1A3, 1A5) were partially sequenced to include this region. A l l 3 clones had C/G at nucleotide 438, indicating that the A/T found at this position in clone 1A6 might be an error due to a base misincorporation during PCR. Three c D N A clones ( F l , F4, F14) representing the 5' end of the mRNA showed identical Sequences when sequenced. The 79 bp overlapping region between the 3' end clones (1A6 and 1A7)  78  and 5' end clones ( F l , F4 and F14) had identical sequences. The presence of a unique restriction site (XmnI) in this overlapping region will facilitate straightforward ligation of the 3' and 5' clones to generate a full length clone. The 1909 bp complete sequence of C B G and the deduced amino acid sequence are shown in Fig.3.8.  Nucleotides 183 - 1721 comprise a single open  reading frame coding for a 513 amino acid protein. Only this reading frame had an open reading frame longer than 130 amino acids. The 5' and 3' untranslated regions of the full length sequence contain 162 and 187 nucleotides respectively. The 3' untranslated region does not contain the conserved polyadenylation signal A A U A A A . This is not surprising since more than 50% of reported plant mRNA sequences lack the canonical polyadenylation motif (Wu et al. 1995). Instead, most plant mRNAs carry A A U A A A - l i k e sequences (Joshi 1987), which is also true for the C B G 3' untranslated region.  CBG-CDNA:  1909bp,  513AA  GGATTTGGACCTGAAAATATCAATTTCAAAGCAATTCCAGAGGGATAACGTGGGATCCTTACCATTACCAAC AACCCACCATTCCGCCCTGCCGACCTCAGGCATATTTTGATTCTATTTAACCATTAATTCATCTGGGCAGTT GTGATTCTGTATAATTCGATCGCTCCGTTTTAGCAGACATGGAGGTGTCTGTGTTGATGTGGGTACTGCTCT M  E  V  S  V  L  M  W  V  L  L  TCTATTCCTTATTAGGTTTTCAAGTGACGACAGCTAGGCTGGACAGGAACAACTTCCCCTCAGATTTCATGT F  Y  S  L  L  G  F  Q  V  T  T  A  R  L  D  R  N  N  F  P  S  D  F  M  TCGGCACAGCCTCTTCAGCGTATCAGTATGAAGGAGCAGTCCGAGAAGATGGCAAGGGTCCTAGCACATGGG F  G  T  A  S  S  A  Y  Q  Y  E  G  A  V  R  E  D  G  K  G  P  S  T  W  ACGCCTTAACACATATGCCTGGTAGAATAAAAGATAGCAGCAATGGAGACGTGGCAGTCGACCAATATCACA D  A  L  T  H  M  P  G  R  I  K  D  S  S  N  G  D  V  A  V  D  Q  Y  H  GATATATGGAAGATATCGAGCTTATGGCTTCACTTGGACTAGATGCCTATAGATTCTCCATATCCTGGTCTC R  Y  M  E  D  I  E  L  M  A  S  L  G  L  D  A  Y  R  F  S  I  S  W  S  GAATCCTTCCAGAAGGAAGAGGTGAAATTAACATGGCTGGGATTGAATATTACAATAATCTGATTGACGCTC R  I  L  P  E  G  R  G  E  I  N  M  A  G  I  E  Y  Y  N  N  L  I  D  A  TTCTGCAAAATGGGATCCAGCCGTTCGTGACATTGTTCCATTTCGATCTTCCCAAAGCACTTGAAGACTCCT L  L  Q  N  G  I  Q  P  F  V  T  L  F  H  F  D  L  P  K  A  L  E  D  S  ATGGGGGATGGCTGAGTCCTCAAATAATTAACGACTTCGAAGCCTATGCAGAGATTTGCTTCCGGGCATTCG Y  G  G  W  L  S  P  Q  I  I  N  D  F  E  A  Y  A  E  I  C  F  R  A  F  GTGACCGTGTCAAATATTGGGCGACAGTGAACGAGCCAAATCTGTTTGTGCCGTTGGGATACACCGTCGGAA G  D  R  V  K  Y  W  A  T  V  N  E  P  N  L  F  V  P  L  G  Y  T  V  G  TATTTCCACCGACGAGGTGTGCTGCCCCTCACGCCAATCCTTTGTGCATGACAGGGAATTGCTCGTCAGCAG I  F  P  P  T  R  C  A  A  P  H  A  N  P  L  C  M  T  G  N  C  S  S  A  AGCCATATCTAGCTGCACATCACGTTTTGCTCGCCCACGCATCTGCAGTGGAGAAATATAGGGAGAAATATC E  P  Y  L  A  A  H  H  V  L  L  A  H  A  S  A  V  E  K  Y  R  E  K  Y  AGAAAATTCAAGGAGGATCTATAGGGTTAGTTATAAGCGCGCCATGGTACGAACCCTTGGAAAATTCTCCAG Q  K  I  Q  G  G  S  I  G  L  V  I  S  A  P  W  Y  E  P  L  E  N  S  P  AAGAGAGATCAGCTGTTGATAGAATTTTATCCTTCAATCTCCGATGGTTTTTGGATCCAATTGTTTTTGGAG E  E  R  S  A  V  D  R  I  L  S  F  N  L  R  W  F  L  D  P  I  V  F  G  ATTATCCACAAGAAATGCGTGAAAGATTAGGATCGCGCTTACCCTCCATATCCTCGGAACTATCTGCGAAAC D  Y  P  Q  E  M  R  E  R  L  G  S  R  L  P  S  I  S  S  E  L  S  A  K  TTCGGGGATCGTTCGACTATATGGGTATTAATCACTATACAACCTTATATGCAACAAGCACTCCTCCCCTTT L  R  G  S  F  D  Y  M  G  I  N  H  Y  T  T  L  Y  A  T  S  T  P  P  L  CCCCCGACCACACGCAATATCTATATCCAGACTCTAGGGTTTATCTGACTGGAGAGCGCCACGGAGTCTCCA S  P  D  H  T  Q  Y  L  Y  P  D  S  R  V  Y  L  T  G  E  R  H  G  V  S  TCGGAGAACGGACAGGGATGGACGGTTTGTTTGTGGTACCTCATGGAATTCAAAAAATAGTGGAGTATGTAA I  G  E  R  T  G  M  D  G  L  F  V  V  P  H  G  I  Q  K  I  V  E  Y  V  AAGAATTCTATGACAACCCGACTATTATTATCGCAGAGAACGGTTATCCAGAGTCTGAGGAATCCTCGTCGA K  E  F  Y  D  N  P  T  I  I  I  A  E  N  G  Y  P  E  S  E  E  S  S  S  CTCTGCAAGAAAATCTAAACGATGTGAGGAGAATAAGGTTTCATGGAGATTGTTTGAGTTATCTCAGTGCAG T  L  Q  E  N  L  N  D  V  R  R  I  R  F  H  G  D  C  L  S  Y  L  S  A  CAATCAAAAATGGCTCAGATGTTCGAGGGTACTTTGTGTGGTCACTTCTGGATAATTTTGAGTGGGCATTTG A  I  K  N  G  S  D  V  R  G  Y  F  V  W  S  L  L  D  N  F  E  W  A  F  GGTATACCATTAGATTTGGTCTTTATCACGTGGATTTCATTTCTGATCAAAAGAGATATCCCAAGCTCTCGG G  Y  T  I  R  F  G  L  Y  H  V  D  F  I  S  D  Q  K  R  Y  P  K  L  S  CTCAATGGTTCAGACAATTTCTTCAGCACGACGATCAGGGAAGTATTAGAAGCAGCAGCAGCATTTAGACTG A  Q  W  F  R  Q  F  L  Q  H  D  D  Q  G  S  I  R  S  S  S  S  I  -  CGTTGTCTATTTGCTAATCAAAGCGCACACATTCCTGCAACTCTACCCAAAATCCTGCAAGCAAATATGTTG TGTTCGGATCTATCCACCGTGAGACACATTAGAAAGAAATCATCAATCTATTCCAAAATGCAGAAAACCCCA TTCAGATGTTCTAGGGAACTGCAGGGTAAGAACATTG  Fig.3.8. Complete nucleotide sequence (1909 nucleotide) of C B G c D N A and its translation.  80  MEVSVLMWVL  LFYSLLGFQV  ALTHMPGRIK DSSNGDVAVD  TTARLDRNNF PSDFMFGTAS  SAYQYEGAVR EDGKGPSTWD  QYHRYMEDIE LMASLGLDAY RFSISWSRIL PEGRGEINMA  GIEYYNNLID ALLQNGIQPF VTLFHFDLPK ALEDSYGGWL SPQIINDFEA YAEICFRAFG • DRVKYWATVN EPNLFVPLGY TVGIFPPTRC AAPHANPLCM TGNCSSAEPY LAAHHVLLAH ASAVEKYREK YQKIQGGSIG LVISAPWYEP LENSPEERSA VDRILSFNLR WFLDPIVFGD YPQEMRERLG SRLPSISSEL SAKLRGSFDY MGINHYTTLY ATSTPPLSPD HTQYLYPDSR VYLTGERHGV SIGERTGMDG LFWPHGIQK IVEYVKEFYD NPTIIIAENG YPESEESSST LQENLNDVRR IRFHGDCLSY LSAAIKNGSD VRGYFVWSLL DNFEWAFGYT IRFGLYHVDF ISDQKRYPKL SAQWFRQFLQ HDDQGSIRSS SSI  Fig.3.9. Deduced amino acid sequence of CBG.  letters in bold - signal peptide,  arrow- predicted cleavage sites, * - predicted N-glycosylation sites, underlineddetermined N-terminal amino acid sequence of the purified enzyme.  81  Primary structure of C B G and similarity to other p-glucosidases: The 1540 nucleotide cDNA reading frame codes for a 513 amino acid protein with a molecular weight of 58.3 kD and a calculated isoelectric point of pH 4.9. The N-terminal amino acid sequence determined for the purified 24kD protein corresponds to amino acids 24 - 41 in the deduced sequence (underlined Fig.3.9). Met35 in the deduced sequence was reported as Thr during N-terminal amino acid sequencing.  This mismatch could be a mistake during amino acid sequencing, or  could represent a polymorphism, since xylem samples used for enzyme purification and c D N A library construction were obtained from 2 different trees. The nascent protein contains a N-terminal signal peptide characteristic of eukaryotic secretory signal sequences for E R targeting. The hydrophobicity plot of the protein (Fig.3.10) clearly demonstrates the hydrophobic character of the signal peptide. Amino acids 2 21 were also identified as a possible transmembrane segment according to the method of predicting membrane spanning segments by Klein et al. (1985).  The "weight  matrix" method of von Heijne (1986) predicts two possible cleavage sites for the signal peptide, one between amino acid Glyl7 and Phel8 and a second between Ala23 and Arg24. Since the earlier determined N-terminal amino acid sequence of the mature protein (Fig.3.9 underlined) begins at Arg 24, it is likely that the cotranslational processing of the signal peptide occurs at the predicted second cleavage site (unless both co-translational and later processing steps are involved).  Amino acid  Fig.3.10. Hydropathy index of CBG amino acid sequence computed using an interval of 11 amino acids, arrow head - predicted signal peptide cleavage sites.  83  The protein contains 2 putative N-asparagine glycosylation sites at Asn223 and Asn447 that match the consensus glycosylation motif of Asn-X-(Ser or Thr), where X could be any amino acid except Pro. At amino acids 34 - 48 it also carries the N-terminal signature sequence F,X,(FYWM),(GSTA),X,(GSTA),X,(GSTA),(GS TA),(FYN),X,E,X(GSTA) characteristic of family 1 glycosyl hydrolases (Henrissat 1991).  Two of the 5 cysteine residues (Table 3.1) found in C B G (Cysl75 and  Cys225) are also conserved in p-glucosidases showing high homology to C B G (Fig.3.11).  They may be involved in forming important intramolecular disulfide  bridges. The derived amino acid sequence of C B G , when compared to other glycohydrolase sequences in the databases showed the strongest similarity to enzymes belonging to family 1 glycosyl hydrolases (Henrissat 1991). This family of glycosyl hydrolases share sequence similarities and likely similar folding characteristics. The P-glucosidases showing the highest similarity to C B G are aligned for maximum homology in Fig.3.11.  P-Glucosidases from plant species Prunus, Hordeum,  Trifolium, Manihot, Sorghum, Avena, Brassica and Arabidopsis showed the highest similarity to C B G (30-50% identity).  Most of these enzymes were cyanogenic  glucosidases active against amygdalin, dhurrin and linamarin, or thioglucosidases acting on glucosinolates.  84  Fig.3.11. Deduced C B G amino acid sequence aligned with p-glucosidase sequences showing high similarity. C B G A A , Pinus contorta coniferin P-glucosidase; L41869, Hordeum P-glucosidase; U26025, Primus amygdalin hydrolase; U39228, Prunus Pglucosidase; X56733, Trifolium cyanogenic P-glucosidase; P26204, Trifolium non cyanogenic P-glucosidase; X94986, Manihot P-glucosidase; S35175, Manihot linemarase; U33817, Sorghum dhurrinase; A48860, Zea p-glucosidase; X78433, Avena P-glucosidase; X89413, Arabidopsis thioglucosidase; S52771, Brassica Pglucosidase;  Q00326,  Brassica thioglucosidase;  X79195, Arabidopsis  thioglucosidase; L11454, Arabidopsis thioglucosidase, LPH3HU, Human L P H sub unit III; A48969, Bacillus p-glucosidase; Q08638, Bacillus B-glucosidase; S45675, Streptomyces B-glucosidase (Bgl3). * - perfectly conserved amino acid, . - well conserved amino acid. The numbers preceding the sequences represent their data base accession numbers.  Fig.  3.11  CBGAA L41869 U26025 U39228 X56733 P26204 xg4986 S35175 U33817 A48860P X78433 X89413 S52771P Q00326 X79195' L11454 LPH3HU A48969P Q08638 S45675  MEVSVLMWVLLFYSLLGF-Q VTTARLDRN N MRSSPVL—LLVIALVAA-AHLAPLECDGPNPNPEIGNTGGLSRQ G T—KLGSLLLCALLLAGF-A-LTNSKAAKTDPPI—HCA-SLNRS S LLLLGF-A-LANTNAARTDPPV--VCA-TLNRT N LLSI-T-TTHIHAFKPLPISFDDFS-DLNRS C M--DFIVAIFALFVISSF-T-ITSTNA--VEASTLLDIG-NLSRS S MASKHSLHLFGLLIVFLV-S-LLLVLTNQATAFDGDFIPLNFSRS Y M LVLFIS-L-LALTRPAMGTDDDDDNIPDDFSRK- Y MALLLASAINHTAHPAGLRSH PNNESFSRHHLCSSPQNISKRRSNLS MAPLLAAAMNHAAAHPGLRSHLVGPNNESFSRHHLPSSSPQSSKRRCNLS MA-LLCSALSNSTH-PSFRSH-IGANSENLW—HLSADPAQKSKRRCNLT MVLQKLPLIGLLLLLT IVASPANAD-GPVCPPSNKLSRAS M KFPLLGLLLLVT LVGSPTRAEEGPVCPKTETLSRAS MKLLHGLALVFLLAAASCK ADEEITCEENNP MKLL-GFALAILLW ATCK PEEEITCEENVP MKLL-MLAFVFLLALATCK GDEFVCEENEP WEKF SSQPKFERDLF M M VVP  29 43 39 29 29 39 44 33 47 50 45 39 37 31 30 29 15 — 1 — 1 — 3  CBGAA L41869 U26025 U39228 X56733 P26204 X94986 S35175 U33817 A48860P X78433 X89413 S52771P Q00326 X79195 L11454 LPH3HU A48969P Q08638 S45675  DFMFGTASSAYQYEGAVRE GFVFGTAASAYQVEGMARQ -LEPGFIFGTASAAYQFEGAAKE -LFPGFTFGTATASYQLEGAANI - — PGFVFGTASSAFQYEGAAFE - — RGFIFGAGSSAYQFEGAVNE DFIFGTATSAYQIEGAANK FPD DFIFGTATSAYQIEGEATA FPD FRPRAQTISSESAGIHRLSPWEIPRRDWFPPSFLFGAATSAYQIEGAWNE FTTRSARVGSQN-GVQMLSPSEIPQRDWFPSDFTFGAATSAYQIEGAWNE LSSRAARISSALESAKQVKPWQVPKRDWFPPEFMFGAASAAYQIEGAWNE FPEGFLFGTATAAYQVEGAINE FPEGFMFGTATASYQVEGAVNE FTCSNTDILSSKN FGKDFIFGVASSAYQIEGGR-FTCSQTDRFNKQD FESDFIFGVASSAYQIEGGR— FTCNQTKLFNSGN FEKGFIFGVASSAYQVEGGR— YHGT FRDDFLWGVSSSAYQIEGAWDA FPSDFKWGVATAAYQIEGAYNE VKK FPEGFLWGVATAS YQIEGS PLA AAQQTATAPDAALT FPEGFLWGSATASYQIEGAAAE  51 65 64 54 51 61 66 55 97 99 95 61 59 64 63 62 41 27 27 39  CBGAA L41869 U26025 U39228 X56733 P2 6204 X94 98 6 S35175 U33817 A48860P X78433 X89413 S52771P Q00326 X79195 L11454 LPH3HU A48969P Q08638 S45675  DGKGPSTWDALTHM-PGRI-KDSSNGDVAVDQYHRYMEDIELMASLGLDA GGRGPCIWDAFVAI-QGMI-AGNGTADVTVDEYHRYKEDVGIMKNMGFDA DGRGPSIWDTYTHNHSERI-KDGSNGDVAVDQYHRYKEDVRIMKKMGFDA DGRGPSIWDAFTHNHPEKI-TDGSNGDVAIDQYHRYKEDVAIMKDMGLDA DGKGPSIWDTFTHKYPEKI-KDRTNGDVAIDEYHRYKEDIGIMKDMNLDA GGRGPSIWDTFTHKYPEKI-RDGSNADITVDQYHRYKEDVGIMKDQNMDS FGRGASVWDTFTHQYPERI-LDHSTGDVADGFYYRFKGDIQNVKNMGFNA KGRAPSVWDIFSKETPDRI-LDGSNGDVAVDFYNRYIQDIKNVKKMGFNA DGKGPSTWDHFCHNFPEWI-VDRSNGDVAADSYHMYAEDVRLLKEMGMDA DGKGESNWDHFCHNHPERI-LDGSNSDIGANSYHMYKTDVRLLKEMGMDA GGKGPSSWDNFCHSHPDRI-MDKSNADVAANSYYMYKEDVRMLKEIGMDS TCRGPALWDIYCRRYPERC-NND-NGDVAVDFFHRYKEDIQLMKNLNTDA GCRGPSLWDIYTKKFPHRV-KNH-NADVAVDFYHRFREDIKLMKKLNTDA -GRGVNVWDGFSHRYPEKAGSDLKNGDTTCESYTRWQKDVDVMGELNATG -GRGLNVWDGFTHRYPEKGGADLGNGDTTCDSYRTWQKDLDVMEELGVKG -GRGLNVWDSFTHRFPEKGGADLGNGDTTCDSYTLWQKDIDVMDELNSTG DGKGPSIWDNFTHT-PGSNVKDNATGDIACDSYHQLDADLNMLRALKVKA DGRGMSIWDTFAHT-PGKV-KNGDNGNVACDSYHRVEEDVQLLKDLGVKV DGAGMSIWHTFSHT-PGNV-KNGDTGDVACDHYNRWKEDIEIIEKLGVKA DGRTPSIWDTYART-PGRV-RNGDTGDVATDHYHRWREDVALMAELGLGA  99 113 113 103 100 110 115 104 146 148 144 109 107 113 112 111 90 75 75 87  FPS FPA FDA FDT FA F P  S  I  H  M  N  CBGAA L41869 U26025 U39228 X56733 P26204 X94986 S35175 U33817 A48860P X78433 X89413 S52771P Q00326 X79195 L11454 LPH3HU A48969P Q08638 S45675  YRFSISWSRILPEGR—GEINMAGIEYYNNLIDALLQNGIQPFVTLFHFD YRFSISWSRIFPDGT—GKVNQEGVDYYNRLIDYMLQQGITPYANLYHYD YRFSISWSRVLPNGKVSGGVNEDGIKFYNNLINEILRNGLKPFVTIYHWD YRFSISWSRLLPNGTLSGGINKKGIEYYNNLTNELIRNGIEPLVTLFHWD YRFSISWPRVLPKGKLSGGVNREGINYYNNLINEVLANGMQPYVTLFHWD YRFSISWPRILPKGKLSGGINHEGIKYYNNLINELLANGIQPFVTLFHWD FRFLISWPRVIPSGTRREGINEQGIEFYNKVINEIINQGMEPFVTIFHWD FRMSISWSRVIPSGRRREGVNEEGIQFYNDVINEIISNGLEPFVTIFHWD YRFSISWPRILPKGTLAGGINEKGVEYYNKLIDLLLENGIEPYITIFHWD YRFSISWPRILPKGTKEGGINPDGIKYYRNLINLLLENGIEPYVTIFHWD YRFSISWPRILPKGTLDGGINHEGIQYYNDLLDCLIENGIKPYITLFHWD FRMSIAWPRIFPHGRKEKGVSQAGVQFYHDLIDELIKNGITPFVTVFHWD LRLSIAWPRIFPHGRMEKGNSKEGVQFYHDLIDELLKNDLTPLVTIFHWD YRFSFAWSRIIPKGKVSRGVNQGGLDYYHKLIDALLEKNITPFVTLFHWD YRFSFAWSRILPKGKRSRGINEDGINYYSGLIDGLIARNITPFVTLFHWD YRFSIAWSRLLPKGKRSRGVNPGAIKYYNGLIDGLVAKNMTPFVTLFHWD YRFSISWSRIFPTGR-NSSINSHGVDYYNRLINGLVASNIFPMVTLFHWD YRFSISWPRVLPQGT- -GEVNRAGLDYYHRLVDELLANGIEPFCTLYHWD YRFSISWPRILPEGT- -GRVNQKGLDFYNRIIDTLLEKGITPFVTIYHWD YRFSLAWPRIQPTGR- -GPALQKGLDFYRRLADELLAKGIQPVATLYHWD  14 7 161 163 153 150 160 165 154 196 198 194 15 9 15 7 163 162 161 139 12 3 12 3 135  CBGAA L41869 U26025 U39228 X56733 P26204 X94986 S35175 U33817 A48860P X78433 X89413 S52771P Q00326 X79195 L11454 LPH3HU A48969P Q08638 S45675  LPKALEDSYGGWLS PQIINDFEAYAEICFRAFGDRVKYWATVNEPNL LPLALHQQYLGWLS PKIVGAFADYAE FC FKV FGDRVKNWFT FNEPRV LPQALEDEYGGFLS-- -PNIVDHFRDYANLCFKKFGDRVKHWITLNEPYT VPQALEEEYGGVLS-- - PRIVYDFKAYAELCYKEFGDRVKHWTTLNEPYT VPQALEDEYRGFLG-- -RNIVDDFRDYAELCFKEFGDRVKHWITLNEPWG LPQVLEDEYGGFLN-- -SGVINDFRDYTDLCFKEFGDRVRYWSTLNEPWV TPQAIEDKYGGFLS-- -ANIVKDYREYADLLFERFGDRVKFWMTFNEPWS TPQALQDKYGGFLS-- -RDIVYDYLQYADLLFERFGDRVKPWMTFNEPSA TPQALVDAYGGFLDEED YKDYTDFAKVCFEKFGKTVKNWLTFNEPET VPQALEEKYGGFLDKSHKSIVEDYTYFAKVCFDNFGDKVKNWLTFNEPQT TPQALADEYKDFLD RRIVKDYTDYATVCFEHFGDKVKNWFTFNEPHS TPQDLEDEYGGFLSE RIVKDFREYADFVFQEYGGKVKHWITFNEPWV MPADLEDEYGGFLSE RVV P D FVE YAN FT FH E YG DKVKNWIT FNE PWV LPQTLQDEYEGFL DRQIIQDFKDYADLCFKEFGGKVKHWITINQLYT LPQSLQDEYEGFL DRTIIDDFKDYADLCFERFGDRVKHWITINQLFT LPQTLQDEYNGFL NKTIVDDFKDYADLCFELFGDRVKNWITINQLYT LPQALQDI-GGW-- -ENPALIDLFDSYADFCFQTFGDRVKFWMTFNEPMY LPQALQDQ-GGW-- -GSRITIDAFAEYAELMFKELGGKIKQWITFNEPWC LPFALQLK-GGW ANREIADWFAEYSRVLFENFGDRVKNWITLNEPWV LPQELENP-GGW PERPTAERFAEYAAIAADALGDRVKTWTTLNEPWC  194 208 210 200 197 207 212 201 24 3 24 3 2 41 20 6 204 210 20 9 20 3 18 5 169 169 181  CBGAA L41869 U2 6025 U39228 X56733 P26204 X94986 S35175 U33817 A48860P X784 33 X89413 S52771P Q00326 X79195 L11454 LPH3HU A48969P Q08638 S45675  FVPLGYTVGIFPPT-RCAAPHANPLCM-TGNCSSAEPYLAAHHVLLAHAS VAALGYDNGFHAPG-RCSK CP-AGGDSRTEPYIVTHNIILSHAA FSSSGYAYGVHAPG-RCSA-WQKLNCT-GGN-SATEPYLVTHHQLLAHAA ISNHGYTIGIHAPG-RCSS-WYDPTCL-GGD-SGTEPYLVTHNLLLAHAA VSMNAYAYGTFAPG-RCSD-WLKLNCT-GGD-SGREPYLAAHYQLLAHAA FSNSGYALGTNAPG-RCSA-SNVAK PGD-SGTGPYIVTHNQILAHAE LSGFAYDDGVFAPG-RCSS-WVNRQCR-AGD-SATEPYIVAHHLLLAHAA YVGFAHDDGVFAPG-RCSS-WVNRQCL-AGD-SATEPYIVAHNLLLSHAA FCSVSYGTGVLAPG-RCSP GVSCAVPTGNSLSEPYIVAHNLLRAHAE FTSFSYGTGVFAPG-RCSP GLDCAYPTGNSLVEPYTAGHNILLAHAE FCGLGYGTGLHAPGARCSA GMTCVIPEEDALRNPYIVGHNLLLAHAE FLHAGYDVGKKAPG-RCSSYVNA KCQDGRSGYEAYLVTHNLLISHAE FSRSAYDVGKKAPG-RCSPYIKDFGHLCQDGRSGFEAYWSHNLLVSHAE VPTRGYAIGTDAPG-RCSP-MVDTKHRCYGGNSSTEPYIVAHNQLLAHAT VPTRGYALGTDAPG-RCSQ-WVDK -RCYGGDSSTEPYIVAHNQLLAHAT VPTRGYALGTDAPG-RCSP-KIDV—RCPGGNSSTEPYIVAHNQLLAHAA LAWLGYGSGEFPPGVK DPGWAPYRIAHTVIKAHAR MAFLSNYLGVHAPGNK DLQLAIDVSHHLLVAHGR VAIVGHLYGVHAPGMR DIYVAFRAVHNLLRAHAR SAFLGYGSGVHAPGRT DPVAALRAAHHLNLGHGL  242 250 25 6 24 6 24 3 251 258 247 28 9 294 28 8 252 253 258 255 254 220 203 203 215  8-7 CBGAA L418 69 U2 6025 039228 X567 33 P2 6204 X94 98 6 S3517 5 U33817 A48860P X78 4 33 X8 9413 S527 71P Q0032 6 X79195. L114 54 LPH3H0 A4 8 969P Q08 638 S4 5 67 5  CBGAA L41869 U26025 U39228 X56733 P26204 X94986 S35175 033817 A48860P X78433 X89413 S52771P Q00326 X79195 L11454 LPH3HO A48969P Q08638 S45675  CBGAA L418 69 02 6025 U39228 X5 67 33 P2 6204 X94 98 6 S35175 033817 A4 88 60P X784 33 X89413 S52771P Q0032 6 X7 9195 L114 54 LPH3HU A4 8 969P Q08 638 S4 5675  AVEKYREKYQKIQGGSIGLVISAPWYEPLENSP-EERSAVDRILSFNLRW AVQRYREKYQPHQKGRIGILLDFVWYEPHSDTD-ADQAAAQRARDFHIGW AVKLYKDEYQASQNGLIGITLVSPWFEPASEAE-EDINAAFRSLDFIFGW AVKLYREKYQASQEGVIGITVVSHWFEPASESQ-KDINASVRALDFMYGW AARLYKTKYQASQNGIIGITLVSHWFEPASKEK-ADVDAAKRGLDFMLGW AVHVYKTKYQAYQKGKIGITLVSNWLMPLDDNSIPDIKAAERSLDFQFGL AVKIYRENYQETQNGKIGITLFTYWFEPLSNSTD-DMQASRTALDFMFGL AVHQYRKYYQGTQKGKIGITLFTFWYEPLSDSKV-DVQAAKTALDFMFGL TVDIYNKYHKG-ADGRIGLALNVFGRVPYTNTFL-DQQAQERSMDKCLGW AVDLYNKHYKR-DDTRIGLAFDVMGRVPYGTSFL-DKQAEERSWDINLGW TVDVYNKFYKG-DDGQIGMVLDVMAYEPYGNNFL-DQQAQERAIDFHIGW AVEAYRK-CEKCKGGKIGIAHSPAWFEAHDLADSQDGASIDRALDFILGW AVDAFRK-CEKCKGDKIGIAHSPAWFEPEDVEGGQR—TVDRVLDFIMGW VVDLYRTKYKF-QKGKIGPVMITRWFLPFDESDPASIEAAERMNQFFHGW VVDLYRTRYKY-QGGKIGPVMITRWFLPYDDTL-ESKQATWRAKEFFLGW AVDVYRTKYKDDQKGMIGPVMITRWFLPFDHSQ-ESKDATERAKIFFHGW VYHTYDEKYRQEQKGVISLSLSTHWAEPKSPGVPRDVEAADRMLQFSLGW AVTLFRE LGISGEIGIAPNTSWAVPYRRTKEDMEACLRVNGWSG-DW AVKVFRE TVKDGKIGIVFNNGYFEPASEKEEDIRAVRFMHQFNNYPL AVQALRDR--LPADAQCSVTLNIHHVRPLTDSEADADAVRRIDALAN-RV  291 299 305 295 292 301 307 296 337 342 336 301 300 307 303 303 270 249 250 262  FLDPIVF-GDYPQEMR ERLGSRLPSISSELSAKLRGSFDY FLDPITN-GRYPSSML KIVGNRLPGFSADESRMVKGSIDY FMDPLTN-GNYPHLMR SIVGERLPNFTEEQSKLLKGSFDF FMDPLTR-GDYPQSMR SLVKERLPNFTEEQSKSLIGSYDY FMHPLTK-GRYPESMR YLVRKRLPKFSTEESKELTGSFDF FMEQLTT-GDYSKSMR RIVKNRLPKFSKFESSLVNGSFDF WMDPITY-GRYPRTVQ YLVGNRLLNFTEEVSHLLRGSYDF WMDPMTY-GRYPRTMV DLAGDKLIGFTDEESQLLRGSYDF FLEPW R-GDYPFSMR VSARDRVPYFKEKEQEKLVGSYDM FLEPW R-GDYPFSMR SLARERLPFFKDEQKEKLAGSYNM FLEPMVR-GDYPFSMR SLVGDRLPFFTKSEQEKLVSS YDF HLDTTTF-GDYPQIMK DIVGHRLPKFTTEQKAKLKASTDF HLDPTTY-GDYPQSMK DAVGARLPKFTKAQKAKLKGSADF YMEPLTK-GRYPDIMR QIVGSRLPNFTEEEAELVAGSYDF FMEPLTK-GKYPYIMR KLVGNRLPKFNSTEARLLKGS YDF FMGPLTE-GKYPDIMR EYVGDRLPEFSETEAALVKGSYDF FAHPIFRNGDYPDTMKWKVGNRSELQHLATSRLPSFTEEEKRFIRATADV YLDPIYF-GEYPKFM LDWYENLGY KPPIVDGDMELIHQPIDF FLNPIYR-GDYPELV LE-FAR-EY LPENYKDDMSEIQEKIDF FTGPMLQ-GAYPEDL VKDTAGLTD WSFVRDGDLRLAHQKLDF  330 338 34 4 334 331 34 0 34 6 335 37 6 381 37 5 34 0 339 34 6 34 2 342 320 2 90 28 9 303  MGINHYTTLYATSTPPLSPDHTQ—YLYPDSRVYLTGERHGV-SIGERTG VGINQYTSYYMKDPGAWNQTPVS--YQ-DDWHVGFVYERNGV-PIGPRAN IGLNYYTTRYASNAPKITSVHA--SYITDPQVNAT-AELKGV-PIGPMAA IGVNYYSARYASAYPEDYSIPTPPSYLTDAYVNVT-TELNGV-PIGPQAA LGLNYYSSYYAAKAPRI--PNARPAIQTDSLINAT-FEHNGK-PLGPMAA IGINYYSSSYISNAPSH--GNAKPSYSTNPMTNIS-FEKHGI-PLGPRAA IGLQYYTSYYAKPNAPYDPNHIR—YLTDNRVTETPYDYNGN-LIGPQAY VGLQYYTAYYAEPIPPVDPKFRR—YKTDSGVNATPYDLNGN-LIGPQAY IGINYYTSTFSKHID-LSPNNSPVLNTDDAYASQETKGPDGN-AIGPPTG LGLNYYTSRFSKNID-ISPNYSPVLNTDDAYASQEVNGPDGK-PIGPPMG VGINYYTSRFAKHID-ISPEFIPKINTDDVYSNPEVNDSNGI-PIGPDVG VGLNYYTSVFSNHLEK--PDPSKPRWMQDSLITWESKNAQ-NYAIGSKPL VGINYYSSFYAKASEK—PDYRQPSWATDSLVEFEPKTVDGSVKIGSQPS LGLNYYVTQYAQPKPNPYPSETHTA-MMDAGVKLTYDNSRGEFL-GPLFV LGLNYYVTQYAHALDPSPPEKL-TA-MTDSLANLTSLDANGQPP-GPPFLGLNYYVTQYAQNNQTIVPSDVHTA-LMDSRTTLTSKNATGHAP-GPPFFCLNTY YSRIVQHKTPRLNPPSYEDD—QEMAEEEDPSWPSTAMNRA IGINYYTSSMNRYNPGEAGGMLSSEAISMGAP KTD VGLNYYSGHLVKFDPDAPAKV SFVERDLP KTA LGVNYYSPTLVSEADGSGTHNSDGHGRSAHSPWPGADRVAFHQPPGETTA  377 384 390 382 377 386 393 382 424 429 423 387 387 394 388 389 365 325 321 353  CBGAA L418 69 U2 6025 U39228 X56733 P2 6204 X94 986 S35175 U33817 A4 8860P X784 33 X8 9413 S52 7 71P Q00326 X7 9195 L11454 , LPH3HU A4 8 969P Q08 638 S4 5675  MDGLFWPH GIQKIVEYVKEFYDNPTII- IAENGYPESE—ESSST GNVS SDWLYIVPW GMNKAVTYVKERYGNPTMI LSENGMDQP SGWLYVYP KGIHDLVLYTKEKYNDPLIY ITENGVDEFN—DPKLS SDWLYVYP KGLYDLVLYTKNKYNDPIMY ITENGMDEFN—NPKIS SSWLCIYP QGIRKLLLYVKNHYNNPVIY ITENGRNSST—INTVSIWIYVYPYMFIQEDFEIFCYILKINITILQFSITENGMNEFN—DATLP SDWFYIFP ESIRHLLNYTKDTYNDPVIY- ITENGVDNQN—NETEP SSWFYIFP KGIRHFLNYTKDTYNDPVIY VTENGVDNYN--NESQP NAWINMYP KGLHDILMTMKNKYGNPPMY ITENGMGDIDKGDLPKP NPWIYMYP EGLKDLLMIMKNKYGNPPIY ITENGIGDVDTKETPLP MYFIYSYP KGLKNILLRMKEKYGNPPIY ITENGTADMDGWGN P-P TAALNVYS RGFRSLLKYIKDKYANPEIM IMENGYGEELGASDSV-A TAKMAVYA AGLRKLVKYIKDRYGNPEII ITENGYGEDLGEKDTDHS FSTPSSENR EDKVNGNSYYYPKGIYYVMDYFKTKYGDPLIY VTENG FSTSGGPIP SKGSYYHPRGMLNVMEHFKTKYGDPLIY VTENG FSTPGDE-D NAASYYYPKGIYYVMDYFKTTYGDPLIY VTENG APW GTRRLLNWIKEEYGDI PI Y I T E N G V G L T N P N T CYNDGLS IGWE IYAEGLYDLLRYTADKYGNPTLY ITENGA AFDDVVS MGWE IVPEGIYWILKKVKEEYNPPEVY ITENGA AFHDYAD MGWA VDPSGLYELLRRLSSDFPALPLV ITENGA  420 4 25 4 33 425 419 4 34 4 36 425 4 69 474 4 67 4 32 4 33 440 430 4 30  CBGAA L41869 U26025 U39228 X56733 P26204 X94986 S35175 U33817 A48860P X78433 X89413 S52771P Q00326 X79195 L11454 LPH3HU A48969P Q08638 S45675  LQENLNDVRRIRFHGDCLSYLSAAIKN-GSDVRGYFVWSLLDNFEWAFGY IADGVHDTVRIRYYRDYITELKKAIDN-GARVAGYFAWSLLDNFEWRLGY MEEALKDTNRIDFYYRHLCYLQAAIKK-GSKVKGYFAWSFLDNFEWDAGY LEQALNDSNRIDYCYRHLCYLQEAIIE-GANVQGYFAWSLLDNFEWSEGY TSRIPF VEEALLNTYRIDYYYRHLYYIRSAIRA-GSNVKGFYAWSFLDCNEWFAGF IQDAVKDGFRIEYHRKHMWNALGSLKFYHVNLKGYFAWSYLDNFEWNIGY IEEALQDDFRISYYKKHMWNALGSLKNYGVKLKGYFAWSYLDNFEWNIGY V—ALEDHTRLDYIQRHLSVLKQSIDLGAD-VRGYFAWSLLDNFEWSSGY MEDALNDYKRLDYIQRHIATLKESIDLGSN-VQGYFAWSLLDNFEWFAGF MTDPLDDPLRIEYLQQHMTAIKEAIDLGRRTLRGHFTWSLIDNFEWSLGY AV-GTADHNRKYYLQRHLLSMQEAVCIDKVNVTGYFVWSLLDNFEWQDGY SV-ALNDHNRKYYHQRHLLSLHQAICEDKVNVTSYFVWSLMDNFEWLDGY -EQAIADYKRIDYLCSHLCFLRKVIKEKGVNVRGYFAWALGDNYEFCKGF FTEAFHDYNRIDYLCSHLCFLRKAIKEKRVNVKGYFVWSLGDNYEFCNGY FEKATADYKRIDYLCSHLCFLSKVIKEKNVNVKGYFAWSLGDNYEFCNGF EDTDRIFYHKTYINEALKAYRLDGIDLRGYVAWSLMDNFEWLNGY LDGRIHDQRRIDYLAMHLIQASRAIED-GINLKGYMEWSLMDNFEWAEGY EDGRVHDQNRIDYLKAHIGQAWKAIQE-GVPLKGYFVWSLLDNFEWAEGY PEGNVNDPERIAYVRDHLAAVHRAIKD-GSDVRGYFLWSLLDNFEWAHGY  4 69 474 4 82 47 4 425 4 83 4 86 4 75 516 523 517 4 80 4 81 489 4 80 4 80 4 45 414 410 4 42  CBGAA L41869 U26025 U39228 X56733 P26204 X94986 S35175 U33817 A48860P X78433 X89413 S52771P Q0032 6 X79195 L11454 LPH3HU A48969P Q08 638 S45675  TIRFGLYHVDFIS-DQKRYPKLSAQWFRQFLQHDDQGS TARFGIVYVDF-N-TLKRYPKDSALWFKNMLSEKKRS TVRFGINYVDYND-NLKRHSKLSTYWFTSFLKKYERSTKEIQMFVESKLE TVRFGINYVDYDN-GLKRHSKLSTHWFKNFLKRSSISKEKIRRCGNNNAR  506 509 531 523 425 493 534 524 553 558 554 523 514 532 521 524 480 449 44 5 478  TVRFGLNFVD TARFGLYYVDYNN-NLTRIPKDSAYWFKAFLN-PENITKTTRTVSWDSRK TSRFGLYYVDYKN-NLTRYPKKSAHWFTKFLNISVNANNIYELTSKDSRK TERFGIVYVDREN-GCERTMKRSARWLQEF NGAAKKVE TERYGIVYVDRNN-NCTRYMKESAKWLKQF NAAKKP— LSRFGIVYIDRND-GCKRIMKKSAKWLKEF NGATKKLN KNRFGLYYVDFKN-NLTRYEKESGKYYKDFLSQGVRPSALKKDE TARFGLYYIDFQN-NLTRMEKESATCS LNSSNRATVRFGLSYVNWEDL-DDRNLKESGKWYQRFIN GTVKNAVKQDFL TVRFGLSYVDFNNVTADRDLKASGLWYQSFLR DTTKNQDIL TVRFGLSYVDFANITGDRDLKASGKWFQKFIN VTDEDSTNQDLL TVKFGLYHVDFNNTNRPRTARASARYYTEVITNNG GMRFGLVHVDYDTL—VRTPKDSFYWYKGVISRGWL-D SKRFGIVYVDYSTQ—KRIVKDSGYWYSNVVKNNGL-E .— SKRFGAVYVDYPTG—TRIPKASARWYAEVARTGVLPT  4 00  3 65 361 393  CBGAA L41869 U26025 U39228 X56733 P26204 X94986 S35175 U33817 A48860P X78433 X89413 • S52771P Q00326 X79195 L11454 LPH3HU A48969P Q08638 S45675  IRSSSS—I HQKFESQMMNKVQSSLAVVV ARKFVYR . 1 AGKF YIM VGKF YVM NNKI LTPAGQLN SKKI LTPA NKILGASSCCSGVTHGGGTA L RSSLSSQS-QKKRFADA RSSLPFKNGDRKSLT RSSVSSKNRDRKSLADA M L D A  513 509 551 531 425 493 541 531 565 566 574 524 514 548 536 541 481 450 446 479  Table 3.1. Deduced amino acid composition of coniferin p-glucosidase  AA  Nb.  Leu Ser Gly Ala Glu He Tyr Arg Asp Pro Phe Val Asn Thr Gin His Lys Met Trp Cys  47 47 38 33 32 31 31 30 30 28 28 27 21 20 17 14 13 11 10 5  % 9.1 9 .1 7.4 6.4 6.2 6.0 6.0 5. 8 5. 8 5.4 5.4 5.2 4.0 3.8 3.3 2.7 2. 5 2 .1 1.9 0. 9  91  C B G contains several sequence elements that are highly conserved among the P-glucosidases compared in Fig.3.11, and among members of family 1 glycosyl hydrolases. These include the N-terminal signature sequence of family 1 glycosyl hydrolases at residues 34 - 48, the sequence -ENG- at residues 24 - 26 within the Cterminal signature of family 1 glycosyl hydrolases, and the sequence -NEP- at residues 189 - 192. The conserved sequence motifs NEP and E N G are thought to be important for enzyme activity (Baird et al. 1990, Withers et al. 1990). Site-directed mutagenesis of the Glu residue in the NEP motif in endo-p-l-4-glucanase of Bacillus sp. resulted in a dramatic loss of enzyme activity (Baird et al. 1990). The authors suggest that this region could be involved in binding of the pyranose ring during catalysis. The N E P motif of both Bacillus endo-P-l-4-glucanase and C B G is flanked by hydrophobic amino acids; next to the signal peptide it is the most hydrophobic region of the C B G enzyme (Fig.3.10). Evidence from inhibitor and site-directed mutagenesis studies strongly suggest that the Glu within the conserved E N G motif is the active site nucleophile (Withers et al. 1990, Trimbur et al. 1992). C B G also contains a conserved aspartate residue (Asp427) which is analogous to Asp374 of Agrobacterium p-glucosidase (cellobiase). This residue has been suggested to play the role of acid-base catalyst during hydrolysis of the glycosidic linkage (Trimbur et al. 1992). "  92  Expression of C B G in E. coli: The pET series of expression vectors used for expressing C B G offers Ncol and Ndel A T G cloning sites to avoid expression of vector-derived sequences at the N-terminus. However, the presence of internal Ncol and Ndel sites within the C B G coding sequence dictated the use of adjacent Nhel as the 5' cloning site. This adds 3 non C B G amino acids (Met, Ala, Ser) to the N-terminus of the expressed protein. To avoid the expression of the His tag of the vector at the 3' end, the native stop codon of C B G was included. Thus, the expressed protein would be identical in sequence to the mature C B G except for 3 additional amino acids at the N-terminus. Soluble proteins and insoluble proteins (inclusion bodies) prepared from induced and noninduced bacterial cells were assayed for coniferin hydrolysis activity, but only the soluble protein fraction of induced cells displayed this activity. The activity in this fraction could be increased up to 2-fold by increasing the IPTG concentration from 0.4 - 1.0 m M , and by reducing the growing temperature from 37°C to 30°C. The protein profiles of induced and uninduced cells were complex and did not reveal a clear-cut protein band as the induced protein.  However, activity  staining of nondenaturing gels using the chromogenic coniferin analogue V R A - G revealed a P-glucosidase active protein band in induced cell extracts. was purified using preparative  Q-Sepharose  anion exchange  This protein  chromatography  followed by an analytical anion exchange step ( Q M A Memsep - Millipore). Throughout purification, coniferin was used as the substrate for monitoring P-  93  glucosidase activity. The purified enzyme from cultures induced with 0.4 m M IPTG at 37°C migrated as a single band (Fig.3.13, lanel) on nondenaturing gels as opposed to the doublet (Fig.3.13 lane2) seen in enzyme purified from cultures induced with 1 m M IPTG at 30°C. Both protein bands in the doublet showed p-glucosidase activity, as assayed by hydrolysis of V R A - G (Fig.3.13B). As shown in Table 3.2, the C B G expressed in E. coli and the enzyme purified from the pine xylem showed very similar substrate specificities.  94  Fig.3.12. Expression of C B G in E.coli and Native-PAGE of soluble protein fraction from E.coli. lane 1 - uninduced cells, lane 2 - following induction of C B G expression with 0.4 m M IPTG at 37°C , lane 3 - coniferin active protein fraction (following a single step Q Sepharose purification) of proteins from lane 2. Arrow indicates C B G protein band.  Fig. 3.13. Native P A G E (A) and V R A - G activity staining (B) of purified coniferin (3glucosidase (using Q Sepharose and QMA-memsep). lane 1 - from 37°C, 0.4 m M IPTG induced cells, lane 2 - from 30°C, I m M IPTG induced cells.  12  96  Table 3.2. Substrate specificity of coniferin P-glucosidase purified from pine xylem and E.coli expressed CBG-cDNA. 100% activity represents 14pKat for coniferin pglucosidase and 22pKat for C B G - c D N A expressed in E.coli. Relative activity substrate  coniferin syringin 4-methyl umbelliferyl-Pglucoside 2-nitrophenyl-P-glucoside 4-nitrophenyl-P-glucoside  coniferin P-glucosidase (coniferin = 100)  C B G - c D N A expressed in E.coli (coniferin =100)  100 51 18  100 65 20  51 30  50 35  97  Discussion: Out of the array of methods potentially available to isolate a c D N A from a library, PCR amplification with degenerate oligonucleotide primer pools based on the N-terminal amino acid sequence and vector sequence-derived  primer proved  successful with C B G . Screening the library with anti-peptide antibodies following induction of fusion protein expression produced undesirably high background. The screening using the chromogenic substrate V R A - G was not pursued exhaustively due to a limited supply of the substrate. The PCR-based approach using gene specificprimers based on the N-terminal amino acid sequence was attractive primarily because of its sensitivity.  Mixed P C R primer pools consisting of different  oligonucleotides with sequences degenerate at ambiguous codon positions provide a powerful tool in cDNA cloning (Lee et al. 1988, Girgis et al. 1988). It has also been shown that primers containing deoxyinosine at degenerate codon positions could be used successfully in PCR. In the design of primers for amplifying C B G - c D N A , the size of the primer pool was reduced by including deoxyinosine at codon positions with complete degeneracy.  The 1909 nucleotide c D N A sequence obtained in this  fashion was confirmed to be that of coniferin P-glucosidase on the basis of its high homology to other plant P-glucosidases, the sequence identity with the N-terminal amino acid sequence of the purified enzyme, and the coniferin hydrolysis activity and substrate specificity displayed by the corresponding protein when expressed in E. coli.  98  The deduced 513 amino acid nascent protein has a calculated molecular mass of 58.3 kD, while the mature protein without the 23 amino acid signal peptide would have a molecular mass of 55.3 kD. The native molecular weight of the C B G enzyme purified from xylem was estimated from gel filtration chromatography to be 60 kD (Chapter 2). The greater estimated molecular weight of the enzyme could be from the carbohydrate residues at the two predicted N-glycosylation sites (Asn 223 and Asn 447) of the protein. The origin of 28kD and 24kD protein bands observed in SDSP A G E gels of purified protein (Chapter 2) can most obviously be attributed to protein degradation at a labile site during SDS-PAGE. Thus, the mature protein should be a 55.3 kD monomeric enzyme. The predicted isoelectric point of pH 4.9 was close to the value determined by chromatofocussing (pH 4.5) of the purified enzyme (Chapter 2). The 23 amino acid N-terminal sequence of C B G shares characteristics with eukaryotic secretory signal peptides.  According to the well-accepted signal  hypothesis (Blobel and Dobberstein 1975), insertion of the protein into the endoplasmic reticulum is achieved through the interaction of the hydrophobic signal peptide with a signal-recognition particle (SRP) and SRP-receptor protein in the E R membrane.  The cotranslational cleavage of the signal peptide at a cleavage site  recognized by the signal peptidase releases the protein into the E R lumen.  These  proteins can then be secreted out of the cell via the constitutive or unregulated pathway (Burgess and Kelly 1987) of the golgi complex. To be retained in the E R or golgi, or to be directed to other cell compartments such as the vacuoles, the protein  99  should contain additional sorting signals which could be located in any region of the sequence or in topogenic information (Crispeels 1991).  Therefore, from the  information at hand , C B G could either be targeted to a vacuolar compartment within the cell or secreted into the cell wall, the latter being analogous to known secretory cell wall-associated proteins such as peroxidases (Lagrimini et al. 1987), cyanogenic P-glucosidases (Hughes et al. 1992) and basic pathogenesis related (PR) proteins (Swoboda et al. 1995). The sequence of C B G displays high homology to enzymes belonging to glycosyl hydrolase family 1.  This family of enzymes is distributed in diverse  organisms and despite their sequence similarities they perform diverse functions. Functions of some of the enzymes showing highest homology to C B G include cyanogenesis (Hughes et al. 1992, Oxtoby et al. 1991), hydrolysis of glucosinolates (Xue et al. 1992, Falk et al. 1992), hydrolysis of short chain oligosaccharides (Leah et al. 1995), and possible hydrolysis of phytohormone conjugates (Brzobohaty et al. 1993). Due to the high sequence similarity, these enzymes are likely to have similar folding characteristics and similar catalytic sites. Despite these similarities, however, their substrate specificities can differ widely as shown by 1-3-P-glucanase and (1-3,14)-p-glucanase, which have very similar 3D structures (Varghese et al. 1994). As suggested by Varghese et al. (1994), amino acid substitutions adjacent to the catalytic sites could be involved in specific substrate recognition and affinities. Interestingly, the Agrobacterium P-glucosidase that was reported to be active against coniferin (Castle et al. 1992) does not show any homology to C B G and is in fact, more  100  homologous to family 3 glycosyl hydrolases (Henrissat 1991). Similarly, plant and bacterial (l-3,l-4)-P-glucanases have identical substrate preferences but proved to have vastly different three-dimensional structures. preferences  Thus, the identical substrate  displayed by these very different proteins might have arisen by  convergent evolution. Unfortunately, there is relatively little known of the detailed substrate preferences and kinetic data for the Agrobacterium coniferin p-glucosidase enzyme. The hydrolytic mechanism of p-glucosidases is considered to be general acid catalysis (Sinnott 1990) with Glu and Asp residues in conserved motifs serving as active site nucleophile and acid catalyst. Glu358 in the E N G motif of Agrobacterium cellobiase (analogous to Glu 408 of CBG) has been identified as the active site catalytic nucleophile by inhibitor binding and site-directed mutagenesis (Trimbur et al. 1992, Withers et al. 1990).  studies  The conserved Asp374 residue of  Agrobacterium cellobiase, which is at a similar spatial distance from the E N G motif of other family 1 members and C B G (Asp 427), is suggested to be the general acid catalyst that protonates the leaving group. The expression of active C B G enzyme in E. coli, and the demonstration that it exhibits identical substrate preferences to the native protein, confirm that the isolated c D N A corresponds to the true coniferin p-glucosidase.  The level of protein  expression obtained in this system was lower than expected, which could be due to accumulation of expressed protein as insoluble inactive inclusion bodies or due to the presence of codons in the C B G sequence that restrict protein synthesis rates in E.coli.  101  The expression protocol could therefore potentially be optimized further.  Codon  usage tables (Grisboskov et al. 1984) show that the codon A G A for Arg is rarely used in E. coli proteins (0 number of occurrences per 1000 codons). The presence of 10 A G A codons in the C B G sequence (Table 3.3), could thus lower the efficiency of expression of C B G in this host. Site-directed mutagenesis could be used to reduce this problem. By reducing the induction temperature it was possible to increase the yield of the soluble active form of C B G to some degree, but under these conditions C B G protein appeared as a doublet of bands on native gels. This could be due to partial degradation, alternate forms of folding, or the synthesis of a truncated protein at the 5' end where C B G has a prokaryotic ribosome binding sequence A G G A G G (Shine-Dalgarno sequence). The latter would result in the synthesis of a polypeptide that is truncated at the N-terminus, as opposed to the full-length polypeptide initiated by ribosome binding to the standard ribosome binding site in the vector.  The  expression of active C B G in E. coli with similar substrate preferences as the enzyme purified from xylem indicates that glycosylation of C B G (Asn 223 and Asn 447) is not essential for display of its catalytic properties and stability.  102  Table 3.3. Deduced codon usage table for Pinus contorta coniferin P-glucosidase c D N A sequence TTT TTC TTA TTG  Phe Phe Leu Leu  14 17 9 15  2 2 1 2  1% 6% 4% 3%  TCT TCC TCA TCG  Ser Ser Ser Ser  7 9 9 14  1 1 1 2  0% 4% 4% 1%  TAT T y r l O TAC T y r 3 TAA 9 TAG 8  1 0 1 1  5% 4% 4% 2%  TGT Cys 13 TGC Cys 7 TGA 10 TGG T r p 17  2.0% 1.0% 1.5% 2.6%  CTT CTC CTA CTG  Leu Leu Leu Leu  4 11 9 15  0 1 1 2  6% 7% 4% 3%  CCT CCC CCA CCG  Pro Pro Pro Pro  17 5 11 3  2 0 1 0  6% 7% 7% 4%  CAT CAC CAA CAG  His His Gin Gin  9 ' 9 8 24  1 1 1 3  4% 4% 2% 7%  CGT CGC CGA CGG  Arg Arg Arg Arg  1.2% 0.6% 1.0% 0.3%  ATT ATC ATA ATG  He He He met  19 29 12 17  2 4 1 2  9% 5% 8% 6%  ACT ACC ACA ACG  Thr Thr Thr Thr  8 8 10 16  1 1 1 2  2% 2% 5% 5%  AAT AAC AAA AAG  Asn Asn Lys Lys  15 8 17 16  2 1 2 2  3% 2% 6% 5%  AGT AGC AGA AGG  Ser 4 Ser 8 A r g 10 Arg 6  0.6% 1.2% 1.5% 0.9%  GTT GTC GTA GTG  Val Val Val Val  6 3 3 8  0 0 0 1  9% 4% 4% 2%  GCT GCC GCA GCG  Ala Ala Ala Ala  6 2 IS 1  0 0 2 0  9% 3% 5% 1%  GAT GAC GAA GAG  Asp Asp Glu Glu  13 3 11 12  2 0 1 1  0% 4% 7% 8%  GGT GGC GGA GGG  Gly Gly Gly Gly  1.7% 0.3% 1.5% 1.5%  8 4 7 2  11 2 10 10  103  CHAPTER 4 In situ localization of lignin biosynthetic enzymes  INTRODUCTION: The work described in the previous chapters has shown conclusively, by biochemical and molecular means, the existence and expression of a coniferinspecific P-glucosidase in Pinus contorta xylem. For a more complete understanding of the metabolic processes involved in these ultimate stages of lignin biosynthesis, determination of the location of the relevant enzymes at the histological and cellular level would be invaluable.  The work described in this chapter examined the  spatiotemporal expression pattern of p-glucosidases and peroxidases within actively growing woody stems. Chromogenic substrates such as indican (indoxyl P-D-glucoside) have been used previously for histochemical localization of P-glucosidase activity (Freudenberg 1964). However, the use of synthetic substrates with no structural resemblance to the native substrate poses the risk of detecting glucosidases not involved in the pathway of interest, and/or not detecting the specific P-glucosidase of interest. V R A - G , the chromogenic analogue of coniferin, was shown to be efficiently hydrolyzed by coniferin p-glucosidase (chapter 2). Hence, I decided to use it for in situ pglucosidase activity detection, even though it has the drawback of producing a soluble and diffusible aglycone upon hydrolysis.  Antibodies against the coniferin P-  glucosidase were not available at the time of this study.  104  The final oxidative polymerization of monolignol into lignin is thought to be mediated by peroxidases (Nakamura 1967, Harkin and Obst 1973, Goldberg et al. 1985) and laccases (Bao et al. 1993). Peroxidases from lignifying cell walls have been shown to have a strong affinity for the synthetic substrate, syringaldazine, (Harkin and Obst 1973). Because of this, and its structural resemblance to lignin precursors, syringaldazine has been promoted as a specific substrate for peroxidases involved in lignification (Goldberg et al. 1985). Since other peroxidase substrates such as diaminobenzidine and guaiacol have been shown not to generate lignificationspecific pattern of peroxidase activity in situ in both my (Chapter 1) and in other investigations (Goldberg et al. 1985), syringaldazine was the substrate of choice for localizing peroxidase activity in the present study. These localization studies were also extended to poplar in order to explore the presence and involvement of P-glucosidases in lignification of angiosperm xylem. To this end, an antibody against poplar bispecific catechol O-methyl transferase (OMT), an enzyme involved in earlier steps of phenylpropanoid metabolism, was also used for parallel immunolocalization studies.  MATERIALS A N D METHODS Plant material: Four-to five-year-old Pinus contorta var. latifolia Engelm. trees from the U.B.C. nursery (south campus, U.B.C.)  were sampled at the beginning of the  105  growing season (late May to early June). One-to two-year old Populus trichocarpa plants from the greenhouse or the growth chamber were sampled during active growth phase.  Detection of lignin: Lignin in the stem sections was detected histochemically according to the method  developed  for  Arabidopsis thaliana using  Basic  Fuchsin-induced  fluorescence or the Maule reaction.(Chapter 1).  In situ enzyme activity staining: To detect p-glucosidase activity in situ, transverse "free hand" sections from a two-year old Pinus contorta stem, or a one-year old Populus trichocarpa stem, were placed in cold degassed 20mM MES, pH 5.5, for 10 min. After incubation in an airtight chamber (to reduce oxidation of phenolics) with the chromogenic substrate V R A - G (2mM) in the same buffer for 30 min without agitation, the sections were transferred onto microscope slides and basified with 0.25M C A P S , pH 9.  The  instantaneous stain response was immediately photographed using a Zeiss Axiophot photomicroscope and Kodak Ektachrome 160 A S A tungsten film. As a control, the non-chromogenic substrate coniferin was used to stain neighboring sections. Peroxidases were detected in situ by equilibrating the tissue sections in citratephosphate buffer, pH 5, for 10 min. followed by the addition of 0.05% alcoholic syringaldazine with, or without, 0.03% H 0 . The color reaction was photographed 2  2  106  as described above.  Immunolocalization: One-year old P.trichocarpa stem sections were fixed  in 2% para-  formaldehyde in p H 7 phosphate buffer for 4 hours. The sections were washed 3-4 times with phosphate buffer, blocked with 2%BSA in PBS for 2 hours and incubated in the 1:400 dilution of anti-poplar O-methyl transferase antibody (provided by Prof. Wilbur H . Campbell, Department of Biological Sciences, Michigan Technology University, Michigan) in blocking solution for 1 hour. Following washing (three times with PBS) to remove excess antibody, the primary immune complexes were visualized by use of FITC-labelled goat anti-rabbit secondary antibody (Sigma F9887) at 1:200 dilution for 1 hour. The sections were washed three times in PBS and mounted in 50% glycerol in PBS for microscopic examination. A MRC-500 confocal laser scanning microscope was used for imaging, as described in chapter 1 (488 nm for FITC). Since pre-immune serum specific for the anti-OMT antibody was not at hand, serum from another non-immunized rabbit. was used for control experiments.  107  Results: The p-glucosidase activity in rapidly growing P. contorta stem tissue is clearly concentrated in the area of differentiating xylem (Fig. 4.1), as indicated by the orange colour generated by the aglycone of the chromogenic substrate, V R A - G . The orange colour is most intense in the region of xylem tracheid maturation, and fades out centripetally as the xylem achieved maturation.  The fully mature xylem,  including both current and previous years' tissue, was devoid of staining. No Pglucosidase activity was detected in the cambial derivatives during their enlargement into xylem cells (i.e. xylem cells close to the cambium), or in the phloem or the cortex. Although the orange reaction product appears to be more prominent on the cell walls of the differentiating xylem, it is not possible to specify the exact cellular or intracellular location of P-glucosidase activity with this technique because of the diffusion of the aglycone product.  Unlike V R A - G , the oxidation products of  syringaldazine are insoluble and could be used as an indicator of the cellular location of peroxidases.  The in situ detection of peroxidase activity using syringaldazine  revealed a tissue distribution pattern similar to that of P-glucosidase (Fig. 4.2) in the differentiating xylem cell walls. The staining of sections with syringaldazine in the absence of H 2 O 2 substrate did not yield a significant colour reaction.  The Basic  Fuchsin-induced lignin fluorescence of the same stem sections (Fig. 4.3) used for enzyme localization demonstrated that both P-glucosidase activity and peroxidase activity are restricted to the most recently lignified/lignifying differentiating xylem.  region of the  108  Fig. 4.1.- 4.3. In situ enzyme activity localization and lignin staining of Pinus contorta stem sections at the beginning of the growing season. Fig. 4.1. Localization of p-glucosidase activity using chromogenic substrate V R A - G (orange colour); Fig. 4.2. Localization of peroxidase activity with syringaldazine/H 0 (purple colour); 2  2  Fig. 4.3. Basic Fuchsin-induced red fluorescence of lignified tissues, xy, xylem; *, cambial zone; ph, phloem; co, cortex.  i<yf  110  The primary vascular bundle and secondary vascular development in young poplar stem is typical of perennial dicot angiosperms.  Initially several primary  vascular bundles are formed, which later became linked by development of vascular cambium/intercalary vascular cambium to form a complete vascular cylinder. Fig. 4.4-4.6 show the histochemical responses in Maule reaction stained stem sections distributed from the apex to the base of an actively growing poplar stem. The brown colour of proto- and meta-xylem vessel elements (primary xylem) in Fig. 4.4 indicate that the lignin in these cell walls is predominantly composed of guaiacyl monomers, where as the pink colour of the secondary xylem and phloem fibers indicates the presence of syringyl-type lignin.  This pattern is very similar to that observed in  Arabidopsis stem sections (Chapter 1). Sections from actively growing poplar stem, when stained for p-glucosidase and peroxidase activity, exhibited a tissue distribution pattern similar to that observed in Pinus (Fig. 4.7 and 4.8). Fig. 4.7A and 4.8A show the pattern obtained in sections closer to the stem apex where differentiation of xylem and phloem are at an earlier stage than in the sections shown in Fig. 4.7B and 4.8B. It is noteworthy that the lignifying phloem fibers in 4.7A and 4.8A also show strong Pglucosidase and peroxidase activity. Phloem fibers in stem sections from the lower part of the stem (4.7B and 4.8B), which are at a later stage of differentiation, show diminished but detectable enzyme activity. Immunolocalization of O M T in poplar stems (Fig. 4.9A to 4.9C) reveals the same spatial pattern during development of the stem. The amount of the enzyme present, as indicated by the intensity of green fluorescence, diminishes as the  Ill  lignification of phloem fibers approaches completion (Fig. 4.9B and 4.9C as opposed to 4.9A). The cytoplasm of most actively lignifying xylem vessels and fibers (closest to cambium) shows intense fluorescence. Fig. 4.9C, which shows a section that has been stained for both lignin (red fluorescence) and OMT (green fluorescence), confirms that O M T is only detectable in the cytoplasm of lignifying cells, i.e. xylem cells located closer to the cambium which do not stain for lignin are also devoid of the O M T fluorescence signal. Occasionally, small local regions of poplar stems show a brown coloration in the pith cell. These cells also show autofluorescence and stain a pink/violet following the Maule reaction (Fig. 4.5). These same pith cells show a positive fluorescence signal in the cytoplasm when probed for O M T (Fig. 4.9A).  112  Fig. 4.4-4.5. Three Maule reaction stained actively growing polar stem sections from apex to the base of the stem, arrow head, phloem fibers; *, lignified pith.  Fig. 4.7A-4.7B. Localization of P-glucosidase activity in actively growing poplar stems using chromogenic substrate V R A - G (orange colour). Section in 4.7A is closer to the apex of the stem than 4.7B. arrow head, phloem fibers.  Fig. 4.8A-4.8B. Localization of peroxidase activity with syringaldazine/H 02 (purple 2  colour), in poplar stem sections similar to 4.7A and 4.7B.  Fig.4.9A-4.9B. Immunolocalization of OMT in poplar stems similar to Fig.4.7 and Fig. 4.8.  Fig. 4.9C. Co-localization of O M T (green fluorescence) and lignin (Basic Fuschininduced red fluorescence) in a poplar stem similar to 4.9B.  114  Discussion:  The in situ P-glucosidase activity staining in sections of rapidly growing P. contorta stem, using V R A - G , showed that the reaction products are restricted to the zone of rapid xylem differentiation and lignification, centripetal to the vascular cambium of the stem. No staining was observed in cambial derivatives enlarging into xylem tracheids, or in differentiating phloem cells, which are very active in cell wall biosynthesis at this stage. Thus, it appears that P-glucosidase activity peaks at the onset of secondary wall formation and lignin deposition. Peroxidases have been implicated in the final step of lignin polymerization, and in situ detection of syringaldazine peroxidases in pine stem reveals a tissue distribution that closely matches that observed for P-glucosidases.  Although V R A - G is not a specific  substrate for coniferin P-glucosidase, the p-glucosidase activity detected in situ was almost exclusively restricted to the lignifying region of the secondary xylem centripetal to the enlarging tracheids, a location where the coniferin concentration is reported to be the highest in conifers (Savidge 1989). The pattern of p-glucosidase activity detected in situ in actively growing poplar stems demonstrated  a strong association with the time and place of  lignification within the stem of this species as well. In contrast to Pinus, poplar also produces lignified cells in the phloem (phloem fibers), and these were also found to display p-glucosidase activity during differentiation.  Developmental timing of P-  glucosidase activity in phloem fibers was consistent with that observed in other  115  tissues; i.e. expression drops off as phloem fibers reach maturity.  P-glucosidase  activity is therefore not unique to xylem but appears to also be present in other lignifying cells. Since V R A - G is not an absolutely specific substrate for coniferin Pglucosidase, I cannot state conclusively that an enzyme analogous to Pinus coniferin P-glucosidase is present in lignifying cells of poplar. However, the expression of Pglucosidase/s in lignifying tissue of poplar stems occurs in a spatially and temporally coordinated manner that is consistent with its involvement in lignin biosynthesis. This conclusion is corroborated by detection of essentially identical spatial/temporal patterns of peroxidase activity and immunolocalization of an enzyme involved in an earlier step of monolignol biosynthesis (OMT). However, further investigations will need to be carried out to prove conclusively that a cinnamyl glucoside/p-glucosidase system is operative during angiosperm lignification. Freudenberg and Harkin (1963) proposed that, during lignin formation, the monolignol precursors are transported to the cell wall as glucosides, and that specific p-glucosidases within the cell wall would regenerate the monolignol aglycones for in situ polymerization. Although the nucleotide sequence of the coniferin P-glucosidase suggests that it is a secretory protein, the present investigation does not provide any conclusive evidence to show that its site of action is the cell wall of lignifying cells. Based on the enzyme purification data, i f it is associated with the cell walls, the association is evidently quite weak. However, the high activity and affinity of the coniferin P-glucosidase enzyme for coniferin, together with the localization of Pglucosidase activity exclusively within the lignifying regions of both gymnosperm  116  and angiosperm stems at the appropriate time of development, suggest that coniferin P-glucosidase is performing a crucial role in cell wall lignification in woody tissues.  117  G E N E R A L DISCUSSION A N D PERSPECTIVES  Vascular lignification in Arabidopsis thaliana: Arabidopsis offers many well documented advantages as a model system for study of plant processes. Even complex issues such as biosynthesis, regulation and mechanism of cell wall assembly can be addressed by both molecular and classical genetic techniques in this versatile species.  Evaluation of the importance and  function of each of the numerous components in cell walls, through isolation of mutants defective in their synthesis and/or assembly can be very valuable, and this for example, is most easily approached using Arabidopsis. The isolation of such mutants also facilitates the cloning of genes involved in cell wall biosynthesis.  Several  mutants in the biosynthesis of cellulose and other cell wall polysaccharides have already been described (Baskin et al. 1992, Potikha and Delmer 1995, von Schaewen et al. 1993, Reiter et al. 1993). A mutant lacking syringyl-type lignin has also been characterized (Chappie et al. 1992). This mutant is unable to convert ferulate into sinapate and has been suggested to carry a defective ferulate-5-hydroxylase gene (Chappie et al. 1994). Before a mutant phenotype can be recovered, however, it is essential to fully characterize the wild type. In the case of vascular lignification, a full description of this phenomenon at both morphological and chemical levels in Arabidopsis would aid in the isolation and evaluation of mutants defective in vascular development, lignification or both. The first sign of lignification observed in this study on Arabidopsis seedling  118  development (chapter 1) occured 48 h after imbibition of seeds. Lignin was evident in annular and spiral thickenings of tracheary elements in the radicle, where lignification proceeded in a coordinated cell-by-cell sequence, following the laying down of the secondary wall thickenings. Although lignin deposition is known to initiate in the cell corners and middle lamella/primary wall regions of cells(Terashima et al. 1993), in cells producing secondary wall thickenings, lignin deposition is exclusively localized to the regions of the cell wall with the patterned cell wall structures.  This could either be due to localization of lignin polymerization  machinery within these areas of the wall or to the targeted transport of monolignols (or their glucosides) following the path of cellulose and other polysaccharide deposition. Microtubules in plant cells show a parallel localization pattern aligned with the cell wall thickenings, and are thought to play a part in directing intracellular vesicular traffic containing noncellulosic cell wall polysaccharides (Reiter 1994). A n analogous mechanism could direct the lignin precursors to the developing secondary wall. The flowering stem in Arabidopsis develops about 15 days following germination.  As the stem extends, several endarch vascular strands with xylem  elements differentiate, showing sequential patterns of secondary wall thickenings. About 6 days following emergence of the flowering stem, clusters of cells located between vascular bundles at the base of the stem begin to develop into fibers. The differentiation of these fibers proceeds acropetally and is correlated with the onset of synthesis and deposition of syringyl-type lignin.  The lignin deposited in the  119  secondary wall-containing cells of the plant up to this point (21 days) is predominantly of the guaiacyl type. A compositional difference between lignin in vessel walls and fiber walls has been observed before in woody angiosperms (Musha and Goring 1975, Hardell et al. 1980), and is also observed in different morphological regions of the cell wall even within the same cell. The first lignin deposited in the middle lamella region is richer in p-hydroxyl- and guaiacyl-type monomer units than the lignin deposited later in the secondary wall (Terashima et al. 1986, Whiting and Goring 1982). Thus, it appears that the temporal pattern of deposition of the different monolignols coincides with the order in which they appear in the phenylpropanoid biosynthetic pathway. This sequential appearance of monolignols in xylem tracheids, vessels and fibers is reminiscent of a recapitulation of the tissue's phylogeny. The spatial and temporal separation of syringyl-type lignin deposition in the flowering stem, and the similarity of its lignin chemical composition to that of other angiosperms, suggests that Arabidopsis would make an excellent model system in which to investigate cell wall and lignification-related issues in plants.  P-glucosidases and lignification: p-Glucosidases catalyze the hydrolysis of the glycosidic bond in aryl and alkyl P-glucosides, and in p-l,4-linked glucans such as cellobiose and cellulose. They are ubiquitous in bacteria, fungi, plants and animals. Plant P-glucosidases have been implicated in a multitude of metabolic and defense-related responses which range from the release of conjugated phytohormones to protection against herbivores  120  and pathogens by liberating cyanide, thiocyanates and coumarins. The glucosides of monolignols have been found to accumulate in the cambial sap of all gymnosperms, and in some of the angiosperms, that have been examined (Freudenberg and Harkin 1963, Terazawa et al. 1984, Savidge 1989). These are considered to be the naturally occurring precursors for lignin biosynthesis, and have been suggested to be the transport, or the storage, form of monolignols. Although their exact role has yet to be defined, their importance in lignin biosynthesis is strongly suggested by the results of monolignol glucoside radiotracer experiments in both gymnosperms and angiosperms. The monolignol glucosides have been shown to be effectively incorporated into protolignin in the proper morphological location ,and they were also found to be the most suitable precursors for specific labeling of lignin in the gymnosperms Pinus and Ginkgo (Terashima et al. 1988, Terashima and Fukushima 1988, Fukushima and Terashima 1991) and in the angiosperms magnolia, lilac, poplar and Oryza (Terazawa et al. 1984, Terashima et al. 1986, Fukushima and Terashima 1990).  However, monolignol glucosides have yet to be detected in  lignifying tissues of the latter two species. UDP-glucose:coniferyl alcohol glucosyltransferase activity has been detected in lignifying tissues (Ibrahim and Grisbach 1976, Ibrahim 1977), consistent with the involvement of monolignol glucosides in lignification.  p-glucosidases capable of  hydrolyzing these monolignol glucosides to release the aglycones have also been reported from a range of plant cell culture systems (Hosel and Todenhagen 1980, Hosel et al. 1982, Campbell and Ellis 1992) and spruce seedlings (Marcinowski and  121  Grisebach 1978). However, until now the presence of a corresponding P-glucosidase in the xylem of plants has not been reported, even though xylem is the most active tissue for lignin biosynthesis in plants. The primary objective of the present study was to identify a coniferin-specific P-glucosidase from lignifying xylem and to clone the corresponding cDNA. The resulting information and tools would be valuable in evaluating the essentiality and regulation of the proposed monolignol glucoside/ pglucosidase system in lignification of woody tissues.  They would provide an  opportunity to attempt the directed modification of lignin in plants by genetic engineering. The differentiating xylem of Pinus was selected as the most appropriate tissue for this study (chapter 2). Anion exchange chromatography of total xylem protein extract, and assay for p-glucosidase activity using a range of synthetic substrates and coniferin revealed two main p-glucosidases in pine xylem. One enzyme was very active against the native substrate coniferin while the other was more active against the synthetic substrates.  The exact function of the latter enzyme has yet to be  determined, but it could be involved in some aspect of cell wall polysaccharide biosynthesis or modification. The purified coniferin P-glucosidase had a high activity and affinity towards coniferin and syringin. The aglycone specificity demonstrated by this enzyme supports the view of Hosel and Conn (1982), that plant P-glucosidases do exhibit high specificity towards their naturally occurring glycosidic substrates. In this sense, they are similar to other enzymes of intermediary metabolism which carry out a specific function in the metabolism of the cell.  These authors further  122  emphasize, that the current understanding that plant P-glucosidases display nonspecific substrate specificities is mainly due to the use of non-physiological substrates and heterogeneous enzyme preparations in many experiments. Both of these points were addressed in the present study by using the native substrate coniferin throughout the enzyme purification process, and by using highly purified enzyme for characterization studies. It is worth noting that coniferin p-glucosidase did utilize both coniferin (Km=0.18mM) and syringin (Km=0.29mM) efficiently as substrates which was also true for the P-glucosidase reported in spruce seedlings (Marcinowski and Grisebach 1978). This pattern is analogous to the specificity of the monolignol glucosyl transferase, which is unable to differentiate between coniferyl alcohol and sinapyl alcohol (Ibrahim and Grisbach 1976). Thus, it appears that these enzymes are unlikely to be involved in regulating the type of lignin deposited (i.e. guaiacyl versus syringyl). The N-terminal amino acid information from the purified coniferin Pglucosidase was used to locate the corresponding cDNA in a library constructed from differentiating pine xylem. The identity of the isolated cDNA was confirmed by it's sequence identity to the N-terminal amino acid sequence, by its homology to other plant P-glucosidases and by the production of active enzyme with appropriate substrate preferences when expressed in a heterologous system. Based on sequence similarity, glycosyl hydrolases have been classified into 35 distinct families (Henrissat 1991). Coniferin P-glucosidase belongs to the family 1, which includes Pglucosidases from diverse organisms. This family of enzymes share common active  123  site motifs and, probably, similar folding characteristics.  Conserved Glu and Asp  residues that could also be identified in the coniferin P-glucosidase sequence have been shown to act as acid catalyst and stabilizing nucleophile during cleavage of the glycosidic bond (Trimbur et al. 1992, Withers et al. 1992). The coniferin Pglucosidase amino acid sequence also includes an N-terminal 23 amino acid sequence that has characteristics of eukaryotic secretory signal peptides, suggesting an extracellular localization for the enzyme. This is analogous to the signal peptide found in cell wall-located pro-peroxidases (Lagrimini et al. 1987) and pro-cyanogenic P-glucosidases (Hughes et al. 1992), which supports the hypothesis that monolignol glucosides are transported to the cell wall and subsequently hydrolyzed within the cell wall before polymerization (Freudenberg and Harkin 1963). However, at this stage we cannot exclude the possibility of coniferin p-glucosidase being targeted to another membrane bound compartment within the cell. The P-glucosidases, and peroxidases which are thought to be involved in the final polymerization of monolignols into lignin, were found to be restricted to the differentiating and lignifying xylem within the actively growing pine stem. Even in these differentiating xylem cells, the enzymes were expressed only during the developmental period that coincides with deposition of the secondary wall and initiation of lignification. Interestingly, this same area of the xylem, a few cell layers beneath the cambium, was also found to contain the highest concentration of coniferin during spring growth (Savidge 1989).  This spatially and temporally coordinated  occurrence of p-glucosidases, peroxidases and coniferin in xylem cells provides  124  support for a specific role and participation of a cinnamyl glucoside/ p-glucosidase system in Pinus stem lignification.  Unlike gymnosperms, angiosperms (except  species belonging to Magnoliaceae and Oleaceae) do not appear to accumulate monolignol glucosides in lignifying stems. This is considered by some as evidence against an essential role for a cinnamyl glucoside/ p-glucosidase system in lignification. Although the present study cannot directly counter the above view, P~ glucosidases and other lignin related enzymes within actively growing poplar stems are localized in a spatial and temporal pattern essentially similar to that seen in Pinus stems.  This, together with the reported efficient incorporation of radiolabeled  monolignol glucosides into lignin in poplar (Fukushima and Terashima 1990), tends to suggest the general participation of p-glucosidases in both gymnosperm and angiosperm xylem lignification.  The analogous temporal expression pattern of P-  glucosidases and other lignification related enzymes observed in the phloem fibers of poplar does also suggest that this model may apply to most, i f not all, lignifying cells in plants. The availability of a full-length coniferin P-glucosidase c D N A will permit not only opportunities for probing the structure and function of the enzyme through sitedirected mutagenesis, but also to down-regulate the expression of coniferin Pglucosidase in order to establish its importance. Such studies can be expected to provide valuable new insights into the regulation and metabolic control of these ultimate stages of lignin biosynthesis. The complete coniferin P-glucosidase gene sequence is also a good potential candidate for yielding a xylem-specific promoter.  125  Together, these resources  should provide new impetus to the quest for directed  modification of lignin in plants by genetic engineering, a goal which has produced largely unpredictable results so far (Campbell and Sederoff 1996).  Cell wall and mode of coniferin P-glucosidase action: The first step in cell wall formation following cell division is the fusion of golgi vesicles to form a cell plate, composed mainly of pectins. In the next step, the primary wall composed mainly of cellulose microfibrils and hemicellulose, is laid down on either side of the cell plate, often accompanied by extension growth of the cell. The cellulose microfibrils and hemicellulose are both deposited at the same time and are tightly associated via hydrogen bonding (Reiter 1994). The final stage of cell-type specific secondary wall deposition occurs in several stages which yield layers termed the SI, S2, and S3 layers in tracheid cell walls. Plant cell walls also contain enzymatic proteins and abundant non-enzymatic structural proteins such as extensins, glycine-rich proteins and arabinogalactan proteins (Showaltter 1993). As opposed to the primary cell wall where cellulose microfibrils seem to be oriented randomly, the secondary wall contains almost regularly oriented microfibrils (Willison and Brown 1978).  The cellulose microfibrils and hemicellulose in  secondary walls have been modelled to have a regular twisted honeycomb structure with 15-30° angle of deviation from layer to layer in tracheary elements (Terashima et al. 1993). At the onset of SI layer deposition, lignification first starts in the cell corners and middle lamella regions.  It then proceeds along the middle lamella,  126  through  the  primary  wall  and  ultimately  into  the  secondary  wall.  Microautoradiographic studies of differentiating fibers have shown that the three monolignols are differentially incorporated into the three cell wall layers, with major lignol occurring the order: p-hydroxyphenyl (SI), guaiacyl (S2) and syringyl (S3):(Terashima and Fukushima 1988, 1989, Fukushima and Terashima 1990). Thus the quality of lignin in different morphological layers of the cell wall is different. Although the end result of cell wall lignification has been well documented, the exact mechanism of cell wall component deposition and assembly still remains unresolved. The cellulose microfibril component is thought to be synthesized and assembled at the cell membrane itself via multisubunit cellulose synthase complexes embedded in the membrane ( reviewed by Delmer and Amor 1995). In algae and plants, the complexes are thought to move two-dimensionally within the membrane, and thus impose the pattern of cellulose deposition.  Microtubules of the cell  cytoskeleton are somehow involved in directing this movement of the complexes, possibly through direct or indirect interactions (Delmer and Amor 1995, Giddings and Staehelin 1990). Other cell wall polysaccharides, however, are synthesized in the endomembrane system and are transported to the cell membrane by vesicles derived from the golgi and rough endoplasmic reticulum (Reiter 1994). The fusion of these vesicles at the cell membrane is again thought to be directed by the microtubules, whose orientation and localization often reflects the orientation of secondary cell wall patterning and cellulose microfibrils (Heath and Seagull 1982, Lloyd 1991). Since lignin synthesis occurs at the cell wall, lignin precursors that are  127  synthesized in the cytoplasm should be transported across the cell membrane for polymerization which could act on either the monolignols or their glucosides. As in the transport of vesicles carrying cell wall polysaccharides, the monolignol precursorcarrying vesicles could again be guided by the same microtubules to regions of the cell active in secondary cell wall deposition and lignification.  In this fashion,  lignification would be closely linked to the previously deposited secondary cell wall patterning. The cellular location of reactions involved in phenylpropanoid metabolism and monolignol synthesis is still uncertain. But due to the membrane association of cytochrome P-450 dependent hydroxylases, there is reason to believe that most of the core reactions of phenylpropanoid metabolism would be at least loosely associated at the surface of membranes such as endoplasmic reticulum. Cinnamate 4-hydroxylase and P A L have been found to reside in the microsomal fraction in cell fractionation studies (Hrazdina and Wagner 1985). This would also facilitate the formation of multienzyme complexes that would aid in metabolic channeling of substrates and products in the pathway , as suggested for the phenyl propanoid pathway by Hrazdina and Jensen (1992) and Hrazdina and Wagner (1985). A large proportion of the monolignols thus formed, at least in gymnosperms, is likely to be glycosylated by UDP-glucose:coniferyl alcohol glucosyltransferase either before, during or following transport into membrane bound compartments (vesicles). These monolignol-carrying vesicles could either fuse with larger vacuoles acting as storage pools, or be directed to the plasma membrane to provide substrates for lignification.  Ultrastructural  128  investigations using radiolabeled phenylalanine and cinnamic acid have revealed abundant vesicle traffic within lignifying tracheids, where the vesicles appeared to be fusing with the plasma membrane (Pickett-Heaps 1968).  Considering the high  concentration of coniferin found in conifer xylem, a considerable amount of coniferin should be accumulating within cells. Experiments conducted with protoplasts made from differentiating pine xylem have shown that most of this coniferin is located within these protoplasts, probably in storage vacuoles (Leinhos and Savidge 1993). Angiosperms that do not show accumulation of monolignol glucosides within their developing xylem could simply have developed an alternative of transport and management of monolignols that avoids large-scale accumulation of the glucosides within lignifying cells. Since coniferin p- glucosidase is a secretory protein, synthesis on ribosomes associated with the endoplasmic reticulum may be followed by glycosylation in the Golgi system.  Enzyme-containing Golgi vesicles could then  directly fuse with the plasma membrane to release the enzyme to the cell wall for monolignol glucoside hydrolysis. Alternatively, they could fuse with monolignol glucoside-loaded vesicles before the latter reach the cell wall. Takabe et al. (1989) observed an increase in the proportion and size of smooth endoplasmic reticulum during active lignification and S3 layer deposition in conifer tracheids. The number of golgi complexes and rough endoplasmic reticulum, on the other hand, had peaked during primary wall deposition and gradually decreased during secondary wall lignification.  This indirectly suggests that smooth endoplasmic reticulum is  associated with lignification and thus could be involved in the synthesis , storage and  129  vesicular transport of monolignols or their glucosides to the plasma membrane. The model discussed above would imply that coniferin P-glucosidase is ultimately deposited and active in the cell wall of lignifying cells. This would not be an unprecedented phenomenon, since the occurrence of other P-glucosidases (e.g. cyanogenic p-glucosidases) in the apoplastic space is well documented (Heyn 1969, Kakes 1985, Frehner and Conn 1987). Such an arrangement would establish a spatial compartmentation of p-glucosidase in the apoplast, separated from the substrate stored in the vacuole. The validity of this model could be tested by examining the subcellular localization of coniferin P-glucosidase using immunocytochemical techniques. Cell culture systems have proven to be very useful in plant secondary metabolism-related studies.  The Zinnia mesophyll cell culture system enables  synchronous induction of tracheary element differentiation to be followed in vitro. The mesophyll cells have been shown to respond to phytohormone manipulation with a shift in their development similar to that seen in wood cells with patterned secondary wall thickenings (Fukuda and Komamine 1982). This is accomplished by coordinated changes in activity of several lignin biosynthetic enzymes (PAL, 4 C L and OMT). This response pattern is analogous to the general lignification observed in elicited Pinus banksiana suspension cultures (Campbell and Ellis 1992) which, in addition, exhibited induced coniferin hydrolytic activity. Thus, these angiosperm and gymnosperm cell culture systems  should be very appropriate  for in-depth  investigations into the mode of action of the monolignol glucoside/ P-glucosidase  130  system during lignification of intact cells.  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