University of Groningen
Unique features of several microbial α-amylases active on soluble and native starch Sarian, Fean Davisunjaya
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Unique features of several microbial α‐amylases active on
soluble and native starch
Fean Davisunjaya Sarian
The work described in this thesis was performed in the research group of the Microbial Physiology of the Groningen Biomolecular Sciences and Biotechnology Institute (GBB), the Aquatic Biotechnology and Bioprocessing Engineering of the Engineering and Technology Institute Groningen (ENTEG) of the University of Groningen, The Netherlands, and the Biochemistry Division, Institut Teknologi Banding Indonesia. This work was financially supported by a Bernoulli scholarship and by Asahi Glass Foundation.
Cover design and layout of the book: F. D. Sarian
Cover front: art of α‐amylase with glucose and maltooligosacharides
Cover back: art of glucose and maltooligosaccharides
Printed by: Ipskamp Printing
ISBN: 978-90-367-8850-2
ISBN: 978-90-367-8849-6 (electronic version)
Copyright © 2016 by Fean Davisunjaya Sarian
All right reserved. No part of this thesis may be reproduced, stored in a retrieval system or transmitted in any form by any means, without permission from the author.
Unique features of several microbial α-amylases active on soluble and native starch
PhD thesis
to obtain the degree of PhD at the University of Groningen on the authority of the Rector Magnificus Prof. E. Sterken and in accordance with the decision by the College of Deans.
This thesis will be defended in public on
Friday 3 June 2016 at 09.00 hours
by
Fean Davisunjaya Sarian
born on 31 July 1981 in Yogyakarta, Indonesia
Supervisors Prof. M.J.E.C. van der Maarel Prof. L. Dijkhuizen
Assessment Committee Prof. G.J.W. Euverink Prof. M.W. Fraaije Prof. B. Svensson Table of Contents
Chapter 1 Introduction ...... 1
Chapter 2 Cold enzymatic hydrolysis of Indonesian roots
and tuber starches ...... 81
Chapter 3 Enzymatic degradation of granular starch
By Microbacterium aurum strain B8.A ...... 99
Chapter 4 Degradation of granular starch by the bacterium Microbacterium aurum B8.A involves a novel modular α‐amylase enzyme system with FnIII and
CBM25 domains ...... 125
Chapter 5 Identification of a novel α‐amylase from marine Bacillus megaterium NL3 representing a new
GH13 subfamily ...... 159
Chapter 6 General Discussion ...... 191
Summary non‐expert ...... 203
Samenvatting ...... 209
Acknowledgments ...... vii
List of publications ...... xi
Chapter
Introduction
Introduction
What is Starch?
Starch is next to cellulose and chitin, the most abundant biopolymer in nature. Plants
Page | 2 utilize the light energy absorbed by chlorophyll to produce simple sugars and oxygen from carbon dioxide and water through a process called photosynthesis. These simple sugars may be used directly in cellular respiration, converted into starch, or used to build cell walls. The starch serves as major energy storage for plants, which can later be converted back to glucose and used in respiration when required. Starch is densely packed in microscopic particles called granules and is extremely insoluble in water at room temperature. The composition, size and shape of these granules are often characteristic for the plant species from which they are extracted (1). Structurally, starch is made up of individual units of glucose, linked together by α(14) and occasional α(16)‐glycosidic linkages (2). Amylose, the minor component, is a relatively long and linear molecule with almost exclusively
α(14)‐glycosidic bonds with number average degree of polymerization (DPn) of 900‐3300 (3). It has a slight degree of branching of 9‐20 branches per molecule (4,
5). Amylopectin is the major component of most starches and has a DPn of 4800‐ 15900 with a molecular mass that is about 5 times bigger than amylose (3). It consists of short α(14) linked chains while branching by α(16)‐glycosidic bonds occurs every 15‐45 glucosyl units (5, 6). Within the starch granule, amylopectin molecules are ordered radially with the ends of non‐reducing chains pointing toward the outer surface (7). Although the exact molecular architecture is still not clear, its structure is often described by a cluster model (Fig. 1.1) (4, 8). Hizukuri (9) and Kobayashi et al. (10) have shown that an amylopectin molecule contains several types of chains (A, B, and C) which differ in their chain length and are present in almost equal proportions. The A‐chains (unbranched) are the shortest chains, linked to B chains and do not carry any other chains, whereas the B chains (B1‐B4, depending on their length and the number of clusters the chain passes through), connect to one or more A chains and/or B chains. B1 chains are short chains that span only one cluster, while B2, B3, and B4 chains are long chains that connect two to four cluster chains in the amylopectin molecule (9). The C‐chain contains the reducing end group of the molecule (11).
Chapter 1
Page | 3
Fig 1.1 Schematic representation of the cluster model of amylopectin. For definition of A, B1‐B2, and C, see text. Straight line, α(14) glycosidic bonds; dashed line, α(16) glycosidic bonds. Adapted from Tester et al. (12), with modifications.
The relative content of amylose and amylopectin varies widely, depending on the botanical origin (Table 1.1). The amylose content of regular starch is typically from 10 to 35% of the total starch. High‐amylose starches contain approximately 35‐70% amylose, while ‘waxy’ (high‐amylopectin) starches have almost no amylose (0% to 4%) (13). Functional properties of starch, such as water‐binding capacity, gelatinization and pasting, retrogradation, and susceptibility to enzymatic degradation are influenced by the amylose content and the placement of the amylose in the amylopectin (14). Both amylose and amylopectin are packed together in complex structures consisting of crystalline and amorphous areas and contain a few phosphate groups mainly at the C‐6 of the glucose residues (15). Phosphorus, present as phosphate monoester and phospholipids, at low amounts significantly affects the functional properties of the starch (16). Apart from these main components, smaller amounts of other components, such as proteins and lipids may also be present in amounts that vary depending on the botanical origin and starch isolation procedure (3).
Introduction
Structure and properties of starch
Native starches are semi‐crystalline structures. The degree of crystallinity of native
Page | 4 starch varies between 15 – 45% depending on the starch source (6, 17). Observed by light microscopy or scanning electron microscopy (after a slight acid or enzyme treatment), starch granules are formed by the deposition of growth rings that consist of alternating semi‐crystalline and amorphous layers (18–20). Irrespective of the source of the starch, the repeat distance of alternating amorphous and crystalline layers is highly conserved at 9 nm (21–23) (Fig. 1.1). The starch granule organization is very complex and depends strongly on the botanical origin.
A schematic representation of the granule’s architecture is given in Fig. 1.2. The outer branches of the amylopectin molecule form double helices that are arranged in crystalline regions. The amorphous part of the starch is believed to consist of amylose that forms single helical structures near the branch points of the amylopectin molecule (24). Jenkins and Donald (21) hypothesized that amylose is predominantly located in the amorphous growth ring based on their investigation of the amylose content of corn, barley and pea. This results in the formation of amorphous and crystalline lamellae arranged in an overall semi crystalline structure. X‐ray diffractometry has been used to reveal the presence and the characteristics of the crystalline structure of starch granules (15). There are three diffraction patterns for starches, the A, B, and C pattern, that explain the difference of packing of the amylopectin double helices (25). The type A pattern is generally found in cereal starches, therefore called the cereal type, while most of tuber and root starches exhibit the typical B type X‐ray pattern (3). A and B patterns differ from each other by the double helix packing arrangement in the unit cells as well as in the amount of associated water (Fig. 1.3). Starch granules of the A‐type are formed from very compactly arranged double helices and shorter average branch chain length of amylopectin. In contrast, the B‐type starches have more open structures with hexagonal channel arrangement that are filled with water molecules and contain longer average branch chain lengths (26, 27). Some of the rhizome and legume starches have the C‐type pattern which is a mixture of A and B types in
Chapter 1
varying proportions rather than a distinct crystalline structure (3, 28). Apart from the chain length (26, 28, 29), other factors are known to influence the crystallinity type of starches: temperature (30), concentration of starch solution, salts (31), and organic molecules (32). Page | 5
Amorphous
Crystalline
Amorphous 9 nm
Crystalline 9 nm Amorphous Granule diameter: Growth ring diameter: 0.5 – 110 µm 120 – 400 nm Cluster/ starch chain
Monomer: 0.55 nm Fig 1.2 Schematic view of the different structural levels within a starch granule. The cluster structure of amylopectin forms ordered double helices in regular arrays to give crystalline lamellae. Crystalline lamellae are separated by amorphous lamellae, giving rise to semi‐crystalline zones. Crystalline‐amorphous repeats, with a periodicity of 9 nm, are called ‘growth rings’ and are organized into spherical blocklets to form granules. Adapted from Buleon et al. (6), with modifications.
Starch granules are synthesized in various plant tissues, such as leaves (33–35), fruits (banana) (36), seeds such as those of corn, soybean (37), and pea (38), different types of stems (sago), roots and tubers (cassava, potato, sweet potato), and even pollen (39, 40). Starches isolated from different botanical sources display a characteristic granule morphology (41). The diameter of starch granules varies depending on the biological origin, and ranges from submicron, such as soybean, to more than 100 µm for edible canna and potato (Table 1.1). In addition, the shape Introduction
ranges from spheres, lenticulars (disks), irregular tubules, polygonals, to ovals (42). Starch granules occur individually or as clusters, and no two granules are identical (5, 43). Potato starch granules have been observed to be both oval and spherical in Page | 6 shape (15). Corn starches are spherical and polygonal in shape (44). Wheat, barley, and rye starches have bimodal size distributions, while rice starches have a unimodal size distribution with a polygonal‐shape (5). Almost all legume starches are oval shape, although spherical, round, and irregular shaped granules have also been found (45). The characteristics attributed to each starch are presented in detail in Table 1.1.
Fig 1.3 Double helix packing arrangement in the A‐ (left) and B‐type (right) of starch. Mixtures of A and B are designated C‐type. Each circle represents a view down the z‐axis of a double helix. In the structure of A‐type starch, for each unit cell, four water molecules (diamond shape) are located between the helices. Adapted from Sarko et al. (25) and Buleon et al. (46), with modifications.
Chapter 1
Table 1.1 Morphology and composition of native starch granules from various sources.
Starch Type Amylose (%) Shape X‐ray Size (µm) Page | 7 diffraction Arrowroot Root 19 – 21 (47) Spherical A 8 – 42 (47) Corn Cereal 22 – 32 (48) Spherical/ A 3 – 23 (49) Polygonal Rice Cereal 1 – 8 (50) Polygonal A 3 – 10 (50) Wheat Cereal 18 – 30 (51) Spherical A 22 – 26 (51) Edible canna Root 28 – 38 (17, Spherical B 10 – 101 (41, 49) 49, 52, 53) Potato Root 18 – 28 (54, Oval/ B 15 – 110 (15, 55) Spherical 54, 55) Banana Fruit 16 – 40 (36) Irregular A, B, or C 20 – 60 (36) Cassava Root 17 – 33 Lenticular/ A or C 7 – 23 (47) Spherical Bean Legume 23 – 39 (56, Oval C 8 – 55 (56, 57) 57) Chickpea Legume 28 – 40 (58) Oval/ C 5 – 35 (58) Spherical Smooth pea Legume 33 – 49 (59) Oval C 2 – 40 (59) Soybean Legume 12 – 16 (37) Spherical/ C 0.7 – 4 (37) Polygonal Sweet potato Root 20 – 25 (47) Polygonal C 7 – 28 (47) Sago Pith 24 – 31 (60) Spherical C 20 – 40 (60) Leaves Leaf 16 – 20 (61) Lenticular n.d. n.d. (tobacco) Leaves Leaf 4 – 34 (62) Lentilcular n.d. 0.2 – 0.3 (62) (Arabidopsis) n.d., no data; (number), reference Introduction
A common feature of all starch granules is that they cannot be degraded directly, they must be hydrolyzed first and the enzymes involved in their degradation process require specific entry points on the structure of granules, in the form of either pores, Page | 8 channels, or zones of attack (63, 64). Some granules as observed by Scanning Electron Microscopy (SEM) and Atomic Force Microscopy (AFM) were found to have many pores, others a few, and some none (65). Pores were observed on the surface of some corn, sorghum, and millet starch granules. Large granules of wheat, barley, and rye starch also have pores along the equatorial groove of the granule (63). Those pores were randomly distributed over the surfaces of starch granules and varied in number per granule. No pores have been observed on granules from arrowroot, cassava, edible canna, or rice, while there is much disagreement as to the presence of pores on the surface of potato starch granules (66–69). The origin of these starch surface pores differs; some pores are created during starch accumulation in the plant and some are formed during thermal or hydrothermal processing when amylose migrates from the granule’s interior to the outer layer and surface (70). Other pores are formed as a result of mechanical damage or granule’s cracks during grain treatment (71, 72). The function of these pores is not well understood but they could be openings to channels that potentially provide direct enzyme access to the granule interior (63).
Starch Degradation
In nearly all types of higher plants, starch is accumulated inside plastids in two forms, namely the short‐term reserve starch, so‐called transitory starch, in photosynthetically active leaves, and the long‐term storage starch in amyloplasts of heterotrophic tissues such as roots, tubers, fruits, or endosperm. Whether both types of starches are developing a similar semi‐crystalline granule of complex structure, the physiological role of these starches is remarkably different (73). Transitory starch along with sucrose is synthesized in the leaves as the primary product of photosynthesis during the day. Transitory starch then accumulates in chloroplasts and sucrose is exported to the sites of growth. During the following night, the starch is degraded to provide a continued supply of carbohydrate for leaf
Chapter 1
cell metabolism and also to provide substrate for sucrose synthesis to allow continued export to non‐photosynthetic parts of the plants (74). In storage organs, starch synthesis occurs over many days or weeks during the development and maturation of the tissue (75). It is degraded rapidly during specific periodic or Page | 9 developmental signals, such as the beginning of spring, and the early ripening of most cultivars (76, 77).
In-vivo degradation of starch granules
Hydrolysis of starch to glucose is important for the maximal utilization of starch to provide energy for animals and plants. It is clear that pathways of starch degradation differ with the organs and species, and that distinct pathways may operate within the same organ (74, 78). In living cells, the process of starch degradation is best understood in leaves, where the transitory starch is degraded by respiration at night (35, 79), while the pathway in other starch containing tissue is poorly understood except in the case of germinating cereal grains (73). To investigate the impact of different day lengths on the amount of starch in leaves, many experiments have been performed in several plants either by growth in low light or in the dark for 24‐48 h (7, 80–84). Leaf starch is converted to maltodextrin by several enzymes, such as β‐amylase (EC 3.2.1.2) and debranching enzyme (EC 3.2.1.33), with maltose and glucose as the main products. These sugars are then exported to the cytosol, and later converted to sucrose that is used to provide energy to the cells or transported to other parts of the plant where it is converted to starch for long term storage (35). In leaves, the amount of starch accumulated during the entire photoperiod does not decrease, because during the night, the starch accumulated during the day is degraded at a relatively constant rate (78). The rate of degradation is regulated in a manner complementary to the rate of synthesis during the day (35).
Starch degradation in chloroplasts differs from that in amyloplasts of germinating cereal grains. Starch degradation is a regulated process and the release of soluble Introduction
α‐glucans from the semi‐crystalline starch granule is initiated by induction of the expression of the hydrolytic enzyme. In leaves, starch degradation starts with the addition of phosphate groups at the C‐6 and C‐3 positions of a small portion of the Page | 10 glucosyl residues by glucan‐water dikinase (GWD, EC 2.7.9.4) and phosphoglucan water dikinase (PWD, EC 2.7.9.4), respectively (85, 86). The phosphate levels in starch granules may disrupt the packing of the amylopectin at the granules surface in such a way as to allow it to become degraded by exo‐amylolytic enzymes (87). α‐Amylase (EC 3.2.1.1) has always been considered to be involved in the conversion of transitory starch to maltodextrin, but recent data indicate that α‐amylase is most likely not required for transitory starch degradation in Arabidopsis (Arabidopsis thaliana) leaves, as evidence for α‐amylase involvement in leaf starch catabolism is lacking (79, 88). Currently, it is considered that the hydrolysis of leaf starch granules is catalyzed mainly by β‐amylase and debranching enzyme (isoamylase 3, EC 3.2.1.68). The Arabidopsis genome contains nine genes encoding β‐amylase in which four of them are localized in the chloroplast (89, 90). Maltose and glucose accumulate during starch degradation, and are exported from the chloroplast to the cytosol. Either for sucrose synthesis, glycolysis, or the oxidative pentose phosphate pathway, conversion of transported maltose and glucose occurs later in the cytosol (79).
During germination of cereals, the solubilisation of starch is initiated by the action of α‐amylase, which is secreted to the endosperm at the early stage of germination (91). The degradation occurs following direct binding of α‐amylase to the starch granules in dead amyloplast cells. In other starch‐storing organs such as tubers and fruits there is no clear evidence for initial attack on the starch granule by α‐amylase. However, for cotyledons the consensus hypothesis is that new α‐amylase needs to be synthesized at the start of starch degradation (92, 93). Studies by Henson and coworkers (94) have suggested that α‐glucosidase (EC 3.2.1.20) is acting together with α‐amylase during degradation of barley grain and perhaps pea seed starch. The complete degradation of soluble starch then proceeds by the action of α‐ and β‐amylase, phosphorylase (EC 2.4.1.7), debranching enzymes (e.g., isoamylase and pullulanase, EC3.2.1.41), and α‐glucosidase. The glucose produced in the endosperm
Chapter 1
is taken up and converted to sucrose, then the sucrose is provided to the growing tissues, such as the young shoot and root tissues (79). Hormonal level and environmental conditions regulate the starch degradation processes. As an example changes in the level of gibberellins, a growth hormone, promotes starch degradation Page | 11 by stimulating the production of α‐amylase and other hydrolyzing enzymes needed for providing soluble sugars into the endosperm during germination and establishment of cereal seedling (95). The pathway of starch degradation is summarized in Fig. 1.4.
Starch granules
GWD and PWD Initial attack α‐amylase
Soluble branched glucans α‐amylase, β‐amylase, Debranching isoamylase, pullulanase
Soluble linear glucans
Degradation of α‐glucosidase linear chains
Glucosyl monomers Fig 1.4 Schematic pathway for starch degradation in plants. The initial attack on granules occurs inside plastids (within chloroplast in leaves, amyloplasts in non‐ photosynthetic tissues). Oligosaccharides released during starch degradation, such as maltose and maltotriose, are hydrolyzed to glucose by α‐glucosidase. GWD, glucan‐water dikinase; PWD, phosphoglucan water dikinase.
The other typical storage starch is found in tubers. Starch degradation in tubers occurs as a result of stress during sprouting (96, 97). In addition, the conversion of starch into sugars can also be initiated by cold‐induced sweetening. Temporary cold sweetening process may be partially reversed since sugar levels decline by reconditioning of the tubers to higher temperature at such periods (98, 99). A series Introduction
of starch‐degrading enzyme activities have been detected in potato tubers, including α‐amylase, isoamylase, β‐amylase, α‐glucosidase, and phosphorylase (99). An indication that starch degradation proceeds differently in cereal endosperms and Page | 12 potato tubers comes from the different appearance of their granules during degradation. Huber and Miller (1997) observed that granules from cereal endosperm have abundant surface pores that are the entrances of channels leading to the interior of the granule (63, 66). During enzymatic degradation, pits in the cereal starch granules become enlarged and deeper. Internal material surrounding the channels is eroded and subsequently more of the granular surface is attacked. In contrast, potato starch granules are highly resistant to enzymatic attack, and no convincing evidence for pores on their surface has been found so far (69, 100). Because of these very different properties, Alvani et al. (2011) suggested that the type of enzyme responsible for the attack on the potato granules may be different from that in cereal endosperm (100).
In order to maintain the cellular integrity and homeostasis, non‐photosynthetic organisms have to take up organic materials, such as starch. Humans, animals, and many other organisms including microbes degrade starch (101–105), as it constitutes a major source of energy. Although the basic enzymology of non‐plant starch degradation is similar, it differs in several aspects from that of plants.
Industrial starch conversion
Among carbohydrate polymers, not only as the main source of food for human, starch is the most important raw material that is widely used in range of a food and non‐food products. Starch is conventionally used in food as a thickener, colloidal stabilizer, gelling agent, encapsulation, coating agent, and water retention agent (106). Some of the non‐food applications include the use in paper manufacturing for surface sizing or in the textile industry as thickening and warp sizing agent (107– 109). Beside the use of starch directly as food source or in industrial products, it can be physically, chemically, or enzymatically modified into maltodextrins or simple
Chapter 1
sugars such as glucose or maltose (110). Glucose syrups then can be used as a substrate in the large‐scale ethanol production (111). The traditional process to convert starch into glucose and other sugars is based either on acid hydrolysis (112) or enzymatic catalyzed hydrolysis. The advantage of enzymatic treatment is a high Page | 13 yield of sugars and minimal formation of by‐products (113).
Enzymatic hydrolysis of starch involves three steps: gelatinization of a 15 – 35% (by weight) slurry starch with steam (114–116), followed by liquefaction by α‐amylases at 90‐105 oC. Finally, saccharification leads to further hydrolysis into glucose by glucoamylase (EC 3.2.1.3) at 60 oC. The latter step is time‐consuming and requires relatively high amounts of energy. To reduce the energy cost for cooking of starchy materials, the no‐cooking conversion has been developed resulting in a reduction of energy consumption by approximately 50% (117). The no‐cooking concept for starch processing, although firstly reported in the 1940’s (118), has only been established at large scale during the last decade.
Starch-Degrading Enzymes
Because of its complex structure, starch requires an appropriate combination of enzymes for its conversion to oligosaccharides and smaller sugars, such as glucose and maltose. A number of enzymes that are known to be involved in starch degradation have been identified and they are basically classified into four distinct groups of amylolytic enzymes, endo‐acting amylases, exo‐acting amylases, debranching enzymes, and glucosyltransferases (Fig. 1.5) (119, 120). Endo‐acting amylases, such as α‐amylase, hydrolyze α(14)‐glycosidic linkages in the inner region of the starch polymer in a random fashion and produce linear and branched oligosaccharides in the α‐configuration. Exo‐acting amylases hydrolyze the substrate from the non‐reducing end, producing only small oligo‐ and/or monosaccharides such as maltose (β‐amylase) or glucose (glucoamylase). Three major types of exo‐ amylases are important for starch hydrolysis; β‐amylase exclusively hydrolyzes α(14)‐glycosidic linkages at the non‐reducing ends, producing maltose and β‐limit Introduction
dextrins. Glucoamylase attacks α(14) linkages, releasing glucose molecules in the β‐configuration; it also attacks α(16) bonds at branching points in the amylopectin molecule, but much more slowly than α(14) bonds. α‐Glucosidase catalyzes the Page | 14 splitting of α‐glycosyl residues from the non‐reducing end of substrates to release α‐glucose (120).
Debranching enzymes catalyze the hydrolysis of α(16)‐glycosidic linkages in amylopectin and related oligosaccharides. These debranching enzymes are also known as pullulanase and isoamylase. To date, there are two main groups of pullulanase based on their substrate specificity and product formation; Pullulanase type I (referred to here as pullulanase), which specifically hydrolyzes the α(16)‐ glycosidic linkages in pullulan as well as in branched oligosaccharides, releasing maltotriose and linear oligosaccharides, respectively. In contrast, isoamylase acts on α(16) branch linkages of amylopectin and glycogen, but it is unable to hydrolyze pullulan. The second group is pullulanase type II or amylopullulanase (neopullulanase, EC 3.2.1.135), acting on both α(16) and α(14) linkages in starch, amylose, amylopectin, and pullulan. Both groups of enzymes are unable to degrade small cyclic oligosaccharides (121). Pullulan is a three α(14)‐linked glucose molecules that are repeatedly connected to each other by α(16)‐ glycosidic linkages on the terminal glucose (122).
Glucosyltransferases (EC 2.4.1.‐) constitute another group of starch‐converting enzymes that hydrolyze the α(14)‐glycosidic linkages in the donor molecule and transfer part of the donor to a glycosidic acceptor, forming a new glycosidic bond (120). Enzymes such as amylomaltases (EC 2.4.1.25) and cyclodextrin glucosyltransferases (CGTases, EC 2.4.1.19) cleave and form α(14)‐glycosidic linkages, while branching enzymes (EC 2.4.1.18) cleave α(14)‐glycosidic linkages and form α(16)‐glycosidic linkages. The CGTases produce cyclodextrins from starch by cyclization of an oligosaccharide of 6, 7, or 8 glucose residues, whilst amylomaltases catalyze transglycosylation to form a linear chain product (123, 124).
Chapter 1
Amylolytic enzymes are abundant in microorganisms, plants, animals, and a number of raw starch‐degrading enzymes have been reported that are able to degrade raw (native) starch (118, 125). These raw starch‐degrading enzymes act on starch in its crystalline form (insoluble substrate). This low‐temperature hydrolysis is being Page | 15 considered a major breakthrough in the starch processing industry, and has been described as granular starch, raw starch, cold, no‐cooking, sub‐gelatinization temperature or nonconventional starch hydrolysis (126, 127). α‐Amylases capable of raw starch degradation have been reported to be produced by a variety of living organisms, ranging from microorganisms including fungi, yeast, bacteria, and archaea, to plants and humans (128). However, only raw starch‐degradation enzymes from microbial origin have gained significant attention, because of their advantages, such as cost/time‐effectiveness, high productivity, and ease of the process of modification and optimization. The data listed in Table S1.1 (supplementary information) represents enzymes that directly degrade raw starch below the gelatinization temperature of starch and includes detailed description of their families and binding modules. The majority of the known enzymes with starch‐ degrading activity belong to the α‐amylase family. They are known as glycosyl hydrolases (GH). These starch degrading enzymes are found in various GH families (13, 14, 15, 31, and 119) (129). GH families are described in more detail in the next section.
Page
|
16
CGTase α-glucosidase/ glucoamylase glucose
cyclodextrin
amylomaltase
α-amylase
maltose β-amylase Debranching
α-limit dextrin Maltose and linear β-limit dextrin Isoamylase Pullulanase II oligosaccharide Pullulanase I maltotriose
maltose
glucose
linear oligosaccharide Fig 1.5 Action of starch‐degrading enzymes. (●) : Reducing α‐glucose residue; (○) : non‐reducing α‐glucose residue; ○ ̶○ : α(14)‐ glycosidic linkages; ○····○ : α(16)‐glycosidic linkages. Adapted from Horváthová et al. (119), with modifications.
Chapter 1
α-Amylases
α‐Amylases are extracellular enzymes that catalyze the random hydrolysis of α(14)‐glycosidic linkages in the inner part of the amylose or amylopectin chains with the retention of the α‐configuration in the products. These enzymes are Page | 17 incapable of hydrolyzing α(16)‐glycosidic linkages present at branch points of amylopectin chains. An exception to this is the α‐amylase produced by Thermactinomyces vulgaris, which can hydrolyze both α(14) and α(16)‐ glycosidic linkages (130, 131). Several models for α‐amylase action pattern have been proposed, such as the random action, also referred to as a single‐chain or multi‐chain attack, and the multi attack action (132). The enzyme binds to the substrate, performs only one hydrolytic cleavage per effective encounter or a number of hydrolytic attacks, dissociates, and binds to a next chain. This mode of action implies that all bonds are equally susceptible to hydrolysis and a rapid decrease in starch polymer mass can be observed (133). Some authors proposed another α‐amylase action pattern, so‐called preferred attack action (134). Like in the random attack action, one bond is hydrolyzed per effective encounter. However, this mode of action presumes that bonds near to the chain ends and the branching points are less susceptible to hydrolysis (134, 135). The multi attack action is generally accepted to explain the differences in action pattern of α‐amylases (136).
Classification of α-amylases
In 1991, Bernard Henrissat proposed a new classification system for glycoside hydrolases (GH) (137). This classification system is based upon amino acid sequence and structural similarities rather than substrate specificity and can be used to reveal evolutionary relationships between different GH families as well as to derive mechanistic information of the enzyme (137, 138). By the end of 2015, 135 sequence‐based families of GHs were available online at the CAZy (Carbohydrate Acting Enzymes) website. Families with many entries have been divided into subfamilies with a reliable prediction of the substrate specificity (139).
Introduction
Starch degrading enzymes are found in just a few of the numerous families of glycoside hydrolases. The majority of the enzymes acting on starch, glycogen, and related oligo‐ and polysaccharides, including enzymes specifically acting on α(11)‐
Page | 18 linkages (trehalose), are found within the family GH13, currently by far the largest family of glycoside hydrolases, including more than 2000 representatives (140). This GH13 family belongs to clan GH‐H which contains also families GH70 and GH77, but in the latter two, no α‐amylase specificity has been found (140). A clan is a higher hierarchical level than the family in the CAZy classification. A group of families from the same clan are thought to share a common ancestor and catalytic machinery. The GH13 family, also known as the α‐amylase family, has been identified very early and groups together enzymes sharing sometimes only very limited sequence identity. This family is the most varied of all GH families, containing many activities, such as hydrolysis, transglycosylation, condensation, isomerization, and cyclization (141). As a consequence, the α‐amylase family has been the subject of numerous analyses in order to derive clear relationships between the sequences and the properties of the enzymes. Fueled by the importance of α‐amylases and related enzymes, many crystallographic studies have been performed on GH13 family enzymes, and currently, of 103 different protein members the three‐dimensional structure (3D) is available (http://www.cazy.org). Besides in family GH13, α‐amylases are also found in families GH57 and GH119 (142).
The family GH13 was subdivided into subfamilies in order to establish robust groups exhibiting an improved correlation between sequences and enzyme specificity (139). The members of the individual GH13 subfamilies share in general a closer relatedness to each other than to the remaining GH13 family members, i.e. higher identity in sequence, structure, specificity, and/or even taxonomy of the host species (143).
The first 3D structure of a GH13 α‐amylase elucidated with X‐ray crystallography was Taka‐amylase A (TAA), the α‐amylase from Aspergillus oryzae (Fig. 1.6) (144).
Chapter 1
Structurally, the GH13 enzymes are characterized by a conserved structural core composed of three domains, referred to as the A, B, and C domains (142). The A‐domain is the most conserved domain in the α‐amylase family and is present at
the N‐terminal end of the protein. It consists of a highly conserved (β/α)8‐ or TIM‐ barrel and can be recognized by 4‐7 conserved amino acid regions containing three Page | 19 active site residues involved in catalysis and substrate binding, which are believed
to represent a common evolutionary origin (142, 145). The (β/α)8 barrel consists of eight inner parallel β‐strands which are surrounded by the outer cylinder composed of eight α‐helices so that the individual β‐strands and α‐helices alternate and are connected by loops (146). This (β/α)8 motif is recognized in at least nine families of glycoside hydrolases (147). Although there is only 10% identity between the amino acid sequences of GH13 α‐amylases from animals, plants, and microbes, all these α‐amylases share four highly conserved regions (148). While the four originally conserved sequence regions contain the invariant catalytic and substrate binding residues, three additional conserved regions were later identified and were shown to be connected to enzyme specificity (140, 149). The B‐domain is an irregular loop of variable length from 44 to 133 amino acid residues inserted between the third
β‐strand and the third α‐helix of the (β/α)8‐barrel and is firmly attached to the A‐domain by a disulphide bond, extending out of the barrel (145, 150). The sequence of this domain varies, and it is the major determinant of substrate specificity. For example, the B‐domain of Bacillus licheniformis α‐amylase is relatively long and folds into a more complex structure (151), while in barley α‐amylase, it folds into a random coil without defined secondary structure elements (152).
The active site is located in a cleft between domains A and B where a triad of catalytic residues performs catalysis. The glutamate residue involved, corresponding to Glu230 of TAA, acts as general acid/base catalyst, performing proton donation to the leaving glycosidic oxygen group and providing a nucleophilic species for the displacement of the glycoside. The acid group of Asp206 (TAA numbering) acts as the catalytic nucleophile, leading to the formation of a covalent intermediate, while the other aspartate residue (Asp297 in TAA) stabilizes the transition‐state (144, 150). The C‐domain, which forms a Greek key motif, is the domain with the lowest
Introduction
degree of conserved sequence in the GH13 family. It has a β‐sheet structure and is linked to the A‐domain by a simple loop. The type and source of the α‐amylase will affect the relative orientation between domain C and domain A (147, 153). The
Page | 20 function of the C‐domain is largely unknown, although various studies demonstrate that it is necessary for catalytic activity and substrate binding (154–156).
B domain A domain
C domain
Fig 1.6 Ribbon representation of the overall structure of TAKA α‐amylase A, α‐amylase from Aspergillus oryzae (1TAA). Domains A, B, and C are represented as green, blue, and red, respectively. The triad of catalytic residues is marked as blue sticks. The structure model was built using the MacPyMOL program (157).
A new family, GH57, containing α‐amylases was established in 1996 (158) after no sequence similarity could be detected between two supposed α‐amylases from the thermophilic bacterium Dictyoglomus thermophilum and the archaeon Pyrococcus furiosus and members of the typical GH13 α‐amylase family. Later both ‘amylases’ were re‐evaluated and characterized as α‐glucanotransferase (159). Nowadays the family GH57 represents a second and smaller α‐amylase family (129). Family GH57 contains more than 1300 members mainly originating from extremophilic prokaryotes and fewer than 25 members have been biochemically characterized. This family exhibits six well‐established enzyme specificities, including α‐amylase
Chapter 1
(160), 4‐α‐glucanotransferase (159, 161), amylopullulanase (162, 163), glycogen branching enzyme (164), and α‐galactosidase (165).
The first 3D structure of the GH57 family was solved by X‐ray diffraction studies Page | 21 with the 4‐α‐glucanotransferase from Thermococcus litoralis (TLGT) (166). Family
GH57 members share a (β/α)7‐barrel catalytic domain, comprised of seven parallel β–strands arranged in a barrel encircled by seven α‐helices. Structural models and mechanistic labelling analysis of mutants allowed identification of two catalytic
amino acid residues in the active site and are located within the catalytic (β/α)7‐ barrel domain, namely a glutamic acid as catalytic nucleophile (Glu123, TLGT numbering) and an aspartic acid acts as general acid/base residue (Asp214) (161, 162). The sequences of GH57 family members vary in length and domain characteristics. Five conserved sequence regions proposed by Zona et al. in 2004 can be used as sequence fingerprints for identifying proteins belonging to the GH57 family (142, 162).
The family GH119 was created based on a study by Watanabe et al. (2006) describing the gene encoding the isocyclomaltooligosaccharide glucanotransferase (igtZ) from Bacillus circulans as an α‐amylase (167). No convincing sequence similarity was found to other glycoside hydrolase families, including GH13, but recently it was shown that GH119 is very closely related in sequence to family GH57 (129, 142). The family GH119 may thus be considered as the third CAZy α‐amylase family, but the family is still very small and until recently only 11 additional hypothetical proteins encoded by ORFs have been added to GH119 (129). No tertiary structure of the GH119 family has been solved; however, a preliminary study suggests that GH119 shares both the conserved (β/α)7‐barrel fold of the catalytic domain and catalytic machinery with family GH57 (142).
Introduction
Physicochemical properties of α-amylases
Many α‐amylases have been characterized in some detail. Table S1.2 presents a selection of the most extensively studied α‐amylases. Most of them are extracellular Page | 22 enzymes, but intracellular α‐amylases also have been reported (168–171). α‐Amylases differ in their physicochemical characteristics, such as the optimum values of temperature and pH, which are essential factors that determine their activities. Despite wide differences between α–amylases, the molecular masses are generally in the range of 45 to 60 kDa (172–174), but smaller or bigger α‐amylases have also been reported. Ratanakhanokchai et al. (1992) reported that the α‐amylase of Chloroflexus aurantiacus has a molecular mass of about 210 kDa (175), while the smallest α‐amylase so far is from Bacillus caldolyticus with a molecular mass of 10 kDa (176).
Most of the α‐amylases are calcium metalloenzymes, in which calcium ion (Ca2+) is required for their activity, structural integrity, or stability (177, 178). The affinity for Ca2+ is much stronger than that for other ions, although calcium can be replaced by other divalent metals such as magnesium, barium, and strontium without major loss in activity (179, 180). The number of bound calcium atoms may vary from one to about ten (181, 182). TAA contains 10 Ca2+ ions but only one atom of calcium is very tightly bound to the protein, while the other nine may be easily removed without any effect on the physical or biological properties of the protein (183). Most of the known α‐amylase structures share a common calcium‐binding site, located at the interface between domains A and B and in close proximity to the active site (151). These calcium ions stabilize the tertiary structure of the enzyme by inducing an ionic bridge between domain A and B, thereby protecting the enzyme from denaturation, heat inactivation, or proteolytic enzymes (184, 185). There are also reports of α‐amylases that do not require Ca2+ for stability and activity (186–188) and even inhibition of the enzyme by Ca2+ has been documented (189). Many metal cations, especially heavy metals such as copper, mercury, silver, and lead, strongly inhibit α‐amylase activity (125).
Chapter 1
The catalytic mechanism of α-amylases
The active site of α‐amylase family GH13 enzyme is localized at the center of the
(β/α)8‐barrel domain and consists of two aspartate (Asp) and one glutamate (Glu) residues, while in many α‐amylases a histidine (His) residue is involved in substrate Page | 23 binding (136, 141, 144). Site‐directed mutagenesis (SDM) and/or X‐ray crystallography studies have shown that these residues are conserved in the α‐amylase of Bacillus subtilis (190, 191), barley (192), human saliva (193), as well as in neopullulanase (194), cyclodextrinase (EC 3.2.1.54) (195, 196), CGTases (197– 199), amylopullulanase (163, 200), and branching enzymes (201, 202). The Glu residue acts as the proton donor, while one of the Asp residues acts as the nucleophile. The role of the second Asp residue has been suggested to be involved in stabilizing the oxocarbenium ion‐like transition state and also in maintaining the Glu in the correct state of protonation for activity (199).
In general, there are two different hydrolytic mechanisms by which glycoside hydrolases cleave a glycosidic linkage: (i) retaining (α α or β β) and (ii) inverting (α β or β α) the anomeric configuration of the substrate (203). Members of the α‐amylase family employ the α‐retaining mechanism. The process of hydrolysis of α‐glycosidic linkages by a retaining mechanism is shown in Fig. 1.7. In the unbound state, the un‐ionized Glu230 (TAA numbering) is hydrogen bonded to the side chain carboxyl of Asp297. When the substrate binds to the active site of the enzyme, the water molecules are displaced from the active site of the enzyme and the substrate breaks the original network of hydrogen bonds. The glycosidic oxygen of the substrate interacts with Asp297 and points with its H1 toward the Asp206 (bottom side of the cleft). The new hydrogen bonds and the hydrophobic interaction between the substrate and the substrate‐binding sites are regenerated and weaken the hydrogen bond between Glu230 and Asp297. Glu230 in the acid form easily protonates the oxygen of the O‐glycosidic linkage and the reducing‐end fragment is released from the cleft. Asp206, the catalytic nucleophile, located at the bottom of the cleft attacks the anomeric center. The transition state promotes the formation of the oxocarbenium ion‐like intermediate and Asp 206 contributes to the stability of the intermediate. The ionization of Glu230 produces electrostatic repulsive forces
Introduction
between Glu230 and Asp297 (197). An ordered water molecule is positioned at the relatively outer side of the cleft to promote formation of a hydrogen bond between Glu230 and Asp297 and leading to reduction of repulsive forces in the active site.
Page | 24 Glu230 acts as a base by accepting the proton from the water molecule to produce an activated hydroxyl ion. The hydroxyl ion of the water then performs a nucleophilic attack on the oxocarbonium ion intermediate leading to the hydrolysis product. The anomeric conformation is maintained, because the nucleophilic water molecule is located on the anomeric side which is close to the proton donor in the position between Glu230 and Asp297. Glu230 and Asp297 are located at the relatively outer side from the anomeric center, while Asp206 is positioned deeper in
the core of the (β/α)8‐barrel (204).
The structural and biochemical information for GH57 is more limited. To date, only four 3D structures out of 1387 identified sequences have been determined (129). Several methods have been employed to characterize the reaction mechanism of GH57, including analysis of the 3D crystal structure of the enzyme. In the GH family members that employ a retaining mechanism, the two catalytic residues are commonly separated by approximately 5.5 Å distance. A larger than 5.5 Å distance usually indicates that the enzyme has an inverting mechanism. The crystal structure of free and ligand‐bound TLGT showed that the distance between Glu123 and Asp214 residues in both states are within the average distance characteristic of the retaining mechanism (166, 205, 206). Further evidence for a retaining mechanism of GH57 enzymes came from NMR analysis of reaction products of the branching enzyme from Thermus thermophilus (164). Therefore, similar to the reaction mechanism adopted by the members of family GH13, the GH57 members act through a retaining mechanism for cleaving the α‐glycosidic linkage.
Chapter 1
In conclusion, enzymes of GH13, 57, and 119 meet the following criteria:
(i) act on α‐O(14)‐, (16)‐ or (11)‐glycosidic linkages and hydrolyze these to produce α‐anomeric monosaccharides or oligosaccharides or form α‐glycosidic linkages by transglycosylation, or a combination of both Page | 25 activities; (ii) contain between four and seven highly conserved regions in the primary structure that contain the amino acids that are essential for the catalytic reaction and substrate‐binding site;
(iii) have a (β/α)8‐ or (β/α)7‐barrel conformation for the catalytic domain; (iv) have the invariant Asp and Glu catalytic residue corresponding to the Asp206, Glu230, and Asp297 of TAA for family GH13 or equivalent to Glu123 and Asp214 of TLGT of GH57 and GH119 members; and (v) employ a retaining reaction mechanism.
Introduction
Asp206 Asp206
δ‐ δ+
Page | 26 δ‐
δ+
Asp297 Glu230 Asp297 Glu230
Asp206 Asp206
Asp297 Glu230 Asp297 Glu230
Asp206 Asp206
δ‐ δ+
δ‐
δ+
Asp297 Glu230 Asp297 Glu230 Fig 1.7 Scheme of retaining reaction mechanism of glycoside hydrolase of family GH13, as explained in the text. Adapted from MacGregor et al. (145), with modifications.
Chapter 1
Starch binding domains
Amylolytic enzymes are often multifunctional proteins composed of distinct domains, a catalytic domain and one or more domains involved in substrate binding or multi‐enzyme complex formation. Various amylolytic enzymes, such as Page | 27 glucoamylases, β‐amylases, cyclodextrin glycosyltransferases and α‐amylases are reported to contain non‐catalytic ancillary domains referred to as carbohydrate‐ binding modules (CBM) (207, 208). CBMs conduct their biological activity by maintaining the interaction of the enzyme with its insoluble substrate and disrupting the crystalline structure of starch granules (209, 210). CBMs have been classified into 71 families based on their sequence similarities, according to the CAZy database (http://www.cazy.org/, last update on 20 October 2015). Less than 5% of CBMs have been characterized experimentally and the sequence identities among families are less than 30% (211). CBMs with affinity for insoluble raw starch are commonly referred to as starch binding domains (SBDs) and the location of SBD can be both, C‐ or N‐ terminal or centrally positioned within the protein (147). All SBDs are approximately 100 amino acids and share several conserved amino acids (typically aromatic amino acids). SBDs are present in about 10% of all α‐amylases and related amylolytic enzymes (212). SBDs are structurally conserved and predicted to be composed of β‐sandwich folds with 7‐8 anti‐parallel β‐strands arranged in two β‐sheets (213). Based on the structures, seven fold types have been observed in CBMs families: β‐sandwich, β‐trefoil, cysteine knot, unique, oligonucleotide/oligosaccharide binding (OB) fold, hevein fold, and hevein‐like fold (214).
To date, amino acid sequence‐based classification categorized SBDs into the 10 following CBM families: 20, 21, 25, 26, 34, 41, 45, 48, 58, and 69 (129, 215). Nuclear magnetic resonance (NMR), X‐ray crystallography, and electron microscopy (EM) have been used to determine the 3D structures of CBMs (216, 217). Except for families CBM45 (218) and CBM69, 3D structures are known for at least one representative of the other families (Fig. 8). Despite the low sequence similarities, all these SBD CBMs have β‐sandwich folds with an immunoglobulin‐like topology (214,
Introduction
219). This suggests a common evolutionary ancestry towards starch recognition by all starch binding domain families (210).
Page | 28 The CBM20 family
The first and best‐characterized SBD of starch degrading enzymes belongs to family CBM20 (210, 220). This family currently has 852 entries: 8 archaea, 616 bacteria, 220 eukaryota, and 8 unclassified. So far, the tertiary structures of 14 members of the family CBM20 are known (http://www.cazy.org/CBM20.html). The CBM20 is generally located at the C‐terminus of amylolytic enzymes from family GH13 – the α‐amylase family, GH14 – the β‐amylases, GH15 – glucoamylases, and GH77 – amylomaltases (207, 210, 221). Exceptions to this rule are a GH57 putative amylopullulanase with three tandemly arranged SBD and several GH77 4‐α‐glucanotransferases (210). The granular raw starch‐binding function of CBM20s has been described, for example, for α‐amylase from Bacillus sp. strain TS‐23 (222, 223), glucoamylase from Aspergillus niger (213), CGTase from Bacillus circulans strain 251 (220, 224), maltogenic α‐amylase from Geobacillus stearothermophilus (156), and β‐amylase from Bacillus cereus var. mycoides (225).
The CBM20 domains are 90‐130 amino acids large and a comprehensive analysis of amino acid sequences showed that there are no absolutely conserved residues in the family (210, 220). However, analysis of a larger set of CBM20s suggested that certain residues appear to be involved specifically in substrate interactions. The structure‐ function relationships of the CBM20 motifs are also quite well understood. The structural information for CBM20 is based on NMR studies of A. niger SBD (213, 226), which is used here as the main representative of CBM20, and the X‐ray crystallography structure of the CBM20 of Bacillus circulans strain 251 CGTase (224, 227). The CBM20 SBD has an open‐sided, distorted β‐barrel structure built by several β‐strands (213, 220). The overall topology shows eight β‐strands arranged into two major β‐sheets (Fig. 1.8A) (228). One is a five‐stranded antiparallel β‐sheet, while the second β‐sheet consists of one parallel β‐strand and one antiparallel
Chapter 1
β‐strand pair. The N‐ and C‐termini are located at opposite ends of the longest axis of the molecule (213). There are two starch binding sites, which differ from each other functionally as well as structurally (226). Binding site 1 is small, accessible, and acts as the initial recognition of the starch molecule. It comprises two conserved tryptophans, Trp543 and Trp590 (A. niger SBD numbering), which are the most Page | 29 important for raw starch‐binding capacity (224, 226, 229, 230). These residues provide surface‐site and hydrophobic stacking interactions to the sugar rings (226). On the other hand, binding site 2 is more extended, flexible, and is suggested as the specific site to lock SBD on the starch surface. The stacking interaction at this site between SBD and the substrate is also governed by hydrophobic effects from the aromatic rings, two tyrosine (Tyr527 and Tyr556, A. niger SBD numbering) and two tryptophan (Trp563 and Trp615, A. niger SBD numbering) residues (226, 229). The latter tryptophan is not strictly conserved in CBM20 (207). The removal of this SBD from A. niger glucoamylase decreased the catalytic activity toward starch granules 100‐fold of the original activity (231). Juge et al. (2006) showed that fusion of this SBD to barley α‐amylase isozyme 1 (AMY1) resulted in a six‐fold enhancement in the affinity and activity for starch granules (232).
The CBM21 family
There are currently 624 entries for the CBM21 family, of which 31 are from bacteria, 592 are from eukaryotes, and 1 is unclassified. The binding sites of CBM21 are very similar to those of CBM20 (233). The CBM21s are approximately 100 residues in length and are positioned at the N‐terminus, in contrast to CBM20s that are normally found at the C‐terminus.
The crystal structure of CBM21 was reported firstly for Rhizopus oryzae glucoamylase (RoSBD). It is composed of eight antiparallel β‐strands forming two major β‐sheets with a distorted barrel structure, similar to the CBM20 structure (Fig. 1.8B) (207, 234). Subsequent studies of RoSBD revealed the presence of two specific ligand‐binding sites: site I (conserved in many starch‐binding CBMs), consisting of Trp47, Tyr83, and Tyr94, and site II, consisting of Tyr32, Phe58, and
Introduction
Tyr67, on the surface of SBD (233, 235). Site I forms a broad, flat and stable hydrophobic environment to interact with the sugar rings, and serves as the major sugar‐binding site (235). Site II is more narrow, forming a protruded binding
Page | 30 environment and is thought to be involved in binding to long‐chain soluble polysaccharides (234). These properties may be important for delivering the substrate from the SBD to the catalytic domain (233). NMR and thermodynamic studies by isothermal titration calorimetry (ITC) of RoSBD have shown that it has two binding sites with high affinity for various maltooligosaccharides (ranging from 61.9 µM for maltotriose to 5.5 µM for maltoheptaose) (235).
The CBM25 and CBM26 families
The CBM25 family has currently about 264 entries in the CAZy database and the majority of characterized entries are found in bacterial α‐amylases (129). This module is present in one or more copies and the motifs adopt a β‐sandwich fold (ten‐strands in this case) (Fig. 1.8C) (236). The CBM25 family was established based on a novel type of SBD in the Bacillus sp. strain 195 α‐amylase (236). CBM25 possesses two sites for binding raw starch with two Trp residues at positions 890 and 930 (Bacillus sp. strain 195 numbering), and His882, whereas the His residue is the only residue that is conserved at the structural and sequence level (Fig. 1.8C) (237).
The structure of the best studied CBM26, that of the maltohexaose‐forming amylase of Bacillus halodurans strain C‐125 was solved by X‐ray crystallography (Fig. 1.8D) (237). CBM25 and CBM26 are structurally related and both are found in the maltohexaose‐forming amylase from B. halodurans (237, 238). CBM26 usually occurs as tandem repeats with one individual starch‐binding site, exhibiting an unusual organization of the SBD related to amylases from family GH13 (239). Tyr25 and Trp36 are responsible for stacking interaction against the pyranose rings of the maltose sugar residues. Tyr23 contributes to hydrogen bond formation with oxygen 6 of the sugar stacking against Tyr25 (237). The starch‐binding function of CBM26
Chapter 1
has also been demonstrated in Lactobacillus α‐amylase containing SBDs arranged in an unusual manner, i.e. five identical modules in L. amylovorus and four in L. plantarum (240) and L. manihotivorans (241). In a previous report, it was shown that the L. amylovorus α‐amylase with no tandem SBDs is incapable of binding and hydrolyzing starch granules even though it still hydrolyzed soluble starch (239, Page | 31 242).
The CBM34 family
The CBM34 family is specific for the GH13 enzymes. Several bacterial maltogenic amylases and cyclodextrin (CD)‐hydrolyzing enzymes have N‐terminal regions that are composed of 120‐140 amino acids which have been classified as CBM34 (129). There are nearly 1450 records in the database of CBM34, of which 17 are from archaea, 1427 are from bacteria, and 5 are unclassified.
The first CBM34 family member to have its structure solved by X‐ray crystallography was the N‐terminal CBM from two Thermoactinomyces vulgaris α‐amylases, TVA I (Fig. 1.8E) and TVA II (243). This module was initially referred to as ‘domain N’. The presence of two glucose molecules bound to this module, supported by functional analysis that revealed the two maltooligosaccharides binding sites, confirming the role of domain ‘N’ as a CBM (244). The crucial amino acid residue involved in raw starch binding in TVA I, Trp65, has no counterpart in TVA II despite the high sequence identity between the two respective domains. TVAI has been shown to have high hydrolyzing and binding capability against starch and pullulan, whereas the preferred substrate of TVAII is more likely to be cyclodextrins (131, 245).
The CBM41 family
Currently, the CBM41 family has more than 1000 entries in the CAZy database, and of 8 entries structural information is available (129). Family 41 CBMs are found mainly in pullulanases or other starch/glycogen debranching enzymes and in
Introduction
general, the modules comprise approximately 100 residues (129). The functionally characterized member is an α‐1,4‐glucan‐specific CBM41 from pullulanase of the marine bacterium Thermotoga maritima (TmPul13) (246). The family 41 CBM from
Page | 32 T. maritima Pul13 has only one main carbohydrate binding site on the protein (247) and the affinities for β‐cyclodextrin, pullulan and amylopectin were in the range of 14 ‐ 42 µM (246). The overall structure of CBM41 is again a distorted antiparallel β‐sandwich fold with three tryptophans, Trp27, Trp29, and Trp73 (T. maritima CBM41 numbering) comprising the binding site of this molecule (Fig. 1.8F) (247). The crystal structure of TmCBM41 in complex with maltotetraose (G4) and α‐D‐glucosyl‐maltotriose (GM3) showed that Trp29 and Trp73 residues formed hydrophobic stacking interactions with the glucose molecule, while Trp27 contributed a single water‐mediated hydrogen bond to the third glucose in the chain (247). The third residue, Trp73, of CBM41 is conserved only at the structural level and has no equivalent in the CBM20 structure (248).
Recently, the N‐terminal domain of the Klebsiella pneumoniae pullulanase (KpCBM41) was identified as a CBM41, using structural comparison. KpCBM41 shares the same topology and binding site as TmCBM41; however, only two of three binding site aromatic residues are conserved (248).
The CBM45 and CBM69 families
No 3D structures have been described for any member of the CBM45 and CBM69 families. CBM45 members are mainly attached to eukaryotic proteins. They are, for example, present as N‐terminal tandem domains in Solanum tuberosum GWD (StGWD) and Arabidopsis thaliana α‐amylase (AtAMY3) and separated by a linker of approximately 200 and 50 amino acids, respectively (249). A detailed bioinformatics and binding analysis of the CBM45 SBD of StGWD and AtAMY3, identified two conserved tryptophan residues as the substrate‐binding site (249, 250). Trp139 and Trp192 (StGWD numbering), appear to be essential for starch binding (249). An interesting observation concerning the StGWD and AtAMY3 SBD is that they have a
Chapter 1
lower‐affinity towards cyclodextrin and starch granules compared to typical SBDs of microbial amylolytic enzymes, suggesting that this SBD is important in the reversible interaction with starch (during starch synthesis and degradation) (249). However, structural elements that are responsible for the low affinity of CBM45 in starch metabolism have not yet been defined. Page | 33
The CBM69 family has only a few members and is exclusively found in enzymes from marine bacteria (215). No structural and almost no functional information is available for this family (129). A SBD was identified in an α‐amylase from a marine metagenomic library (AmyP) and deletion of this SBD did not affect the ability of the α‐amylase to degrade raw starch. However, it was observed that the deletion mutant had significantly lower activity for soluble starch than the wild type AmyP (215).
The CBM48 family
The family CBM48 is often found associated with enzymes from the GH13 family (251), except in the non‐amylolytic SEX4 proteins of plants and green algae, where CBM45 domain of SEX4 proteins are positioned at the C‐terminus (252). They are specifically found in a number of enzymes that act on branched substrates, i.e. isoamylase, pullulanase, and branching enzymes (251). These CBMs were originally referred to as a (CBM20 + CBM21)‐related SBD group (221), but some were later classified into the family CBM48 (129). It was previously suggested to name them glycogen‐binding domains (GBDs) instead of CBMs, because several members of this family have a role in glycogen metabolism (251). Secondary and tertiary structure‐ based comparison of CBM20 and CBM48 revealed the close relatedness of these SBDs and, in some cases GBDs. The families CBM20 and CBM48 probably have a common ancestral form (251).
So far 28 members of CBM48 have been structurally characterized (251), and the first CBM48 module that interacts with the catalytic domain to be identified and characterized was that of the GH13 maltogenic amylase of Staphylothermus marinus
Introduction
(SMMA) (253). The crystal structure of SMMA revealed that the CBM48 domain contains a long loop that extends into the catalytic site and participates in substrate binding at the catalytic site. Only one binding site has been identified in this CBM48
Page | 34 comprising of three aromatic residues, Phe95, Phe96, and Tyr99 (Fig. 1.8G) (253).
The CBM58 family
The CBM58 family has 14 members. The domains are about 120 amino acids in length. CBM58 shares most structural similarity with the starch‐binding proteins CBM26 of B. halodurans maltohexaose‐forming amylase (PDB code 2C3G) and CBM41 of T. maritima pullulanase Pul13 (PDB 2J71) (254). To date, the only crystal structure for this family is that of SusG from Bacteroides thetaiotamicron that has been solved by X‐ray crystallography to a resolution of 2.2 Å (254). The crystal structure of CBM58 of SusG in complex with maltoheptaose revealed that Tyr260, Trp287, and Trp299 bind to the backbone of this sugar molecule [233]. It also showed that CBM58 has a different topology of the β‐strands compared to other CBM families, comprising nine β‐strands organized into two opposing four‐stranded and five‐stranded antiparallel β‐sheets and two additional parallel β‐strands that form a small β‐sheet (Fig. 1.8H).
Other Binding Sites
The carbohydrate binding domains have been shown to be important for adsorption to starch and degradation of starch granules. However, many α‐amylases lacking of SBD are still able to bind and degrade starch granules (207). Non‐catalytic carbohydrate binding, however, is not only observed for CBMs (255–257). A growing number of structural studies on various GHs have revealed the presence of other additional non‐catalytic carbohydrates binding modules, such as fibronectin‐ like domains or surface binding sites. The presence of a SBD may not be the major requirement for degradation of raw starch, but it is still not clear why some α‐amylases contain SBD whereas others do not (147).
Chapter 1
A B C
Page | 35
D E F
G H
Fig 1.8 The structures of the SBDs from individual CBM families. The tryptophan and tyrosine residues involved in stacking interactions with the substrate in binding site 1 are colored magenta and in binding site 2, if present, blue. (A) CBM20 of Aspergilus niger glucoamylase, PDB code: 1AC0 (226); (B) CBM21 of Rhizopus oryzae glucoamylase, PDB code: 2V8L (233); (C) CBM25 of Bacillus halodurans α‐amylase, PDB code: 2C3V; (D) CBM26 of Bacillus halodurans α‐amylase, PDB code: 2C3G (237); (E) CBM34 of Thermoactinomyces vulgaris α‐amylase (TVA I), PDB code: 1JI1 (243); (F) CBM41 of Thermotoga maritima pullulanase (Pul13), PDB code: 2J71 (247); (G) CBM48 of Staphylothermus marinus maltogenic amylase (SMMA), PDB code: 4AEE (253); (H) CBM58 of Bacteroides thetaiotaomicron α‐amylase (SusG), PDB code: 3K8K (254). The figures were drawn by using the MacPyMOL program (157).
Introduction
Fibronectin III modules
Fibronectin III modules (FnIII) are one of three types of common constituents of animal plasma proteins that act as a linker and mediate protein‐protein interactions Page | 36 (258). Several animal FnIII structures have been elucidated by X‐ray crystallography and NMR spectroscopy (259, 260). They contain approximately 90 residues and fold into an immunoglobulin‐like β‐sandwich motif. The secondary structure consists of seven β‐strands, forming two antiparallel β‐sheets (258, 261, 262). FnIII modules have been found only in extracellular bacterial α‐amylases and pullulanases (263, 264). The existence of bacterial FnIIIs was first reported for chitinase A1 from Bacillus circulans WL‐12 (265). This FnIII domain is not directly involved in chitin binding but seems to have an important role in the hydrolysis of chitin (266). Another study also reported that FnIII supported the hydrolytic ability of cellobiohydrolase (267). Very little information is available on the function of FnIII modules in starch‐degrading enzymes. So far the domain has been found in the α‐amylase of an alkaliphilic Gram‐positive bacterium (263), and Microbacterium aurum B8.A that also contains CBM25 (264), and in the amylopullulanase of Thermoanaerobacterium saccharolyticum NTOU1 (268). Truncation of the FnIII domain of M. aurum B8.A α‐amylase (264) and amylopullulanase from T. saccharolyticum NTOU1 did not alter the function of the enzyme (268). Therefore, it was suggested that FnIII may not be required for the catalytic reaction, but it may serve as a linker between the other domains (discussed in more detail in Chapter 4).
Surface Binding Sites (SBSs)
The existence of carbohydrate surface binding sites (SBSs), sometimes referred to as secondary binding sites, was first described more than three decades ago for glycogen phosphorylase from plants (269, 270). SBSs occur either on the surface of the catalytic site or on the outside of the enzyme, in contrast to the non‐catalytic binding in CBMs, which are often attached to the enzyme by a flexible linker. Although SBSs are different from CBMs, they seem to have an important function for activity towards polysaccharide substrates, equal to CBMs (255, 271). In a recent
Chapter 1
review, SBSs have been described for several polysaccharide processing enzymes (272). Examination of the SBSs showed that they frequently co‐occur with CBMs, suggesting that SBSs have a complementary role to CBMs rather than being an alternative to CBMs (273, 274). Approximately half of the known carbohydrate SBSs have been found within several GH13 subfamilies (1‐11, 14, 21, 24, 31) (273). The Page | 37 best‐known examples are the ones of the α‐amylase isozymes from barley (AMY1 and AMY2) and Saccharomycopsis fibuligera, both of subfamily GH13_6 (275, 276). The crystal structures of AMY1 and AMY2 revealed two SBSs, originally named “the starch granule‐binding surface site” and ‘the pair of sugar tongs’, and now referred to as SBS1 and SBS2, respectively (255). The crystal structure and kinetic properties of the individual replacement of Tyr380 of SBS2 at the ‘sugar tongs’ by alanine showed that these mutations reduced the affinity for starch and destroyed oligosaccharide binding (255, 277). Analysis of a triple SBS mutation with Trp278, Trp279, and Tyr380 replaced by alanine revealed complete loss of the affinity of the resulting AMY1 mutant to starch granules and only 0.2% hydrolytic activity remaining compared to the wild‐type towards starch granules (255, 271, 278). The Trp278, Trp279, and Tyr380 thus appear to be the functional residues for substrate binding (Fig 1.9).
These findings confirmed that SBSs play a variety of crucial roles to possess enzymatic activity towards starch, as compensating for a possible lack of CBMs. The exact mechanism involved in raw starch degradation still remains largely unknown and none of those three residues were conserved when comparing with other subfamilies. Further studies thus are required to elucidate the presence and functions of SBS.
Introduction
Page | 38
Fig 1.9 Representation of the crystal structure of barley α‐amylase isozyme 1 (AMY1) highlighting the surface binding sites. The picture shows the presence of the ‘sugar tongs’ surface binding site in different surface regions of the AMY1 in complex with maltoheptaose (PDB code: 1RP8). The residues defining the surface binding sites are colored in magenta and five sugar rings of maltoheptaose (red) bound at SBS1 and SBS2 can be seen. On the right, schematic representations are shown of the oligosaccharides bound at the secondary binding sites of AMY1. The diagrams of protein‐substrate interactions were generated using MacPymol (157) and ChemDraw based on Xavier et al. (279).
Chapter 1
Outline of this thesis
The susceptibility of starch granules to degradation by α‐amylases depends on their botanical origin and on the α‐amylase source (280). Considerable progress has been made in elucidating the nature, type, and mechanism of α‐amylases. This thesis Page | 39 focuses on the enzymatic degradation of starch, especially granular starch, by different types of microbial enzymes derived from several different places and conditions. It is suggested that some of the newly discovered aspects play a key role in the degradation process, such as the enzymatic mechanism. The introductory chapter (Chapter 1) reviews the current status of understanding of the structures and organization of starches, the regulation of degradative pathways of starch hydrolysis, and the structure‐function relationships of the α‐amylase enzymes involved.
The commercial enzyme preparation StargenTM 002 (DuPont BioScience, The Netherlands) enables the hydrolysis of native starch granules without the need for starch gelatinization. It consists of Aspergillus kawachi α‐amylase expressed in Trichoderma reesei and a glucoamylase from T. reesei, working synergistically to hydrolyze starch granules in a single step below liquefaction temperatures. The direct conversion of starch granules from tropical plants using StargenTM 002 was evaluated in Chapter 2. The susceptibility of starch granules of six different plants (arrowroot, corn, edible canna, potato, sago, and sweet potato) to StargenTM 002 was investigated. It was found that StargenTM 002 degrades sago and sweet potato at the same rate as corn starch, while the other starches were much less degraded. Sweet potato and sago therefore are recommended as alternative raw material sources for glucose production by the starch industry in tropical regions.
Chapter 3 describes the raw starch degrading α‐amylase from M. aurum strain B8.A, which was isolated from the waste water treatment facility of a potato‐processing plant. M. aurum B8.A was grown in minimal medium supplemented with potato starch, because α‐amylase synthesis is known to be induced by starch or its hydrolytic products (174, 281). A zymogram prepared using the cell‐free
Introduction
supernatant showed several protein activity bands. The potential raw starch degradation ability of M. aurum B8.A was investigated by monitoring the extent of hydrolysis of various types of raw starch granules. It turned out that M. aurum B8.A
Page | 40 is able to degrade raw cassava, wheat, and potato starch.
Furthermore, analysis of the whole M. aurum B8.A genome revealed an intriguing modular family GH13 α‐amylase (MaAmyA) containing two CBM25 and four FnIII modules arranged in tandem at the C‐terminal end. Chapter 4 explores further investigations into the function of CBM25 and FnIII domains in MaAmyA of M. aurum B8.A. The effects of deleting the CBM25, FnIII, and the unknown C‐terminal tail of MaAmyA were studied. Deletion of the CBM25s resulted in a loss of enzyme activity to raw starch. It was also shown that the deletion of the FnIII modules did not affect the MaAmyA’s hydrolytic activity or its binding to starch granules.
In Chapter 5, characterization of a novel α‐amylase from Bacillus megaterium NL3, BmaN1, isolated from Kakaban landlocked marine lake in Indonesia, is presented. Based on the amino acid sequence identity and a phylogenetic analysis, it was found that BmaN1 belongs to a new subfamily of GH13. Of the three catalytic residues conserved in GH13, Asp203 and Glu221 were present in BmaN1, while the third catalytic residue, Asp279 (Geobacillus thermoleovorans α‐amylase numbering) was replaced by a His. In spite of this, the BmaN1 amylase was active towards soluble substrates, amylopectin, cyclodextrins (CDs), and pullulan, as well as raw starch.
The results of this study are summarized and discussed in Chapter 6. A perspective of future research on the topic of starch degradation mechanism is also included in this chapter.
Chapter 1
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324. Kawaguchi, T., Nagae, H., Murao, S., and Arai, M. (1992) Purification and some properties of a haim-sensitive α-amylase from newly isolated Bacillus sp. No. 195. Biosci. Biotechnol. Biochem. 56, 1792–1796
325. Lo, H.-F., Lin, L.-L., Chen, H.-L., Hsu, W.-H., and Chang, C.-T. (2001) Enzymic properties of a SDS-resistant Bacillus sp. TS-23 α-amylase produced by recombinant Escherichia coli. Process Biochem. 36, 743–750
326. Paquet, V., Croux, C., Goma, G., and Soucaille, P. (1991) Purification and characterization of the extracellular α-amylase from Clostridium acetobutylicum ATCC 824. Appl. Environ. Microbiol. 57, 212–218
327. Horinouchi, S., Fukusumi, S., Ohshima, T., and Beppu, T. (1988) Cloning and expression in Escherichia coli of two additional amylase genes of a strictly anaerobic thermophile, Dictyoglomus thermophilum, and their nucleotide sequences with extremely low guanine-plus-cytosine contents. Eur. J. Biochem. 176, 243–253
328. Coronado, M. J., Vargas, C., Mellado, E., Tegos, G., Drainas, C., Nieto, J. J., and Ventosa, A. (2000) The α-amylase gene amyH of the moderate halophile Halomonas meridiana: Cloning and molecular characterization. Microbiology.
Chapter 1
146, 861–868
329. Coronado, M.-J., Vargas, C., Hofemeister, J., Ventosa, A., and Nieto, J. J. (2000) Production and biochemical characterization of an α-amylase from the moderate halophile Halomonas meridiana. FEMS Microbiol. Lett. 183, 67–71 Page | 73 330. Giraud, E., Brauman, A., Keleke, S., Lelong, B., and Raimbault, M. (1991) Isolation and physiological study of an amylolytic strain of Lactobacillus plantarum. Appl. Microbiol. Biotechnol. 36, 379–383
331. Liu, Y., Lei, Y., Zhang, X., Gao, Y., Xiao, Y., and Peng, H. (2012) Identification and phylogenetic characterization of a new subfamily of α-amylase enzymes from marine microorganisms. Mar. Biotechnol. 14, 253–260
332. Onishi, H., and Sonoda, K. (1979) Purification and some properties of an extracellular amylase from a moderate halophile, Micrococcus halobius. Appl. Envir. Microbiol. 38, 616–620
333. Freer, S. N. (1993) Purification and characterization of the extracellular α-amylase from Streptococcus bovis JB1. Appl. Environ. Microbiol. 59, 1398–1402
334. Shimizu, M., Kanno, M., Tamura, M., and Suekane, M. (1978) Purification and some properties of a novel α-amylase produced by a strain of Thermoactinomyces vulgaris. Agric. Biol. Chem. 42, 1681–1688
335. Najafi, M. F., and Kembhavi, A. (2005) One step purification and characterization of an extracellular α-amylase from marine Vibrio sp. Enzyme Microb. Technol. 36, 535–539
336. Planchot, V., and Colonna, P. (1995) Purification and characterization of extracellular α-amylase from Aspergillus fumigatus. Carbohydr. Res. 272, 97–109
337. Zaferanloo, B., Bhattacharjee, S., Ghorbani, M. M., Mahon, P. J., and Palombo, E. A. (2014) Amylase production by Preussia minima, a fungus of endophytic origin: Optimization of fermentation conditions and analysis of fungal secretome by LC- MS. BMC Microbiol. 14, 55–67
338. de Freitas, A., Escaramboni, B., Carvalho, A., de Lima, V., and de Oliva-Neto, P. (2014) Production and application of amylases of Rhizopus oryzae and Rhizopus microsporus var. oligosporus from industrial waste in acquisition of glucose. Chem. Pap. 68, 442–450
Introduction
339. Wanderley, K. J., Torres, F. A. G., Moraes, L. M. P., and Ulhoa, C. J. (2004) Biochemical characterization of α-amylase from the yeast Cryptococcus flavus. FEMS Microbiol. Lett. 231, 165–169
Page | 74 340. Galdino, A. S., Silva, R. N., Lottermann, M. T., Alvares, A. C. M., de Moraes, L. M. P., Torres, F. A. G., de Freitas, S. M., and Ulhoa, C. J. (2011) Biochemical and structural characterization of Amy1: An α-amylase from Cryptococcus flavus expressed in Saccharomyces cerevisiae. Enzyme Res. 2011, 1–7
341. Spencer-Martins, I., and Uden, N. (1979) Extracellular amylolytic system of the yeast Lipomyces kononenkoae. Eur. J. Appl. Microbiol. Biotechnol. 6, 241–250
342. Takeuchi, A., Shimizu-Ibuka, A., Nishiyama, Y., Mura, K., Okada, S., Tokue, C., and Arai, S. (2014) Purification and characterization of an α-amylase of Pichia burtonii isolated from the traditional starter “Murcha” in Nepal. Biosci. Biotechnol. Biochem. 70, 3019–3024
343. Zoltowska, K. (2001) Purification and characterization of α-amylases from the intestine and muscle of Ascaris suum (Nematoda). Acta Biochim. Pol. 48, 763–773
344. Bertoft, E., Andtfolk, C., and Kulp, S.-E. (1984) Effect of pH, temperature, and calcium ions on barley malt α-amylase isoenzymes. J. Inst. Brew. 90, 298–302
345. Rodenburg, K. W., Juge, N., Guo, X.-J., Sogaard, M., Chaix, J.-C., and Svensson, B. (1994) Domain B protruding at the third β-strand of the α/β barrel in barley α- amylase confers distinct isozyme-specific properties. Eur. J. Biochem. 221, 277– 284
346. Walker, G. J., and Hope, P. M. (1963) The action of some α-amylases on starch granules. Biochem. J. 86, 452–462
Supplementary information
Table S1.1 List of raw-starch degrading enzymes from different species.
Organism Type Source Raw Starch Activity Binding GH (sub) References Domain Family Aspergillus awamori var. glucoamylase n.d. corn CBM20 GH15 (282, 283) kawachi Aspergillus awamori KT-11 α-amylase, air in Bogor, cassava, corn, CBM20 GH13 (284, 285) glucoamylase Indonesia potato, rice, (act together) sago, sweet potato, wheat Aspergillus fumigatus α-amylase soil corn, pea, n.d. n.d. (280, 286) wheat Bacillus amyloliquefaciens α-amylase Turkish soil corn, potato, no SBD GH13_5 (287) rice, sweet potato, wheat Bacillus aquimaris α-amylase soft coral cassava, corn, no SBD GH13_36 (288) Sinularia sp. potato, rice Bacillus cereus β-amylase n.d. corn CBM20 GH14 (225, 289, 290) Bacillus circulans α-amylase soil n.d. CBM25 GH119 (167, 291) Bacillus firmus CGTase soil cassava, rice CBM20 GH13_2 (292, 293) n.d., no data; CBM, carbohydrate-binding module; SBD, starch-binding domain; GH, glycoside hydrolase Page
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Table S1.1 continued. List of raw-starch degrading enzymes from different species.
Organism Type Source Raw Starch Activity Binding GH (sub) References Domain Family Bacillus sp. no. 195 α-amylase soil corn, potato, rice, CBM25 GH13_32 (236) sweet potato, wheat Bacillus sp. strain TS-23 α-amylase soil corn CBM20 GH13 (223, 294) Corticium rolfsii glucoamylase tomato stem cassava, corn, CBM20 GH15 (220, 295) potato, rice, sago, sweet potato Cryptococcus flavus α-amylase culture collection cassava CBM20 GH13_1 (296) Curculigo pilosa (flower β-amylase tubers corn, potato, rice, n.d. n.d. (297) plant) wheat Glycine max (soybean) β-amylase n.d. rice no SBD GH14 (298, 299) Homo sapiens (human) maltase- small intestinal banana, cassava, no SBD GH31 (101, 300) glucoamylase corn, rice, wheat Hordeum vulgare (barley) α-amylase germinated barley no SBD GH13_6 (301, 302) barley Klebsiella pneumoniae AS- CGTase n.d. cassava, corn, CBM20 GH13_2 (303, 304) 22 potato, rice, wheat Lactobacillus amylovorus α-amylase Human saliva corn no SBD n.d. (242) Lactobacillus plantarum A6 α-amylase retted cassava cassava CBM26 GH13_28 (240, 305) n.d., no data; CBM, carbohydrate-binding module; SBD, starch-binding domain; GH, glycoside hydrolase
Table S1.1 continued. List of raw-starch degrading enzymes from different species.
Organism Type Source Raw Starch Activity Binding GH (sub) References Domain Family Marine metagenomics library α-amylase subsurface rice, wheat CBM69 GH13_37 (215, 306) sediments in South China sea Microbacterium aurum B8.A α-amylase potato plant corn, wheat, CBM25 GH13_32 (264) waste water potato Populus x canadensis α-amylase leaf and wood corn, potato n.d. n.d. (307, 308) Moench Rhizopus oryzae glucoamylase commercial corn CBM21 CBM15 (309, 310) Sitophilus oryzae (insect) α-amylase n.d. amaranth, pea no SBD CBM13_15 (311) Streptococcus bovis 148 α-amylase cattle rumen corn CBM26 GH13 (312) Streptomyces α-amylase culture cassava, corn, n.d. n.d. (313) thermocyaneoviolaceus collection rice, sago, IFO14271 sweet potato, potato n.d., no data; CBM, carbohydrate-binding module; SBD, starch-binding domain; GH, glycoside hydrolase
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Table S1.2 Physicochemical properties of α‐amylases
Source Molecular mass Optimum pH Optimum Metal References (kDa) temperature (oC) requirement Archaea Flavobacteriaceae sinomicrobium 52 6.0 50 Ca2+ (314) Haloarcula japonica 73 6.5 45 no (171) Methanococcus jannaschii 33 7.0 120 no (315) Pyrococcus furiosus 76 4.5 90 no (196, 316) Pyrococcus woesei 70 5.5 100 no (317) Thermococcus profundus 43 5.5 80 Ca2+ (318) Bacteria Bacillus acidocaldarius 68 3.5 75 Ca2+, Mg2+ (319) Bacillus amyloliquefaciens 52 6.0 55 Ca2+ (287) Bacillus caldolyticus 10 5.4 70 Ca2+ (176, 320) Bacillus licheniformis 58 6.0 90 Ca2+ (321) Bacillus megaterium 59 5.8 70 Ca2+ (322) Bacillus stearothermophilus 52.7 6.0 65 Ca2+ (323) Bacillus sp. no.195 60 7.0 45 n.d. (324) Bacillus sp. strain TS-23 65 9.0 65 Co2+, Mn2+, Fe2+ (294, 325) Chloroflexus aurantiacus 210 7.5 71 Ca2+ (175) n.d., no data
Table S1.2 continued. Physicochemical properties of α‐amylases
Source Molecular mass Optimum pH Optimum Metal References (kDa) temperature (oC) requirement Clostridium acetobutylicum ATCC 84 5.6 45 no (326) 824 Dictyoglomus thermophilum 67 5.5 80 n.d. (327) (AmyB) Dictyoglomus thermophilum 59 5.5 70 n.d. (327) (AmyC) Geobacillus thermoleovorans 59 5.0 80 no (188) Halomonas meridiana 49 7.0 37 Ca2+ (328, 329) Lactobacillus plantarum 104.7 5.0 55 n.d. (330) Marine metagenomic library 90 6.5 50 n.d. (331) Micrococus halobius 89 6.5 50-55 Ca2+ (332) Streptococcus bovis JB1 77 6.5 35 no (333) Streptococcus bovis 148 79 5.5 55 no (312) Streptomyces 49 6.5 40 Ca2+ (313) thermocyaneoviolaceus IFO14271 Thermotoga maritima MSB8 50 7.0 70 Ca2+ (170) Thermoactinomyces vulgaris 67 5.0 40 Ca2+ (130, 334) n.d., no data
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Table S1.2 continued. Physicochemical properties of α‐amylases
Source Molecular mass Optimum pH Optimum Metal References (kDa) temperature (oC) requirement Vibrio sp. 52.5 6.5 60 Ca2+, Co2+, Mg2+, (335) Mn2+, Fe2+ Fungi Aspergillus fumigatus 65 5.5 40 Ca2+ (minor) (336) Preussia minima 70 9.0 25 Ca2+, Mn2+ (337) Rhizopus oryzae 48 5.0 50 Ca2+ (338) Yeasts Cryptococcus flavus 67 5.5 50 no (339, 340) Lipomyces kononenkoae 34 5.5 40 Ca2+ (341) Pichia burtonii 51 5.0 40 Ca2+ (342) Eukaryotes (other) Ascaris suum (nematoda) 59 8.2 30 and 50 Ca2+ (343) Hordeum vulgare (barley) Amy1 45 5.5 70 Ca2+ (344, 345) Hordeum vulgare (barley) Amy2 45 5.0 65 Ca2+ (344, 345) Human saliva 55.2 6.9 25 Ca2+ (346) Populus canadensis Moench (leaf) 44 6.5 30 Ca2+, Mg2+ (307, 308) Sitophilus oryzae (insect) 53 5.0 40 no (311) n.d., no data
Chapter
Cold Enzymatic Hydrolysis of Indonesian Root and Tuber
Starches
Fean D. Sarian, Hans Leemhuis, Zeily Nurachman, Marc J.E.C. van der Maarel, and
Dessy Natalia
To be submitted for publication Cold Enzymatic Hydrolysis
Abstract
In most tropical countries, carbohydrate‐based agricultural products occur in large quantities. Wider utilization of local starch crops will offer various economic and ecological advantages. Local starch crops can be a catalyst for rural industrial development and eventually open up new markets. The potential glucose yield from the typical Indonesia starches (edible canna, arrowroot, sago, and sweet potato) using the cold‐starch hydrolyzing enzyme preparation StargenTM 002 was compared Page | 82 to that of corn and potato. Sago and sweet potato gave similar glucose yields as corn, while arrowroot and edible canna gave much lower amounts. This study demonstrates that sago and sweet potato starches provide interesting alternatives to corn starch in a cold hydrolysis conversion process of starch to glucose.
Chapter 2
Introduction
Starch is found in the seeds, roots and tubers of many plants and serves as energy storage for the seedling. Starch is a mixture of amylose and amylopectin, two polymers of α(14) linked glucose units that differ in size and degree of α(16) branching. Amylose has virtually no α(16) branches and has a degree of polymerization of a few hundred to a few thousands whereas amylopectin has on average 5% α(16) linkages and is very large, with as many as several thousand Page | 83 anhydroglucopyranose residues (1). In general, starch contains 20‐30% amylose and 70‐80% amylopectin, though there are exceptions such as waxy (amylopectin) starches and high amylose starches. These two molecules are packed within small starch granules of 1 to 110 µm, depending on the botanical origin in alternative layers of crystalline and amorphous regions (1).
Starch is a raw material for the industrial production of glucose syrups, which in turn is converted into high fructose corn syrup, a sweetener for soft drinks and filling agent of many (liquid) food products. Glucose syrups also serve as feedstock for large‐scale fermentative production of fine chemicals, organic acids and ethanol. To produce glucose syrup, the starch slurry is first gelatinized at high temperatures, followed by liquefaction with a thermostable ‐amylase and finally saccharification with glucoamylase or amyloglucosidase (2). Liquefaction and saccharification are commonly performed at high temperatures to reduce the viscosity of the slurry and prevent microbial spoilage (3). The glucose obtained can then be converted to ethanol via fermentation. One way of reducing the amount of energy needed to produce ethanol from starch is to skip the gelatinization step and combine the saccharification and fermentation process in one vessel, using enzymes that are able to degrade native starch granules (4–7).
A few years ago a raw starch degrading (RSD) enzyme cocktail specially developed for cold raw material processing, namely StargenTM 001, was introduced on the market by Genencor, now Dupont Industrial Biosciences. Robertson et al. (8) Cold Enzymatic Hydrolysis
estimated that up to 20% energy saving is possible by using a RSD type enzyme cocktail. An additional advantage is that the recovery of value‐added products from the cold hydrolysis process is more efficient (9, 10). StargenTM 002 is a second‐ generation enzyme released commercially by Genencor International, BV to complement StargenTM 001 in hydrolyzing grain crops. This enzyme is an optimized blend of Aspergillus kawachi α‐amylase and a glucoamylase from Trichoderma reesei (11). To date, most research has been conducted on StargenTM 002 to improve the liquefaction and saccharification steps in the glucose and bioethanol production Page | 84 from grain crops such as corn and rice starch (12–14). No experimental information is available on the enzymatic saccharification of tropical starches using StargenTM 002.
The main raw material for bioethanol production is still corn starch, yielding almost 300 billion liters of bioethanol per year (15). By far the main sources of starch for the production of bioethanol in Asia are cassava, wheat, and sweet potato (16). Several studies have explored the potential use of alternative starch containing crops for biofuel production, thereby increasing the range of starch sources to be used (17–20). Besides cassava and corn, Indonesia has several other biomass resources that can be developed and utilized as energy sources such as arrowroot, edible canna, sago, and sweet potato.
Sago palm (Metroxylon sagu) grows mainly in Southeast Asia and Melanesia, and is one of Indonesia’s main natural, renewable resources. More than half of all Asian sago palm trees grow in Indonesia, almost exclusively in its Eastern part, especially in Mollucan and Papua. However, the use of sago palm starch has received little attention compared to the starch of typical root plants such as cassava, despite the ability of the sago palm to grow on constraint soils such as wet lowland, acidic, brackish, and peat soils without any soil improvements (21). Sweet potato (Ipomoea batatas L.) is cultivated in over 100 developing countries and is an important root/tuber crop in Indonesia. Sweet potato production in Indonesia ranks fourth in the world with 2.4 million metric tons in 2013 (22). Among the tropical tuber crops, Chapter 2
sweet potato has a higher starch yield per unit land cultivated than grains and comes next to cassava in starch content (23). Arrowroot is a starch‐containing root crop, native to the Andean region in South America. However, in recent years the plant has been cultivated and used as an alternative source of carbohydrate in several regions of Indonesia (24). There are several types of arrowroot plants cultivated, including West Indian arrowroot (Maranta arundinacea), or arrowroot, and Queensland arrowroot (Canna edulis), or edible canna (25). The arrowroot and edible canna tubers contain more than 75% starch on a dry weight basis (25, 26). Page | 85
This communication reports the direct conversion of tropical granular starches into glucose via a cold enzymatic process using the commercial enzyme cocktail StargenTM 002. Cold Enzymatic Hydrolysis
Results and Discussion
Numerous starch degrading enzymes have been described, however, only a small percentage of these are capable of degrading starch in its native crystalline state and carried out below gelatinization temperature (cold hydrolysis) (28–30). One of the commercial enzyme preparations that hydrolyzes native granular starch is StargenTM 002 from Dupont Industrial Biosciences (USA). This is a blend of a fungal α‐amylase and a glucoamylase (11). Page | 86
100
80
60
40
20
0 arrowroot sweet sago potato edible relative amount of glucose (%) potato canna starch source Fig 2.1 Glucose production from several tropical starches (40% dry solids, w/v) relative to the amount of glucose produced from corn starch (100% value is expressed as 89.3 grams of glucose equivalent/L ) after 24 h of hydrolysis at 30 oC. 20 IU of StargenTM 002 (0.5 IU per gram of starch) were used. The values recorded are the mean values from three experiments, each in triplicate.
Enzymatic hydrolysis
Industrial starch hydrolysis is done at high dry solid concentrations of up to 35% (w/v) starch (31, 32). The efficiency at which native starch is degraded depends not only on the enzyme employed, but also on the botanical origin of the starch (33, 34). The different tropical starches were incubated with StargenTM 002 at 40% (w/v) starch and the amount of glucose formed after 24 h was measured (Fig. 2.1). The Chapter 2
amount of glucose produced from sago and sweet potato starches by StargenTM 002 were almost equal to that of corn starch and considerably higher than that of arrowroot, edible canna or potato starches. The relative order of glucose yield after 24 h was as follows: corn > sago > sweet potato > arrowroot > potato > edible canna (Fig. 2.1).
100 arrowroot canna potato Page | 87 75 corn sweet potato sago
50
25 Degree of Hydrolysis (%) 0 01234567 incubation time (h)
Fig. 2.2 The time‐course curve of StargenTM 002 activity on the hydrolysis of different granular starches at 30 oC. Reaction mixtures contained 15% (w/v) of various starch granules and 20 IU of StargenTM 002. Data shown are average values of three different experiments.
To evaluate the effect of incubation time on the extent of starch granule hydrolysis, the different starches (15%, w/v) were incubated with StargenTM 002 for 7 h (Fig. 2.2). Initially, sago and sweet potato starch were degraded faster than corn, while arrow root starch was degraded much slower and potato and canna starch were not degraded at all (Fig. 2.2). The observation that the hydrolysis rate continued to increase indicates that the enzymes remained active throughout the course of this degradation study. Depending on the degradation conditions, the hydrolysis rate can either plateau after prolonged time periods, or remain essentially linear. Based on effectiveness and production of glucose, it is clear that sago and sweet potato will be Cold Enzymatic Hydrolysis
the most effective source of starch to use in the StargenTM 002 cold enzymatic hydrolysis process.
Scanning Electron Microscopy
To explain the variation in the effectiveness of enzymatic degradation, the granules were analyzed by SEM after incubation with or without addition of 20 IU StargenTM Page | 88 002. Cold hydrolyzed sweet potato starch yielded the highest percentage of hydrolysis (degree of hydrolysis, DH), about 86.5%, followed by corn, sago, arrowroot, potato, and edible canna. As a comparison, the DH of starch granules after incubating with acid was 100% (complete hydrolysis).
The arrowroot, canna, corn, potato, sago, and sweet potato granules varied considerably in size and shape (Fig. 2.3A – 3F). The surface of the granules from corn (Fig. 2.3A) and sago (Fig. 2.3C) appeared to be less smooth than arrowroot (Fig. 2.3E), edible canna (Fig. 2.3F), potato (Fig. 2.3D), and sweet potato (Fig. 2.3B). A magnified view of corn and sago granules revealed small pores and cracks that were randomly distributed on the surface (see arrows in Fig. 2.3A and 2.3C, respectively). Such pores have been observed on the granular surfaces of corn, sorghum, millet, wheat, rye, barley, and sago (35), while granules of tapioca, rice, oat, canna, and arrowroot appeared to have no pores (36). The origin of these surface pores varies. It has been suggested that pores are a result of the drying processes of starch granules or that they are simply artefacts caused by sample preparation for electron microscopy. Pores may also have formed by the action of endogenous amylases during starch synthesis or during the treatment of native starch (36). Fannon et al. (36) proposed that the presence of pores is a natural feature of many starch granules and that they do not appear as a response to mechanical treatment. However, their study also showed that amylolysis and retrogradation also could lead to an increase in the size and number of pores (36).
Chapter 2
After 48 h of incubation, two modes of enzyme action were observed. Enzyme treated corn (Fig. 2.3A) and sago (Fig. 2.3C) showed deep holes and a more porous structure compared to the other starches. Corn starch granules displayed homogenously distributed large cavities while the degradation did not occur uniformly in sago starch granules. Some degraded sago starch granules displayed rough and highly perforated surfaces while some other granules were still intact (Fig. 2.3C). The pores of degraded starches gradually widened, but the size, depth, and width were different for each pore. For all starch granules studied, as soon as Page | 89 the interior was reached, the inner layers were hydrolyzed, resulting in “Swiss cheese” pattern. In contrast, the StargenTM 002 treated potato starch granule samples have a rough surface with small pores (Fig. 2.3D). SEM of enzyme‐treated sweet potato, arrowroot, and edible canna granules (Fig. 2.3B, E, F, respectively) indicated that the mode of attack on those starches was different than on corn starch. The granules of sweet potato and arrowroot starches had more or less roughened surfaces, while edible canna starch granules exhibited disc‐like depressions on the surface.
Edible canna and potato starch were far less susceptible to enzymatic hydrolysis, which is in agreement with the results of Kimura and Robyt (37) who studied glucoamylase activity with various starch granules. The number and size of granules and the structure of the starches apparently influences the rate of enzymatic degradation, but this is not yet fully understood (33, 37). Edible canna and potato starches have smooth and even surfaces, while the surface of corn and sago starch granules have cracks and pores going deep in the interior of the granule, enabling a more easy penetration of amylase enzymes into the granules. According to Juszczak et al. (38) and Kimura and Robyt (37), the natural pores present on starch surfaces could enhance enzyme penetration into starch granules during enzymatic attack resulting in degradation from inside (37–39). Cold Enzymatic Hydrolysis
A B C
- B - - C
Page | 90 C + + + D D E F
- E - -
F + + + Fig 2.3 SEM observations of various kind of starches after incubation with or without StargenTM 002 for 48 h at 30 oC. Starch granules (5% (w/v)) incubated without (‐) or with 20 IU of StargenTM 002 (+) from corn (A, magnification 4,000 x and 4,500 x; below and above, respectively) sweet potato (B, 6,000 x), sago (C, magnification 1,000 x and 1,500 x; below and above, respectively), potato (D, 7,500 x and 2,000 x; below and above, respectively), arrowroot (E, 3,500 x and 4,500 x; below and above, respectively), and edible canna (F, 1,400 x and 750 x; below and above, respectively). Arrows in SEM images indicate the presence of pores and some cracks on the (untreated) granule surface.
Chapter 2
The penetration of starch granules by hydrolyzing enzymes is restricted and only possible through pores or channels (37, 38). This explains the high rate of amylase activity that is associated with corn and sago starch granules.
Enzymatic hydrolysis at longer incubation times
With longer incubation times (up to 125 h), corn starch gave the highest glucose Page | 91 yield (Fig. 2.4A). After about 25 h the rate of glucose formation differed considerably. Compared to corn starch, sweet potato and sago gave less glucose over 125 h of incubation but this was still much more than potato, arrowroot, and edible canna (Fig. 2.4). TLC analysis of the products of all incubations clearly showed that StargenTM 002 hydrolyzed starch granules to glucose only, as oligosaccharides were not visible in the final reaction (data not shown).
The observation that StargenTM 002 is highly active towards sweet potato and sago, indicates that these abundantly present tropical starches can be used in addition to corn starch in the manufacturing of glucose. The high level of glucose obtained from sago and sweet potato starch incubated with StargenTM 002 under low‐temperature hydrolysis conditions justifies the use of these raw materials in addition to corn starch as renewable resources in the production of bioethanol in a simultaneous saccharification and fermentation process. This is potentially possible at a relatively small scale already, thus requiring little investment capital, opening routes towards local/rural industrial activities in developing countries. Cold Enzymatic Hydrolysis
D A 160 40 140 120 30 100
80
20 60 Glucose (mg/mL) Glucose 40 (mg/mL) Glucose 10 20 0 0 0 25 50 75 100 125 0 25 50 75 100 125 time (h) time (h)
B 120 E
100 Page | 92 100
80 75
60 50 40 Glucose (mg/mL) Glucose (mg/mL) Glucose 25 20
0 0 0 25 50 75 100 125 0 25 50 75 100 125 time (h) time (h)
C 120 F 40
100 30 80
60 20
40 Glucose (mg/mL) 10 Glucose (mg/mL) Glucose 20
0 0 0 255075100125 0 25 50 75 100 125 time (h) time (h) Fig 2.4 Time course of enzymatic hydrolysis of various kind of starches during incubation with StargenTM 002. The reaction was conducted with StargenTM 002 at 20 IU and a 15% (w/v) starch granule mixture in 50 mM acetate buffer at 30 oC and at pH 4.0 for the indicated time. A. corn; B. sweet potato; C. sago; D. potato; E. arrowroot; and F. edible canna. The graphs and the trend lines were drawn using OriginPro software.
Acknowledgments
FDS received a grant of the Bernoulli Sandwich PhD Program of the University of Groningen. We would like to send our gratitude to Lubbert Dijkhuizen for his valuable discussion and critical review of this manuscript.
Chapter 2
Materials and Methods
Materials
StargenTM 002 is a commercial enzyme mixture marketed by DuPont Industrial Biosciences (Palo Alto, USA) and is a blend of Aspergillus kawachi glucoamylase and Trichoderma reesei α‐amylase (11). The sample used in this study was a gift from B. Koops (DuPont Industrial Biosciences, Leiden, The Netherlands). Corn (Zea mays), Page | 93 edible canna (Canna discolor syn. C. edulis), arrowroot (Maranta arundinacea), potato (Solanum tuberosum), and gandola’s sweet potato (Ipomoea batatas) starch were purchased from a local market in Bandung, West Java (Indonesia). Sago (Metroxylon sagu) starch was a generous gift from Mr Y. Mandik (Papua, Indonesia). All other chemicals were from Sigma (St. Louis, USA) and were reagent grade.
Starch isolation
Fresh edible canna, arrowroot, potato, and sweet potato were washed, chopped, and then grounded in a blender. The starch fraction was separated from the pulp by sieving through bolting cloth. The filtrate was allowed to settle for starch sedimentation and the supernatant was decanted. The white starch was collected and washed with ethanol (30%) at least three times to remove fat. The starch was dried in an oven at 50oC for about 48 h. Finally the dried starch was grounded into powder and stored at room temperature.
Granular starch hydrolysis
Granular starch‐degrading activity was measured by following the degree of hydrolysis with the GOPOD method (Megazyme, Ireland). Briefly, 5‐40% (w/v) granular starch was mixed with 1 mL of the appropriately diluted StargenTM 002 1X (20 IU/mL) in 50 mM acetate buffer pH 4.0. The enzyme activity units are given as provided by the enzyme manufacturer. Enzymatic reactions were performed in Cold Enzymatic Hydrolysis
triplicate at 30 oC with agitation at 200 rpm. Progress of the reaction was followed in time by taking samples in triplicate; the reaction was stopped by centrifugation at 12,000 x g for 10 min and analyzing the glucose content in the supernatant. The absorbance at 510 nm was taken as a measure of glucose (dextrose) equivalent (DE). Glucose (0.01 – 0.1 mg/mL) was used as a standard. The total amount of glucose was assayed by the same method after acid hydrolysis (6 M HCl for 120 min at 100 oC) . All values are averages of three determinations. DE was calculated as follows:
Page | 94 DE = x 100%
The degree of hydrolysis (DH) was defined by the following equation (27):
DH = x 100%
Scanning Electron Microscopy
To visualize the starch degrading activity of StargenTM 002, starch granules were incubated with this enzyme preparation for 2 days. 15% (w/v) granular starch was mixed with 1 mL of the appropriately diluted StargenTM 002 1X (20 IU/mL) in 50 mM acetate buffer pH 4.0. Incubation was at 30 oC with agitation at 200 rpm. Then the starch granules were washed twice with water‐ethanol (1:1) and dried by heating in an oven at 37 oC for one day. The samples (dried starch powder) were mounted on Scanning Electron Microscopy (SEM) stubs with double‐sided adhesive tape and coated with gold. Scanning electron micrographs were taken using the JSM Scanning Microscope (JEOL) in the PPPGL (Geology Research and Development Centre), Bandung, Indonesia.
Chapter 2
References
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9. Gibreel A, Sandercock JR, Lan J, Goonewardene L a., Zijlstra RT, Curtis JM, Bressler DC. 2009. Fermentation of barley by using Saccharomyces cerevisiae: Examination of barley as a feedstock for bioethanol production and value-added products. Appl Environ Microbiol 75:1363–1372.
10. Gohel V, Duan G. 2012. No-cook process for ethanol production using indian broken rice and pearl millet. Int J Microbiol 2012:1–9.
11. Genencor. 2008. Product information sheet-STARGEN 002 Technical Bulletin.
12. Adams JMM, Teunissen PJM, Robson G, Dunn-Coleman N, Donnison IS. 2011. Scanning electron microscopy and fermentation dtudies on delected known maize starch mutants using StargenTM enzyme blends. Bioenergy Res 5:330–340. Cold Enzymatic Hydrolysis
13. Kuhn RC, Mazutti MA, Foletto EL, Prá VD, Zimmermann E, Souza M, Foletto VS, Maleski TPS, Lunelli FC, Sfalcin P. 2015. Glucose obtained from rice bran by ultrasound-assisted enzymatic hydrolysis. Ing e Investig 35:61–66.
14. Chu-Ky S, Pham T-H, Bui K-LT, Nguyen T-T, Pham K-D, Nguyen H-DT, Luong H-N, Tu V-P, Nguyen T-H, Ho P-H, Le T-M. 2015. Simultaneous liquefaction, saccharification and fermentation at very high gravity of rice at pilot scale for potable ethanol production and distillers dried grains composition. Food
Page | 96 Bioprod Process 98:79–85. 15. Rajagopal D, Sexton SE, Roland-Holst D, Zilberman D. 2007. Challenge of biofuel: Filling the tank without emptying the stomach? Environ Res Lett 2:1–9.
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18. Pandey A, Soccol CR, Mitchell D. 2000. New developments in solid state fermentation: I-bioprocesses and products. Process Biochem 35:1153–1169.
19. Rowe RL, Street NR, Taylor G. 2009. Identifying potential environmental impacts of large-scale deployment of dedicated bioenergy crops in the UK. Renew Sustain Energy Rev 13:271–290.
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21. Sato T, Yamaguchi T, Takamura T. 1979. Cultivation, harvesting, and processing sago palm (in Japanese). Japanese J Trop Agric 23L:130–136.
22. FAOSTAT. 2015. Food and Agriculture Organization of the United Nations. Agric data (last Updat June, 2015).
23. Duvernay WH, Chinn MS, Yencho GC. 2013. Hydrolysis and fermentation of sweetpotatoes for production of fermentable sugars and ethanol. Ind Crops Prod 42:527–537.
24. Hermann M, Heller J. 1997. Andean roots and tubers Ahipa, Arracacha, Maca, and Yacon. Volume 21 of promoting the conservation and use of underutilized and neglected crops. IPGRI, 1997. Chapter 2
25. Odeku OA. 2013. Potentials of tropical starches as pharmaceutical excipients: A review. Starch - Stärke 65:89–106.
26. Soni PL, Sharma H, Srivastava HC, Gharia MM. 1990. Physicochemical properties of Canna edulis starch — comparison with maize starch. Starch - Stärke 42:460–464.
27. Wang WJ, Powell AD, Oates CG. 1995. Pattern of enzyme hydrolysis in raw sago starch: Effects of processing history. Carbohydr Polym 26:91–97.
28. Sun H, Zhao P, Ge X, Xia Y, Hao Z, Liu J, Peng M. 2010. Recent advances in Page | 97 microbial raw starch degrading enzymes. Appl Biochem Biotechnol 160:988– 1003.
29. Oates CG. 1997. Towards an understanding of starch granule structure and hydrolysis. Trends Food Sci Technol 8:375–382.
30. Bertoldo C, Antranikian G. 2002. Starch-hydrolyzing enzymes from thermophilic archaea and bacteria. Curr Opin Chem Biol 6:151–160.
31. Lee SH, Shetty JK, Strohm BA. 2015. Liquefaction and saccharification of granular starch at high concentration. US 2015/0037844 A1. United States.
32. Myat L, Ryu G-H. 2014. Effect of thermostable α-amylase injection on mechanical and physiochemical properties for saccharification of extruded corn starch. J Sci Food Agric 94:288–295.
33. Gallant DJ, Bouchet B, Buléon A, Pérez S. 1992. Physical characteristics of starch granules and susceptibility to enzymatic degradation. Eur J Clin Nutr 46 Suppl 2:S3–16.
34. Tester RF, Karkalas J, Qi X. 2004. Starch structure and digestibility enzyme- substrate relationship. World Poul Sci J 60:186–195.
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36. Fannon JE, Hauber RJ, Bemiller JN. 1992. Surface pores of starch granules. Cereal Chem 69:284–288.
37. Kimura A, Robyt JF. 1995. Reaction of enzymes with starch granules: kinetics and products of the reaction with glucoamylase. Carbohydr Res 277:87–107. Cold Enzymatic Hydrolysis
38. Juszczak L, Fortuna T, Krok F. 2003. Non-contact atomic force microscopy of starch granules surface. Part II. Selected cereal starches. Starch - Stärke 55:8–16.
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Page | 98 Chapter
Enzymatic Degradation of Granular Potato Starch
by Microbacterium aurum strain B8.A
Fean D. Sarian, Rachel M. van der Kaaij, Slavko Kralj, Dirk J. Wijbenga†, Doede Binnema, Marc J.E.C. van der Maarel, and Lubbert Dijkhuizen
Applied Microbiology and Biotechnology (2012) : doi:10.1007/s00253‐011‐3436‐7 Enzymatic Degradation by M. aurum B8.A
Abstract
Microbacterium aurum strain B8.A was isolated from the sludge of a potato starch‐ processing factory on the basis of its ability to use granular starch as carbon‐ and energy source. Extracellular enzyme hydrolyzing granular starch were detected in the growth medium of M. aurum B8.A, while the type strain M. aurum DSMZ 8600 produced very little amylase activity, and hence was unable to degrade granular starch. The strain B8.A extracellular enzyme fraction degraded wheat, tapioca, and potato starch at 37 oC, well below the gelatinization temperature of these starches. Starch granules of potato were hydrolyzed more slowly than of wheat and tapioca, probably due to structural differences and/or surface area effects. Partial hydrolysis of starch granules by extracellular enzymes of strain B8.A resulted in large holes of irregular sizes of wheat and tapioca and many smaller pores of relatively Page | 100 homogenous size in case of potato. The strain B8.A extracellular amylolytic system produced mainly maltotriose and maltose from both granular and soluble starch substrates; also, larger malto‐oligosaccharides were formed after growth of strain B8.A in rich medium. Zymogram analysis confirmed that a different set of amylolytic enzymes was present depending on the growth conditions of M. aurum B8.A. Some of these enzymes could be partly purified by binding to starch granules.
Chapter 3
Introduction
Starch is the major carbohydrate reserve in plants. It consists of amylopectin, a large α‐(1,4) linked anhydroglucose polymer with α‐1,6‐ linked side chains, and amylose, a linear α‐(1,4) linked anhydroglucose polymer. Regular starch contains on average 20‐25% amylose and 75‐80% amylopectin (1). In nature, starch in plants occurs in the form of inert granules that differ in size and structure between species (2).
Starch is one of the major caloric sources in the human diet. It is also used in many non‐food applications, e.g. in paper making, adhesives, and oil drilling fluids. The last few years have seen a considerable interest in starch as a raw material for the production of bioethanol as an automotive fuel supplement. For most of these Page | 101 applications, starch needs to be solubilized, degraded or modified to change its properties, which can be performed either physically, chemically or enzymatically. A well‐established industrial conversion process is the hydrolysis of starch into maltodextrins, maltose and glucose syrups; a process which is performed with acids or specific enzymes. Acid hydrolysis of starch had widespread use in the past but is now largely replaced by enzymatic processing (1, 3, 4). The industrial enzymatic conversion of starch consists of a heating step to disrupt the granular structure (gelatinization), liquefaction using a thermostable α‐amylase and subsequent saccharification by glucoamylase and pullulanase enzymes. To reduce processing cost and save energy, enzyme preparations have been developed that hydrolyze granular starch below the gelatinization temperature (5, 6). A key property of ‐amylases acting on starch granules is the presence of one or more Starch Binding Domain (SBD). Such auxiliary domains bring the biocatalyst into close and prolonged vicinity with starch granules, allowing hydrolysis of the insoluble substrate (7–9). Carbohydrate‐binding domains have been classified into 64 different families, based on their amino acid similarity and fold (see http://www.cazy.org/Carbohydrate‐Binding‐Modules.html for more information). Nine of these families contain different types of SBDs. CBM20 is one of the best described types; it is related to CBM21, 48, and 53, and occurs in a range of different enzymes with activity on starch including cyclodextrin glucanotransferase, β‐amylase, and phosphorylating enzymes (10). Another clan is formed by CBM25 Enzymatic Degradation by M. aurum B8.A
and 26, which often occur in tandem repeats (11).
Amylases are widespread in nature and are produced by all kind of organisms ranging from microorganisms, plants, insects, to mammals (12–14). The main function of theses amylases is the degradation of starch and glycogen to support the organisms’ growth. Most amylases belong to the glycoside hydrolase family 13 (see http://www.cazy.org/GH13.html for a detailed review) or glycoside hydrolase family 57 (http://www.cazy.org/GH57.html) (15). Amylases and related enzymes are retaining enzymes using a double displacement mechanism with the formation of a covalent enzyme‐substrate intermediate (16).
Page | 102 Various microorganisms have been found to produce granular starch degrading‐ amylolytic enzymes; many of them are fungal species. Ueda et al. (1974) reported that the fungi Aspergillus awamori (17) and Aspergillus oryzae (18) produce glucoamylases that adsorb to the granules while degrading them. There are also several reports of granular starch‐degrading enzymes from bacteria. For example, Hayashida et al. (1988) reported that an α‐amylase produced by Bacillus subtilis strain 65 is able to degrade potato and corn starch granules (19). A similar ability to degrade granular starch was found in Bacillus circulans, Bacillus amyloliquefaciens and Bacillus cereus (20, 21). Within the Actinomycetes, Streptomyces sp. no. 4 produces ‐amylases which hydrolyzes cassava starch granules (22) and Streptomyces limosus produces an α‐amylase that hydrolyzes corn starch granules with the release of maltose (23).
Starch granules are relatively rigid structures and several factors, such as the crystalline structure, presence of cracks and pores, and the granule sizes contribute to the relative resistance of granules to enzymatic degradation (24–26). Large starch granules have a relatively smaller surface area than smaller granules thus, reducing the chance for enzymes to become absorbed. Therefore large granules are generally less easily degraded than smaller granules (27–29). Chapter 3
Most of the known granular starch‐degrading enzymes show a reasonable hydrolyzing activity on corn, maize and cassava starch granules, but only few enzymes are able to degrade potato starch granules. Enzymes that are capable of degrading potato starch granules are attractive for increasing the range of starch sources suitable for enzymatic hydrolysis. Their detailed characterization may reveal specific protein structural features important for starch granule degradation.
We have isolated a bacterial species from the sludge of a potato starch‐processing factory, which was identified as Microbacterium aurum strain B8.A. It was shown to be capable of degrading potato starch granules, in contrast to the type strain M. aurum DSMZ 8600 which showed only minor amylase activity. In this paper, we report characteristics of starch degradation by M. aurum B8.A. It was found to Page | 103 employ an amylolytic enzyme system that initiates granular potato starch degradation by making numerous small pores in the starch granules. Enzymatic Degradation by M. aurum B8.A
Results
Selection and identification of M. aurum B8.A
Samples of several mixed liquid cultures containing granular starch‐degrading activity were plated out and covered with a thin agar layer containing potato starch granules. Between 1 and 5% of all colonies that appeared produced a clear halo in the turbid top layer, indicating granular starch‐degrading activity. Pure cultures were isolated from several halo‐forming colonies by repeatedly transferring a single colony to a new agar plate. These isolates were divided into six groups according to their properties, e.g., colony appearance, pigmentation and type of degradation of the starch granules (pore formation or peeling of layers of the granule). One of these groups, containing identical colonies, was selected for further analysis. The selected Page | 104 strain, B8.A, was a Gram‐positive, aerobic, non‐motile, coccoid rod‐shaped (0.6 x 0.8 µm in size) bacterium, which did not form spores. Growth of the strain occurred within a pH range of 6.0 to 9.0. The strain was deposited with the BCCM/LMG culture collection of the University of Gent, Belgium under the deposit number LMG S‐26033.
Analysis of the 16S rDNA gene of the B8.A isolate revealed highest similarity with Microbacterium species. A phylogenetic tree based on the 16S rDNA sequences of several Microbacterium species (Fig. 3.1) showed that strain B8.A has the highest sequence similarity with M. aurum strain DSMZ 8600 (GenBank accession no. GU441767.1) (99%), Microbacterium sp. KV‐492 (GenBank accession no. AB234028) (99%), Microbacterium schleiferi (GenBank accession no. Y17237) (98%) and Microbacterium oxydans (GenBank accession no. AB365061) (97%). As the newly isolated microorganism was most closely related to M. aurum DSMZ 8600, the isolated strain was named M. aurum B8.A. For comparison, the M. aurum DSMZ 8600 type strain was included in all experiments reported in this study. Chapter 3
Fig 3.1 Bootstrapped phylogenetic tree of several Microbacterium species based on 16S rDNA sequences. The sequence alignment and construction of the phylogenetic tree were performed with MEGA version 3.1 using default settings (30). The bar indicates 0.2% difference by nucleotide substitution or deletion. Bootstrap values are shown at the nodes. Page | 105
Granular starch‐degrading activity
Granular starch‐degrading activity was analyzed by growing M. aurum strains B8.A and DSMZ 8600 on MMTV agar containing granular starch. M. aurum B8.A colonies showed clear zones (halos) after iodine staining (data not shown), indicating that degradation of starch granules occurred. No halos appeared around colonies of M. aurum DSMZ 8600.
Enzymatic Degradation by M. aurum B8.A
Page | 106
Fig 3.2 Scanning electron microscopy pictures of starch granules incubated with M. aurum cell free supernatants (10 g/ml protein). Starch granules were incubated with cell free supernatants of M. aurum B8.A for 72 h (C, potato) and 24 h (D, tapioca) or cell free supernatants of M. aurum DSMZ 8600 for 7 days (E, potato; F, tapioca) compared to untreated control samples (A and B, potato and tapioca, respectively). Potato starch granules incubated for 72 h with PYE cell free supernatants of M. aurum B8.A (G) or M. aurum DSMZ 8600 (H). Chapter 3
In subsequent experiments, cell‐free supernatants were used as a source of extracellular enzymes. Cell‐free supernatants of both M. aurum strains, grown on MMTV medium, were incubated with several types of starch granules. Scanning Electron Microscopy (SEM) (Fig. 3.2) was used to visualize the enzymatic degradation. Cell‐free supernatant of M. aurum B8.A partially degraded starch granules from wheat (data not shown), tapioca and potato origin. It produced numerous small pores in potato starch granules (Fig. 3.2C) and fewer but much larger pores in tapioca starch granules (Fig. 3.2D). Supernatant of the type strain DSMZ8600 did not show visible degradation of any of the granular starches.
Page | 107
Fig 3.3 Enzymatic degradation of starch granules. Degradation of starch granules (5% w/v) by cell free supernatants obtained from cultures of M. aurum B8.A and DSMZ 8600 grown on MMTV with 1% potato starch granules. Cell free supernatants of M. aurum B8.A incubated with starch granules: ■ , Tapioca; ▲ , Wheat; ⧫, Potato; , cell free supernatants from M. aurum DSMZ 8600 incubated with potato starch granules.
The ability of extracellular enzymes produced by M. aurum B8.A to degrade starch granules was quantified by measuring the total carbohydrate released from granular starch. After 7 days incubation, the cell‐free supernatant of M. aurum B8.A had degraded wheat, tapioca, and potato starch granules by 77%, 76%, and 50% respectively, while cell‐free supernatant of strain DSMZ 8600 had only released 3% of the total amount of carbohydrate from potato starch granules (Fig. 3.3). Enzymatic Degradation by M. aurum B8.A
G1
G2
G3
G4
G5 G6 G7
M 1 2 3 4 5 (‐)
Page | 108 Fig 3.4 TLC analysis of hydrolysis products from starch granules produced by cell free supernatants of M. aurum B8.A. Cell free supernatants of M. aurum B8.A (10 g/ml protein) grown on MMTV media incubated with 5% (w/v) starch granules at 37 oC for various periods. M, Standard (G1, Glucose; G2, Maltose; G3, Maltotriose; G4, Maltotetraose; G5, Maltopentaose; G6, Maltohexaose; G7, Maltoheptaose); cell free supernatants incubated with starch granules on 37 oC for: 1, 1 h; 2, 2 h; 3, 24 h; 4, 48 h; 5, 96 h; (‐), negative control (starch granules in buffer solution).
Product formation from starch
Cell‐free supernatants of both strains B8.A and DSMZ 8600 were incubated with different types of starch followed by TLC analysis of the products formed. Enzymes in the cultures of M. aurum B8.A grown on MMTV (Fig. 3.4) or PYE (Fig. 5) media initially produced maltotriose followed by production of maltose after 2 h incubation with starch granules (Fig. 3.4) and soluble starch (Fig. 3.5). When grown on TSYE, a rich medium lacking starch, less activity was observed in strain B8.A supernatants and mostly larger maltooligosaccharides (but not maltose) were formed (Fig. 3.5). Cell‐free supernatants of M. aurum DSMZ 8600 did not show activity towards starch granules and produced only very small amounts of maltotriose from all soluble starch substrates (Fig. 3.5). Chapter 3
G1 G2 G3 G4 G5 G6 G7
A B C A B C A B C A B C A B C M M 1 2 3 4 5 Fig 3.5 TLC analysis of hydrolysis products from several substrates (1% w/v) produced by cell free supernatants (10 g/ml protein) of M. aurum strains. M, Standard (G1, Glucose; G2, Maltose; G3, Maltotriose; G4, Maltotetraose; G5, Maltopentaose; G6, Maltohexaose; G7, Maltoheptaose); A, Amylose; B, Amylopectin; C, Soluble starch; 1, negative control, 2, cell free supernatants of M. aurum B8.A grown on PYE; 3, cell free supernatants of M. aurum B8.A grown on TSYE; 4, cell free supernatants of M. Page | 109 aurum DSMZ 8600 grown on PYE; 5, cell free supernatants of M. aurum DSMZ 8600 grown on TSYE.
Purification of Granular Starch‐Degrading Enzymes
The presence of amylolytic enzymes in cell‐free supernatants of M. aurum B8.A and DSMZ 8600 was analyzed by SDS‐PAGE containing amylopectin, involving separation and renaturation of extracellular enzymes, and allowing visualization of enzyme activity. Proteins with amylase activity appeared as a clear area in a blue background after staining gels with iodine (Fig. 3.6). Cell‐free supernatants of M. aurum B8.A grown on MMTV showed seven major activity bands (Fig. 3.6, lane 1). When grown on PYE, M. aurum B8.A produced a clearly different set of proteins with amylase activity. M. aurum DSMZ 8600 did not produce any protein with visible amylase activity (data not shown). Enzymatic Degradation by M. aurum B8.A
Starch‐binding enzymes from M. aurum B8.A cell‐free supernatant were partially purified by binding to starch and subsequent elution at elevated temperature. After this procedure, fewer bands with amylase activity were observed on SDS‐PAGE. The main activity band was observed from a protein with an apparent mass of approx. 95 kDa (Fig. 3.6, lane 2).
M A B kDa
170 130 100 70 55
40 Page | 110 35 25 Fig 3.6 Activity gel from M. aurum B8.A on SDS‐polyacrylamide gel electrophoresis containing amylopectin. Symbols: M = Marker; A = MMTV cell free supernatants of M. aurum B8.A; B = Supernatants after Starch Binding Purification. The gel was incubated overnight at 37oC in 50 mM Tris, pH 6.8 and stained with an iodine solution. Chapter 3
Discussion
In this paper, we present the characteristics of the first M. aurum strain with the ability to degrade granular potato starch. The B8.A strain was allocated to the genus Microbacterium in accordance with its morphological and physiological properties and its phylogeny according to the 16S rDNA. It was designated M. aurum B8.A.
To characterize and investigate its starch‐degrading properties, cell‐free supernatant of M. aurum B8.A was incubated with different types of granular starch. The partly hydrolyzed wheat and tapioca starch granules showed many holes of different sizes, depths and width on the surface of wheat and tapioca starch. On potato starch, numerous smaller‐sized pores were found. No visible degradation of Page | 111 starch granules occurred with the cell‐free supernatants derived from M. aurum DSMZ 8600 even though the growth rate and extracellular protein concentration of both strains was similar. Apparently, a large difference in amylolytic potential between these closely related strains exists.
Amylase activity resulting in pore‐formation in maize starch granules has been reported for Bacillus subtilis and barley α‐glucosidase (19, 31). Other studies have reported enzyme systems which degraded the granules by peeling off layers from the surface, without apparent erosion of their internal regions (no pores detected) (20).
As shown in Fig. 3.3, potato starch granules were degraded slowly compared to wheat and tapioca starch granules. This quantitative assessment confirmed the observation that larger holes were formed in wheat and tapioca starch granules compared to potato starch granules. It is quite apparent that starch granules from different botanical sources have widely different susceptibilities to enzymatic hydrolysis. Potato is generally less susceptible to enzymatic degradation than other starches. It has been proposed that larger granules are less easily degraded than smaller granules (24–26), partly explaining why potato starch granules could be less Enzymatic Degradation by M. aurum B8.A
susceptible (2). In addition, potato starch and other tuber starches have a typical B‐crystalline structure, and contain more water, thicker and larger growth rings (crystalline‐amorphous portion) and longer average amylopectin branch chain length than other types (32). It has been reported that this crystalline form is more resistant to enzymatic hydrolysis than the A‐type (33).
The extracellular amylolytic enzymes in M. aurum B8.A mainly produced maltose as their smallest product, while no glucose was formed in detectable amounts. Apparently, no or little glucosidase or glucoamylase type of activity is present, although these enzymes normally play an important role in starch degradation by microorganisms (34). Previous examinations of granular starch‐degrading enzymes have shown that such enzymes often contain starch‐binding domains, allowing Page | 112 efficient binding to starch granules. This property was used to purify and select for amylolytic enzymes with the capacity to bind to granules. The partially purified enzyme preparations displayed fewer activity bands than the original cell‐free supernatant (Fig. 3.6): mostly the larger enzymes were retained, indicating that indeed enzymes containing extra domains like SBDs to bind granules had been purified. One main activity band was purified, but it is not clear whether more than one enzyme in the culture supernatant is involved in pore formation. When searching the CAZy database (http://www.cazy.org) for enzymes with similar specificities, like binding to raw starch and production of maltotriose, two enzymes were found, both from Gram+ organisms with a high GC content like M. aurum (15, 35, 36). Both these enzymes contain an SBD of family CBM20, suggesting that the M. aurum raw starch‐degrading amylase may have a similar structure. On the other hand, the lack of similar enzymes with other types of CBMs in the databases might also be representative of the fact that only a small percentage of predicted enzymes have been characterized biochemically, biasing such predictions.
According to the 16S rDNA sequence of both organisms characterized in this paper are M. aurum species; however a clear difference was observed in their ability to degrade starch granules. In all experiments performed in this study, either involving agar‐grown colonies (data not shown) or cell‐free supernatant of liquid grown Chapter 3
cultures on various media, M. aurum DSMZ 8600 showed little or no amylolytic activity. Granular starch‐degrading activity was only found in the M. aurum B8.A strain. The apparent incompleteness of the amylolytic system of M. aurum, with the absence of a glucoamylase or glucosidase activity, may suggest that its starch‐ degrading ability originates from a single enzyme, which may have been acquired by horizontal gene transfer (37–40). Although horizontal gene transfer could theoretically explain this striking difference between very closely related strains, it is not very likely in this case as no other strains with similar granule degrading ability were isolated from the wastewater source. The only other strain isolated for its ability to degrade granules from the same screening, identified as a Bacillus firmus/lentus strain, did this by “peeling off” layers of the surface of the granules, which is likely to require a different set of enzymes (41). This suggests that genes encoding amylolytic, pore forming, enzymes were not widely present in the Page | 113 microbial gene pool, not even in the environment where granular starch is an abundantly available substrate. An alternative explanation is that the type strain, M. aurum DSMZ 8600, has lost its starch‐degrading ability during evolution. We are currently isolating the amylase genes of the B8.A strain to allow a further study of its complex of amylolytic enzymes that plays a role in granular (potato) starch degradation.
Acknowledgments
F.D. Sarian was financially supported by a Bernoulli PhD student bursary from the University of Groningen. The authors thank the Netherlands Organization for Applied Scientific Research (TNO) for providing M. aurum B8.A. We also thank Itje Stokroos and Jeroen Kuipers (Cell Biology and Electron Microscopy Department, University Medical Centre Groningen, the Netherlands) for taking the SEM pictures. Enzymatic Degradation by M. aurum B8.A
Material and Methods
Materials
Potato, wheat and tapioca starches were a kind gift of TNO – Quality of Life, (Zeist, The Netherlands). Soluble potato starch Paselli SA2 was obtained from AVEBE (Foxhol, The Netherlands). Oligosaccharides, amylose and amylopectin were purchased from Sigma Chemie GmbH (Germany).
Strain Isolation and Characterization
Sludge from a wastewater treatment unit of AVEBE, a potato starch‐processing Page | 114 factory in The Netherlands, was resuspended in demineralized water, stirred for 3 h, and filtered over glass wool. The filtrate was inoculated into medium containing 20
mM KH2PO4, 11 mM K2HPO4, 0.8 mM MgSO4.7H2O, 6.7 mM KCl, 0.07 mM FeCl3.6H20,
23.5 mM NaNO3, pH 7.2, and 1% (w/v) native potato starch granules (sterilized by gamma radiation). Cultures were incubated at 37 oC while shaken. Cultures containing granular starch degrading microorganisms, as judged by the degradation of starch granules observed with light microscopy, were plated on medium
containing 18 mM KH2PO4, 40 mM K2HPO4, 0.8 mM MgSO4.7H2O, 6.7 mM KCl, 0.07
mM FeCl3.6H20, 23.5 mM NaNO3 and 2% (w/v) agar, pH 7.2. A top layer was added containing 0.9% (w/v) agar and 2% (w/v) native potato starch granules in suspension, added after cooling the agar to 55 oC. Colonies forming a clearing zone in the opaque top layer were selected, and pure cultures were isolated by repeatedly inoculating single colonies on nutrient agar. Morphological, physiological, and biochemical studies on the bacterium were carried out according to Bergey’s Manual of Determinative Bacteriology (42).
Genomic DNA of the isolated granular starch degrading strain was extracted using the Gen Elute Bacterial Genomic DNA Kit (Sigma‐Aldrich, US). 16S ribosomal DNA (rDNA) was amplified using the primers B3F (5’‐ GGT TAC GTT GTT ACG ACT T‐3’) Chapter 3
and 1492 R (5’‐AGA GTT TGA TCA TGG CTC AG‐3’) (Eurogentec, The Netherlands). The amplification products were purified with Gen‐Elute PCR Cleaning Kit (Sigma).
Cloning of the PCR product was done by standard methods (43). The amplified product was ligated into vector XL‐TOPO (Invitrogen, The Netherlands) and the recombinant plasmid was sequenced by using T7 and M13‐RP primers at GATC Biotech (Germany). Similarity searches with M. aurum B8.A DNA sequences were performed with nucleotide collection (nr/nt) sequences at the NCBI website (http://www.ncbi.nlm.nih.gov) using the available BLAST tools (44). A phylogenetic tree was produced with MEGA version 3.1 using default settings (45). The 16S rRNA gene sequence was deposited with GenBank and has the accession number GU441767.1. Page | 115
Bacterial Cultures and Media
M. aurum strain B8.A was maintained on a medium containing 20 mM KH2PO4, 20 mM K2HPO4, 0.75 mM MgSO4.7H2O, 7 mM KCl, 0.05 mM FeCl3.6H2O, 0.2% (w/v) trypton, 1% (w/v) potato starch granules with 1% (v/v) trace elements and 1% (v/v) vitamins (MMTV) (41). The composition of the trace element solution was: 3.5 mM MnCl2, 0.5 mM Na2B4O7.10H2O, 70 M ZnSO4.7H2O, 30 M CuCl2.2H2O, 0.3 mM
H3BO3, 12 M Na2MoO4.2H2O, 8 M VSO5.5H2O, 3.5 M CoSO4.7H2O, 25 mM H2SO4. The vitamin solution consisted of 40 M thiamine‐HCl, 20 M biotin, 10 M pyridoxine‐HCl 15 M riboflavin, 35 M p‐aminobenzoic acid, 40 M nicotinic acid, 10 M D‐Ca‐pantothenate, 1 M folic acid, 0.05 M cyanocobalamine, sterilized by ultrafiltration. The M. aurum type strain (DSMZ 8600) was obtained from the DSMZ collection (Germany). Precultures were grown in shake flasks containing Nutrient Broth (NB) medium (41) for 24 h at 30 oC on a rotary shaker at 200 rpm. Main cultures were grown in shake flasks containing MMTV medium for 48 h at 30 oC and 200 rpm. Routinely a 1% (v/v) inoculum was used. Cultivation was also performed in rich media containing either 3% (w/v) Trypticase Soy Broth and 0.3% Yeast Extract (TSYE), or 3% potato starch granules and 0.3% Yeast Extract (PYE) or only 0.3% Yeast Extract (YE). Starch granules were sterilized by gamma‐radiation. Enzymatic Degradation by M. aurum B8.A
Screening for starch degradation was performed by inoculating bacteria on MMTV agar with 1% (w/v) granular potato starch. After incubation for 48 h at 30 oC, a
solution of 1% KI/0.25% I2 (w/v) was added, visualizing starch‐degrading colonies by a clear halo.
Production and harvest of extracellular enzymes
Following 3 days of batch cultivation of Microbacterium strains, the fermentation broths were collected and centrifuged at 10,000 x g for 10 min at 4 oC. The resulting cell free supernatants containing extracellular enzymes were collected for further investigation. Protein concentrations were estimated by the Bradford method (Bio‐ Rad Protein Kit, Bio‐Rad Lab. GmbH, Germany) (46). Cell free supernatants were Page | 116 diluted to a protein concentration of 10 μg ml‐1 with 50 mM Tris Buffer pH 6.8 before use in activity assay experiments.
Purification of starch binding proteins
Granular starch‐degrading enzymes were partially purified from cell free supernatants by binding to starch granules. Thus, 5% (w/v) potato starch granules were added to supernatants prepared as described above, and the suspensions were stirred gently at 4 oC. After 2 h, the suspensions were centrifuged at 5,000 x g for 20 min, and the precipitated granular starch was washed with ice‐cold 50 mM Na‐ phosphate buffer (pH 6.8). Adsorbed enzymes were eluted from starch granules by stirring gently in 50 mM Na‐phosphate (pH 6.8) at 45 oC for 1 h followed by centrifugation at 5,000 x g for 20 min. Supernatant containing partially purified starch binding enzymes was collected for further analysis.
Sugar analysis
Cell free supernatants obtained from cultures grown on MMTV containing 10 μg ml‐1 protein, 100 μg ml‐1 of ampicillin and 100 μg ml‐1 of kanamycin were incubated with Chapter 3
1% (w/v) starch granules at 37 oC with agitation at 200 rpm. Samples were removed after various time intervals. Reaction products were separated from granules by centrifugation at 5,000 x g for 20 min. Reaction products were analyzed by Thin
Layer Chromatography (TLC) (Silica gel 60 F254, Merck, Germany) by spotting 5 μl from each time point. Products were separated with a solvent system of 1‐ butanol/ethanol/water (5:5:3, v/v/v) overnight. After running, the plates were dried and sprayed with 50% (v/v) sulphuric acid in methanol and left to develop at 110 oC (47). The reaction products from soluble substrates were also investigated by incubation of 1% (w/v) substrate for 4 h with cell free supernatants of both the M. aurum strains B8.A and DSMZ 8600, diluted to a protein concentration of 10 μg ml‐1 obtained from cultures grown on various rich media. Formation of products was determined qualitatively by TLC as described above.
Page | 117
To estimate granular starch‐degrading activity, cell free supernatants were incubated with granular starch, and the total carbohydrate released was measured with the Anthron method. Briefly, 5 ml 10% (w/v) granular starch suspension was mixed with 5 ml cell‐free supernatant obtained from M. aurum strains grown on MMTV, with a final concentration of 10 μg ml‐1 protein and containing 100 μg ml‐1 of ampicillin and 100 μg ml‐1 of kanamycin. Incubations were performed in duplicate at 37 oC with agitation at 200 rpm. Samples were taken in duplicate after several time intervals up to 7 days (148 h); the reaction was stopped by centrifugation. Subsequently, 0.5 ml supernatant was added to 5 ml Anthron solution [0.2% (w/v)
Anthron powder in 200 ml of concentrated H2SO4 and 60 ml water, 15 ml 95% ethanol] and incubated at 100 oC for 10 min. The absorbance at 620 nm was taken as a measure of total carbohydrate present. Maltose (20‐100 mM) was used as a standard.
Scanning electron microscopy
The incubation of supernatant samples with starch granules was performed as described above (Anthron test). After incubation, starch granules were washed twice with water‐ethanol (1:1) and dried by heating in an oven at 55 oC for 1 d. The Enzymatic Degradation by M. aurum B8.A
samples (dried starch powder) were mounted on SEM stubs with double‐sided adhesive tape and coated with gold. Scanning electron micrographs were taken using the JSM‐6301F Scanning Microscope (JEOL) in the Department of Cell Biology and Electron Microscopy, Faculty of Medical Sciences, University Medical Centre Groningen, The Netherlands. The accelerating voltage (2.0 kV) and the magnification are shown on the micrographs.
Protein Electrophoresis
Analysis of starch‐hydrolyzing activity on sodium dodecyl sulphate polyacrylamide gels (SDS PAGE) was conducted according to Zeeman et al. (1998) (48). Protein samples (final concentration 30 ng ml‐1) were separated on 7.5% (w/v) SDS PAGE Page | 118 containing 0.5% (w/v) amylopectin at a constant voltage of 150 V per gel for 55 min. After electrophoresis, gels were washed twice with distilled water and once with 50 mM Tris buffer (pH 6.8) and subsequently incubated overnight in the same buffer at 37 oC. Protein bands containing starch‐hydrolyzing activity were visualized by
staining with a solution of 1% KI/0.25% I2 (w/v). Chapter 3
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Enzymatic Degradation by M. aurum B8.A
Page | 124 Chapter
Degradation of granular starch by the bacterium
Microbacterium aurum B8.A involves a novel modular
α ‐amylase enzyme system with FnIII and CBM25 domains
Vincent Valk, Wieger Eeuwema, Fean D. Sarian, Rachel M. van der Kaaij, and Lubbert
Dijkhuizen
Applied and Enviromental Microbiology (2015) : doi:10.1128/AEM.01029‐15 Degradation of granular starch involves FnIII and CBM25 domains
Abstract
The bacterium Microbacterium aurum strain B8.A, originally isolated from a potato plant waste water facility, is able to degrade different types of starch granules. Here we report the characterization of an unusually large, multi‐domain M. aurum B8.A α‐amylase enzyme (MaAmyA). MaAmyA is a 1417 amino acid protein with a predicted molecular mass of 148 kDa. Sequence analysis of MaAmyA shows that its catalytic core is a family GH13_32 α‐amylase with the typical ABC domain structure, followed by a Fibronectin (FnIII) domain, two Carbohydrate Binding Modules (CBM25) and another three FnIII domains. Recombinant expression and purification yielded an enzyme with the ability to degrade wheat and potato starch granules by introducing pores. Characterization of various truncated mutants of MaAmyA revealed a direct relation between the presence of CBM25 domains and the ability of MaAmyA to form pores in starch granules, while the FnIII domains most likely function as stable linkers. At the C‐terminus, MaAmyA carries a novel 300 aa domain Page | 126 which is uniquely associated with large multi‐domain amylases; its function remains to be elucidated.
We conclude that M. aurum B8.A employs a novel multi‐domain enzyme system to initiate degradation of starch granules via pore formation. Chapter 4
Introduction
Starch is an excellent carbon‐ and energy source for many micro‐organisms, which employ a dedicated set of proteins for extracellular hydrolysis of this polysaccharide, uptake of shorter oligosaccharides into the cell, and further degradation into glucose. Most studies on degradation of starch by microbial enzymes have focused on soluble starch. This has resulted in identification and characterization of a large variety of enzymes, cleaving either α(14) or α(16) linkages in amylose and amylopectin. Most of these enzymes belong to glycoside hydrolase family 13 (GH13) (1). Sequence diversity is such that at the moment family GH13 counts a total of 40 subfamilies (1). Most of the new members in subfamilies are identified in DNA sequencing projects and biochemical information about the activity and specificity of these potentially new enzymes is strongly lagging behind.
Page | 127 Many plants produce starch in a granular form for the storage of carbohydrates. The crystallinity of such granules varies with the plant source. Potato starch granules have a relatively high degree of crystallinity making them notoriously resistant to bacterial and fungal degradation (2–4). Nevertheless, some micro‐organisms have been reported to employ enzymes able to digest granular starch (5, 6).
Amylases found to be involved in granular starch degradation are often multi‐ domain enzymes that include one or more Carbohydrate Binding Modules (CBMs) which aid in the binding of the enzyme to the granular substrate (7–10).
In previous work we have isolated various bacteria able to grow on potato starch granules as carbon source and evaluated their enzymatic degradation mechanism. Initially this resulted in identification of an enzyme mechanism involving peeling off layer after layer of the starch granules in Paenibacillus granivorans (11).
Degradation of granular starch involves FnIII and CBM25 domains
In this paper we focus on Microbacterium aurum strain B8.A that originally has been isolated from a potato plant waste water facility. It is able to degrade different types of starch granules as carbon and energy sources. Recently we reported that it attacks and degrades granular potato starch by an alternative mechanism initially involving pore formation in wheat, tapioca and also potato starch granules (12).
This paper reports the characterization of a novel and unusually large M. aurum B8.A α‐amylase enzyme (MaAmyA) that is able to form pores in starch granules in vitro. It belongs to the glycoside hydrolase (GH) family 13, subfamily 32. MaAmyA is about 2 times larger than other GH13_32 members with a single catalytic domain. It has two CBM domains and it is the only known GH13_32 member with FnIII domains. Multiple deletion constructs of MaAmyA were expressed and characterized to study the roles of the different domains in degradation of both soluble and granular starch. A direct relation between the presence of CBM25 domains in Page | 128 MaAmyA and its ability to form pores in starch granules was observed. Chapter 4
Results
Gene identification
In previous work (12) we reported that culture fluid of M. aurum B8.A degraded granular starches by introducing pores initially. To identify the starch acting enzyme(s) involved, a M. aurum B8.A DNA library of 70,000 clones was screened for α‐amylase activity. The clone with the highest activity carried a vector with an insert of 11.6 kb. DNA sequencing revealed a complete open reading frame of 4227 nt encoding a large, multi‐domain (putative) α‐amylase enzyme of 1409 amino acids (with a predicted mass of 148 kDa), including a signal sequence of 34 aa. The sequence of this unusually large α‐amylase, designated MaAmyA, was confirmed when the full genome of M. aurum B8.A was sequenced recently (Valk et al., manuscript in preparation).
Page | 129 The catalytic domain of MaAmyA shows similarity to glycoside hydrolase family 13, subfamily 32 (GH13_32), which currently includes 109 members and is mainly found in bacteria (1). The catalytic domain contains the A, B and C regions typical for the α‐amylase superfamily (24) as well as all catalytic residues and regions generally conserved in α‐amylases (25). The characterization of MaAmyA revealed that it was enzymatically active with soluble starch but also degraded starch granules by introducing pores (see below).
Domain organization
A CDD (13) search revealed a total of 7 domains in MaAmyA: the N‐terminal GH13 catalytic domain is followed by 1 FnIII domain, 2 CBM25 domains, and 3 highly similar FnIII domains (>95% identity) (Fig. 4.1). The C‐terminal tail of MaAmyA of 331 aa does not match with any known domain. A BLAST search nevertheless returned 26 hits for similar 300 aa long fragments (34‐60% identity, 48‐72% similarity). Of the 26 hits, 21 are part of predicted multi‐domain α‐amylases. An additional four hits are genomically linked to a nearby ORF encoding an α‐amylase, further indicating a relation between this domain and α‐amylases. None of these Degradation of granular starch involves FnIII and CBM25 domains
putative amylases belong to subfamily GH13_32; instead 21 belong to GH13_28. The 300 aa fragment is usually located C‐terminally of the catalytic domain, either at the C‐terminus or in between other domains. None of these multi‐domain α‐amylases have been characterized biochemically.
Fig 4.1 Schematic representation of the domain organization of MaAmyA and the different truncated derivatives, MaAmyA2, MaAmyA4, MaAmyA7, and the two related GH13_32 amylases of Bacillus sp. No. 195 and Kocuria varians ATCC 21971. Domains Page | 130 are indicated as follows: White box: signal sequence; Dark box: GH13 catalytic domain; Dotted box: FnIII domain; Vertically lined box: CBM25 domain; Dashed box with horizontal lines: unknown domain. Predicted molar masses are indicated.
Phylogenetic tree of subfamily GH13_32
In a MaAmyA full length BLAST search (26), the GH13_32 α‐amylases of Bacillus sp. no. 195 (GenBank BAA22082) and Kocuria varians ATCC 21971 (GenBank BAJ52728) came up as highest similarity hits. The latter enzymes consist of a catalytic domain and two CBM25 domains (Fig. 4.1). With a predicted mass of 77 kDa they are much smaller than the 148 kDa MaAmyA. Both enzymes have been described as granular starch degrading GH13_32 amylases but their mechanism of action on starch granules has not been investigated (7, 8, 27–30). A phylogenetic tree based on catalytic AB regions of MaAmyA and all other GH13_32 members (1) shows the diversity within this subfamily (Fig. 4.2). MaAmyA closely clusters with 5 other members, in a subgroup of in total 52 GH13_32 members. Surprisingly, MaAmyA is the only protein in this subfamily with FnIII domains and the C‐terminal tail of 300 aa. Interestingly one blade of the tree contains GH13_32 enzymes with a Chapter 4
second GH13_14 catalytic domain (pullulanase), and several binding domains. Only one of these large enzymes has been described in literature (31).
Members of GH13_32 are richly decorated with diverse starch binding domains (Fig. 4.2). Approximately 66% of all GH13_32 subfamily members contain one or two CBM20 domains. CBM25 domains are relatively rare among the single catalytic domain enzymes (present in 7 members of GH13_32 including MaAmyA). However, none of the other currently known GH13_32 enzymes in CAZy or in the NCBI database containing non‐redundant protein sequences possesses FnIII domains or the 300 aa C‐terminal tail (data not shown). This makes MaAmyA an exceptional member of GH13_32.
To obtain more insight into the evolutionary origin of the CBM25 domains in Page | 131 MaAmyA, phylogenetic analysis was performed (Fig. 4.3). The phylogenetic tree shows clustering of the CBM25 domains present as tandems in MaAmyA, the two other closely related GH13_32 enzymes of Bacillus and K. varians, and a third enzyme from Jonesia denitrificans DSM 20603 (GenBank ACV09568.1) with similar global domain organization. The latter enzyme has not been allocated to a specific GH13 subfamily in CAZy but it clearly clusters with other GH13_32 amylases (Fig. 4.2).
Degradation of granular starch involves FnIII and CBM25 domains
Z Y
Page | 132
Fig 4.2 Phylogenetic tree of all 109 GH13_32 members in CAZy, Jonesia denitrificans amylase (GenBank ACV09568.1) and MaAmyA. The tree is based on the alignment of α‐amylase catalytic domains (regions AB) obtained from DbCAN (aa 74‐362 for MaAmyA). Domain organization was based on combined CDD and DbCAN data. For comparison, mixed selections of the related subfamilies GH13_5, GH13_6, GH13_7, GH13_27, and GH13_28 are also included. At Y and Z, 32 and 5 sequences which were similar to others in that branch were collapsed to improve the readability of the tree. The GH13 subfamilies are indicated through the background color of the protein names. Domain colors and shapes are explained in the legend. Chapter 4
Page | 133
Fig 4.3 Phylogenetic tree of all 153 CBM25 domains in CAZy with sequences in GenBank and MaAmyA. The tree is based on the CBM25 domains obtained from CDD (aa 604‐681 and 707‐783 for MaAmyA); therefore, proteins with multiple CBM25 domains also have multiple entries in the tree. Domain organization was based on CDD data. At Y, a branch containing 34 sequences of single CBM25 domains was collapsed to improve the readability of the tree. The GH (sub)families are indicated through the background color of the protein names, proteins without catalytic domain are not colored. Domain colors and shapes are explained in the legend.
Degradation of granular starch involves FnIII and CBM25 domains
Expression and His-Tag purification of MaAmyA
MaAmyA was successfully expressed in E. coli as full length protein and in truncated forms MaAmyA2, MaAmyA4 and MaAmyA7 (Fig. 4.1), with His‐Tags at both the N‐ and C‐termini of all constructs. Studies with proteins carrying a single His‐Tag showed that only the C‐terminal His‐Tags were functional in metal‐affinity chromatography purification; the presence of the N‐terminal His‐Tag resulted in higher expression levels (data not shown). Removal of the predicted N‐terminal signal sequence did not improve protein yields (data not shown). Expression levels were low for all forms. Different expression strains, vectors, and conditions, including pBAD vector based constructs, different E. coli strains and a Rhodococcus expression system, were tested to improve expression without noticeable improvement (data not shown). The amount of protein obtained after purification of MaAmyA and the truncated derivatives was relatively low; no protein bands were visible after SDS‐PAGE analysis with silver staining. In gel activity‐staining (after Page | 134 SDS‐PAGE and washing steps) of the MaAmyA protein and truncated versions, using soluble potato starch, confirmed the presence of all MaAmyA protein derivatives with the expected masses (Fig. 4.4). The full length MaAmyA protein of 148 kDa, and the MaAmyA7 truncation of 114 kDa, lacking the C‐terminal tail, both showed a minor additional activity band between 200 and 300 kDa, which may be the result of protein dimerization. MaAmyA also showed a band at the same height as the MaAmyA7 truncation which may indicate early expression termination in E. coli and (partial) loss of the C‐terminal tail. A similar effect is seen with the MaAmyA4 truncation of 83 kDa, which shows a lower band of approximately 73 kDa, corresponding to the expected mass of MaAmyA4 without the second CBM25 domain. Storage of the enzymes for 5 months at 4 °C did not affect these results, indicating that the observed fragments were not formed due to enzyme instability (data not show). M. aurum culture fluid showed a single activity band at the same height as MaAmyA. No background E. coli amylase activity was detected (empty vector construct).
Chapter 4
Fig 4.4 Activity staining on SDS‐PAGE with soluble starch of heterologously expressed, His‐Tag purified M. aurum B8.A proteins. MW = marker, A = empty vector; B = MaAmyA2 (62 kDa); C = MaAmyA4 (83 kDa); D = MaAmyA7 (114 kDa); E = MaAmyA (148 kDa); F = M. aurum B8.A culture fluid. Marker masses are indicated. The main activity bands correspond with the expected masses of the expressed proteins.
Enzyme activity Page | 135
The activities of the purified MaAmyA and truncated enzyme derivatives were determined on both soluble potato starch and granular wheat and potato starch (Table 4.1). The MaAmyA activity was set at 100 % for each substrate. The activity on granular potato starch was too low to be determined. MaAmyA4, MaAmyA7, and MaAmyA have similar activity ratios for both soluble and granular starch (Table 4.1). The truncated enzyme MaAmyA2 clearly holds activity with soluble starch (79%) but its activity on granular starch is strongly reduced compared to the full enzyme. Especially the two CBM25 domains thus play a crucial role in the activity of MaAmyA on granular starch (see Figure 4.1).
Degradation of granular starch involves FnIII and CBM25 domains
Table 4.1 Relative activities of MaAmyA and truncated derivatives on 0.4% soluble potato starch and 3.3% granular wheat starch. Incubations with soluble starch were performed in triplicate, with granular starch in duplicate. An empty vector construct was used as negative control.
Relative activity Relative activity on on soluble starch wheat starch granules (%) (%)
MaAmyA2 79 ± 3 9 ± 1
MaAmyA4 99 ± 2 94 ± 2
MaAmyA7 97 ± 4 83 ± 10
MaAmyA 100 ± 3 100 ± 10
M. aurum B8.A culture fluid 123 ± 5 122 ± 16 Page | 136 Negative control <2 <5
Granular starch degradation
To study the roles of the different MaAmyA domains on pore formation in granular starch, wheat starch granules were incubated with MaAmyA and truncated MaAmyA2, MaAmyA4 and MaAmyA7 enzymes.
During the first 6 h of incubation, MaAmyA, MaAmyA7, MaAmyA4 as well as the M. aurum B8.A culture fluid showed similar rates of degradation of granular starch, whereas MaAmyA2 and the negative control were much slower (Fig. 4.5). To exclude a possible effect of the FNIII domain present in MaAmyA2, the experiment was also performed with this domain removed which gave similar results (data not shown). After longer incubation times, MaAmyA, MaAmyA7, and MaAmyA4 continued to show similar curves but their degradation rates had decreased over time and Chapter 4
reached a threshold at about 20%. The M. aurum culture fluid was most active in degrading granular starches, reaching about 80% degradation of wheat starch after 72 h.
When fresh MaAmyA4 enzyme sample (300 U, CNPG3 assay) was added at t = 24 h and t=48 h to MaAmyA4 incubations with wheat starch granules, the degradation rate increased again, reaching 40% degradation after 72 h (data not shown). This indicated that the rate of degradation decreased at least partly due to loss of enzyme activity within the first 48 h. The granular substrate itself thus was further degradable by MaAmyA4.
Page | 137
Fig 4.5 Degradation of granular wheat starch in time by M. aurum B8.A culture fluid, MaAmyA, and truncated MaAmyA enzymes. In each reaction, 300 U of enzyme activity (CNPG3 assay) was added. His‐tag purified empty vector E. coli extract was added as a negative control. The total carbohydrate concentration in the supernatant was determined (Anthrone) and the percentage of granular starch degraded was calculated.
Degradation of granular starch involves FnIII and CBM25 domains
Page | 138
Fig 4.6 SEM images of wheat starch granules following incubation with the different M. aurum B8.A enzyme samples (300 U, CNPG3 assay) at 5,000x magnification. All images show granules that where incubated for 48 h unless indicated otherwise. A was incubated for 6 h with M. aurum culture fluid; B with MaAmyA; C with negative control; D with MaAmyA2; E with MaAmyA4; F with MaAmyA7. A 1 µm scale bar is shown on the images.
A selection of samples with similar degradation percentage was used for SEM imaging analysis (Fig. 4.5, 4.6). After 48 h of incubation the wheat starch granules treated with M. aurum B8.A culture fluid had lost their granular structure and only debris remained (data not shown). Chapter 4
Fig 4.7 Pore sizes in wheat starch granules after incubation with different MaAmyA enzymes, or M. aurum culture fluid (CF). The images used to determine the pore sizes were lower magnification images (750x) taken in the same region as the images shown in Fig. 6. All samples had been incubated for 48 h except the CF samples which were incubated for 6 h. CF treated samples showed also a set of large pores (CF LP) which were included as a separate group.
Page | 139
The SEM images were used for analysis of the pore sizes using imaging software (Fig. 4.7). Incubations with MaAmyA clearly resulted in homogeneous pore formation (Fig. 4.6B). Wheat starch granules degraded with M. aurum B8.A culture fluid (Fig. 4.6A) additionally showed a set of large pores (LP) that were approx. 20 times larger and absent in the MaAmyA treated granules (Fig. 4.7, see LP). The truncated proteins MaAmyA7 and MaAmyA4 also introduced homogeneous pore sizes, although these pores were approximately 3 times smaller than those formed by MaAmyA (Fig. 4.7). This indicated a possible role for the C‐terminal protein tail in formation of larger pores. MaAmyA2 (Fig. 4.6D) does not show pore formation and these granules look similar to the negative control samples (Fig. 4.6C). Apparently, the presence of the CBM25 domains in MaAmyA4 is sufficient for pore formation in granular starches. The incubations and subsequent image analysis were also performed with granular potato starch, which gave similar results: pore formation in granules was observed for the truncated MaAmyA constructs that included the CBM25 domains but was absent in granules incubated with MaAmyA2 and negative control (data not shown). Degradation of granular starch involves FnIII and CBM25 domains
Discussion
This paper reports the characterization of MaAmyA, a multi‐domain α‐amylase enzyme of Microbacterium aurum B8.A, carrying four FnIII and two CBM25 domains, plus a novel C‐terminal domain of 300 aa. The MaAmyA enzyme degrades granular starches by initially introducing pores. Deletion of the additional domains did not affect activity on soluble starch. The data clearly show that the CBM25 domains are essential for activity with granular starch and for pore formation. CBM25 domains previously have been shown to be involved in granular starch degradation (7, 8), but a role in pore formation specifically has not been reported before.
The MaAmyA enzyme thus allows M. aurum B8.A to degrade granular starches directly, without the need for starch gelatinization. This may be a more general situation in the natural environment. Extracellular bacterial α‐amylases degrading Page | 140 starch granules by peeling off layer after layer (11), or by introducing pores in starch granules (12, 32, 33) (this paper), may occur more widespread than currently described in literature.
On basis of its catalytic domain MaAmyA belongs to the CAZy GH13_32 subfamily, but its length and the presence of FnIII domains and a C‐terminal tail make it an exceptional member of this subfamily (Fig. 4.2). To study the roles of the different domains, MaAmyA as well as four C‐terminal truncated proteins were successfully expressed in E. coli, purified and characterized. Low levels of expression were achieved; therefore the activity with the CNPG3 substrate was used to standardize the amount of protein used in further experiments. We assumed that the presence or absence of additional domains had no effect on the activity with CNPG3, in view of the small size of this substrate, and that a direct relation exists between CNPG3 activity and enzyme concentration. However, this assumption can only be made for MaAmyA and its truncated versions, but may not hold for M. aurum culture fluid which may contain additional enzymes. The M. aurum culture fluid shows a very high ratio of granular starch activity over CNPG3 activity (Fig. 4.5), compared to Chapter 4
MaAmyA. It is therefore likely that the culture fluid contains one or more additional enzymes with low activity on CNPG3 but high activity on starch granules.
FnIII domains
MaAmyA has four FnIII domains, which is an unusually high number compared to other GH13 enzymes. α‐Amylases generally lack FnIII domains (none found in the other 109 GH13_32 members) and when present there are rarely more than two. The three adjacent FnIII domains in MaAmyA are almost identical and may be the result of recent duplications, suggesting that they provide a strong evolutionary advantage. In eukaryotes FnIII domains are widely spread and well known for their protein‐protein interactions (34). In prokaryotes FnIII domains were initially only found in carbohydrate acting enzymes (35) but more recently they have been identified in a wide variety of bacterial proteins (36, 37). Little is known about the function(s) of FnIII domains in amylases. Published reports about FnIII domains in Page | 141 carbohydrate enzymes do not show a clear common function, with results pointing at possible roles in substrate binding or in enzymatic activity (38–43). So far there are no reports of FnIII domains in prokaryotes with a function in protein‐protein interactions.
In the present study, no direct effect of the FnIII domains on substrate binding or enzyme activity was observed (Figs. 4.5‐4.8, Table 4.1). Previously, FnIII domains were suggested to function as stable linkers between separate domains in bacterial cellobiohydrolase (42) and chitinase enzymes (41). Also the FnIII domains in MaAmyA may function as stable linkers, between the catalytic domain and the CBM25 domains, and the C‐terminal tail.
CBM25 domains
Enzymes acting on insoluble carbohydrates like granular starches often possess carbohydrate binding modules (CBM) (5). When functioning in starch binding, they Degradation of granular starch involves FnIII and CBM25 domains
are known as starch binding domains (SBD). Currently SBDs have been identified in CBM families 20, 21, 25, 26, 34, 41, 45, 48, 53, 58, and 69 (1). The functions of CBM25 domains have been described previously and their 3D‐structures have been resolved (44–46). It also has been demonstrated that a CBM25‐26 tandem in the maltohexaose forming amylase from Bacillus halodurans C‐125 has a 50 fold stronger binding affinity to granular corn starch than each of these single domains (44). A similar effect, though much lower, has been observed for a CBM25 tandem found in the two homologs closely related to MaAmyA: in the halophilic K. varians α‐ amylase the binding affinity of the CBM25 tandem was only 20% higher than that of a single CBM25 (8). Despite this low increase in binding affinity, the homologous enzyme in Bacillus sp. no. 195 showed a 2 to 4 fold increase in degradation rate of different granular starches, compared to a single CBM25 domain attached to the C‐ terminus (8). This work shows that CBM25 domains greatly enhance the ability of MaAmyA to degrade granular starch through the formation of pores in the granules (Figs. 4.6‐4.8), while they have little effect on activity on soluble substrates (Table Page | 142 4.1). Phylogenetic analysis revealed that the CBM25 domains of MaAmyA are most closely related to each other, which suggests that a duplication of this CBM25 domain occurred. The same is true for the other GH13_32 amylases with two CBM25 domains (Fig. 4.2, 4.3). The CBM25 domains tree (Fig. 4.3) suggests that an ancestral GH13_32 amylase acquired a single CBM25 domain, which duplicated only recently in the different enzymes independently, resulting in a tandem of CBM25 domains. These duplication events may have improved binding capability and the granular starch degradation rates of the enzymes (8, 28).
Another remarkable feature of MaAmyA is the presence of a single FnIII domain between the catalytic core and the 2 CBM25 domains, even though both the CBM25 (Fig. 4.3) and catalytic domain (Fig. 4.2) cluster with two related GH13_32 amylases lacking FnIII domains. In view of the recent duplications of the CBM25 and FnIII domains we conclude that the M. aurum MaAmyA is able to easily acquire and include other domains and duplicate them.
Chapter 4
C-terminal tail
The presence of three FnIII domains between the preceding CBM25 domains and the C‐terminal tail, potentially functioning as stable linkers, suggests that the C‐terminal tail of MaAmyA has a specific functional role. Homologs were found in 21 other large multi‐domain α‐amylases. The 300 aa C‐terminal tail thus appears to represent a novel domain that is often part of a large multi‐domain α‐amylase. The pores formed by full length MaAmyA on granular wheat starch are approximately 3 times larger than pores formed by AmyA4 and AmaA7 lacking the tail (Fig. 4.6, 4.8). At present, the mechanism by which the C‐terminal protein tail has an effect on granule pore size is unknown; deletion had no apparent effect on enzyme activity (Table 4.1, Fig. 4.5). Further research is needed to elucidate the precise role of this novel domain.
Comparison of MaAmyA to M. aurum B8.A culture fluid
Page | 143 The M. aurum B8.A culture fluid degraded wheat starch up to 60% further than incubation with only MaAmyA. Part of this difference can be explained by instability of MaAmyA. Still, the sharp increase in degradation rate (between 6‐12 h, Fig. 4.6A) could not be achieved with MaAmyA alone. Furthermore, the granules degraded by culture fluid show a different pore pattern than those degraded by MaAmyA indicating the presence of additional enzymes in the culture fluid. The recently obtained genome sequence of M. aurum B8.A (Valk et al., manuscript in preparation) revealed 14 other family GH13 enzymes, one of them in close proximity to MaAmyA. In previous work with M. aurum B8.A culture fluids, multiple activity bands were visible in gels with soluble starch (12). The current study shows that most of the sizes of these activity bands correspond to the sizes of the four truncated enzymes. Therefore it is likely that the SDS‐PAGE gel analysis procedure resulted in the loss of one or more domains from MaAmyA. When pre‐cast THX‐gels were used, these artifacts were strongly reduced (Fig. 4.4). However, the culture fluid sample showed a second weak activity band of approx. 130 kDa in the smear below the main band. This band does not correspond to any of the truncated MaAmyA masses. Also in view of the difference between CNPG3 activity and granular starch activity of the Degradation of granular starch involves FnIII and CBM25 domains
culture fluid, it appears likely that M. aurum B8.A harbors one or more additional enzymes involved in granular starch degradation.
Acknowledgements
This study was (partly) funded by the Top Institute Food & Nutrition (project B1003) and the University of Groningen.
The SEM studies have been performed at the UMCG Microscopy and Imaging Center (UMIC), Groningen, The Netherlands , which is sponsored by NWO grants 40‐00506‐98‐9021 and 175‐010‐2009‐023.
Page | 144 Chapter 4
Materials and Methods
Bacterial strains, media, and plasmids
Microbacterium aurum strain B8.A was isolated from a waste water treatment plant of a potato starch processing factory. Isolation and growth conditions have been described previously (12). Escherichia. coli TOP10 and BL21(DE3) were cultivated at 37 oC overnight in LB with orbital shaking (220 rpm). When required, ampicillin or kanamycin was added to a final concentration of 100 or 50 µg·mL‐1, respectively. Vector pZERO‐1 (Invitrogen) was used to construct a genomic library of M. aurum B8.A in E. coli Top10. The pCR‐XL‐TOPO vector (Sigma) was used for sequencing of the MaAmyA encoding gene; pET‐15b (Novagen) was used as expression vector for the amyA gene constructs in E. coli BL21(DE3).
Bioinformatic tools Page | 145 All BLAST searches were performed with NCBI BLASTP using standard settings. To find all sequences related to the catalytic domain of MaAmyA, aa57‐504 were used as query in BLAST searches, using standard settings but with the maximum target sequences increased to 1000 (instead of the default 100). Conserved domains were detected using both the NCBI conserved domain finder (13) with forced live search, without low‐complexity filter, using the conserved domain database (CDD), and dbCAN (14) with standard settings. Signal sequence was predicted with SignalP4.1 (15) using standard settings. Catalytic AB regions of the GH13 domain sequences were extracted from the full length protein sequences based on dbCAN domain information. CBM25 domain sequences were extracted from the full length protein sequences based on CDD information. Alignments were made of the extracted domain sequences with Mega6.0 (16) using its build‐in muscle alignment with standard settings. Alignments were checked for the correct positions of conserved residues and manually tuned when needed. Phylogenetic trees were made with Mega6.0 using the maximum Likelihood with gaps/missing data treatment set on “partial deletion” instead of “full deletion”. Trees were visualized with Interactive Tree Of Life v2 (17). The protein domain annotations shown in the trees are based Degradation of granular starch involves FnIII and CBM25 domains
on combined data from CDD and dbCAN. Information about GH13 subfamilies was obtained from the CAZy database (1). An initial tree with a diverse selection of all GH13 subfamilies was constructed to select the closest related subfamilies for display in the final tree. The selection included multiple amylases from each of the 40 defined subfamilies, which varied in domain organization and origin. The final tree based on catalytic domain region AB, included MaAmyA, all GH13_32 members present in the CAZY database as well as a selection of the most closely related proteins based on the initial tree. The tree based on CBM25 domain sequences included the CBM25 domains from MaAmyA as well as 153 CBM25 domains listed in CAZy with a GenBank link.
Genomic DNA library and gene identification
Chromosomal DNA isolated of M. aurum B8.A was partially digested with Sau3A Page | 146 restriction enzyme (NEB), cloned into the pZERO‐1 vector (Invitrogen) and transformed to E. coli Top10. Transformants were grown on LB agar plates containing red amylopectin to screen for α‐amylase activity (18, 19). Colonies producing the largest halo sizes were isolated and the plasmid inserts sequenced.
Expression constructs of MaAmyA in E. coli
To easily create C‐terminal truncations of MaAmyA, pBAD‐VV was constructed by insertion of a synthesized multiple cloning site (containing removable N‐ and C‐terminal His‐Tags, a stop codon after the C‐terminal His‐Tag, and various restriction sites (NdeI, SpeI, NotI, PstI, and EcoO109I) into pBAD/Myc‐His B, using NcoI and EcoRI. The existing NdeI site in pBAD/Myc‐His B was disrupted with site directed mutagenesis. Full length amyA gene was obtained using PCR with the FW primer: gatgcatgatatcatatgtatccgaaaggaacaggcgca and RV primer: gctactctagaggatccttaacaccttggggtgggtgtgtggacta (restriction sites are underlined). The resulting PCR product was ligated into pCR‐XL‐TOPO vector and then inserted into pBAD‐VV using NdeI and SpeI restriction (Fig. S1). The final constructs were made through single restriction and self‐ligation of pamyApre and subsequently Chapter 4
transferred to pET15b using NdeI and EcoRI. The amyA construct was obtained through PstI restriction, the amyA2 construct through NotI restriction, and amyA4 through EcoO109I restriction (Fig. S4.1). For the amyA7 construct, an additional PstI site was inserted in the pamyApre construct through site directed mutagenesis, which was then cut with PstI, self‐ligated and transferred to pET15b (Fig. S4.1). All products were confirmed through sequencing (GATC‐Biotech). Final constructs all had an N‐terminal His‐Tag from the pET15b vector and a C‐terminal His‐Tag from pamyApre. The terminator present in pET15b was removed. All constructs were prepared with both a N‐ and C‐terminal His‐Tag.
Page | 147
Fig S4.1. Schematic overview of pamyApre and the different constructs prepared from it. AmyApre was created by cloning the amyA gene from M. aurum B8.A into pBAD‐VV, a modified pBAD/Myc‐His B vector. Final constructs were produced through restriction with the underlined restriction enzyme and self‐ligation. * = Inserted PstI site to create amyA7# = NotI, PstI and EcoO109I sites. Degradation of granular starch involves FnIII and CBM25 domains
Protein expression
Recombinant E. coli strains were grown as 500 ml cultures in 3 l flasks, with 100 µg·ml‐1 ampicillin and 50 µM (final concentration) IPTG for 6 h at 30 °C (220 rpm) and then for 40 h at 18 °C (220 rpm). Cells were collected by centrifugation at 4250 x g for 20 min at 4 °C (Thermo Lynx 4000). Pellets were resuspended in 50 mM Tris‐
HCl buffer pH 6.8 containing 10 mM CaCl2. Protease inhibitors (Mini EDTA‐free Protease Inhibitor, Roche) were added and cells were broken by sonication (15 sec at 10,000 , 30 sec cooling, 7 x). Cell debris and intact cells were removed by centrifugation at 15,000 x g for 20 min at 4 °C. Resulting cell free extracts were immediately used for His‐Tag purification of recombinant proteins.
His-Tag purification
Page | 148 The pH of E. coli cell lysates was adjusted to pH 7.5 and after mixing with Ni‐NTA (Sigma‐Aldrich), they were left to bind for 1 h at 4 °C. The column was washed 5
times with 2 column volumes of 50 mM Tris buffer pH 8.0 containing 10 mM CaCl2, 250 mM NaCl, 20 mM imidazole. Then the column was eluted 3 times with 2 column
volumes of 50 mM Tris buffer pH 8.0 containing 10 mM CaCl2, 250 mM NaCl, 500 mM imidazole. Elution fractions were desalted using a 5 ml desalting column (Amersham Pharmacia Biotech) and stored at 4 °C in standard assay buffer (50 mM
Tris‐HCl buffer pH 6.8 containing 10 mM CaCl2).
M. aurum B8.A culture fluid production
M. aurum B8.A was grown as a pre‐culture overnight at 37 °C at 220 rpm in LB medium. MMTV medium (12) containing 1% (w/v) granular potato starch (200 ml in a 1 l flask) was inoculated with 200 µl preculture and grown for 48 h at 30 °C, 220 rpm. The golden yellow cultures were harvested by centrifugation at 4250 x g for 20 min at 4 °C (Thermo Lynx 4000). The resulting cell free culture fluid containing the extracellular enzymes was collected, 100 µg·ml‐1 ampicillin and 100 µg·ml‐1 kanamycin were added and the culture fluid was stored at 4 °C. Chapter 4
Standard assay buffer
A 50 mM Tris‐HCl buffer pH 6.8 containing 10 mM CaCl2 was used as standard assay buffer for enzymatic incubations. Unless indicated otherwise all incubations were performed at 37 °C.
Activity staining on SDS-PAGE
SDS‐PAGE analysis was used to determine protein masses. The locations of starch‐ acting enzymes were visualized in the gels on basis of their activity. SDS‐PAGE was performed as described by Laemmli (20) using precast THX gels (Bio Rad). Use of THX gels prevented appearance of the additional bands of protein fragments which were observed previously (12) (see discussion). After loading and running, gels were washed 3 times for 5 min in MilliQ to remove SDS and then incubated for 2 h in Page | 149 standard assay buffer containing 0.5% soluble potato starch (Sigma‐Aldrich) (7).
After incubation the gel was stained with Lugol's iodine (2.5% I2 / 5% KI) to visualize active protein bands. After imaging, gels were partly destained by washing in MilliQ and then stained with Bio‐Safe™ Coomassie Stain (Bio Rad) to visualize proteins. The Fermentas PageRuler prestained marker was included in each gel.
Activity assay and activity unit determination using CNPG3 as substrate
The CNPG3 compound with a 2‐chloro‐4‐nitrophenol (CNP) group coupled to maltotriose (G3) is a suitable substrate to determine α‐amylase activity (21). The assay is based on the detection of the released CNP group from the CNPG3 substrate by α‐amylase activity. The enzyme solution (10 µl) to be tested was prepared in a 96‐well microtiter plate. Prewarmed substrate (100 µl 2 mM CNPG3) was added and the reaction was followed through absorbance reading at 405 nm for 10 min in a microtiter plate reader (Spectramax plus, Molecular Devices, Sunnyvale CA, USA) set at 37 °C. A calibration curve was prepared using 0.03‐0.15 mM CNP (Sigma) in assay buffer. The activity was calculated as µmole CNP released min‐1·µl‐1 of enzyme Degradation of granular starch involves FnIII and CBM25 domains
solution. One unit was defined as the amount of enzyme needed to release 1 µmole CNP·min‐1 under standard assay conditions.
Enzymatic activity on soluble potato starch
Hydrolysis of soluble potato starch was determined with the DNS method by measuring the increase of reducing ends in time (22). A 0.4% soluble potato starch (Sigma‐Aldrich, catalog no. S2004) solution in assay buffer was preheated at 37 °C in a heating block (VWR). Then, 2.6 U·ml‐1 (CNPG3 assay) of enzyme preparation (in assay buffer) was added at T = 0 h, in a total volume of 500 µl. Samples of 50 µl were taken at T= 0, 7, 14, 21, 28 min. The reaction was stopped by adding 50 µl DNS reagent (22) and immediate incubation at 100 °C for 7 min in a heating block. Afterwards the samples were left to cool down and 400 µl of MilliQ water was added. Absorbance was measured at 540 nm in a spectrophotometer (Spectramax Plus). A standard curve containing 833 – 8333 µM glucose, as well as a blank sample Page | 150 to which no enzyme was added, were included. Determination of activity (Table 4.1) was based on the hydrolysis during the first 6 h of incubation.
Granular starch degradation
The degradation of granular starch was followed by measuring the release of soluble carbohydrates from the granules, as determined with the Anthrone method (23). Two different granular starches were used: milled potato starch (AVEBE) and wheat starch (Sigma‐Aldrich, catalog no. S5127). Granules were sterilized through gamma irradiation (Synergy Health). The moisture content of the starches was determined with a dry‐matter balance (Sartorius). For calculations of the percentage of starch degradation, the dry‐matter content of the starches was considered to be 100% carbohydrate, since other components like protein form a minor part of the contents (<2%). Ampicillin and kanamycin (100 µg·ml‐1 final concentrations) were added to prevent bacterial growth. His‐Tag purified E. coli cell free extract blank (empty vector) was included as a negative control.
Chapter 4
Granular starch substrate (2.3 ml 3.3%, w/v) was prepared in a 15 ml tube including 300 U (CNPG3 assay) of the enzyme to be tested. Samples were incubated at 37 °C in a rotating wheel and gently mixed. At T= 0,1,2,3,4,5,6,14,24,48 h 200 µl samples were taken from the suspension and centrifuged for 20 min at 14,000 x g. Supernatant (50 µl) was transferred to clean glass test tubes. Samples containing more than 8 mg·ml‐1 carbohydrate were diluted with assay buffer before analysis. MilliQ (200 µl) and 2 ml Anthrone reagent (23) were added and mixed. Samples were incubated in a boiling water bath for 10 min, cooled down in a cold water bath and the absorbance measured at 620 nm in a spectrophotometer (Spectramax Plus). A calibration curve of 1.66‐8.33 mg·ml‐1 glucose was included.
Pelleted starch granules were dried for 48 h at 48 °C and stored in a desiccator for at least 1 day. The dried starch granules were transferred to SEM stubs with double‐ sided adhesive tape and coated with gold. Scanning electron micrographs (SEM)
were taken using the JSM‐6301F Scanning Microscope (JEOL at the UMCG Microscopy Page | 151 and Imaging center (UMIC), Groningen, The Netherlands). The accelerating voltage (3.0 kV) and the magnification are shown on the micrographs. For each wheat starch sample three images where recorded of the same area at 750x, 2000x and 5000x magnification. For each potato starch sample 2 images where recorded of the same area at 2500x and 25000x magnification.
Determination of pore surface sizes was performed using the digitally recorded SEM images with lowest magnification. Pictures where loaded in Adobe Photo shop (version 6.0) in grayscale picture modus, zoomed to true pixel. Pores were selected with “the magic wand” selection tool at following settings: size 1 pixel, tolerance 40, contiguous, no Anti‐Alias. A total of 10 pores were randomly chosen and the number of pixels selected by ‘the magic wand’ was recorded for each pore. The average and standard deviations were calculated. The scale bar was used to determine the conversion of pixel numbers to µm2.
Nucleotide sequence accession number Degradation of granular starch involves FnIII and CBM25 domains
The sequence for the novel amylase gene amyA, isolated from M. aurum B8.A has been deposited into GenBank database under accession no. KP901246.
Page | 152 Chapter 4
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28. Yamaguchi R, Arakawa T, Tokunaga H, Ishibashi M, Tokunaga M. 2012. Distinct characteristics of single starch-binding domain SBD1 derived from tandem domains SBD1-SBD2 of halophilic Kocuria varians alpha-amylase. Protein J 31:250–8.
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Degradation of granular starch involves FnIII and CBM25 domains
Page | 158 Identification of Novel α‐amylase from B. megaterium NL3
Chapter
Identification of a Novel -Amylase from Marine Bacillus megaterium NL3 Representing a New GH13 Subfamily
Fean D. Sarian, Tjaard Pijning, Ihsanawati, Zeily Nurachman, Marc J.E.C. van der
Maarel, Lubbert Dijkhuizen and Dessy Natalia
Page | 159
To be submitted for publication Identification of Novel α‐amylase from B. megaterium NL3
Abstract
Microbes employ a large diversity of α‐amylase to degrade starch as growth substrate. Most of these α‐amylase have been classified in Glycoside Hydrolase family 13 (GH13) and their three‐dimensional structures and reaction mechanisms have been studied in detail. New variants however are still identified from diverse natural environments. Here we report the characterization of a novel α‐amylase enzyme from Bacillus megaterium NL3, a bacterial strain isolated from Kakaban landlocked marine lake, Derawan Island, East Kalimantan, Indonesia. The mature recombinant BmaN1 consisting of 505 amino acid residues shares about 90% amino acid sequence similarity with the putative α‐amylases of Bacillus flexus, Bacillus aryabhattai, B. megaterium strain DSM319, and strain WSH‐002. Phylogenetic analysis clustered BmaN1 and its homologs as a separate branch in family GH13, representing a new subfamily. Sequence alignments revealed that members of this subfamily lack one of the conserved aspartate residues in their catalytic region, however BmaN1 still possessed normal α‐amylase activity. Modelling of the BmaN1 active site suggested that members of this new subfamily are likely to use a histidine instead of an aspartate residue as transition state stabilizer.
Page | 160 Chapter 5
Introduction
α‐Amylases are ubiquitously present in nature. In humans, α‐amylases are synthesized primarily in the salivary glands and the pancreas (1, 2). The digestion of starch in humans begins with the action of salivary α‐amylase and is completed by the pancreatic α‐amylase converting starch into glucose (3). α‐Amylases are also produced by many plants and microorganisms, in the latter mostly as extracellular enzymes to degrade glycogen and starch to smaller oligosaccharides and finally into glucose that serves as an energy source.
α–Amylases act on the α(14) glycosidic linkages in glycogen, starch, and related α‐glucans. The vast majority of α‐amylases belongs to glycoside hydrolase (GH) family 13 (4), constituting 20 different reaction and product specificities including, glycoside hydrolases (EC 3.2.1.x), glucanotransferase enzymes (EC 2.4.1.x and EC 2.4.99.16), and isomerases (EC 5.4.99.11; EC 5.4.99.15 and EC 5.4.99.16), all sharing a conserved structural scaffold (4). The first experimentally determined three‐ dimensional (3D) structure of α–amylase was Taka α‐amylase A (TAA), the amylase from Aspergillus oryzae (5). The crystal structure of TAA revealed that α‐amylases Page | 161 have three characteristic domains: A, B, and C (6). The A domain containing the
catalytic residues is the most conserved domain, with a typical (β/α)8‐ or TIM‐ barrel motif comprised of eight stranded parallel β‐sheet surrounded by eight α‐helices. Domain B is found inserted between the third β‐strand and the third
α‐helix of the (β/α)8‐barrel and varies in length and structure. The C domain folds into eight antiparallel β‐strands, is connected to the A domain by loops and seems to be an independent domain with unknown function (6, 7). Despite low similarity between the amino acid sequences of α‐amylases from animals, plants, and microbes, the GH13 enzymes share four highly conserved regions that are involved in the formation of the catalytic site. The α‐amylase active site is located in an open cavity between the A and B domains, and contains the invariably carboxylic acid Asp206, Glu230 and Asp297 (TAA numbering) that are essential for catalysis, acting as nucleophile, general acid/base and transition state stabilizer, respectively (5).
Identification of Novel α‐amylase from B. megaterium NL3
Recently, several α‐amylase and related enzymes composed of a (β/α)7‐barrel (an irregular TIM‐barrel domain) have been classified into families GH57 and GH119 (8, 9). Both of these enzyme families are at present considerably smaller than GH13 and only few have been characterized in detail (10). The first determined 3D structure of GH57 was that of the 4‐α‐glucanotransferase from Thermococcus litoralis (TLGT). X‐ray crystallography supported by site‐directed mutagenesis of TLGT revealed that it has two catalytic residues, Glu123 and Asp214, acting as the catalytic nucleophile and the general acid/base, respectively (11). No 3D structure is currently available for GH119 members. In addition to the structural differences between GH13 and GH57‐GH119 family members, there are also distinctive conserved regions between these families (8). The GH57 and GH119 families possess their own five conserved sequence regions (12).
We screened several microbial strains isolated from a unique land‐locked marine lake located in Kakaban island for the production of α‐amylases. Kakaban island is part of the Derawan Islands, East Kalimantan, Indonesia. Kakaban lake is also known as Jellyfish lake or Halimeda lake. It was originally the lagoon of an atoll, formed by Page | 162 corals over a period of two million years. As a result of movements in the earth's
crust the coral reef was raised above the sea level, trapping 5 km2 of seawater within a 50 meter high ridge, effectively creating a land‐locked marine lake (13). It has a pH of 7.6‐7.8, a salinity of 23‐26% and a tidal range of 11 cm. There is only one other similar ecosystem known which is found on Rock Island‐Palau in Micronesia (14).
The present paper reports the characterization of a novel α‐amylase produced by Bacillus megaterium strain NL3 isolated from a sea anemone from the Kakaban land‐ locked marine lake. B. megaterium strains have been reported as producers of extracellular α‐amylases (15). Analysis of the full genome sequences of B. megaterium strain DSM319 and strain WSH‐002 has revealed that they encode several putative amylolytic enzymes (16, 17). From the B. megaterium NL3 isolate, a gene coding for an α‐amylase, BmaN1 was identified. Of the three conserved catalytic residues in GH13 α‐amylases, only two catalytic amino acids were
Chapter 5
identified in BmaN1, being Asp203 and Glu231. In spite of this, BmaN1 was shown to be active towards soluble starch. Phylogenetic analysis clustered BmaN1 and its homologs as a separate branch in family GH13, representing a novel subfamily.
Page | 163
Identification of Novel α‐amylase from B. megaterium NL3
Results
Screening and characterization of bacteria producing extracellular α‐amylases
Twenty bacterial isolates from Kakaban landlocked marine lake were tested for production of ‐amylase by the starch hydrolysis test. Eight bacterial isolates grown on MB plates containing red‐dyed amylopectin or raw starch, produced clear zones around their colonies. The isolate NL3 showed the largest clearing zone on starch agar plates, indicating a relatively high ‐amylase activity and was selected for further study. A 16S rDNA sequence analysis showed that the sea anemone associated bacterial strain NL3 was most closely related to Bacillus megaterium. This result was in agreement with biochemical and physiological properties (data not shown) and hence the selected isolate was designated as B. megaterium NL3. ‐ Amylase activity was detected in culture supernatant of strain NL3 by measuring its activity towards soluble starch. The extracellular ‐amylase enzyme was partially purified by ammonium sulphate precipitation at a 50‐80% concentration, as this fraction contained most of the α‐amylase activity. The 50‐80% ammonium sulphate sample of the supernatant was run on SDS‐PAGE gels and by activity staining, a Page | 164 single protein band with molecular mass of approximately 55 kDa was found (data not shown).
Molecular identification of α-amylase
Using degenerate ‐amylase specific primers and inverse PCR, a DNA fragment of 2.3 kb was obtained. Analysis of the nucleotide sequence of this fragment showed that an open reading frame of 1515 bp with clear α‐amylase sequence similarity, with an ATG as the initial codon and TAG as the stop codon was present. This gene was designated as bmaN1. According to the prediction, the putative ribosomal binding site (RBS) corresponds to the AGGAGG sequence located 12 nucleotides upstream of the start codon. A probable catabolite responsive element (CRE) was found together with possible ‐10 (TATAAT) and ‐35 (TTAACA) regions (Fig. 5.1). The CRE sequence showed only one mismatch in the last position when compared to
Chapter 5
the consensus sequence (TGT/AAANCGNTNA/TCA) (26). The ORF downstream from bmaN1, had lacI as predicted product and is oriented in the same direction from bmaN1.
The sequence of bmaN1 was then used to perform a homology search using BLASTN software against the GenBank nucleotide collection (nr/nt) database (27). On the basis of multiple sequence alignments, the bmaN1 gene exhibits the highest identity (97%) with the putative ‐amylase of B. megaterium strain DSM319, and 94% identity with the putative ‐amylase of the B. megaterium strain WSH‐002. The full‐ length DNA sequence of the putative ‐amylase gene of B. megaterium NL3, bmaN1, has been deposited in the GenBank database under the accession no. AGT45938.
CRE TCAACTTATTTAACAATAATGTAAGCGTTAACGTTATAATGGAAACAAAAGTGAGTTTGAAACAGTTC
RBS GTCTATAGGAGGGGAATAGATGAAATGAAATGGAAA
Fig 5.1 Sequence analysis of the 5’ region of the B. megaterium NL3 bmaN1 gene. The putative ‐10 and ‐35 promoter regions (underlined), a putative CRE (overlined), the Page | 165 ribosome‐binding site (double‐underlined), and the start codon (boldface) are shown.
The BmaN1 polypeptide deduced is 505 amino acid residues in length with a clear putative signal peptide sequence of 23 residues preceding the mature enzyme, as predicted by SignalP 4.0 Server (28). The molecular weight and pI of BmaN1 were predicted using ExPASy server (http://web.expasy.org/protparam) as 56934 Da and 9.05, respectively. The deduced amino acid sequence of BmaN1 was subjected to BLASTP in GenBank revealing that BmaN1 is considerably similar to the glycoside hydrolase family 13 (GH13). BmaN1 shared the highest identity (39%) to ‐amylase from Geobacillus thermoleovorans (GTA, GenBank accession number: 4E20_A) (29) of all ‐amylases characterized functionally and biochemically. Furthermore, BmaN1 shared 95, 94, 94, and 92% amino acid sequence identity with the ‐amylase catalytic regions, I to IV, of B. flexus (GenBank accession number: WP_050691052),
Identification of Novel α‐amylase from B. megaterium NL3
B. megaterium DSM319 (GenBank accession number: ADF37524), B. aryabhattai (GenBank accession number: WP_043980530), and B. megaterium WSH‐002 (GenBank accession number: AEN91476), respectively. These ‐amylases have not yet been characterized experimentally yet.
To identify the conserved regions and the catalytic triad of BmaN1, the amino acid sequence of BmaN1 was aligned with α‐amylases from archaea and bacteria that have been classified into various GH13 subfamilies (Table 5.2). The conserved regions II, III, and IV are clearly present in BmaN1. However, the region corresponding to region I is different from any of the α‐amylases. Regions I‐IV are the well‐known conserved sequence regions in GH13 α‐amylases; region II, III, and IV contain the three carboxylic residues (Asp, Glu, and Asp, respectively) that together function as a catalytic triad. Surprisingly, two of these residues are absent in BmaN1 (Table 5.2 and supplementary data). First, at the position of the conserved nucleophilic aspartate residue, BmaN1 has a lysine residue (Lys202). Interestingly however, directly downstream of this lysine residue is an aspartate residue (Asp203). Second, the aspartic acid residue that stabilizes the covalent transition Page | 166 state is not present in BmaN1; it is replaced by a histidine residue (His294). Alignment with the sequences from B. flexus, B. megaterium DSM319, B. aryabhattai, and B. megaterium WSH‐002 (supplementary data) showed that also in these sequences, a lysine and a histidine are present at the positions of the nucleophilic and the transition state stabilizing residues, respectively. Also the aspartate residue directly downstream of the lysine residue is conserved in these sequences.
Chapter 5
Table 5.2 Comparison of amino acid sequences of the four conserved regions of B. megaterium NL3 BmaN1 and various ‐amylases from selected members of GH13 subfamilies
Accession GH13 Organisms Region I Region II Region III Region IV numbera subfamily
BmaN1 EFPLTI GYYVKDIDQ GEING FLDDVH T1SIF2 new
B. amyloliquefaciens DVVLNH GFRIDAAKH AEYWQ FVENHD P00692 5
Pyrococcus woesei DVVINH GWRFDYVKG GEYWD FVANHD Q7LYT7 7
Hordeum vulgare DIVINH AWRLDFARG AEVWD FVDNHD P00693 6
Homo sapiens DAVINH GFRLDASKH QEVID FVDNHD P04746 24
D. melanogaster DVVFNH GFRVDAAKH QEVID FVDNHD P08144 15
Streptomyces limosus DSVINH GFRIDAAKH QEAIH FVDNHD P09794 32
Xanthomonas campetris DVVFNH GFRVDAAKH GEVIT FAVTHD Q56791 27
Bacillus subtilis DAVINH GFRFDAAKH GEILQ WVESHD P00691 28
G. thermoleovorans DFVANH GYRLDTVRH GEVWS FLDNHD G8N5V2 ‐ Page | 167
Aspergillus oryzae DVVANH GLRIDTVKH GEVLD FVENHD P0C1B3 1
Vibrio fischeri DGVFGH GWRLDQAYQ AEIWN MLGNHD B5FDL2 37
D. thermophilum DLVVNH GFRLDAAKH GEVWD FLRNHD P14899 36
Escherichia coli DVVMNH GFRVDTAKH GEAWG YLSSHD P25718 19
CONSENSUS D***NH G*R*D***H GE*** F**NHD
The catalytic residues are shown in shadow and bold. Representatives of the 13 subfamilies of GH13 that showed α‐amylase specificity were compared. Organisms: B. amyloliquefaciens = Bacillus amyloliquefaciens; D. melanogaster = Drosophila melanogaster; G. thermoleovorans = Geobacillus thermoleovorans; D. thermophilum = Dictyoglomus thermophilum. The available gene sequences are aligned by ClustalW. aAccession numbers from the protein sequence database.
Identification of Novel α‐amylase from B. megaterium NL3
B A Domain A Domain A Domain B Domain B
Domain C Domain C
C
Page | 168
Fig 5.2 Structural comparison of BmaN1 of B. megaterium NL3 and GTA of Geobacillus thermoleovorans. (A) The BmaN1 model structure, (B) The structure of BmaN1 (orange) superimposed on GTA (grey) structure, (C) Active‐site region in a superposition of BmaN1 with GTA including the acarbose bound in subsites ‐2 to +2 (white carbon atoms). Active‐ center residues of BmaN1 (orange) and GTA (grey) are given as stick models and labeled in orange (BmaN1) and black (GTA).
A topological alignment of BmaN1 and the putative ‐amylase of B. megaterium DSM319, Bacillus sp. 278922, B. flexus, B. aryabhattai, B. megaterium WSH‐002, and GTA was made (supplementary data). Almost all β‐strands and α‐helices of the TIM Chapter 5
barrel in domain A, and the Greek key motif in domain C, are conserved in these ‐amylases. A model of the three‐dimensional structure of BmaN1 was generated by the PHYRE server (25) and visualized by the MacPymol software (30). The BmaN1 protein displayed 40% homology (100% confidence, 85% sequence coverage) with the X‐ray crystal structure of Geobacillus thermoleovorans α‐amylase (GTA, PDB code: 4E2O) (29) which was used as a template for the modelling. The comparison between the model and the GTA crystal structure revealed that the global topology is almost the same (Fig. 5.2B). The BmaN1 protein model folds into three distinct domains: a central A domain of 366 residues harboring a (/)8 barrel, with an irregular loop domain of 37 residues (domain B) connecting the third β‐sheets strand and the third α‐helix of the barrel. The C domain of 79 residues has an eight‐ stranded anti‐parallel β‐sandwich‐like fold (Fig. 5.2A).
Superposition of acarbose‐bound GTA with the BmaN1 model demonstrates that of three catalytic residues found in GH13 ‐amylases, only residue Glu231 of BmaN1 superimposes with the corresponding residue in GTA (Glu246), and presumably is the general acid/base in BmaN1. As already concluded from sequence alignments, two of the three catalytic residues are not conserved in the BmaN1 model. Lys202 Page | 169 replaces the catalytic aspartate (Asp217 of GTA); however, Asp203 directly downstream of the lysine is positioned nearby and has its carboxylic acid side chain pointing into the presumed substrate binding groove. Furthermore, at the position corresponding to the nucleophile, His294 replaces the transition‐state stabilizing aspartate residue (Asp314) found in ‐amylases.
To elucidate the evolutionary status of BmaN1, a phylogenetic tree of α‐amylases was constructed using representatives from each of the GH13 subfamilies as well as the B. megaterium NL3 α‐amylase, BmaN1 and the putative amylase of B. megaterium DSM319, Bacillus sp. 278922, B. flexus, B. aryabhattai, B. megaterium WSH‐002. To date, a total of 40 subfamilies of GH13 can be found in the Carbohydrate‐Active enzyme (CAZy) database (http://www.cazy.org/), with the α‐amylase specificity present in the subfamilies 1, 5, 6, 7, 15, 19, 24, 27, 28, 32, 36,
Identification of Novel α‐amylase from B. megaterium NL3
and 37 (31). The unrooted distance tree was based on the alignment of the sequences of the conserved regions, I to VII, of α‐amylases. BmaN1 does not cluster with any of the previously defined GH13 subfamilies (Fig. 5.3). This characterized BmaN1 forms a cluster together with the uncharacterized α‐amylases from B. flexus, B. megaterium DSM319, B. aryabhattai, and B. megaterium WSH‐002, and this cluster is most closely related to the GH13_x group comprised of several Bacillus α‐amylases (32) (Fig. 5.3). The amino acid identity of the sequences in the uncharacterized group with BmaN1 was in the range of 90‐95%, while similarity of these sequences with the nearest neighbor (GTA group) was below 30%.
At present, a few amylolytic enzymes with no sequence similarity to family GH13 have been classified in families GH57 and GH119. Some GH57 members are known to lack one or even both catalytic residues (12). Since only two corresponding residues to the conserved catalytic site of α‐amylase are identified in BmaN1, a comparative analysis of BmaN1 to GH57 and GH119 families was done. In phylogenetic analysis, BmaN1, 8 amylases of GH57 and 2 amylases of GH 119 formed three separate clusters (data not shown). Page | 170
Fig 5.3 Phylogenetic relationship of BmaN1 with ‐amylases of GH13 subfamilies. The tree was constructed using the neighbor‐joining method based on the sequence alignment of ‐amylases conserved regions (I‐VII) of a selection of GH13 α‐amylases performed using Clustal X embedded within the MEGA4 software. The GH13 ‐amylases are divided into subfamilies. Protein database numbers are shown following the sources of the ‐amylases. Bootstrap values, expressed as a percentage of 1000 replications, are given at the branching nodes. The scale bar at the lower left corner refers to the substitution rate (substitution/site).
Chapter 5
GH13_5
GH13_7
GH13_6
GH13_24
GH13_15
GH13_32
GH13_27
GH13_28
GH13_x
GH13_ Page | 171
GH13_1
GH13_37
GH13_36
GH13_19
Identification of Novel α‐amylase from B. megaterium NL3
Characterization of BmaN1
The gene encoding BmaN1 was cloned in vector pMM1525 and this recombinant plasmid was transformed to B. megaterium MS941. A transformant with clear ‐amylase activity, as detected on starch plates by iodine staining, was selected and grown in liquid medium. The culture medium was saturated with 50‐80% concentrations of ammonium sulphate to purify the BmaN1 ‐amylase enzyme. The molecular weight of the partially purified BmaN1 was estimated to be 55 kDa as judged from activity staining after protein renaturing on SDS‐PAGE gels (Fig. 5.4, lane 2). In contrast, no band was observed in the culture supernatant of B. megaterium MS941 carrying pMM1525 without any insert (Fig. 5.4, lane 1).
(kDa) 2 1 M 1 2
70
53
41 Page | 172
Fig 5.4 SDS‐PAGE(left) and activity staining after renaturation (right image) of BmaN1 purified from B. megaterium MS941. M, molecular weight marker; lane 1, pMM1525 empty vector; lane2, BmaN1 protein. Activity staining was performed by soaking the starch‐ containing gel in 50 mM maleate buffer ph 6.0 and incubating at 55 oC for 4 h. The clear bands after iodine solution treatment indicate amylolytic activity.
Amylolytic activity of BmaN1 was measured spectrophotometrically by incubating it with soluble starch and measuring the increase in the amount of reducing sugars released over a period of 40 min (Fig. 5.5). A clear increase in reducing ends was
Chapter 5
observed, resulting in an activity of 8.4 U/mg of protein. BmaN1 was found to be most active at 55 oC and pH 6.0. The main end products formed from soluble starch were glucose and maltose. Minor amounts of longer maltooligosaccharide were also found (Fig. 5.6).
6
4
2 reducing ends (mM) ends reducing
0 0 10203040 Incubation period (min)
Fig 5.5 Incubation of BmaN1 (open squares) with soluble starch; open circles: control (empty vector). 1% (w/v) soluble starch and 12.5 µg/mL of the BmaN1 protein were incubated for various time at 55 oC. Each data point represents the means of triplicate Page | 173 experiments.
G1
G2
≥>G3G3
Fig 5.6 HPLC analysis for hydrolyzed products of soluble starch by BmaN1. A reaction mixture containing 1% (w/v) soluble starch, 12.5 µg/mL of BmaN1, 50 mM maleate buffer pH 6.0 in a total volume of 0.1 mL was incubated at 55 oC. G1, glucose; G2, maltose; ≥G3, maltooligosaccharides.
Identification of Novel α‐amylase from B. megaterium NL3
Discussion
In the present study, Bacillus megaterium NL3 was found to produce an extracellular α‐amylase, BmaN1. Several novel structural properties of the BmaN1 protein were found using bioinformatics tools and homology modelling. The secondary structure
prediction of BmaN1 showed a typical (/α)8 barrel structure, which is a common feature of α‐amylase glycoside hydrolase family 13. The overall structure of BmaN1 can be divided into three domains (see Fig. 5.3): the first domain is composed of 339 residues (1‐139 and 177‐377) corresponding to domain A in the α‐amylase family enzymes, containing an eight‐stranded parallel ‐barrel and eight α‐helices
surrounding the ‐barrel (33). This barrel is quite similar to the (/α)8‐barrel found in maltogenic amylase from Pseudomonas saccharophila and Bacillus stearothermophilus (34), that both have an additional helix between Aα6 and A7, which is a three‐turn helix lying nearly parallel with the Aα6 strand. The B‐domain
with residues 140‐176 is formed by a single α‐helix interrupting the (/α)8‐barrel between the third β–sheet and the helix, and the C‐terminal domain with residues
378‐505. The members of GH13 possess related (/)8 barrels with a small irregular
loop B domain, while GH57 members contain the catalytic (/α)7 barrel and the Page | 174 GH119 have a yet uncharacterized motif. On the basis of this information we conclude that BmaN1 belongs to family GH13. In sequence alignments, the BmaN1 protein showed more than 90% similarity to the putative B. flexus, B. megaterium DSM319, B. aryabhattai, and B. megaterium WSH‐002 α‐amylases. Phylogenetic analysis of BmaN1 together with these five homologous putative α‐amylases against other GH13 family enzymes from the NCBI database indicated the uniqueness of these 6 proteins. They have their own separate branch that constitutes a new GH13 subfamily (Fig. 5.3).
GH13 ‐amylases have four conserved regions that are important for catalytic activity or substrate binding (6). Sequence alignments revealed that the conserved GH13 regions I‐IV of BmaN1 are remarkably different. The sequence of region II (199GYYKDID205 in BmaN1) suggests that Asp203 may function as the nucleophile, although it is one position downstream of the expected position, where in BmaN1 a
Chapter 5
lysine residue is found. In region III (229GEING233), the general acid/base residue, Glu230, is conserved. In region IV (289FLDDVH294), a histidine (His294) replaces the canonical aspartate that functions as a transition‐state stabilizer (Fig. 5.2). Sequentially similar motifs have also been recognized in the 5 BmaN1 homologs representing unclassified putative α‐amylases, of various Bacillus species. They have been added to GH13 because part of their sequence exhibits similarity to α‐amylase (http://www.cazy.org).
A three‐dimensional structure of BmaN1 was built by homology modelling using Quick Phyre. The secondary and tertiary structures of the BmaN1 model are similar to that of the template enzyme (GTA, PDB code: 4E2O). Superimposition of the active site of BmaN1 on GTA revealed several differences as well as similarities in the active site structure. In particular, there is an insertion of a lysine residue in the catalytic region around Asp203 (BmaN1 numbering). As a consequence, Asp203 seems to be shifted in position, although still positioned proximate to the substrate. In the absence of an experimental 3D structure, it is unclear whether or not it can perform the nucleophilic attack. In addition, residue Asp314 in GTA, providing hydrogen bonds to the sugar moiety in subsite ‐1, is substituted by His294 in Page | 175 BmaN1. Although different from aspartate, a histidine side chain is also able to form hydrogen bonds; this residue thus may act as alternative transition state stabilizer.
α‐Amylase activity was observed following expression of BmaN1 in B. megaterium expression host. The specific activity of the BmaN1 (8.4 U/mg from a DNS assay) is in the moderate range of α‐amylase activity (www.brenda‐enzymes.info). Glucose and maltose were the dominant products of BmaN1 incubated with soluble starch (Fig 5.6). This mode of action is very similar to that of the amylase from Bacillus sp. IMD 435 that releases glucose and maltose as the major products on hydrolysis of both soluble starch and raw corn starch (37). Interestingly, with regards to the amylolytic activity, our results show that despite the substitution of a negatively charged aspartate residue by a positively charged histidine (His294), BmaN1 still has an α‐amylase activity. The reaction mechanism of BmaN1 thus is essentially
Identification of Novel α‐amylase from B. megaterium NL3
similar to the general catalytic reaction mechanism of α‐amylases. Further experiments are needed to validate the role of the proposed catalytic residues, i.e by constructing and characterizing mutant enzymes with different residues at this position, and/or determining the 3D structure, preferably in complex with a substrate.
Conclusion
An extracellular α‐amylase of the new bacterial isolate B. megaterium NL3, BmaN1, has been identified and produced in a B. megaterium expression host. Based on amino acid sequence similarity and phylogenetic analysis, the BmaN1 and five unclassified homologous α‐amylases belong to a new GH13 subfamily. Interestingly, the highly conserved and catalytic aspartic acid residue in conserved region IV is replaced by a histidine in BmaN1. Taken together, this paper provides evidence that BmaN1 is a novel α‐amylase with an unusual catalytic triad.
Acknowledgements Page | 176 This research was funded by Asahi Glass Foundation, Institut Teknologi Bandung. We thank Prof. F. Meinhardt, University of Munster, Germany, for B. megaterium MS941 strain and pMM1525 plasmid. We are grateful to Dr. Lily Eurwilaichitr and Dr. Sutipa Tanapongpipat for allowing us to do the preliminary experiments on BIOTEC, NSTDA, Thailand.
Chapter 5
Material and Methods
Materials
All chemicals used were reagent grade and were obtained from either Fermentas (Maryland, USA) or Difco Laboratories (New Jersey, USA).
Bacterial strains, plasmids, and growth conditions
Twenty microbial strains (gift of Prof. Ocky Karna Radjasa of Diponegoro University, Indonesia) that had been isolated from Kakaban landlocked marine lake (Derawan Islands, East Kalimantan, Indonesia) were screened for extracellular α‐amylase activity. The isolates were cultured in marine broth (MB) medium containing 0.25% (w/v) yeast extract, and 0.5% (w/v) peptone in filtered sea water (Seaworld, Ancol, Jakarta, Indonesia) at 30 oC. B. megaterium MS941 (MoBiTec, Germany) and Escherichia coli TOP10 were grown at 37 oC in LB medium (1% (w/v) Bacto‐ tryptone, 1% (w/v) NaCl and 0.5% (w/v) yeast extract). Ampicillin and tetracyclin were used at concentrations of 100 g/ml and 12 g/ml, respectively. The medium was autoclaved at 120 oC for 30 min prior to adding the antibiotics. Plasmid pGEM‐T Page | 177 (Promega, USA) was used for PCR product cloning, whereas pMM1525 (MoBiTec, Germany) was used as expression vector.
Screening of α‐amylase producing bacteria
Bacterial isolates were inoculated on MB agar plates supplemented with 1.0% (v/v) red‐dyed amylopectin (18) and then incubated at 30 oC for 24 hrs. The appearance of a clear zone against a red background was indicative for the production of α‐amylase activity. The positive isolates were then subjected to a second screening round using MB agar plates containing 1.0% (w/v) potato or wheat starch granules. A clearing zone around the bacterial colony indicated that the starch granules was hydrolyzed and thus that the isolate produced extracellular α‐amylase activity.
Identification of Novel α‐amylase from B. megaterium NL3
Bacterial identification
Isolate NL3 showing the largest clearing zone on starch‐agar plates was identified by 16S rDNA sequencing. Chromosomal DNA of strain NL3 was isolated using Wizard Genomic DNA Purification (Promega). The 16S rDNA gene was amplified by PCR using universal primers UniB1 and BactF1 (Table 5.1). The resulting 1.4 kb fragment was sequenced using the dideoxy‐chain termination method (Macrogen, South Korea). Bacterial isolates were identified by aligning their 16S rDNA sequence with those of known bacteria using NCBI BLASTn (http://www.ncbi.nlm.nih.gov). The 16S rDNA gene sequence was submitted to GenBank (accession number KU862862)
Cloning of the α‐amylase‐encoding gene and plasmid construction
Two degenerate primers (Table 5.1) were designed based on the amino acid sequences of the well‐conserved regions (region VI‐VII) of α‐amylases from several Bacilli. The first α‐amylase gene fragment was amplified by polymerase chain reactions (PCR), using chromosomal DNA from B. megaterium NL3 as a template and the two degenerate primers. The PCR products were inserted into pGEM‐T vector Page | 178 (Promega) and transformed into E. coli TOP10. Plasmid DNA of the transformed E. coli TOP10 was isolated and the nucleotide sequence of the inserted DNA was determined using the dideoxy‐chain termination method (Macrogen). The resulting nucleotide sequence was used to design a set of primers, NL3_SP8‐invF1 and NL3_SP8‐invR1 (Table 5.1), to amplify parts of α‐amylase gene beyond the conserved region. The chromosomal DNA was partially digested with EcoRV and then self‐ligated using T4 DNA ligase (Fermentas). Inverse PCR (iPCR) (Ochman et al. 1988) was performed with Dream Taq polymerase (Fermentas) and the primers listed in Table 5.1 using the self‐ligated DNA fragment as a template. Analysis of sequence data and sequence similarity searches was performed using the BLAST program of the National Center for Biotechnology Information (NCBI). Primers pMM‐NL3‐F and pET/MM‐NL3‐R were used to amplify the complete open reading frame of the α‐amylase gene which was designated as bmaN1.
Chapter 5
Table 5.1 Primers used in amplification by PCR
Name Sequence Used
BactF1 AGAGTTTGATC(A/C)TGGCTCAG 16S rDNA
UniB1 GGTTAC(G/C)TTTGTTACGACTT 16S rDNA
bmD_reg2F GACGGATATTACGTTAAAGATATC PCR of region VI‐VII of bmaN1
bmD_reg6F GGATTTACCGCTGTTTTATTAACTCC PCR of region VI‐VII of bmaN1
bmD_reg3R ATTTCTCCTATTAATAGAAAAGAAGG PCR of region VI‐VII of bmaN1
bmD_reg7R ATAAAAAACGATTGGTACACTCG PCR of region VI‐VII of bmaN1
NL3_SP8‐invF1 ACTGCTGCTACRTTTAGC iPCR of bmaN1
NL3_SP8‐invR1 TATATKRCGCTCATGTGC iPCR of bmaN1
pMM‐NL3‐F GGATTCAGCAAGATCATAAAGATATACG PCR of bmaN1 Page | 179
pET/MM‐NL3‐R CGGCCGCTAGTGATGATGGTGATGGTGCGACGCGCTGT PCR of bmaN1 CCTTTTTACG
Transformation of B. megaterium
The recombinant plasmid containing the α‐amylase gene, pMM1525‐bmaN1, was transformed into the expression host, B. megaterium MS941. The transformation procedure was essentially conducted as described by Puyet et al. (1987) with some modifications (19). A 0.5 ml protoplast suspension was added to a tube containing 5.0 g DNA and 1.5 ml PEG‐P (40% (w/v) PEG6000 in 1x SMMP) for each transformation and incubated for 2 min at room temperature. SMMP medium contains 3.5% (w/v) AB3 (Antibiotic medium no. 3, Difco), 1 M sucrose, 40 mM disodium maleic acid and 40 mM MgCl2 (pH adjusted to 6.5 before autoclaving for 12
Identification of Novel α‐amylase from B. megaterium NL3
min) and prepared freshly before use. To the mixture, 5.0 ml SMMP was added and mixed by rolling the tube carefully. Cells were harvested by centrifugation at 2,700 x g for 10 min at room temperature and the supernatant was poured off immediately. The pellets were resuspended with 0.5 ml SMMP and incubated at 37 oC for 90 min with gentle shaking or rolling of tubes (max. 100 rpm). Then, 50 to 200 l of cells were added into top agar and mixed gently by rolling the tube. The mixture was poured on a pre‐warmed plate of LB containing 12 g/ml of tetracycline and incubated at 37 oC overnight.
Expression and partial purification of recombinant α‐amylase BmaN1
α‐Amylase BmaN1 was produced by growing the B. megaterium MS941 harboring recombinant plasmids in 20 ml of LB medium supplemented with 12 g/ml tetracycline at 37 oC with shaking. The overnight culture was transferred into fresh media and incubated until the 546‐nm absorbance reached 0.8‐1.0. Subsequently, expression was induced by adding 1% (v/v) xylose, and the culture was incubated at 18 oC with constant shaking at 150 rpm for 24 hrs. Cells were removed by
Page | 180 centrifugation (6000 x g, 10 min) and the resulting supernatant was subjected to ammonium sulphate precipitation at a saturation value up to 80%. The precipitate was dissolved and dialyzed against 50 mM maleate buffer pH 6.0 at 4 oC. This partially purified α‐amylase was used for further studies.
Gel electrophoresis and activity staining
SDS‐PAGE was carried out as described by Laemmli (1970) and proteins in gels were then stained with Coomassie Brilliant Blue (Bio‐Rad). For α‐amylase activity test, the protein samples were separated by SDS‐PAGE (20) containing 1% soluble starch. After electrophoresis, SDS was removed by washing the gels with water followed by 10 min incubation at room temperature. This was repeated twice. The gels were then immersed in the enzyme reaction buffer (50 mM maleate buffer pH
o 6.0) for 4.0 h at 55 C and then stained with KI/I2 solution for 10 min, followed by
Chapter 5
rinsing with water. The α‐amylase activity was detected as a clear zone against a purple background.
Enzyme assay
DNS assay. The α‐amylase activity assay was conducted using the 3,5‐ dinitrosalicylic acid (DNS) method described by Miller (1955) with a slight modification (21). Briefly, the assay was performed by adding 25 l (~12.5 µg/mL) of enzyme sample into 25 l of 1% (w/v) soluble starch (Fermentas, USA) in 50 mM of the appropriate buffer and then incubated at 55 oC for 20 min. To the reaction mixture, 50 l of DNS reagent was added. The absorbance at 500 nm was measured and then the amount of reducing sugar‐end was calculated using a glucose standard curve (1‐2.5 mM). One unit of α‐amylase activity was defined as the amount of enzyme that releases 1 µmol of reducing sugars per min under the assay conditions. The protein concentration was determined using the Lowry method (Lowry, et al. 1951) and bovine serum albumin as the standard.
Page | 181 Iodine assay. The activity of α‐amylase was also assayed by the iodine method (22) using soluble starch as substrate. A 50 l of enzyme was mixed with 50 l of 0.125% (w/v) soluble starch (Fermentas, USA) dissolved in 50 mM maleate buffer pH 6.0. After 20 min of incubation at 55 oC, 20 l of 1 M HCl was added to stop the enzymatic reaction, followed by the addition of 100 l of iodine solution (0.5%
(w/v) KI and 0.15% (w/v) I2). The blue color was read at 600 nm. One unit of the α‐amylase activity was defined as the amount of enzyme that produced 10% reduction in starch‐iodine staining after 20 min of incubation under the assay conditions. All enzyme assays were conducted in triplicate.
Analysis of sugars
The starch hydrolysis products were analyzed by high‐performance liquid chromatography (HPLC). Aliquots of 100 l of enzyme solution (12.5 µg/mL) were
Identification of Novel α‐amylase from B. megaterium NL3
incubated at 55 oC in the presence of 1% (w/v) soluble starch, maleate buffer 50 mM pH 6.0. After specific time intervals, samples were withdrawn and hydrolysis was stopped by incubation at 90 oC for 10 min. After centrifugation at 12000 x g for 10 min at 4 oC, the hydrolysates were then analyzed by HPLC (Aminex® HPX‐87H
system). H2SO4 0.008 N served as mobile phase, the flow rate being 1 mL/min at 35 oC. The injection volume of either hydrolysate was 10 µL. Calibration of the peak area was performed based on standard glucose, maltose, and purified maltooligosaccharide (Sigma). The separated hydrolysis products were identified and calculated by comparison with the standards.
Bioinformatics
A phylogenetic tree of family GH13 ‐amylases and BmaN1 was constructed using the conserved regions I‐VII of representatives of the following ‐amylase subfamilies (GH13_1, GH13_5, GH13_6, GH13_7, GH_15, GH13_19, GH13_24, GH13_27, GH13_28, GH13_32, GH13_36, and GH13_37). This set covers all GH13 subfamilies with well‐defined (GH13 subfamilies 1, 5, 6, 7, 15, 19, 24, 27, 28, 32, and
Page | 182 37) and putative (GH13_36) α‐amylase activities. The sequence alignment was performed using the Clustal W program (23) embedded within MEGA4 (24). The three‐dimensional structure of the BmaN1 protein was predicted by using the Quickphyre structure program server (http://www.sbg.bio.ic.ac.uk/phyre) (25). Structural analysis of the BmaN1 protein was performed based on superimposition with the crystal structure of the α‐amylase from Geobacillus thermoleovorans [PDB access code: 4E20]. The generated BmaN1 structures were displayed and drawn by MacPymol (http://pymol.org/educational).
Chapter 5
References
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15. David M-H, Günther H, Röper H. 1987. Catalytic properties of Bacillus megaterium amylase. Starch - Stärke 39:436–440.
16. Eppinger M, Bunk B, Johns MA, Edirisinghe JN, Kutumbaka KK, Koenig SSK, Creasy HH, Rosovitz MJ, Riley DR, Daugherty S, Martin M, Elbourne Page | 184 LDH, Paulsen I, Biedendieck R, Braun C, Grayburn S, Dhingra S, Lukyanchuk V, Ball B, Ul-Qamar R, Seibel J, Bremer E, Jahn D, Ravel J, Vary PS. 2011. Genome sequences of the biotechnologically important Bacillus megaterium strains QM B1551 and DSM319. J Bacteriol 193:4199–4213.
17. Liu L, Li Y, Zhang J, Zou W, Zhou Z, Liu J, Li X, Wang L, Chen J. 2011. Complete genome sequence of the industrial strain Bacillus megaterium WSH-002. J Bacteriol 193:6389–6390.
18. Jørgensen S, Vorgias CE, Antranikian G. 1997. Cloning, sequencing, characterization, and expression of an extracellular α-amylase from the hyperthermophilic archaeon Pyrococcus furiosus in Escherichia coli and Bacillus subtilis. J Biol Chem 272:16335–16342.
19. Puyet A, Sandoval H, López P, Aguilar A, Martin JF, Espinosa M. 1987. A simple medium for rapid regeneration of Bacillus subtilis protoplasts transformed with plasmid DNA. FEMS Microbiol Lett 40:1–5.
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regions of a novel Anoxybacillus α-amylase. Sci Rep 4:5850–5858.
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35. Nielsen JE, Borchert T V. 2000. Protein engineering of bacterial α-amylases. Biochim Biophys Acta - Protein Struct Mol Enzymol 1543:253–274.
36. Ke T, Ma XD, Mao PH, Jin X, Chen SJ, Li Y, Ma LX, He GY. 2007. A mutant α-amylase with only part of the catalytic domain and its structural implication. Biotechnol Lett 29:117–122.
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Page | 186
Chapter 5
β1
VI
α1 β2 α2
Page | 187
I Domain B V
α3 β3
Identification of Novel α‐amylase from B. megaterium NL3
II III
α4 α5 α6 β4 β5 β6
IV
α7 β7 β8
Page | 188
α8
Chapter 5
Page | 189
Supplementary data. Topological alignment of the BmaN1 primary and secondary structure with GTA (PDB 4E20) and other putative amylases. Alignment was prepared based on the super positioned models. Secondary structure: α‐helix ( ), TM‐helix ( ), and β‐sheet ( ); residues of the catalytic site (▲). Conserved secondary structures are named.
Identification of Novel α‐amylase from B. megaterium NL3
Page | 190
Chapter
General Discussion
General Discussion
Starch: structure, accessibility, and reactivity
Starch is one of the most abundant polysaccharides in nature next to cellulose and chitin and occurs as an insoluble storage polymer (Chapter 1). It is widely distributed in the form of granules of different sizes and shapes. Starch is composed of two polymers of glucose that have different composition, structure, and properties: amylose and amylopectin (1, 2). The distribution and amount of amylose and amylopectin define the structure of the starch granule, which in turn affects the physicochemical and functional properties (2).
Starch is widely used as a raw material for the production of a large variation of products that find their way in many daily life commodities. Numerous studies have been devoted to understanding the starch degradation process, due to the importance of starch as an industrial commodity and staple food for humans and animals. In general, native unmodified starch granules are much more difficult to degrade then soluble starch. Modifications such as gelatinization, phosphorylation, or oxidation, lead to an increased susceptibility of the granule for enzymatic degradation (3). Gelatinization disrupts the crystalline structure resulting in free amylose and amylopectin, increasing the starch susceptibility to amylolysis (4). The enzymatic degradation of native starch granules and its regulation are poorly Page | 192 understood (Chapter 1), except for the conversion of transitory starch present in leaves or the degradation of starch in germinating cereal seeds (5–8). The mode of enzymatic degradation depends both on the starch and enzyme used (9). The molecular architecture of starch can vary depending on the botanical source, which is presumably a major factor contributing to the α‐amylase accessibility. Differences in the resistance to enzyme hydrolysis can also be attributed to the starch granule’s physical and chemical characteristics, such as their polymorphism, porosity, amylose/amylopectin ratio, and granule size and distributions that vary among starches from various botanical origins (1). Starch granules are made of crystalline (ordered) and amorphous (disordered) parts in layers as can clearly be seen with polarized light or wide‐angle X‐ray diffraction (10, 11).
Chapter 6
Starch granules need to be seen as inert substrates when it comes to enzymatic degradation. Parallels can be drawn with the enzymatic degradation of plant cell walls containing (hemi)cellulose and pectin or the exoskeleton of insects and crustaceans, containing chitin (12). The way Nature overcomes the complexity of insoluble substrates has mostly been studied by mimicking enzymatic degradation in‐vitro. As discussed in Chapter 2, not all of the tropical starches tested were hydrolyzed to the same extent. Various tropical starches were treated in‐vitro with the commercial enzyme preparation StargenTM 002, which is used to convert native no‐cook starch to glucose at moderate temperatures. Differences in enzyme susceptibility were observed with sweet potato, sago, and corn having the highest degree of hydrolysis when incubated with StargenTM 002 for 7 h. Native edible canna and potato starches were more resistant to hydrolysis (Fig. 2.2). This result is in agreement with work by Sarikaya et al. (2000) that showed that the degree of hydrolysis depends on the botanical source of the starch granule (13). Rice and corn starches were found to be easily hydrolyzed, while potato starch was reported to be more resistant to degradation by α‐ and β‐amylases. Similar results have also been reported for the α‐amylase of Bacillus sp. IMD370 (14).
The high crystallinity of starch gives an ordered and tightly packed structure which limits α‐amylases to penetrate these organized structural entities. Potato starch granules are among the most difficult starch granules as they have a B‐type Page | 193 crystallinity and no pores or cracks on the granule’s surface (Chapter 1). An efficient native potato starch degrading bacterial strain was isolated from the waste‐ water treatment plant of a potato starch processing factory (Chapter 3). This isolate was identified as a new Microbacterium aurum strain, named M. aurum B8.A. Where the type strain DSMZ8600 did not degrade native potato starch granules, this new isolate did by drilling small pores in the starch granule (Fig. 3.2). Native potato starch degrading activity was detected in the culture medium of M. aurum B8.A grown on potato starch granules, while the type strain DSMZ8600 did not produce any potato starch degrading enzyme activity (Fig. 3.3). Hoover and Zhou (2003) also reported that normal cereal starch granules (A‐type) are most susceptible to amylolytic degradation compared to granules of potato, high amylose starches
General Discussion
(B‐type), or legume starch granules (C‐type) (15). Normal cereal starches have a smaller granular size than tuber or legume starches and in addition have a less perfect crystalline structure (16). A‐type starch granules also show more pores on the surface than the B‐type starch granules (16, 17). The presence of pores on the surface of starch granules enables α‐amylases to diffuse into the granules. According to Oates (1997), degradation of A‐type corn starch starts at susceptible areas followed by pore formation (18). The smooth surface structure of arrowroot and potato starch limits the enzyme diffusion, and hence, the degradation takes place on the surface (exocorrosion). After enzyme treatment, corn and sago starch granules were extensively damaged and showed the most porous structures while arrowroot or potato granules were slightly scratched or pitted (Fig. 2.3). The variation in hydrolysis efficiency is most likely correlated to the crystallinity of the starch and morphology of the granular surface.
Starch Hydrolyzing Enzyme Systems
Like starch, cellulose possesses a crystalline structure, which makes it highly resistant to enzymatic attack (19). Cellulose is composed of two different crystal forms co‐existing within a single microfibril (20). There are two well studied cellulolytic systems: the non –complex and the complex system. The non‐complex Page | 194 cellulolytic system is mainly found in aerobic bacteria and filamentous fungi (21, 22). This system is comprised of at least 3 types of free cellulases, that act sequentially and synergistically for effective cellulose hydrolysis into sugars (23, 24). Anaerobic cellulolytic bacteria produce complex cellulase systems that consists of large cell wall‐bound extracellular multi‐enzymes complexes, called cellulosomes (20, 24–27). Cellulosomes usually are attached to the cell surface and mediate efficient uptake of the hydrolysis products (20, 27, 28). Cellulosomes consist of several different cellulases and related enzymes which are assembled by unique scaffoldin subunits each containing a dockerin domain (20, 27). Cellulose degradation is initially mediated by binding of the enzyme to the cellulose substrate by the cellulose‐binding domain (CBD) (20). The presence of CBD was first discovered for the Trichoderma reesei cellobiohydrolase, showing binding activity to
Chapter 6
cellulose and functioning independently of the catalytic domain (29–31). Later on, the name of this module was changed to carbohydrate‐binding module (CBM), as it was found that similar modules were also present in other carbohydrate‐active enzymes.
As starch has a complex crystalline structure, the combined action of various different enzymes is required to convert starch polymer into soluble sugars. Many microbial amylolytic enzymes have one or more CBMs that interact with the crystalline part of starch (32, 33). The CBMs found in glycoside hydrolase (GH) family 13 enzymes have been classified into the families 20, 21, 25, 26, 34, 41, 45, 48, 58, and 69, and are commonly known as starch‐binding domains (SBDs) (34).
An extracellular α‐amylase of the Microbacterium aurum B8.A isolate (MaAmyA) that is described in Chapter 3, is unusually large with 1409 amino acids and a predicted mass of 148 kDa (Fig 4.1). The MaAmyA protein consists of 7 domains: 1. One catalytic domain, having a clear similarity to subfamily GH13_32; 2. Four fibronectin type III domains (FnIII); 3. Two CBM25 domains; and 4. A C‐terminal domain of 331 amino acids of which the function is not known (Chapter 4). The effects of truncation of the CBMs of the MaAmyA on its activity with potato starch granules was investigated. This α‐amylase initiates degradation of potato starch Page | 195 granules by making small pores into the granules (Fig. 4.6). The truncated MaAmyA mutant showed almost no activity towards starch granules (insoluble starch), while it was still active on soluble starch. This suggests that the CBMs clearly play a role in the adsorption of the enzyme onto starch granules. In addition, it was found that the CBMs also play a clear role in the pore formation (Fig. 4.6). Several other investigations have also shown that removal of the CBMs of α‐amylases reduces their hydrolytic activity on starch granules (35–37). The absence of the CBM even led to the complete inactivation of Lactobacillus amylovorus α‐amylase (35). Removal of the CBM of the Aspergillus niger glucoamylase reduced the enzyme activity to about 90% of the original activity (38).
General Discussion
The roles of fibronectin III (FnIII) domain in carbohydrate enzymes are still unclear. Besides being important for mediating protein‐protein interactions (42), FnIII has been shown to be important for hydrolysis of chitin of Bacillus circulans WL‐12 chitinase A1 (39). Truncation of FnIII of MaAmyA, however, did not abolish activity against insoluble starch (Chapter 4). It was suggested that FnIII domains are a stable linker between separate domains in cellobiohydrolase (40) and chitinase enzymes (41). The FnIII domains in MaAmyA may play a similar role. FnIII type domains also seem to play a role in protein‐protein interaction. It can be speculated that the FnIII domains in MaAmyA help this enzyme to stick to other extracellular amylases and in this way form a modular, multi‐enzyme structure, an amylosome, similar to the cellulosome. M. aurum B8.A has at least two extracellular α‐amylases, of which one is MaAmyA; the other α‐amylase is also a multi domain protein with at least one FnIII domain (van der Maarel, personal communication).
α‐Amylase Mechanism
Degradation of starch requires the action of several enzymes, among which α‐amylases (EC. 3.2.1.1) play a key role. α‐Amylases are endoacting enzymes that catalyze the hydrolysis of starch and related carbohydrates into simple sugars. α‐Amylase is found abundantly in saliva and the small intestine, playing a pivotal Page | 196 role in the conversion of starch to glucose (40). Many plants and microorganisms also produce various α‐amylases (41, 42). In most plant tissues, α‐amylase is produced to utilize transitory starch as a carbon source, which is synthesized and stored during photosynthesis (43). α‐Amylases are also found in tubers, roots, and seeds producing glucose for the young seedling. Bacteria and fungi secrete α‐amylases into their environment to convert starch into the oligosaccharides and glucose that are then taken up to support metabolism and growth (44).
Most reported α‐amylases are classified into family GH13 based on amino acid similarities, whereas a minority of α‐amylases belongs to families GH57 and GH119.
GH13 amylases are characterized by a (β/α)8‐barrel structure with a deep active site
Chapter 6
channel harboring the conserved catalytic machinery, consisting of the nucleophilic residue Asp206, the acid/base residue Glu230, and transition‐state stabilizer residue Asp297 (Taka amylase A, TAA, numbering). Several studies have reported that α‐amylase mutants lacking one of the three catalytic residues barely hydrolyzed starch (45, 46).
A new type of α‐amylase was isolated from Bacillus megaterium NL3, a strain that lives on a sea anemone in the landlocked lake Kakaban, Indonesia. BmaN1 has a characteristics (β/α)8‐barrel and exhibits high sequence similarity with α‐amylases from family GH13 (Chapter 5). Together with five other putative α‐amylases from various Bacilli, BmaN1 represents a new subfamily of GH13 (Fig. 5.3). However, BmaN1 as well as the other five putative α‐amylases have two peculiar features. The first is that at the position of the nucleophilic residue Asp206, BmaN1 has a lysine, coming from a completely different structural element. One position downstream of this lysine, an aspartate residue is present, very likely representing the nucleophilic residue in BmaN1. The lysine insertion very probably affects the active‐site topology and could have an effect on the enzyme’s activity. The second peculiar feature was a big surprise, at the position of the transition‐state stabilizing residue Asp297 (TAA numbering) a histidine (His 219, BmaN1 numbering) was present. This was not only found in BmaN1 but also in the five putative α‐amylases from other Bacilli. 3D modelling of the BmaN1 protein structure confirmed the position of the His at the Page | 197 position where in all other α‐amylases the transition‐state stabilizing Asp is present. Resolving the crystal structure of the BmaN1 protein, ideally with a bound model substrate, may shed light on this surprising finding.
The question remains what the actual substrate is for BmaN1, as (granular) starch is almost certainly not present in the Kakaban lake. Many organisms, including bacteria, synthesize glycogen as an energy and carbon storage compound when carbon compounds are abundantly available. Glycogen is, similar to amylopectin, a polymer of glucose with α(14) and α(16)‐O‐glycosidic linkages. To degrade glycogen, bacteria use α‐amylases and debranching enzymes. B. megaterium NL3
General Discussion
was isolated from a sea anemone of Kakaban lake. It is not known whether sea anemones contain glycogen but this appears very likely. Zierer et al. (47, 48) found that the glycogen of the marine sponge Aplysina fulva has an unusual glycogen as this contains sulphate groups at half of the non‐reducing ends. They stated that “these sulphate groups render the sponge glycogen more resistant to the degradative enzymes of microorganisms that inhabit the sponge canals” (47). It can be speculated that the sea anemone also has a sulphated‐glycogen similar to the A. fulva and that the BmaN1 is excreted by B. megaterium NL3 to hydrolyze this sulphated‐glycogen to glucose and maltose (Chapter 5) that subsequently are used by the NL3 strain for growth. A sulphated‐glycogen as substrate for BmaN1 may also explain the unusual architecture of the active site, with the nucleophilic Asp shifted one position, and the His at the position of canonical Asp that normally acts as a transition‐state stabilizer. Further studies clearly are needed here.
Page | 198
Chapter 6
References
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46. Zierer MS, Vieira RP, Mulloy B, Mourão PAS. 1995. A novel acidic glycogen extracted from the marine sponge Aplysina fulva (porifera-demospongiae). Carbohydr Res 274:233–244.
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Summary
For the non‐expert
Summary for non‐expert
Photosynthesis is unambiguously a fundamental process that supports almost all life forms on Earth. During photosynthesis, green plants produce oxygen and two simple sugar molecules, fructose and glucose, from carbon dioxide and water. These simple sugars are used to form various types of insoluble carbohydrates, such as cellulose or starch, or used directly to provide energy to the cells. Starch is one of the insoluble natural polymer found abundantly in plant storage organs, such as those of cereals (corn, bean), roots (cassava, sweet potato), tubers (potato), fruits (banana, mango), and pith (sago). Starch has found its way into many commercial applications, including food, pharmaceutical, and non‐food uses. In recent years, many efforts have been dedicated to the conversion of starch into ethanol as biofuel and starting material for various chemicals.
Chemically, starch comprises two glucose polymers: amylose and amylopectin, deposited as alternating amorphous and crystalline layers in the form of granules. Because of its partial crystalline structure, starch granules are insoluble in water at moderate temperatures (below the melting temperature of the starch) and difficult to degrade enzymatically (Chapter 1). But when starch is heated in the presence of an excessive amount of water, known as cooking or gelatinization, starch granules swell and amylose leaches out, improving the availability of starch for enzymatic degradation.
In Nature, the primary degradation of starch material is performed by enzymes, Page | 204 known as α‐amylases. They catalyze the breakdown of the glucosidic bonds between the polymer units in starch releasing small oligosaccharides and glucose. The interaction of α‐amylases with starch granules is severely limited, due to the low accessibility of the enzyme to the substrate. The enzymatic susceptibility of starch granules has been the subject of many investigations over the past decades. Differences in the resistance to enzymatic degradation can be attributed to the physical and chemical characteristics of starch, including granule size and shape, type of crystallinity, and surface porosity (Chapter 1). Understanding how these factors influence the accessibility of starch to α‐amylases is an important key for
Summary for non‐expert
utilization of starch in industrial applications and for starch digestion by humans and animals. This PhD thesis addresses some of these aspects by investigating the enzymatic degradation of various insoluble starches by several types of microbial enzymes. The action of α‐amylases on different types of starch granules was analyzed by determining the extent of the degradation of these starches in their native insoluble state by spectrophotometrically monitoring the concentration of products released, identifying products using chromatography, and observing the granule degradation by Scanning Electron Microscopy (SEM).
The extent of degradation of several tropical starches was compared to that of corn starch by estimating the remaining starch content and the increase in the amount of glucose (Chapter 2). Using a commercial blend of α‐amylase and glucoamylase, StargenTM 002, more than 70% degradation of sago and sweet potato starch granules was observed. This was comparable to degradation of corn starch (75%). In contrast, arrowroot, edible canna, and potato starch granules were only degraded for 25% or less. The ability of StargenTM 002 to degrade starch granules was confirmed by observing the residue left after enzymatic degradation using SEM. The SEM images showed extensive erosion of the treated corn and sago starch granules. In spite of the larger number of plants able to produce starch, at present, the major commercial starches for industrial processing are obtained from seeds, particularly corn, wheat, and rice, and from tubers and roots (potato and cassava). The results of Chapter 2 show that sago and sweet potato starch can be used as raw materials for a sustainable glucose production and as alternative substrates in the ethanol fermentation. Page | 205
Bacteria isolated from starch rich materials can be promising sources of new amylases. Amylase producing microorganism have been isolated from places such as soil around mills, markets, cassava farms, and processing factories. During this study, a potato starch‐degrading bacterium was isolated from a starch‐rich environment, namely the waste water treatment plant of a potato processing factory (Chapter 3). The new isolate was identified as Microbacterium aurum and was
Summary for non‐expert
designated M. aurum B8.A. An enzyme with α‐amylase activity was partially purified from M. aurum B8.A cultures (Chapter 3). This enzyme was able to degrade various insoluble starches including wheat and potato starch.
A further study focused on the cloning, characterization, and the architecture of the M. aurum B8.A α‐amylase as well as its contribution to starch degradation (Chapter 4). Analysis of the amino acid composition of the M. aurum B8.A α‐amylase, MaAmyA, demonstrated that the protein is unusually large and contains starch‐ binding domains (CBM25), fibronectin‐like domains (further details can be found in Chapters 1 and 4), and also possesses an extension of 300 amino acid residues at the C‐terminus, which does not possess significant homology to any known α‐amylase domain. Whether this domain functions as an extension of the binding domain or is involved in starch degradation is yet to be understood. The importance of the CBM25 domains was studied by making mutations that removed the domain at C‐terminus, resulting in a loss of enzyme activity to raw starch.
Since starch is absent in the marine environment, the presence of α‐amylases in bacteria living in this niche is rather surprising. Moreover, the fact that α‐amylases are constitutively synthesized by some marine bacteria suggests that these enzymes probably have an important metabolic role in these organisms. Out of twenty bacterial isolates from Kakaban landlocked marine lake, Indonesia, a total of eight bacterial cultures showed significant starch‐degrading activity (Chapter 5). Page | 206 Therefore, these isolates were selected for further analysis. It was found that the isolate Bacillus megaterium NL3 displayed the highest α‐amylase activity. It was also found that the novel B. megaterium NL3 α‐amylase, hereafter BmaN1, exhibited special features that are found in five unclassified homologous α‐amylases (Chapter 5). Based upon comparisons of the amino acid sequence of α‐amylases available to date and homology protein model analysis, BmaN1 was found to lack one of the conserved aspartic acid residues involved in catalysis. Naturally occurring α‐amylases which lack one of the three catalytic residues are usually not enzymatically active. However, our results showed that BmaN1 is still active in
Summary for non‐expert
breaking down starch to glucose and oligosaccharides (Chapter 5). This finding strongly suggest that BmaN1 and its homologs represent a new type of α‐amylase, of which further studies are warranted, including mutagenesis analysis and crystallization of the protein for structure‐function studies.
In addition to identifying new types of α‐amylases, the research presented in this PhD studies revealed the distribution of extracellular enzymes in different environment samples, as well as an evaluation of the biotechnological potential of the bacteria present in Nature.
Page | 207
Summary for non‐expert
Page | 208
Samenvatting
voor de leek
Samenvatting
Fotosynthese is zonder twijfel één van de meest fundamentele processen welke het leven op Aarde voedt. Groene planten maken tijdens fotosynthese zuurstof en twee eenvoudige suikers, fructose en glucose, uit water en kooldioxide. Deze eenvoudige suikers worden of direct gebruikt voor het genereren van energie of worden omgezet in verschillende soorten onoplosbare koolhydraten zoals cellulose en zetmeel. Zetmeel is een onoplosbaar natuurlijk polymeer dat veelvuldig voorkomt in granen (mais, boon), wortels (tapioca, zoete aardappel), knollen (aardappel), fruit (banaan, mango) en merg (palm). Zetmeel wordt gebruikt in allerlei voedingsmiddelen, pillen, tabletten en een scala aan alledaagse producten zoals behangplaksel, tegellijm en verf. Veel energie en tijd is de afgelopen tien jaar gestoken in de omzetting van zetmeel in ethanol dat als biobrandstof of als basis voor allerlei chemicaliën gebruikt wordt.
Vanuit een chemisch perspectief bestaat zetmeel uit twee glucose polymeren: amylose en amylopectine. Deze twee polymeren worden gedeponeerd in een korrel in de vorm van alternerende amorfe en kristallijne delen. De kristallijne structuur zorgt ervoor dat zetmeel korrels bij gematigde temperaturen niet oplosbaar zijn in water en daarom zijn de korrels moeilijk enzymatisch af te breken (Hoofdstuk 1). Wanneer zetmeel korrels in aanwezigheid van een overmaat aan water worden verwarmd, zwellen ze en lekt amylose uit de korrel. Op deze manier wordt het zetmeel gevoelig voor enzymatische afbraak.
In de Natuur wordt zetmeel door enzymen met de naam α‐amylase afgebroken. Deze enzymen verbreken de glucosidische binding tussen de glucose eenheden van het polymeer waarbij korte oligosachariden en glucose gevormd worden. α‐Amylasen hebben een beperkte interactie met de zetmeel korrel als gevolg van de fysische en Page | 210 chemische eigenschappen van de korrel en het zetmeel. Enzymatische afbraak van zetmeel korrels is de afgelopen tientallen jaren intensief bestudeerd. Verschillen in de afbraak van de korrel zijn het gevolg van het type en de morfologie van de korrel, de mate van kristalliniteit en de aan/af‐wezigheid van poriën en kanalen (Hoofdstuk 1). Kennis van de factoren die van invloed zijn op de toegankelijkheid
Samenvatting
van de zetmeel korrel voor α‐amylasen is van belang voor het gebruik van zetmeel in allerlei industriële toepassingen en voor het vergroten van ons inzicht in de vertering van zetmeel door mens en dier. Dit proefschrift gaat in op verschillende van deze aspecten en beschrijft de resultaten van onderzoek aan de enzymatische afbraak van verschillende typen zetmelen door verschillende soorten microbiële enzymen. De actie van α‐amylasen op verschillende typen zetmeel werd geanalyseerd door het bepalen van de mate van afbraak van deze zetmelen met behulp van spectrofotometrie, vloeistof chromatografie en scanning elektronen microscopie (SEM).
De mate waarin enkele tropische zetmelen werden afgebroken, werd vergeleken met de mate van afbraak van mais zetmeel door de hoeveelheid zetmeel welke overbleef en de hoeveelheid glucose welke werd gevormd, te bepalen (Hoofdstuk 2). Met het commercieel verkrijgbare enzym preparaat StargenTM 002, een mengsel van α‐amylase en glucoamylase, werd meer dan 70% afbraak van sago en zoete aardappel zetmeel korrels gevonden. Deze waarden waren vergelijkbaar met die van mais zetmeel korrels (75%). In tegenstelling hiermee was de mate van afbraak van pijlwortel (arrow root), eetbare canna en aardappel zetmeel korrels (25% of minder). De korrels die overbleven na enzymatische afbraak werden bestudeerd met behulp van SEM. Hieruit bleek dat met StargenTM 002 behandelde mais en sago zetmeel korrels een sterke erosie van het oppervlakte was opgetreden. Een heel beperkt aantal planten welke zetmeel produceren worden op dit moment gebruikt als grondstof voor industriële verwerking; mais, tarwe en rijst (zaden) en aardappel en tapioca (wortels en knollen). De resultaten van Hoofdstuk 2 laten zien dat ook sago en zoete aardappel zetmeel gebruikt kan worden voor de duurzame productie van glucose en daarmee ook als grondstof voor de productie van ethanol.
Page | 211 Nieuwe bacterie stammen geïsoleerd van plekken rijk aan zetmeel kunnen veelbelovende bronnen zijn van nieuwe amylasen. Amylase‐producerende micro‐ organismen zijn geïsoleerd van allerlei plekken zoals de bodem rondom graan molens, traditionele boeren markten, boerderijen en zetmeel fabrieken. In dit
Samenvatting
onderzoek werd een aardappel zetmeel‐afbrekende bacterie stam geïsoleerd uit de afvalwater zuivering van een aardappelzetmeel fabriek van AVEBE. (Hoofdstuk 3). Deze nieuwe stam bleek te behoren tot de soort Microbacterium aurum en kreeg de naam M. aurum B8.A. Een α‐amylase kon gedeeltelijk gezuiverd worden uit een kweek van M. aurum B8.A (Hoofdstuk 3). Het bleek dat dit enzym verschillende soorten zetmeelkorrels (tarwe en aardappel) goed kon afbreken.
In een vervolg onderzoek werd het gen dat codeert voor het amylase van M. aurum B8.A opgespoord en in kaart gebracht (Hoofdstuk 4). Gedetailleerde analyse van de aminozuur volgorde van het corresponderende eiwit, MaAmyA, liet zien dat dit een ongewoon groot eiwit is dat naast een typisch katalytisch domein ook nog verschillende zetmeel bindingsdomeinen (CBM25), fibronectine‐achtige domeinen alsook een 300 aminozuur groot C‐terminaal deel bevat (verdere details zijn in Hoofdstuk 1 en 4 te vinden). Het C‐terminale domein heeft geen enkele overeenkomst met bekende α‐amylase domeinen. Het is nog niet duidelijk welke rol dit C‐terminale domein speelt; het kan een verlenging zijn van het bindingsdomein maar ook net zo goed betrokken zijn bij de zetmeel afbraak. De rol van de CBM25 domeinen werd nader bestudeerd door deze domeinen te verwijderen uit het enzym. Dit leidde tot verlies van enzym activiteit (Hoofdstuk 4).
De aanwezigheid van α‐amylasen in bacteriën uit zee is verrassend omdat er geen zetmeel in zee te vinden is. Acht van de twintig bacteriële isolaten uit het Jelly Fish meer op Kakaban eiland, Indonesië, een van zee afgesloten zout meer, hadden een duidelijke zetmeel‐afbrekende activiteit (Hoofdstuk 5). Eén van deze isolaten, Bacillus megaterium NL3, welke van een zee anemoon afkomstig is, vertoonde de hoogste α‐amylase activiteit. Dit α‐amylase, aangeduid als BmaN1, heeft net als vijf Page | 212 andere Bacillus amylasen, twee afwijkende kenmerken (Hoofdstuk 5). Op basis van een vergelijking van de aminozuur volgorde van BmaN1 en de vijf andere Bacillus α‐amylasen met bekende α‐amylasen en de vergelijking van een 3D eiwit model van BmaN1 met een bekende α‐amylase eiwit 3D structuur werd geconcludeerd dat in BmaN1 één van de twee geconserveerde aspartaat residuen welke betrokken is bij
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de katalyse niet aanwezig was. Normaal zijn α‐amylasen welke één of meerdere van de drie katalytische aminozuur residuen missen enzymatisch niet actief. De resultaten in Hoofdstuk 5 van dit proefschrift laten zien dat BmaN1 in staat is om zetmeel af te breken tot glucose en oligosachariden. Ook suggereren deze resultaten dat BmaN1 en de andere vijf nauw verwante Bacillus α‐amylasen een nieuwe sub‐ familie van de α‐amylasen vertegenwoordigen. Verdere studies aan deze enzymen, inclusief mutagenese en kristallisatie van het eiwit in combinatie met 3D structuur analyse zullen meer licht moeten werpen op deze nieuwe α‐amylasen.
In dit proefschrift wordt naast de identificatie van nieuwe type α‐amylasen de verspreiding van extracellulaire amylasen in verschillende milieus alsook de biotechnologische toepassing van amylase‐producerende bacteriën beschreven.
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Acknowledgements
Alhamdulillahirrabbil’alamin
First and foremost, I would like to express my gratitude to the one and only, the al‐ Mighty Allah SWT, the Omnipotent, the Benevolent, the Sovereign, for having blessed me and giving me these valuable experiences during my PhD period.
This PhD thesis is the end of long journey, starting at Rijkuniversiteit Groningen (RuG) and then at Institut Teknologi Bandung. Then it has ending at the Rijkuniversiteit Groningen (RuG). There are a lot of memorable experiences that cannot describe with words and many people have been involved to complete this thesis. It would not have been possible without their countless help and support. Therefore, I would like to thank all those who were involved directly or indirectly.
I would like to express my hearty thanks to my supervisor and promotor Prof. Marc J. E. C. van der Maarel for his guidance, constructive suggestions and understanding throughout the past two years. I have benefited tremendously from working with and learning from you. I am extremely grateful for your patience and ability in helping to improve my English skills in writing and conversation. Thank you for translating the Dutch summary for me, I (literally) could not have done it without your helping. Thanks to my co‐promotor, Prof. Lubbert Dijkhuizen who invited me to be part of Microbial Physiology group in my first two years of my program. I sincerely thank Dr. Dessy Natalia and Dr. Zeily Nurachman for their support and encouragement, especially to Dr. Ihsanawati for the assistance and encouragement when I was in difficult times at the end of my PhD period. Thank you all for your understanding and patience in guiding me over the years.
I would like to sincerely thank to reading committee – Prof. Gert‐Jan W. Euverink, Prof. Marco W. Fraaije, and Prof. Birte Svensson‐ for their precious time and for valuable input. Your interest in evaluation of this thesis as well as valuable comments and insight are very much appreciated.
My special thanks go to members and staffs in ABBE group, both former and present, visiting and permanent, for a pleasant working atmosphere and interesting discussion on various matters. In addition to my colleagues, Marta Martinez‐Garcia and Delicia Yunita Rahman, who helped me in the Galdieria work. This is the first time I’ve been involved in microalgae work, it is a very impressive and exciting experience, thank you for discussion in between the busy times of experiments. To all others colleagues in the ABBE and PPBE groups: Martin Velthuis, Selvaraj Chinnatambi, Xue‐Wen Zhang, Spyridon Achinas, and whom their names are not mentioned here; thank you for the wonderful environment. Many thanks to the
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technical staff who keeps the lab running, Alle van Wijk and the office staff, Inge Meijerink‐Verver who was so helpful in organizing office life in Groningen.
My fellow graduate student in ABBE provided great advice and inspiring conversation, Otto Schepers. Whether it was setting up the bioreactors; counting the microalgal cells; doing assay; or cataloging equipment used, I appreciate all the help rendered. Thanks also extended to Leonardo Silva, Theresa Passath, and Thomas Riche for their valuable suggestion and comments on manuscripts and thesis preparation, your participation keeps me on track and helps a lot. I am also grateful to Marta Martinez Garcia and Fikri Zul Fahmi for being my paranymphs.
I would also like to thank several member and ex‐members of Microbial Physiology group Pieter Grijpstra, Hans Leemhuis, Marta Palomo i Reixach, and Munir Anwar. Thank you for the friendly atmosphere that helped me a lot at beginning of my PhD in Haren.
Dear Pipit, Herfita Agustiandari, you were my first Indonesian friend back when we were in Biological Centre, Haren. Thank you for all the nice moments together, all the fun in general, we shared many laughs. Thank you also for keeping the contact when we are so far away from each other. I would like to thank my friends out of the lab for their support and making life more fun. In particular I’d like to thank Syarif Riyadi and Desti Alkano, Muhammad Insanu, A’ Nandang Mufti, teh Rahmatun Nisa, mbak Puri Handayani, teh Uyung (Yunia Sribudiani), especially to Guntur Fibriansah‐Intan Widyasari (also Nadya and Azka) family who helped me a lot at the beginning of my study in Groningen. For Arramel‐Puti Syahrani, thanks for offering help for a lot of stuffs inside and outside the university.
I am blessed that I was surrounded by wonderful and special people. Special thanks to Adhyatmika and Nur Rochmah Hidayati, mas Didik Setiawan and family (mbak Roselina Wulandari, Nafi, and Najwa), Kurnia Sari Setio Putri and Raditya Iswandana for being close friends, for traveling around Europe or Netherlands, and for playing capsa (for sure… hahahaha), M. Zainal Abidin and family (Ayu Nuzulia and Aira), Erlangga H. Roesdyoko, Ali Syari’ati‐Liany Septiany who created such a great atmosphere at home (NS74 family), M. Nazmuddin and Guntur D. Putra for spending so many hours with me in the office during thesis writing, as well as in many social activities. Mbak Tiurma P. Allagan, mbak Sri Awalia Febriana, and Anisha Maharani for Pathe’s craziness (hahahaha). Genggong woy ngumpul woy, (Masyitha Ambarwati, Dina Maniar, Rasyida Noor, Yusran A. Muthahari,
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Caecilia A. Satyawan, Natasha Witto, Florence Husada, and Elisabet Keisia) and Muh. Insan Nugraha are thanked for being good friends and arranging off‐sciences activities out of office time. They all have made my stay in Groningen very special and unforgettable. I have spent amazing times and had lots of fun here during all these years. I like to thank all member of Chem‐ITB in Groningen, and mbak Rosewitha Irawaty, you helped me on my summary as well as for being good housemate while in GS38. I also thank mbak Nur Qomariyah and Pak Asmoro family for many different things.
Dear Ayu Anggraini, thank you for the friendship we have, and also for being there, for helping me to fight, to listen to my complaints, and to make me feel better. Sari Dewi Kurniasih, Sofi Siti Shofiyah and bu Fernita Puspasari, thank you for fruitful discussion. Jehan, I’m very glad to have such a nice friend, and thanks for your wise advices.
My appreciations also go to the Indonesian student community and Indonesia moslem society in Groningen (PPIG and deGromiest, respectively) for their kindness so that I never felt alone in the Netherlands. I hope I will be forgiven if I am forgetting someone.
In addition, my gratefulness to my auntie Sri Susanti and my uncle oom Ilsyafri for your valuable support, encouragement and love throughout my life.
I wish to express my deepest appreciation and sincere gratitude to most my beloved parents, Papa Sarian F. Sulaiman and Mama Sri Widayanti, your care, support, endless love, and never‐ending prayers are the reason I am here. My success is only for you. To my little brother, Fedo R. Sarian, you always support me whenever you can. Our personalities are very different but that makes our relationship colorful. Thanks for taking care of Papa and Mama, while I have been away. I love you all, without you I would never be able to finish this work!
Last but not least, special thanks goes to all of everyone who I forgot to mention, thank you for your help and support. Life is nothing without friends, wherever you are I will always remember you, InsyaAllah. At some points during my stay here, I may have been anxious to get work done and thus could have offended people without knowing. I use this medium to sincerely apologizing and beg for full forgiveness. x
List publications
This thesis is based on the following publications:
I. Sarian, Fean D., Leemhuis, Hans, Nurachman, Zeily, van der Maarel, Marc J. E. C., and Natalia, Dessy. (2016). Cold Enzymatic Hydrolysis of Indonesian Root and Tuber Starches (submitted) II. Sarian, Fean D., van der Kaaij, Rachel M., Kralj, Slavko, Wijbenga, Dirk J., Binnema, Doede, van der Maarel, Marc J. E. C., and Dijkhuizen, Lubbert. (2012). Enzymatic degradation of granular potato starch by Microbacterium aurum strain B8.A. Appl Microbiol Biotechnol 93 (2): 645‐654. III. Valk, Vincent, Eeuwema, Wieger, Sarian, Fean D., van der Kaaij, Rachel M., and Dijkhuizen, Lubbert. (2015). Degradation of granular starch by the bacterium Microbacterium aurum B8.A involves a novel modular α‐amylase enzyme system with FnIII and CBM25 domains. Appl Environ Microbiol 81 (19): 6610‐6620. IV. Sarian, Fean D., Ihsanawati, Nurachman, Zeily, van der Maarel, Marc J. E. C., Dijkhuizen, Lubbert, and Natalia, Dessy. (2016) Identification of a novel α‐ amylase from marine Bacillus megaterium NL3 representing a new GH13 subfamily (manuscript)
Additional publication and patent application not included in this thesis:
V. van der Maarel, Marc, J. E. C., Martinez‐Garcia, Marta, and Sarian, Fean D. Natural blue photopigments, method for producing them and to uses thereof as colorant. European patent application filled on 16 December 2014, EP141198287. VI. Sarian, Fean D., Rahman, Delicia Y., Scheppers, Otto, and van der Maarel, Marc J. E. C. (2016) Effects of oxygen limitation on the biosynthesis of photo pigments in the red microalgae Galdieria sulphuraria strain 074G. PLoS One
11 (2): e0148358. doi:10.1371/journal.pone.0148358
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VII. Rahman, Delicia Y., Sarian, Fean D., and van der Maarel, Marc J. E. C. (2016) Heterotrophic growth of Galdieria sulphuraria strain 074G using complex sugar and granular starches as carbon source (in preparation)
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