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Exploring glycoside hydrolase family 5 (GH5) enzymes Yang Wang Licentiate Thesis in Biotechnology Stockholm, Sweden 2013 ©Yang Wang School of Biotechnology Royal Institute of Technology AlbaNova University Center SE-106 91 Stockholm Sweden Printed by Universitetsservice US-AB TRITA-BIO Report 2013:8 ISSN 1654-2312 ISBN 978-91-7501-719-8 Abstract In 1990, the classification of carbohydrate-active enzymes (CAZymes) was introduced by the scientist Bernard Henrissat. According to sequence similarity, these enzymes were separated into families with conserved structures and reaction mechanisms. One interesting class of CAZymes is the group of glycoside hydrolases (GHs) containing more than 138000 modules divided into 131 families as of February 2013. One of the most versatile and the largest of these GH families, containing enzymes with numerous biomass-deconstructing activities, is glycoside hydrolase family 5 (GH5). However, for large and diverse families like the GH5 family, another layer of classification is required to get a better understanding of the evolution of diverse enzyme activities. In Paper I, a new subfamily classification of GH5 is presented in order to sort the family members into distinct groups with predictive power. In total, 51 subfamilies were defined. Despite the fact that several hundred GH5 enzymes have been characterized, 20 subfamilies lacking biochemically characterized enzymes and 38 subfamilies without structural data were identified. These highlighted subfamilies contain interesting targets for future investigation. The GH5 family includes endo-β-mannanases catalyzing the hydrolysis of the β-1,4-linked backbone of mannan polysaccharides, which are common hemicelluloses found as storage and structural polymers in plant cell walls. Mannans are commonly utilized as raw biomaterials in food, feed, paper, textile and cosmetic industries, and mannanases are often applied for modifying and controlling the property of mannan polysaccharides in such applications. The overwhelming majority of characterized mannanases are from microbial origin. The situation for plant mannanases is quite different, as the catalytic properties for only a handful have been determined. Paper II describes the first characterization of a heterologously expressed Arabidopsis β-mannanase. Key words: GH5, glycoside hydrolase, subfamily classification, mannans, polysaccharides, mannanases, cell wall 3 Sammanfattning År 1990 introducerade forskaren Bernard Henrissat en klassificering av kolhydrataktiva enzymer (CAZymer), enligt vilken enzymerna - baserat på sekvenslikhet - delades in i familjer med konserverade strukturer och reaktionsmekanismer. En intressant CAZym-klass är glykosidhydrolaserna (GH), en klass som i februari 2013 innehöll fler än 138000 katalytiska moduler indelade i 131 olika familjer. En av de största och mest varierade av GHfamiljerna är glykosidhydrolasfamilj 5 (GH5), vilken innehåller en mångfald av identifierade enzymaktiviteter relevanta för nedbrytning av biomassa. För stora och diversifierade familjer som GH5 krävs det dock ytterligare en klassificeringsnivå för att bättre förstå evolutionen och uppkomsten av de många förekommande enzymaktiviteterna. I manuskript I presenteras en ny uppdelning av GH5 enzymer i subfamiljer med syfte att dela upp familjemedlemmarna i distinkta grupper som representerar olika funktioner. Utifrån denna klassificering kan sedan ett enzyms funktion förutsägas baserat på vilken subfamilj det tillhör. Totalt definierades 51 subfamiljer. Trots att hundratals GH5 enzymer har karaktäristerats så visade det sig att 20 av subfamiljerna helt saknar biokemiskt karaktäriserade enzymer och 38 av dem saknar publicerade proteinstrukturer. Dessa subfamiljer är särskilt intressanta för framtida studier. GH5-familjen inkluderar endo-β-mannanaser som katalyserar hydrolysen av den β-1,4länkade huvudkedjan i mannanpolysackarider. Dessa växtpolymerer som ingår i hemicellulosagruppen är vanligt förekommande i cellväggarna, där de fungerar som energilagringsmolekyler eller har en strukturell funktion. Mannaner används ofta som råmaterial för industriell livs- och djurfodersproduktion, papper, textilier och kosmetika. I dessa processer behövs ofta mannanaser för modifiering och kontroll av egenskaperna hos dessa polysackarider. Den överväldigande majoriteten av alla karaktäriserade mannanaser kommer från mikroorganismer. Endast för ett fåtal växtmannanaser har de katalytiska egenskaperna analyserats. Manuskript II beskriver den första karaktäriseringen av ett heterologt uttryckt β-mannanas från Arabidopsis. 4 Sökord: GH5, glykosidhydrolas, subfamiljklassificering, mannaner, polysackarider, mannanaser, cellvägg 5 List of publications I Aspeborg H, Coutinho PM, Wang Y, Brumer H, Henrissat B. (2012) Evolution, substrate specificity and subfamily classification of glycoside hydrolase family 5 (GH5). BMC Evolutionary Biology 2012, 12:186. II Wang Y, Vilaplana F, Brumer H, Aspeborg H. (2013) Enzymatic characterization of a GH5_7 mannanase from Arabidopsis thaliana. Manuscript 6 Table of Contents 1 Introduction ....................................................................................................................... 8 1.1 The plant cell wall ............................................................................................................ 8 1.2 Mannan-based polysaccharides ...................................................................................... 14 1.3 Mannanases .................................................................................................................... 19 1.4 Glycoside hydrolase family 5 (GH5).............................................................................. 22 2 Present investigation ..................................................................................................... 26 2.1 Aim of present investigation........................................................................................... 26 2.2 Materials and methods .................................................................................................... 26 2.2.1 Recombinant protein expression by Gateway technology ....................................... 26 2.2.2 Dinitrosalicylic acid assay (DNS assay) .................................................................. 27 2.3 Results and discussion .................................................................................................... 28 2.4 Concluding remarks ........................................................................................................ 34 3 Acknowledgements ........................................................................................................ 35 4 References ......................................................................................................................... 36 7 1 Introduction 1.1 The plant cell wall All plant cells are surrounded by a polysaccharide-rich matrix known as the cell wall, a highly dynamic structure having various roles during plant growth, development and defense responses. Signal molecules enclosed in the early cell wall guide the further development of the cell [1]. The primary cell wall is formed at the initial stage of cell differentiation. When extra structural support is needed, a secondary cell wall is produced on the inner side of the primary cell wall. These cell-wall layers are distinctly diverse in structure, organization and composition, reflecting different functions. Generally, the walls from different species are highly organized assemblies of various kinds of polysaccharides, phenylpropanoids, and structural proteins [2, 3]. These polymers represent the major part of plant biomass which is utilized by mankind as a source for timber, paper, fuel, food and pharmaceuticals. 1.1.1 Plant cell wall structure and application 1.1.1.1 Primary cell wall The primary cell walls of plants are formed by growing cells under the pectin-rich middle lamella which is formed after cell division (Fig. 1). The thickness and morphology of primary cell walls are dependent on the source. In most cell types, they are architecturally simple and thin structures (between 0.1 and 1.0 μm thick), while the primary walls of collenchyma or epidermal cells may be multilayered and thicker [4]. Primary walls have many important functions such as provision of structural and mechanical support, protection against pathogens and dehydration, maintainance and determination of cell shape and regulation of internal turgor pressure. Moreover, primary walls provide plasticity allowing cell expansion and cell division during growth. Primary cell walls are classified into three types according composition: Type I, II and III primary walls. Type I walls, found in dicots and the noncommelinoid monocots, consist of approximately equal amounts of cellulose and crosslinking xyloglucan [5]. The cellulose–xyloglucan framework is embedded in a pectin-rich matrix in which three types of pectin (homogalacturonans, rhamnogalacturonan I and 8 rhamnogalacturonan II) are covalent interlocked with each other. Type II walls, found in grass species such as rice [6], are characterized by cellulose microfibrils with the same structure as Type I walls but using glucuronoarabinoxylans as the key glycans to cross-link these microfibrils by hydrogen bonds. Type II walls also contain a low amount of xyloglucan without arabinose and fucose decorations, and pectin with the similar structure as those in dicots. The recently described Type III walls, found typically in many ferns, have mannans as the major cross-linking glycan, occupying more than 40 mol% of the cross-linking glycan containing fractions from the examined samples [7]. Similar to Type II walls, Type III walls have low content of pectin polysaccharides. There is a strong interest in primary wall structure and organization from plant scientists, nutritionists and the food industry. For instance, primary walls are the fundamental textural component of plant-derived foods, especially abundant in plant-derived beverages [8]. Fig. 1: Plant primary cell wall structure. Adapted from: [9] 1.1.1.2 Secondary cell wall The secondary cell walls of plants are formed after cell expansion and elongation between plasma membranes and primary walls. They surround highly specialized cells such as vessel elements or fiber cells. With the formation of secondary walls, cell walls become thicker and 9 stronger. In fact, their contribution to plant biomass is much higher than primary walls. Usually, secondary walls are divided into three layers known as S1, S2 and S3 [10] (Fig. 2), where the cellulose microfibrils are strictly ordered in parallel orientation with different angles. The S1 layer, as the thinnest layer, is approximately 0.1–0.35 µm thick with a microfibril (MF) angle of 60º–80º. The S2 layer is the thickest (1–10 µm) and most dominant layer in a secondary wall. The MF angle of this layer is between 5º and 30º, which significantly affect the physical and mechanical properties of the cell. The S2 layer provides mechanical support to the cell for resisting external stress. Compared to the S2 layer, the S3 layer is relatively thin (0.5–1.10 µm) with a MF angle in the range of 60º–90 º [11]. Secondary walls from hardwoods (woody angiosperms) and softwoods (conifers) are very important raw materials for the production of paper, textiles, timber and fuel [12, 13]. Fig. 2: General three dimensional structure of secondary cell wall. The MF angles of cellulose microfibrils are drawn by black lines. Adapted from: [11] 1.1.2 Plant cell wall components Primary walls extracted from higher plant cells typically consist of 20-30% cellulose, 30-70% hemicelluloses, 5-35% pectins, up to 5% glycoproteins [14] and a tiny amount of other components such as phenolic esters (Table 1) [15, 16]. Secondary walls of hardwoods and softwoods generally contain 35‐50% cellulose, 20-50% hemicelluloses, up to 20% lignin, and 10 a small amount of pectins, proteins and extractives (Table 1) [15, 16]. However, the components and their proportion in secondary cell walls may vary among plant species, different tissues in the same species and even in cell type varieties of the same plant. For example, poplar wood is composed by 48% cellulose, 27% hemicelluloses and 21% lignin, but pine wood consists of 41% cellulose, 27% hemicelluloses and 29% lignin [17]. Table 1: The composition of typical plant cell walls by dry mass. Composition (% dry mass) Primary cell wall Secondary cell wall Polymer Dicot Grass Dicot Grass 15-30 20-30 45-50 35-45 Cellulose Hemicellulose Xyloglucan 20-25 1-5 Minor Minor Mannan and Glucomannan 5-10 Minor 3-5 Minor Xylan 5 20-40 20-30 40-50 Mixed-linkage Glucan Absent 10-30 Absent Minor 20-35 5 0.1 0.1 Pectins Minor 1-5 Minor 0.5-1.5 Phenolic esters Minor Minor 7-10 20 Lignin 1.1.2.1 Cellulose Cellulose is the most abundant and characteristic polysaccharide in the plant cell wall. This carbohydrate is synthesized at the plasma membrane with the cellulose microfibrils being deposited directly into the extracellular matrix [18]. Cellulose and cellulose microfibrils are chemically stable, insoluble and resistant to enzymatic digestion. The cellulose molecule is a linear β-1,4-linked glucan chain including 500-14,000 glucose units. Each chain is stabilized by intramolecular hydrogen bonds, and about 40 cellulose chains connect to each other by intermolecular hydrogen bonds between hydroxyl groups in overlapping parallel arrays to form the highly ordered cellulose microfibril. The microfibril is a crystalline or semicrystalline lattice, usually a few micrometers long and 4–10 nm wide [19], although in algae the width of some microfibrils can reach up to 30 nm [20]. The microfibrils are able to support a highly tensile and compressive force, in a manner that is comparable to the functionality of steel. These microfibrils are consequently assembled to form larger fibers [21]. 11 1.1.2.2 Hemicellulose Hemicellulose polysaccharides are found in both plant primary walls and secondary walls, in which they usually tightly bind onto the surface of cellulose via hydrogen bonds. Galactoglucomannan, glucomannan, arabinoglucoronoxylan, glucuronoxylan and xyloglucan (XG) are examples of common and abundant plant cell wall hemicelluloses [17]. In primary walls of dicotyledonous plants, XG is the main hemicellulose, while glucuronoarabinoxylan and mixed linkage β-glucans are the principle hemicelluloses of grasses. Glucuronoxylan is the dominant hemicellulose in the secondary cell wall of both dicotyledonous plants and grasses, whereas galactoglucomannan is the major secondary wall hemicellulose in gymnosperms [22]. Hemicelluloses are characterized from a β-1,4-linked backbone. They are often branched polymers constructed from various 5- and 6-carbon sugars. Generally, glucose, xylose, mannose, and galactose are the major monosaccharides found in the backbone of hemicellulose, whereas arabinose, glucuronic acid, and galactose are located in the side chain. Virtually, all hemicelluloses are more soluble in alkali solutions than in water. Those including acids in their side chains are slightly charged and hydrophilic, and the ones containing hexoses and uronic acid are more easily digested by bacterial enzymes than the other hemicelluloses [23]. XG is one of the most abundant hemicelluloses and consciously the most well-studied, it is formed in the Golgi [24]. Xyloglucans (XGs) cross-link cellulose microfibrils and therefore have a stabilizing function [25]. Moreover, XG is also the primary storage polysaccharide in certain seeds [26]. XGs share a β-1,4-glucan backbone in which the 6-O-position of glucosyl residues is linked with xylosyl side chains in a regular pattern of three substituted glucosyl residues followed by one unsubstituted glucose unit at the reducing end. 1.1.2.3 Pectin Pectin is a highly hydrophilic polysaccharide and is the main component of middle lamella. This polysaccharide is also abundant in the Type I primary wall. It is generally believed that pectins and hemicelluloses associate with each other to form a hydrated gel in which cellulose 12 microfibrils are embedded [27]. Pectins have many functions, for example to limit wall porosity and regulate pH. Additionally, they act as recognition compounds in plant defense and symbiotic systems [28]. The backbone of pectins is generally formed from α-1,4-linked D-galacturonic acid units. Pectic polysaccharides are divided into three groups: homogalacturonan (HG), rhamnogalacturonan-I (RG-I) and substituted galacturonans (SG) [29, 30]. HG is a partly methyl esterified linear polysaccharide with a backbone of 1,4-linked α-D-galactopyranosyluronic acid (GalpA) residues. Additionally, the C-3 or C-2 positions of some HG polymers are O-acetylated in certain plant species [31]. RG-I is a highly branched polysaccharide with the repeating disaccharide [→4)-α-D-GalpA-(1→2)-α-L-Rhap-(1→] as the main chain which may have O-acetylated GalpA residues at C-2 and/or C-3 positions [32]. Like most other types of pectins, SGs, a diverse group of polysaccharides, have linear 1,4linked α-D-GalpA backbones but with substituted galacturonan side chains [29]. Rhamnogalacturonan-II and xylogalacturonans are examples of two well-known pectin polysaccharides in the SG group. 1.1.2.4 Lignin Lignins are highly complex and hydrophobic molecules found in both the middle lamella and the secondary wall. These hard and hydrophobic polymers are space fillers in the cell wall in between the other wall components, making the cell wall waterproof. The major function of lignin is to make the wall rigid and durable in order to protect the wall against physical or biological attacks [33]. Lignins are built up from three different polyphenolic precursors (coumaryl alcohol, coniferyl alcohol, and sinapyl alcohol) through covalent and noncovalent bonds [34]. Plant materials with different proportions of lignins are used in different commercial processes. For example, the durable and long-lasting wood with high percentage of lignins is a very suitable building material, while low lignified materials are ideally utilized in paper production and in biomass saccharification for producing ethanol [35]. 13 1.1.2.5 Protein In addition to carbohydrates, plant cell walls accommodate a wide variety of proteins. Probably several hundreds of different proteins are cell wall-localized [36]. These wall proteins are divided into two groups: structural proteins and functional proteins [37]. Structural proteins are found throughout the plant cell wall and especially abundant in the primary wall. These proteins have been divided into major classes based on their characteristic amino acid composition: hydroxproline-rich glycoproteins, glycine-rich proteins, proline-rich proteins and so on [38]. Compared to structural proteins, functional proteins such as oxidative enzymes and hydrolytic enzymes are more involved in cell development, pathogen protection, cell expansion and cell wall maturation [39]. 1.2 Mannan-based polysaccharides Mannan polysaccharides are found in diverse organisms such as bacteria, fungi and plants. These important hemicellulosic carbohydrates are located in the cell wall of different types of cells and tissues such as root, tuber, bulb and seed of various plants [40]. Mannans include homogeneous and heterogeneous polymers consisting of different sugars such as D-mannose, D-galactose and D-glucose. According to their sugar unit composition, they are classified into four groups: pure mannans, galactomannans, glucomannans, and galactoglucomannans. These mannan types mainly function as seed storage and/or structural components [41]. In the endosperm cell wall of legume, galactomannan as a storage component occupies approximately up to 30% of the seed dry weight [42]. In the Type III primary cell wall of many ferns, mannans function as a key glycans replacing xyloglucans and glucuronoarabinoxylans as the cellulose cross-linking hemicellulose [43]. In the secondary wall of softwoods, galactoglucomannan as the principal hemicellulose binds cellulose to form a polysaccharide network, which is embedded in lignin [44]. Besides their function as storage and structural saccharides, mannans also have an important role as signaling molecules in plant cell differentiation [45]. Currently, there is great interest in mannan polysaccharides. This interest comes not only from the important role of these polysaccharides in plant cell 14 wall formation [46, 47], but also because these polysaccharides are a largely unexploited resource of raw material, which could find use in many industrial applications such as feed, textile and mining [48]. 1.2.1 Pure Mannan In pure mannan, the linear chain contains only β-1,4 cross linked D-mannosyl residues (Fig. 3). This type of mannan is usually found in the seed endosperm of Palmae, green coffee bean and the seed of some Umbelliferae species [49, 50], where they are used to protect the seed endosperm from mechanical damage [51]. These polysaccharides are hard and highly insoluble in water at neutral pH. In ivory nut, pure mannan is the principle component of the seed endosperm cell wall. According to the feature of crystalline polymorphism, the linear mannan extracted from ivory nut is divided into two families: mannan I (also called mannan A) and mannan II (also called mannan B). The major differences between the two fractions are: solubility, degree of polymerization (DP) [52, 53] and morphology [53, 54]. Mannan I is highly crystalline with low molecular mass. This mannan polymer can be dissolved in 6% (w/w) sodium hydroxide solution [55, 56]. In contrast, mannan II is less dense and crystalline but with a high molecular mass [57]. Mannan II is insoluble even in high concentrated sodium hydroxide solution [55, 58]. H HO ......O H H H OH O OH H OH H O HO H H O OH O...... H H n Fig. 3: Pure mannan structure. 1.2.2 Galactomannan In galactomannan, the backbone is formed by β-1,4-linked D-mannosyl substituted by α-1,6linked galactosyl as side group [59] (Fig. 4). This type of mannan serves as a carbohydrate reserve in the seed endosperm of leguminous plants (Leguminosae), but is also present in the 15 seeds of a few non-leguminous plants [60]. In contrast to pure mannan, galactomannan is soluble in water, but the solubility is affected by the proportion of mannose to galactose. Three of the most commercially important galactomannans are carob galactomannan from carob or locust bean tree (Ceratonia siliqua), guar gum extracted from guar bean (Cyamopsis tetragonoloba or Cyamopsis psoraloides) and tara gum isolated from tara tree (Caesalpinia spinosa). Carob galactomannan, known as carob/locust bean gum (E-number: 410), is mainly extracted from the seed endosperm of carob tree which has its natural habitat in the Mediterranean region. Normally, 100 g/kg seeds (pod weight) contain roughly 320-400 g/kg highly purified carob galactomannan. They are utilized as common raw materials in food industry (e.g. ice cream) to replace fat. The typical carob galactomannan has a molecular weight of approximately 310 kDa and a mannose/galactose substitution level at 4:1 [61]. It is viscous and relatively stable in different pH solutions. Guar gum (E-number: E412), the major replacement of carob galactomannan, is isolated from the seed of guar tree, a tree abundantly cultivated in northwestern India, Pakistan and the US. The yield of guar is approximately 150 000 tons per year in the world, and about two thirds originate from Pakistan. Since the price of commercial guar is cheaper than carob, it is broadly used in cattle feeding and gum production to coat/stiffen paper. Guar gum has a mannose:galactose substitution level of 2:1 and a determined molecular weight of approximately 220 kDa [61]. Tara gum (E-number: E417) is obtained from the tara bush which is grown mainly in Ecuador, Peru and East Africa. This mannan is commonly utilized in food industry to control ice crystal growth in frozen dessert and to improve the gel structure in meat based product. The viscosity of tara gum is similar to that of carob and guar gum in cold solution, but it has a higher viscosity compared to these two kinds of galactomannan in heated solution. Typically, tara gum has a mannose:galactose ratio of 3:1 and a very high molecular mass [62]. 16 OH HO H HO H H H HO ......O H H H H OH O H O H OH O H O HO H H O OH H H OH HO O H H H H H OH H O HO OH O H O OH H OH O...... H H Fig. 4: Galactomannan structure. 1.2.3 Glucomannan Glucomannan is a linear chain composed of D-mannosyl and D-glucosyl residues which are connected to each other by β-1,4-linkages, and are often acetylated (Fig. 5). Many of these polysaccharides are water soluble with a mannose:glucose ratio ranging from 4:1 to below 1: 1 [40] and a DP value higher than 200. Glucomannan has been detected in various tissues of plant such as the tuber of the konjac plant (e.g. A. konjac plant C. Koch), which is widespread in the Far East and Southeast Asia. Between 60–80% of the konjac tuber is composed of glucomannan. Since konjac glucomannan is a water soluble and abundant material, it has been widely used in food industry and pharmaceutical research. Konjac glucomannan (E-number: 425) has a molecular weight 200 kDa-2000 kDa [63] and a mannose:glucose ratio at approximately 1.5:1 [64]. Usually, 5–10% of the backbone residues of konjac glucomannan are connected with acetyl groups by ester bonds [65]. These extra chemical modifications significantly affect konjac glucomannan properties such as solubility and gelation. For example, at elevated temperature, a higher degree of acetylation weakens the network structure of the konjac glucomannan gel [66]. OAc H ..... O H HO H H H O OH H HO O H OH H H H O OH OH H O HO H H H O OH H HO O H H H H H O...... OH O OH Fig. 5: Acetylated glucomannan structure. 17 1.2.4 Galactoglucomannan The backbone of galactoglucomannan consists of β-1,4-linked D-mannosyl and D-glucosyl residues decorated with α-1,6-linked galactosyl side chains (Fig. 6). Sometimes, the backbone mannopyranose residues are acetylated at the hydroxyl groups of the C2 or C3 positions with an estimated acetylation ratio of 1:3-4. Generally, acetylated galactoglucomannan extracted from the lignified secondary wall has a DP range from 100 to 150, and two groups can be distinguished. The first group is rich in galactose (5-8 w/w% dry weight) and water soluble with a ratio of mannose:glucose:galactose residues determined to be 3:1:1. The second group is poor in galactose (10-15w/w% dry weight) and aqueous alkali soluble with mannose:glucose:galactose residue ratio at 3:1:0.1 [17]. Galactoglucomannan is an abundant hemicellulose in many species of softwoods such as Norway spruce (Picea abies), which contains approximately 10-20% of O-acetylated galactoglucomannans [17, 67]. The economically important Norway spruce is grown throughout Nordic countries and is mainly used for timber, or in the pulp and paper industry. Spruce galactoglucomannan, one of the most characterized and extensively studied galactoglucomannans, has a mannose:glucose:galactose ratio of 3.5-4.5:1:0.5-1 [68] and an approximate molecular weight between 20 kDa and 78 kDa [69, 70]. In addition to softwoods, galactoglucomannan has been found in other tree species, for example Populus monilifera, and also in clubmoss [71], blackberry [72], and fern [73]. OH HO H HO H H H AcO ......O H H H OH O OH H H O H OH O H O HO H H H O OH H HO O H H Fig. 6: Acetylated galactoglucomannan structure. 18 OH H H H OH O OH H O HO H H O OH O...... H H 1.3 Mannanases For a complete digestion of mannan polysaccharides, the degrading enzyme system usually involves endo-β-1,4-mannanase (EC 3.2.1.78), endo-β-1,4-mannosidase (EC 3.2.1.130), exoβ-1,4-mannosidase (EC 3.2.1.25), β-glucosidase (EC 3.2.1.21), α-galactosidase (EC 3.2.1.22) and acetyl mannan esterase. Endo-β-1,4-mannanases hydrolyze the backbone of mannan polysaccharides into oligosaccharides, then the other enzymes degrade these oligosaccharides into monosaccharides, as exemplified in the degradation of O-acetyl-galactoglucomannan (Fig. 7). Fig. 7: Overview of the of the enzyme portfolio required for the complete digestion of Oacetyl-galactoglucomannan. 1.3.1 Origin and function Mannanases have been found in various organisms including bacteria [74], fungi [75], plants and animals [76, 77]. Microbial mannanases are mostly secreted extracellular enzymes used together with other glycoside hydrolases to degrade plant biomass providing simple sugars as energy for the microorganism. Many plant mannanases are involved in seed development. In Arabidopsis thaliana, three endo-β-mannanases genes are expressed during seed germination [78]. In some plants, mannanases play a role in the degradation of endosperm in order to provide energy for seedling growth [79]. For example, GmMAN1 from soybean digests the βmannan-rich cell wall during the period of soybean seedling establishment [80]. Beyond their involvement in seed development, plant mannanases also participate in the process of tissue 19 softening during fruit ripening [81, 82] and the development of pollen and anther during flower differentiation [76]. 1.3.2 Biochemical activities Mannanases belong to the glycoside hydrolase (GH) class of enzymes. Based on amino acid sequence similarities, mannanases have been classified into the GH-families: GH5, GH26 and GH113 in carbohydrate-active enzymes database (CAZy) [83]. Through a retaining mechanism, mannanases are responsible for catalyzing the hydrolysis of the β-1,4-linked backbone within different mannan polysaccharides. Generally, the complete digestion of mannan polysaccharides by mannanases produces manno-oligosaccharides such as mannotetraose, mannotriose and mannobiose, and the proportion of glucose and galactose residues in certain substrates can greatly affect the result of digestion. Biochemically, the optimal temperature and pH of examined mannanases are usually in the range of 35-70ºC and 3.0-7.5 respectively. One interesting exception is the Thermotoga neapolitana 5068 mannanase with an optimal temperature of 92ºC [84]. In addition to their hydrolytic activity, some mannanases from these three GH families have been reported to have transglycosylation activity (Fig. 8). LeMAN4a from tomato is one example of a mannanases with both hydrolytic and transglycosylation activities [85, 86]. H HO HO Glu(acid/base) OH H ......O HO H H HO H O OH H O HO O H H - O H H ROH H O...... OH O OH O H H H H O ..... OH Glu(acid/base) R=H O O Hydrolysis - OH O OH H .....O HO H H O ROH OH H OH H .....O HO H OH H H H R=Suger .....O O O Transglycosylation HO Glu(nucleophile) O OH OH H H O OH OSugar H Glu(nucleophile) Fig. 8: An overview of the hydrolysis and transglycosylation mechanisms of mannanase. 1.3.3 Three dimensional structure of mannanase Although mannanases from different GH families show no significant protein sequence similarities, they are most similar in the spatial arrangement. All mannanases share the (β/α) 8 20 barrel fold structure with relatively conserved amino acids (e.g. Two Glu: as acid/base and nucleophile) located at the active site, and cleave the glycosidic bond of mannosides between subsites -1 and +1 [87]. Most of them require at least five substrate-binding sites to be able to perform efficient hydrolysis [74, 88, 89]. However, the interaction within the enzymesubstrate complex may vary. For example, the GH5 Thermobifida fusca mannanase (TfMan5) binds mannotriose at subsites -2, -3, and -4 [90], whereas the GH26 Cellvibrio japonicas mannanases A (CjMan26A) uses subsites -1, -2, and -3 for binding to the same mannooligosaccharide [91]. Another specificity difference between GH5 and GH26 mannanases was recently revealed. Both Bacillus agaradhaerens mannanase (BaMan5A) and Bacillus subtilis mannanase (BsMan26A) from GH5 and GH26 respectively can accommodate mannose at the negative binding sites; but BaMan5A also has the capacity to bind glucose in subsites -2 and +1 [74]. Thus, the flexible substrate binding cleft of BaMAN5A allows the enzyme to hydrolyze glucomannan with a variable backbone of mannose and glucose units. Table 2: Summary of GH5, GH26 and GH 113 mannanases. Characterized enzyme GH family Organism EC number with manganese activity No GH5 Bacteria 3.2.1.78 31 14 3.2.1.78; 3.2.1.4 4 9 3.2.1.78; 3.2.1.73 1 1 3.2.1.78/73/4/8 1 N 3.2.1.78 41 7 3.2.1.78; 2.4.1- 1 1 3.2.1.78 30 13 3.2.1.- 1 4 3.2.1.78; 3.2.1.73 1 N 3.2.1.78/4/115 1 N 3.2.1.78/73/4/8 1 N Eukaryota 3.2.1.78 7 N Bacteria 3.2.1.78 1 1 Eukaryota GH26 GH113 PDB No Bacteria 21 1.3.4 Mannanase application There are many great potential benefits related to the application of mannanases in the industrial field. Some examples are given in below. In the pulp and paper industry, mannanases can be used in softwood pulp bleaching for lignin removal from wood fibers. Compared to alkaline pretreatment of pulp, enzymatic pretreatment is environmentally friendly reducing the use of harsh chemicals. In the detergent industry, mannanases have a role as stain removal agents because they are applied to digest mannan polymers into smaller water-soluble fragments. In food industries, the degradation of coffee mannans is often performed by different mannanases preparations. This process can efficiently decrease the high viscosity of coffee extract in order to improve the quality and technical process of instant coffee. Moreover, mannanases are utilized in the degradation of mannan and galactomannan in enzymatic oil extraction from coconut meat, producing high quality coco oil. 1.4 Glycoside hydrolase family 5 (GH5) 1.4.1 Classification GH5, previously known as cellulase family A, is quantitatively a large GH family and belongs to Clan GH-A. This family comprises a diversity of enzymes which are found in prokaryotes (archaea and bacteria), eukaryotes (fungi, plants and animals) and viruses. Interestingly, no human enzyme has been reported in this family. At the time of writing, more than three thousand GH5 enzyme sequences have been identified in the CAZy database. Recently, about 80% of the public sequences were classified into 51 subfamilies (GH5_1–GH5_53, excluding GH5_3 and GH5_6 which have been merged into GH5_4 and GH5_5 respectively) [92]. 1.4.2 Mode of action GH5 consists of enzymes with approximately 20 different identified enzyme activities (Table 3). They share a classical retaining mechanism following a canonical double-displacement of glycosyl transfer involving a covalent glycosyl-enzyme intermediate in order to retain the anomeric configuration [93, 94] (Fig. 9). Because GH5 enzymes are retaining glycoside 22 hydrolases, they all have the potential to act as both hydrolytic and transglycosylating enzymes. At the active site, a pair of conserved Glu, as a general acid/base and a catalytic nucleophile, is responsible for driving the overall hydrolysis reaction in the anomeric center [95]. The catalytic nucleophile is responsible for the attack of the anomeric carbon forming a substrate-enzyme intermediate [96]. The acid/base serves to protonate the glycosidic oxygen hydrolyzing the intermediate [97]. Besides these two catalytic amino acids, there are additionally at least 5 conserved residues located at the -1 subsite of GH5 enzymes [98]. Table 3: A variety of specificities of GH5 enzymes. Enzyme Activity Chitosanase β-mannosidase Cellulase Glucan 1,3-β-glucosidase Licheninase Glucan endo-1,6-β-glucosidase Mannan endo-β-1,4-mannosidase Endo-β-1,4-xylanase Cellulose β-1,4-cellobiosidase β-1,3-mannanase Xyloglucan-specific endo-β-1,4-glucanase Mannan transglycosylase Endo-β-1,6-galactanase Endoglycoceramidase β-primeverosidase β-glucosylceramidase Hesperidin 6-O-α-L-rhamnosyl-β-glucosidase Exo-β-1,4-glucanase / cellodextrinase EC 3D number Subfamily Organism structure 3.2.1.132 2 bacteria No 3.2.1.25 7 bacteria Yes 3.2.1.4 1,2,4,5,8,22,26 all Yes 3.2.1.58 9,14 Eukaryota No 3.2.1.73 2,26,36,37 bacterial, Eukarayota No 3.2.1.75 9,15 Eukaryota No 3.2.1.78 7,8,10,17,25,36 bacterial, Eukarayota Yes 3.2.1.8 21 bacteria No 3.2.1.91 2 bacterial, Eukarayota No 3.2.1.31,34,39,48 bacterial, Eukarayota Yes 3.2.1.151 4 bacterial Yes 2.4.1.7 Eukaryota Yes 3.2.1.164 16 Eukaryota No 3.2.1.123 27,28,29 bacterial, Eukarayota Yes 3.2.1.149 23 Eukaryota No 3.2.1.45 12 Eukaryota No 3.2.1.168 23 Eukaryota No 3.2.1.74 37,52,53 bacteria No 23 Aacid/Base O O Aacid/Base O H - ‡ O + O OR1 δ O H Oδ R1 HOR2 - δ - O O O O HOR1 Nucleophile O Transition state Aacid/Base O O H Aacid/Base O - O + OR2 O H O ‡ - O - O R2 O Nucleophile δ O H δ O O Aacid/Base Nucleophile Glycosyl-enzyme intermediate R2 - δ O - O O O Nucleophile Nucleophile Fig. 9: Mechanism of retaining glycosidase [99]. 1.4.3 Structure property To date, there are 38 reported GH5 three dimensional (3D) structures. GH5 enzymes contain an amino acid chain which forms a (β/α) 8 fold with a classical cleft topology of the active site (Fig. 10A) in which the catalytic nucleophile Glu and acid/base Glu are positioned at the C terminal of β-strand 7 and β-strand 4 respectively (Fig. 10B). The first solved 3D structure of GH5 enzymes was a 343 amino acid endoglucanase from Clostridium thermocellum (Cel C) [100]. The crystal structure of Cel C determined at 2.15 Å resolution revealed a typical (α/β) 8 barrel structure with an active site having a cleft topology [100] (Fig. 10A). The active site of Cel C is located at the C-terminal of the barrel and involves several important conserved amino acids such as the catalytic residues (Glu 280 and Glu 140) but also Trp 313, His 198, Tyr 200 and Arg 46 [100]. The formation of the observed deep active site is possible because of an insertion of a subdomain consisting of four α helices and two β strands which are located between α-helix 6 and β-strand 6, enlarging the top of α/β barrel on one side [100]. 24 (A) (B) Fig. 10: The three dimension structure of CelC (PDB: 1CEC). (A) The overview structure with a distinct cleft topology of the active site. (B) Secondary structure with two catalytic Glu residues. 1.4.4 Modular architecture of GH5 enzymes Many GH5 enzymes consist of a single catalytic domain which is responsible for the hydrolysis of glycoside substrate. However, a great number of GH5 family proteins (e.g. proteins form GH5 subfamilies 7 and 8) contain, in addition to the catalytic module, one or several carbohydrate binding modules (CBMs) which can be located at both the N- or Cterminal of the complete protein [92]. CBMs direct the catalytic module to its substrate enhancing the glycoside hydrolytic efficiency. In the CAZy database, CBMs have been divided into 66 families based on sequence similarity [101]. Based on substrate binding specificity, they have been grouped into three types which are: type A (surface-binding: long insoluble polysaccharides), type B (glycan chain-binding: medium-sized soluble polysaccharides) and type C (small sugar-binding) [101]. 25 2 Present investigation 2.1 Aim of present investigation Specific aim of Paper I: The aim of Paper I was to divide the large Glycoside hydrolase family 5 (GH5) into subfamilies. Specific aim of Paper II: The aim of Paper II was to clone, heterologously express and characterize a GH5 endo-βmannanase from Arabidopsis thaliana in detail. 2.2 Materials and methods 2.2.1 Recombinant protein expression by Gateway technology Recombinant protein expression is widely used in life science and biotechnology to generate adequate amount of protein for further study, for example to investigate the physical and biochemical properties of the target protein. Expression systems using various host organisms, both prokaryotic and eukaryotic, have been developed from bacteria, yeast, insects, mammals and plants. Cell-free protein expression methods have also been developed using in vitro translation. Two of the most well-known and commonly used hosts are the gram-negative bacterium Escherichia coli (E. coli) and the methylotrophic yeast Pichia pastoris (P. pastoris). Protein expression using these hosts is easily performed in large scale, and the organisms are cultivated at low cost. For the two expression systems, it is possible to efficiently produce the desired protein either intracellularly or extracellularly. Another advantage of using these systems is that the whole process of expression can be regulated by inducible promoters (e.g., T7 promoter and alcohol oxidase promoter), which are activated by inducing reagents, such as isopropyl β-D-1-thiogalactopyranoside (IPTG), or methanol. However, there are several important differences between the two hosts. For example, although E. coli growth is much faster than that of P. pastoris, the production of eukaryotic proteins in E. coli may be hampered, as those expressed proteins are frequently misfolded, 26 denatured and insolubly aggregated. These problems can be overcome by using the P. pastoris system, which is capable of performing most posttranslational modifications needed to produce eukaryotic proteins with correct folding and high solubility [102]. Gateway® cloning technology is designed for efficient transfer of the desired DNA sequence from a single entry clone into any Gateway® expression vector of interest. This recently developed method supports many available expression choices by using multiple destination vectors such as vectors with His-tags and GST-tags, which makes the Gateway® system very convenient for protein expression. Generally, the whole process is handled by two reactions: the LR reaction and the BP reaction. The LR reaction, as the major pathway, contributes to transfer the desired gene fragment from entry clone to destination vector in order to form an expression clone (Fig. 11A). The BP reaction, as the reverse pathway of LR reaction, is responsible for recombining the ideal DNA fragment from an expression clone with a donor vector creating a new entry clone (Fig. 11B). (A) (B) Fig. 11: (A) The LR reaction. (B) The BP reaction. Adapted from: the protocol of GATEWAY™ Cloning Technology. 2.2.2 Dinitrosalicylic acid assay (DNS assay) The hydrolysis of carbohydrate substrates catalyzed by glycoside hydrolases releases reducing 27 sugars. Formation of reducing sugars can be used to determine the activity of glycoside hydrolases such as amylases, pectinases mannanases, and xyloglucanases. Two of the most popular and widely used reducing sugar assays are the Nelson-Somogyi (NS) assay and the 3,5-dinitrosalicylic acid (DNS) assay. Other methods have also been developed but are rarely used. The DNS assay was introduced in 1925 to determine the amount of reducing sugars in urine [103]. The principle of the DNS assay is to measure free carbonyl groups which are released during hydrolysis reactions catalyzed by glycoside hydrolases. Under alkaline conditions, the aldehyde functional group of sugars (e.g. glucose and mannose) is oxidized to carboxyl group, and simultaneously 3,5-dinitrosalicylic acid is reduced to 3-amino,5-nitrosalicylic acid leading to a color shift in the reaction solution from yellow to orange or brown (Fig. 12), which can be measured spectrophotometrically at 540 nm. Oxidation O C H OH O OH O R Reduction + Aldehyde group O 2N NO2 3,5-Dinitrosalicylic acid O R C OH OH + O 2N Carboxyl group OH NH2 3-amino,-5nitrosalicylic acid Fig. 12: Principle reaction of DNS assay. 2.3 Results and discussion 2.3.1 Paper I Glycoside hydrolase family 5 (GH5) is a glycoside hydrolase family containing diverse enzymes with various enzymatic activities involved in the degradation of carbohydrates and glycoconjugates (Table 3), which previously have been grouped into 10 subfamilies (A1A10). However, the number of GH5 sequence deposited in the CAZy database has rapidly increased in recent years (even though some have been reclassified as GH30 enzymes) and 28 numerous GH5 enzymes cannot be classified into these ten original subfamilies. With a new GH5 subfamily classification, a larger portion of the enzymes could be assigned into subfamilies, and this could be used to aid functional prediction, and guide enzyme discovery and structure-function studies, while providing fruitful insights into the complex evolutionary relationships of GH5 enzymes. Our large-scale bioinformatic analysis of GH5 included all public available family sequences (approximately 2300 catalytic module sequences with highquality), and we were able to define 51 GH5 subfamilies including those previously described. Among these classified subfamilies, 31 subfamilies had at least one enzyme with a determined activity, while 20 subfamilies contained only uncharacterized enzymes. Approximately 20% of the analyzed sequences could still not be assigned into a subfamily, even though some of them have been functionally characterized. The main reason for this was that these sequences lacked sufficient sequence similarity with other GH5 sequences, so that the five-membered criterion we had specified for a subfamily could not be met. 2.3.1.1 Monospecific and polyspecific subfamilies When the subfamily classification was supplemented with available biochemical data, 17 subfamilies were found to be monospecific, whereas 14 subfamilies were polyspecific (Table 4). A monospecific subfamily is defined as a subfamily containing characterized enzymes with a single activity (one EC number). Particularly, if the subfamily includes several identified enzymes with the same specificity, it can be confidently described as a monospecific subfamily. However, if a monospecific subfamily contains only one biochemically reported enzyme, chances are that other activities will be eventually discovered as more members will be characterized and the subfamily will become polyspecific. In GH5_5, the largest monospecific subfamily, only enzymes with endo-β-1,4-glucanase activity (EC 3.2.1.4) have been reported. One example of a endo-glucanase with reported crystal structure comes from the thermophilic fungus Thermoascus aurantiacus [104]. In contrast to a monospecific subfamily, a polyspecific subfamily is comprised of members with two or more identified activities (several EC numbers). The largest polyspecific 29 subfamily in GH5 is GH5_2 which contains a number of enzymes with β-1,4-glucan cleaving activities. For example, Streptomyces griseus HUT 6037 is a chitosanase (EC 3.2.1.132) with additional transglycosylation activity [105], and the Fibrobacter succinogenes S85 endoglucanase (EC 3.2.1.4) can use both oat spelt xylan and carboxymethyl cellulose (CMC) as substrate [106]. Table 4: A summary of monospecific and polyspecific subfamilies in GH5. Monospecific subfamily Sequence No. Characherized enzyme No. 3D structure No. GH5_5 123 44 0 GH5_8 71 23 4 GH5_10 19 6 1 GH5_14 15 1 0 GH5_15 10 9 0 GH5_16 10 2 0 GH5_17 5 3 0 GH5_21 10 4 0 GH5_22 12 1 0 GH5_27 5 2 0 GH5_28 2 1 GH5_29 1 0 GH5_31 5 1 0 GH5_34 5 1 1 GH5_39 7 1 0 GH5_52 6 2 0 GH5_53 6 1 0 Polyspecific subfamily Sequence No. Characherized enzyme No. 3D structure No. GH5_1 133 29 2 GH5_2 245 90 7 GH5_4 160 50 0 GH5_7 8 5 GH5_9 107 22 1 GH5_12 42 3 0 GH5_23 5 2 0 GH5_25 5 2 GH5_26 17 7 1 GH5_36 23 2 1 GH5_37 19 6 1 GH5_38 10 0 0 GH5_46 15 0 0 30 2.3.1.2 Subfamilies with no identified activities We also defined a large number of GH5 subfamilies completely composed of members lacking a determined activity (EC number). This category includes GH5_11, GH5_13, GH5_18 - GH5_20, GH5_24, GH5_30, GH5_32, GH5_33, GH5_40 - GH5_45, GH5_47, and GH5_49 - GH5_51. A detailed study of these enzymes may exhibit valuable insights regarding their hidden activities. Hints of the enzymatic function for bacterial enzymes may be derived from carbohydrate binding modules (CBMs) or operons, but also from transcriptomics and functional metagenomics. 2.3.1.3 GH5 subfamily 7 (GH5_7) Since paper II describes the characterization of a GH5_7 mannanase, this subfamily is presented in more detail below. Subfamily GH5_7, previously known as subfamily A7, contains β-1,4-mannan-cleaving enzymes from various species of eukaryotic, archaeal and bacterial origin. In this subfamily, the identified enzymes reveal three different specificities which are endo-β-1,4-mannanase activity (EC 3.2.1.78; e.g. MAN5C), mannan transglycosylase activity (EC 2.4.1.-; e.g. LeMAN4a) and exo-β-mannosidase activity (EC 3.2.1.25; e.g. CmMan5A). The Man5C from Vibrio sp. Strain MA-138, an endo-mannanase, hydrolyses the β-1,4-linked backbone within mannan-based saccharides [107]. The tomato LeMAN4a, involved in fruit development, is the first characterized plant mannanase which has the ability to act both as a mannan hydrolase (MEH) and as a mannan transglycosylase (MET) in vitro [85, 86]. The CmMan5A is an exo-β-1,4-mannanase or a β-mannosidase from Cellvibrio mixtus, which catalyzes a hydrolytic reaction releasing mannose from the nonreducing end of manno-substrates [108]. The main difference between CmMan5A and the other GH5 endo-mannanases is that CmMan5A has three loops (closed to the active site) with distinct different lengths (Fig. 13), which probably explains the evolved exo-activity [108]. The most important distinction is the extended loop (378-412) inserted between β-strand 8 and α-helix 8, which forms a double steric barrier closing the substrate binding cleft at subsite -1 [108]. Noticeably, all plant mannanases belong to GH5_7, and besides this subfamily, enzymes with mannanase activity have also been identified in GH5_8, GH5_10, GH5_17, 31 GH5_25, and GH5_36. Fig. 13: The 3D structure of CmMan5A (PDB: 1UUQ). The arrows mark three loops with different lengths compared to other GH5 endo-mannanases. 2.3.2 Paper II Mannanases, as central enzymes in the degradation or modification of mannan polysaccharides, are of biotechnological interest for both basic and applied research. Endo-βmannanase genes belonging to GH5_7 have been identified in Arabidopsis thaliana, a small flowering plant that is the premiere model organism for plant biology. But so far no enzymology study of the gene products has been reported. Based on a plant GH5_7 phylogeny, four Arabidopsis enzymes distributed across the phylogenetic tree were selected for characterization. Thus, as a first step to increase the understanding of the function of Arabidopsis mannanases, one of these identified targets, AtMan5-1, was cloned, the recombinant protein was expressed and the enzymatic property was investigated. The recombinant AtMan5-1 was successfully expressed in E. coli and P. pastoris. Results from enzyme activity measurements using the DNS assay and additional HPLC product analysis, indicated that the enzyme efficiently degraded carob galactomannan, konjac glucomannan and spruce galactoglucomannan as well as mannopentose. However, AtMan5-1 32 was less adapted to depolymerize guar gum, probably due to the high number of galactose side chains found in guar gum. There was no indication of transglycosylation activity from the product analysis. Although recombinant AtMan5-1 was expressed in two different hosts, the obtained temperature and pH optima were similar for both AtMan5-1 variants. Like LeMAN4a and GmMAN1, the pH optimum of AtMan5-1 is close to 5 [80, 86]. One significant difference was nevertheless observed: the recombinant protein expressed in P. pastoris (AtMan5-1p) was more stable at higher temperatures compared to the E. coliproduced (AtMan5-1e) enzyme. Since the ATMAN5-1p protein was glycosylated, and glycosylation is known to influence protein stability, it seems likely that this modification contributed to the higher stability of that particular protein. Finally, the kinetic analysis revealed that the catalytic efficiency of AtMan5-1e was a slightly higher than that of AtMan51p. 33 2.4 Concluding remarks Paper I presented in this thesis describes a global bioinformatic analysis of GH5 family, which allowed the assignment of 4/5 of all analyzed family members to either one of the 43 newly defined or the 8 previously described subfamilies. The introduction of this additional hierarchical level of GH5 enzyme classification combined with biochemical data provided: (1) novel evolutionary insights of this family; (2) improved functional prediction; and (3) an opportunity to highlight interesting targets in unexplored subfamilies. The outcome of this work is available to the scientific community at the CAZy website (www.cazy.org). The analysis in Paper I revealed that for an overwhelming majority of GH5 carbohydrases enzyme activities have not been determined. The proportion of characterized plant GH5 enzymes is even less. Thus, to achieve a detailed understanding of these enzymes “hardcore”, biochemical characterization is sorely needed, especially for plant GH5 enzymes. The investigation of the catalytic properties of an Arabidopsis mannanase belonging to GH5 subfamily 7 presented in Paper II is an example of such a crucial study – the first report of a characterized Arabidopsis mannanase. The enzyme AtMan5-1 was heterologously expressed in both E. coli and P. pastoris, and was shown to efficiently hydrolyze most tested mannan substrates. The analysis also included kinetic constants determination, enzyme stability investigation and studies of pH, temperature and metal ion effect on enzyme activity. The next objective will be to compare the catalytic properties of AtMan5-1 with additional selected Arabidopsis mannanases. 34 3 Acknowledgements First of all, I would special thank Prof. Harry Brumer and Assoc. Prof. Ines Ezcurra as my main supervisors and Prof. Vincent Bulone, for making it possible to accept me becoming a member of Glycoscience. I would like to thank my co-supervisor Dr. Henrik Aspeborg for guiding me into the magic world of science and giving me the opportunity as a Ph.D. student in his research group. Although I missed the time for announcement, you know how exciting and feeling lucky I am! Because that is my dream when I was a little girl and it never changed with the time going on. And Dr. Oliver Spadiut as my former supervisor, for organizing my diploma work and nicely teach me how to develop as a young researcher. Many thanks to Dr. Gustav Sundqvist, Dr. Vaibhav Srivastava, Dr. Chunlin Xu and Dr. Francisco Vilaplana for giving the technique support. With the help from you, the work becomes much smoother and easier. Thanks very much to Assoc. Prof. Ines Ezcurra and Dr. Laura Grenville-Briggs for kindly helping with the improvement of my thesis and Dr. Erika Groth for the translation of abstract into Swedish! Thanks to all my colleagues and friends at plan 2, present and past, for helping me in the lab and creating a greatly active place for development. Last but not least, I would like to thank my family, especially my grandparents, my parents and Zhihui Chen. Your loves are the fountainhead for giving me power and confidence. 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