Download Exploring glycoside hydrolase family 5 (GH5) enzymes

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. No
matter what decision I make, you always support me at the first time and guide me how to
make it better. 谢谢亲爱的家人， 我爱你们！
35
4 References
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
16.
17.
18.
19.
20.
21.
22.
36
Carpita NC, Gibeaut DM: Structural models of primary cell walls in flowering plants:
consistency of molecular structure with the physical properties of the walls during growth.
The Plant journal : for cell and molecular biology 1993, 3(1):1-30.
Popper ZA, Fry SC: Xyloglucan-pectin linkages are formed intra-protoplasmically, contribute
to wall-assembly, and remain stable in the cell wall. Planta 2008, 227(4):781-794.
Roppolo D, De Rybel B, Tendon VD, Pfister A, Alassimone J, Vermeer JE, Yamazaki M, Stierhof
YD, Beeckman T, Geldner N: A novel protein family mediates Casparian strip formation in
the endodermis. Nature 2011, 473(7347):380-383.
Leroux O: Collenchyma: a versatile mechanical tissue with dynamic cell walls. Annals of
botany 2012, 110(6):1083-1098.
Fry SC: Primary cell wall metabolism: tracking the careers of wall polymers in living plant
cells. New Phytol 2004, 161(3):641-675.
McCann MC, Carpita NC: Designing the deconstruction of plant cell walls. Current opinion in
plant biology 2008, 11(3):314-320.
Silva GB, Ionashiro M, Carrara TB, Crivellari AC, Tine MAS, Prado J, Carpita NC, Buckeridge MS:
Cell wall polysaccharides from fern leaves: Evidence for a mannan-rich Type III cell wall in
Adiantum raddianum. Phytochemistry 2011, 72(18):2352-2360.
Harris PJ, Smith BG: Plant cell walls and cell-wall polysaccharides: structures, properties and
uses in food products. Int J Food Sci Tech 2006, 41:129-143.
McCann M, Roberts K: Architecture of the primary cell wall. The cytoskeletal basis of plant
growth and form 1991:109-129.
Timell TE: Compression wood in Gymnosperms. Volume 3. Ecology of compression wood
formation, silviculture and compression wood, mechanism of compression wood action,
compression wood in the lumber and pulp and paper industries, compression wood
induced by the balsam woolly aphid, opposite wood: Springer-Verlag; 1986.
Plomion C, Leprovost G, Stokes A: Wood formation in trees. Plant physiology 2001,
127(4):1513-1523.
Pauly M, Keegstra K: Plant cell wall polymers as precursors for biofuels. Current opinion in
plant biology 2010, 13(3):305-312.
Wang HZ, Dixon RA: On-off switches for secondary cell wall biosynthesis. Molecular plant
2012, 5(2):297-303.
Taiz L, Zeiger E: Plant Physiology . 2002. Massachusetts: Sinauer Associates Inc Publishers
2002.
O’Neill MA, York WS: The composition and structure of plant primary cell walls. The plant
cell wall 2003:1-54.
Vogel J: Unique aspects of the grass cell wall. Current opinion in plant biology 2008,
11(3):301-307.
Timell TE: Recent progress in the chemistry of wood hemicelluloses. Wood Sci Technol 1967,
1(1):45-70.
Somerville C: Cellulose synthesis in higher plants. Annual review of cell and developmental
biology 2006, 22:53-78.
Mccann MC, Wells B, Roberts K: Direct Visualization of Cross-Links in the Primary Plant-Cell
Wall. J Cell Sci 1990, 96:323-334.
Preston RD: The physical biology of plant cell walls: London.: Chapman & Hall; 1974.
Brett C, Waldron K: The molecular components of the wall. Physiology and Biochemistry of
Plant Cell Walls 1996:4-43.
Pauly M, Keegstra K: Cell-wall carbohydrates and their modification as a resource for
biofuels. The Plant journal : for cell and molecular biology 2008, 54(4):559-568.
23.
24.
25.
26.
27.
28.
29.
30.
31.
32.
33.
34.
35.
36.
37.
38.
39.
40.
41.
42.
43.
44.
45.
Southgate DA, Branch WJ, Hill MJ, Drasar BS, Walters RL, Davies PS, Baird IM: Metabolic
responses to dietary supplements of bran. Metabolism: clinical and experimental 1976,
25(10):1129-1135.
Zhang GF, Staehelin LA: Functional compartmentation of the Golgi apparatus of plant cells :
immunocytochemical analysis of high-pressure frozen- and freeze-substituted sycamore
maple suspension culture cells. Plant physiology 1992, 99(3):1070-1083.
Waldron K, Faulds C: Cell wall polysaccharides: composition and structure. Comprehensive
glycoscience: analysis of glycans/polysaccharide functional properties 2007, 2:181-201.
Piens K, Henriksson AM, Gullfot F, Lopez M, Faure R, Ibatullin FM, Teeri TT, Driguez H,
Brumer H: Glycosynthase activity of hybrid aspen xyloglucan endo-transglycosylase
PttXET16-34 nucleophile mutants. Organic & biomolecular chemistry 2007, 5(24):3971-3978.
Darley CP, Forrester AM, McQueen-Mason SJ: The molecular basis of plant cell wall
extension. Plant molecular biology 2001, 47(1-2):179-195.
Buchanan BB, Gruissem W, Jones RL: Biochemistry & molecular biology of plants: American
Society of Plant Physiologists Waldorf^ eMD MD; 2000.
O'Neill M, Albersheim P, Darvill A: The pectic polysaccharides of primary cell walls. Methods
in plant biochemistry 1990, 2:415-441.
Visser J, Voragen A: Progress in Biotechnology 14: Pectins and Pectinases. In: Proceedings of
an International Symposium, Wageningen Elsevier, Amsterdam Science BV, Amsterdam: 1996.
Ishii T: O-acetylated oligosaccharides from pectins of potato tuber cell walls. Plant
physiology 1997, 113(4):1265-1272.
Komalavilas P, Mort AJ: The Acetylation at O-3 of Galacturonic Acid in the Rhamnose-Rich
Portion of Pectins. Carbohydrate research 1989, 189:261-272.
Li X, Chapple C: Understanding Lignification: Challenges Beyond Monolignol Biosynthesis.
Plant physiology 2010, 154(2):449-452.
Grima-Pettenati J, Goffner D: Lignin genetic engineering revisited. Plant Sci 1999, 145(2):5165.
Asp NG: Dietary Fiber - Definition, Chemistry and Analytical Determination. Mol Aspects
Med 1987, 9(1):17-29.
Showalter AM: Introduction: plant cell wall proteins. Cell Mol Life Sci 2001, 58(10):13611362.
Showalter AM: Structure and Function of Plant-Cell Wall Proteins. The Plant cell 1993,
5(1):9-23.
Ringli C, Keller B, Ryser U: Glycine-rich proteins as structural components of plant cell walls.
Cell Mol Life Sci 2001, 58(10):1430-1441.
Vaaje-Kolstad G, Westereng B, Horn SJ, Liu ZL, Zhai H, Sorlie M, Eijsink VGH: An Oxidative
Enzyme Boosting the Enzymatic Conversion of Recalcitrant Polysaccharides. Science 2010,
330(6001):219-222.
Meier H, Reid J: Reserve polysaccharides other than starch in higher plants. In: Plant
Carbohydrates I. Springer; 1982: 418-471.
Matheson NK: Mannose-based polysaccharides. Methods in plant biochemistry 1990, 2:371413.
Buckeridge MS: Seed cell wall storage polysaccharides: models to understand cell wall
biosynthesis and degradation. Plant physiology 2010, 154(3):1017-1023.
Rodriguez-Gacio Mdel C, Iglesias-Fernandez R, Carbonero P, Matilla AJ: Softening-up
mannan-rich cell walls. J Exp Bot 2012, 63(11):3976-3988.
Rowell RM, Pettersen R, Han JS, Rowell JS, Tshabalala MA: Cell wall chemistry. Handbook of
wood chemistry and wood composites 2005:35-74.
Benova-Kakosova A, Digonnet C, Goubet F, Ranocha P, Jauneau A, Pesquet E, Barbier O,
Zhang ZN, Capek P, Dupree P et al: Galactoglucomannans increase cell population density
and alter the protoxylem/metaxylem tracheary element ratio in xylogenic cultures of
zinnia. Plant physiology 2006, 142(2):696-709.
37
46.
47.
48.
49.
50.
51.
52.
53.
54.
55.
56.
57.
58.
59.
60.
61.
62.
63.
64.
65.
66.
67.
68.
38
Whitney SEC, Brigham JE, Darke AH, Reid JSG, Gidley MJ: Structural aspects of the
interaction of mannan-based polysaccharides with bacterial cellulose. Carbohydrate
research 1998, 307(3-4):299-309.
Handford MG, Baldwin TC, Goubet F, Prime TA, Miles J, Yu X, Dupree P: Localisation and
characterisation of cell wall mannan polysaccharides in Arabidopsis thaliana. Planta 2003,
218(1):27-36.
Maier H, Anderson M, Karl C, Magnuson K, Whistler RL: Guar, locust bean, tara, and
fenugreek gums: Academic Press: New York; 1993.
Wolfrom M, Laver M, Patin D: Carbohydrates of the Coffee Bean. II. Isolation and
Characterization of a Mannan1. The Journal of Organic Chemistry 1961, 26(11):4533-4535.
Hopf H, Kandler O: Characterization of Reserve Cellulose of Endosperm of Carum-Carvi Asa
Beta-(1-4)-Mannan. Phytochemistry 1977, 16(11):1715-1717.
Reid J, Edwards M: Galactomannans and other cell wall storage polysaccharides in seeds.
FOOD SCIENCE AND TECHNOLOGY-NEW YORK-MARCEL DEKKER- 1995:155-155.
Aspinall G, Hirst E, Percival E, Williamson I: 635. The mannans of ivory nut (Phytelephas
macrocarpa). Part I. The methylation of mannan A and mannan B. J Chem Soc 1953:31843188.
Meier H: On the structure of cell walls and cell wall mannans from ivory nuts and from
dates. Biochimica et biophysica acta 1958, 28:229-240.
Charrier M, Rouland C: Mannan-degrading enzymes purified from the crop of the brown
garden snail Helix aspersa Muller (Gastropoda Pulmonata). The Journal of experimental
zoology 2001, 290(2):125-135.
Chanzy HD, Grosrenaud A, Vuong R, Mackie W: The Crystalline Polymorphism of Mannan in
Plant-Cell Walls and after Recrystallization. Planta 1984, 161(4):320-329.
Chanzy H, Perez S, Miller DP, Paradossi G, Winter WT: An Electron-Diffraction Study of
Mannan .1. Crystal and Molecular-Structure. Macromolecules 1987, 20(10):2407-2413.
Mackie W, Sellen D: The degree of polymerization and polydispersity of mannan from the
cell wall of the green seaweed< i> codium fragile</i>. Polymer 1969, 10:621-632.
Millane RP, Hendrixson TL: Crystal-Structures of Mannan and Glucomannans. Carbohyd
Polym 1994, 25(4):245-251.
Mccleary BV, Clark AH, Dea ICM, Rees DA: The Fine-Structures of Carob and Guar
Galactomannans. Carbohydrate research 1985, 139(Jun):237-260.
Reid J: Galactomannans. Biochemistry of storage carbohydrates in green plants 1985:265288.
McCleary B, Matheson N: Galactomannan structure and β-mannanase and β-mannosidase
activity in germinating legume seeds. Phytochemistry 1975, 14(5):1187-1194.
Dea IC: Conformational origins of polysaccharide solution and gel properties. Industrial
Gums 1993:21-52.
Nishinari K, Williams PA, Phillips GO: Review of the Physicochemical Characteristics and
Properties of Konjac Mannan. Food Hydrocolloids 1992, 6(2):199-222.
Dey PM, Dixon R: Biochemistry of storage carbohydrates in green plants: Academic Press;
1985.
Maekaji K: Relationship between Degree of Deacetylation and Gelation of Konjac Mannan Kinetic Study on Gelation of Konjac Mannan .3. J Agr Chem Soc Jpn 1978, 52(11):513-517.
Penroj P, Mitchell JR, Hill SE, Ganjanagunchorn W: Effect of konjac glucomannan
deacetylation on the properties of gels formed from mixtures of kappa carrageenan and
konjac glucomannan. Carbohyd Polym 2005, 59(3):367-376.
Willför S, Sundberg A, Hemming J, Holmbom B: Polysaccharides in some industrially
important softwood species. Wood Sci Technol 2005, 39(4):245-257.
Sundberg A, Holmbom B, Willfor S, Pranovich A: Weakening of paper strength by wood resin.
Nord Pulp Pap Res J 2000, 15(1):46-53.
69.
70.
71.
72.
73.
74.
75.
76.
77.
78.
79.
80.
81.
82.
83.
84.
85.
86.
Hannuksela T, du Penhoat CH: NMR structural determination of dissolved O-acetylated
galactoglucomannan isolated from spruce thermomechanical pulp. Carbohydrate research
2004, 339(2):301-312.
Hannuksela T, Holmbom B, Mortha G, Lachenal D: Effect of sorbed galactoglucomannans
and galactomannans on pulp and paper handsheet properties, especially strength
properties. Nord Pulp Pap Res J 2004, 19(2):237-244.
Willför S, Sundberg K, Tenkanen M, Holmbom B: Spruce-derived mannans–A potential raw
material for hydrocolloids and novel advanced natural materials. Carbohyd Polym 2008,
72(2):197-210.
Cartier N, Chambat G, Joseleau JP: Cell-Wall and Extracellular Galactoglucomannans from
Suspension-Cultured Rubus-Fruticosus Cells. Phytochemistry 1988, 27(5):1361-1364.
Bailey R, Pain V: Polysaccharide mannose in New Zealand ferns. Phytochemistry 1971,
10(5):1065-1073.
Tailford LE, Ducros VM, Flint JE, Roberts SM, Morland C, Zechel DL, Smith N, Bjornvad ME,
Borchert TV, Wilson KS et al: Understanding how diverse beta-mannanases recognize
heterogeneous substrates. Biochemistry 2009, 48(29):7009-7018.
Van Zyl WH, Rose SH, Trollope K, Görgens JF: Fungal β-mannanases: Mannan hydrolysis,
heterologous production and biotechnological applications. Process Biochemistry 2010,
45(8):1203-1213.
Filichkin SA, Leonard JM, Monteros A, Liu PP, Nonogaki H: A novel endo-beta-mannanase
gene in tomato LeMAN5 is associated with anther and pollen development. Plant
physiology 2004, 134(3):1080-1087.
Xu B, Hagglund P, Stalbrand H, Janson JC: endo-beta-1,4-Mannanases from blue mussel,
Mytilus edulis: purification, characterization, and mode of action. Journal of biotechnology
2002, 92(3):267-277.
Iglesias-Fernandez R, Rodriguez-Gacio MC, Barrero-Sicilia C, Carbonero P, Matilla A: Three
endo-beta-mannanase genes expressed in the micropylar endosperm and in the radicle
influence germination of Arabidopsis thaliana seeds. Planta 2011, 233(1):25-36.
Nonogaki H, Morohashi Y: An endo-beta-mannanase develops exclusively in the micropylar
endosperm of tomato seeds prior to radicle emergence. Plant physiology 1996, 110(2):555559.
Lin JY, Pantalone VR, Li GL, Chen F: Molecular Cloning and Biochemical Characterization of
an Endo-beta-mannanase Gene from Soybean for Soybean Meal Improvement. J Agr Food
Chem 2011, 59(9):4622-4628.
Bewley JD, Banik M, Bourgault R, Feurtado JA, Toorop P, Hilhorst HW: Endo-beta-mannanase
activity increases in the skin and outer pericarp of tomato fruits during ripening. J Exp Bot
2000, 51(344):529-538.
Brummell DA, Dal Cin V, Crisosto CH, Labavitch JM: Cell wall metabolism during maturation,
ripening and senescence of peach fruit. J Exp Bot 2004, 55(405):2029-2039.
Cantarel BL, Coutinho PM, Rancurel C, Bernard T, Lombard V, Henrissat B: The CarbohydrateActive EnZymes database (CAZy): an expert resource for Glycogenomics. Nucleic acids
research 2009, 37(Database issue):D233-238.
Duffaud GD, McCutchen CM, Leduc P, Parker KN, Kelly RM: Purification and characterization
of extremely thermostable beta-mannanase, beta-mannosidase, and alpha-galactosidase
from the hyperthermophilic eubacterium Thermotoga neapolitana 5068. Applied and
environmental microbiology 1997, 63(1):169-177.
Bourgault R, Oakley AJ, Bewley JD, Wilce MC: Three-dimensional structure of (1,4)-beta-Dmannan mannanohydrolase from tomato fruit. Protein science : a publication of the Protein
Society 2005, 14(5):1233-1241.
Schroder R, Wegrzyn TF, Sharma NN, Atkinson RG: LeMAN4 endo-beta-mannanase from
ripe tomato fruit can act as a mannan transglycosylase or hydrolase. Planta 2006,
224(5):1091-1102.
39
87.
88.
89.
90.
91.
92.
93.
94.
95.
96.
97.
98.
99.
100.
101.
102.
103.
104.
40
Davies GJ, Wilson KS, Henrissat B: Nomenclature for sugar-binding subsites in glycosyl
hydrolases. The Biochemical journal 1997, 321 ( Pt 2):557-559.
Le Nours J, Anderson L, Stoll D, Stålbrand H, Leggio LL: The structure and characterization of
a modular endo-β-1, 4-mannanase from Cellulomonas fimi. Biochemistry 2005,
44(38):12700-12708.
Zhang YL, Ju JS, Peng H, Gao F, Zhou C, Zeng Y, Xue YF, Li Y, Henrissat B, Gao GF et al:
Biochemical and Structural Characterization of the Intracellular Mannanase AaManA of
Alicyclobacillus acidocaldarius Reveals a Novel Glycoside Hydrolase Family Belonging to
Clan GH-A. Journal of Biological Chemistry 2008, 283(46):31551-31558.
Hilge M, Gloor SM, Rypniewski W, Sauer O, Heightman TD, Zimmermann W, Winterhalter K,
Piontek K: High-resolution native and complex structures of thermostable beta-mannanase
from Thermomonospora fusca - substrate specificity in glycosyl hydrolase family 5.
Structure 1998, 6(11):1433-1444.
Hogg D, Woo EJ, Bolam DN, McKie VA, Gilbert HJ, Pickersgill RW: Crystal structure of
mannanase 26A from Pseudomonas cellulosa and analysis of residues involved in substrate
binding. Journal of Biological Chemistry 2001, 276(33):31186-31192.
Aspeborg H, Coutinho PM, Wang Y, Brumer H, Henrissat B: Evolution, substrate specificity
and subfamily classification of glycoside hydrolase family 5 (GH5). Bmc Evol Biol 2012, 12.
Sinnott ML: Catalytic Mechanisms of Enzymatic Glycosyl Transfer. Chem Rev 1990,
90(7):1171-1202.
Zechel DL, Withers SG: Glycosidase mechanisms: Anatomy of a finely tuned catalyst.
Accounts Chem Res 2000, 33(1):11-18.
Yuan JS, Yang XH, Lai JR, Lin H, Cheng ZM, Nonogaki H, Chen F: The endo-beta-mannanase
gene families in Arabidopsis, rice, and poplar. Funct Integr Genomic 2007, 7(1):1-16.
Jenkins J, Leggio LL, Harris G, Pickersgill R: Beta-Glucosidase, Beta-Galactosidase, Family-a
Cellulases, Family-F Xylanases and 2 Barley Glycanases Form a Superfamily of Enzymes with
8-Fold Beta/Alpha-Architecture and with 2 Conserved Glutamates near the CarboxyTerminal Ends of Beta-Strand-4 and Beta-Strand-7. Febs Lett 1995, 362(3):281-285.
Henrissat B, Callebaut I, Fabrega S, Lehn P, Mornon JP, Davies G: Conserved catalytic
machinery and the prediction of a common fold for several families of glycosyl hydrolases
(vol 92, pg 7090, 1995). Proceedings of the National Academy of Sciences of the United States
of America 1996, 93(17):9302-9302.
Rosengren A, Hagglund P, Anderson L, Pavon-Orozco P, Peterson-Wulff R, Nerinckx W,
Stalbrand H: The role of subsite +2 of the Trichoderma reesei beta-mannanase TrMan5A in
hydrolysis and transglycosylation. Biocatal Biotransfor 2012, 30(3):338-352.
Rye CS, Withers SG: Glycosidase mechanisms. Current opinion in chemical biology 2000,
4(5):573-580.
Dominguez R, Souchon H, Spinelli S, Dauter Z, Wilson KS, Chauvaux S, Beguin P, Alzari PM: A
Common Protein Fold and Similar Active-Site in 2 Distinct Families of Beta-Glycanases. Nat
Struct Biol 1995, 2(7):569-576.
Boraston AB, Bolam DN, Gilbert HJ, Davies GJ: Carbohydrate-binding modules: fine-tuning
polysaccharide recognition. The Biochemical journal 2004, 382(Pt 3):769-781.
Pokoj S, Lauer I, Fotisch K, Himly M, Mari A, Enrique E, Miguel-Moncin Mdel M, Lidholm J,
Vieths S, Scheurer S: Pichia pastoris is superior to E. coli for the production of recombinant
allergenic non-specific lipid-transfer proteins. Protein expression and purification 2010,
69(1):68-75.
Sumner JB: A more specific reagent for the determination of sugar in urine. Journal of
biological chemistry 1925, 65(2):393-395.
Lo Leggio L, Larsen S: The 1.62 A structure of Thermoascus aurantiacus endoglucanase:
completing the structural picture of subfamilies in glycoside hydrolase family 5. Febs Lett
2002, 523(1-3):103-108.
105.
106.
107.
108.
Tanabe T, Morinaga K, Fukamizo T, Mitsutomi M: Novel chitosanase from Streptomyces
griseus HUT 6037 with transglycosylation activity. Bioscience, biotechnology, and
biochemistry 2003, 67(2):354-364.
Zverlov VV, Velikodvorskaya GA, Schwarz WH: A newly described cellulosomal
cellobiohydrolase, CelO, from Clostridium thermocellum: investigation of the exo-mode of
hydrolysis, and binding capacity to crystalline cellulose. Microbiology 2002, 148(Pt 1):247255.
Tanaka M, Umemoto Y, Okamura H, Nakano D, Tamaru Y, Araki T: Cloning and
Characterization of a beta-1,4-Mannanase 5C Possessing a Family 27 Carbohydrate-Binding
Module from a Marine Bacterium, Vibrio sp Strain MA-138. Biosci Biotech Bioch 2009,
73(1):109-116.
Dias FM, Vincent F, Pell G, Prates JA, Centeno MS, Tailford LE, Ferreira LM, Fontes CM, Davies
GJ, Gilbert HJ: Insights into the molecular determinants of substrate specificity in glycoside
hydrolase family 5 revealed by the crystal structure and kinetics of Cellvibrio mixtus
mannosidase 5A. The Journal of biological chemistry 2004, 279(24):25517-25526.
41