University of Groningen Unique features of several microbial -amylases active on soluble and native starch Sarian, Fean Davisunjaya IMPORTANT NOTE: You are advised to consult the publisher's version (publisher's PDF) if you wish to cite from it. Please check the document version below. Document Version Publisher's PDF, also known as Version of record Publication date: 2016 Link to publication in University of Groningen/UMCG research database Citation for published version (APA): Sarian, F. D. (2016). Unique features of several microbial -amylases active on soluble and native starch [Groningen]: University of Groningen Copyright Other than for strictly personal use, it is not permitted to download or to forward/distribute the text or part of it without the consent of the author(s) and/or copyright holder(s), unless the work is under an open content license (like Creative Commons). Take-down policy If you believe that this document breaches copyright please contact us providing details, and we will remove access to the work immediately and investigate your claim. Downloaded from the University of Groningen/UMCG research database (Pure): http://www.rug.nl/research/portal. For technical reasons the number of authors shown on this cover page is limited to 10 maximum. Download date: 18-06-2017 Chapter Introduction Introduction What is Starch? Starchisnexttocelluloseandchitin,themostabundantbiopolymerinnature.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,whichcanlaterbeconvertedbacktoglucoseandusedinrespirationwhen required. Starch is densely packed in microscopic particles called granules and is extremelyinsolubleinwateratroomtemperature.Thecomposition,sizeandshape ofthesegranulesareoftencharacteristicfortheplantspeciesfromwhichtheyare 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).Ithasaslightdegreeofbranchingof9‐20branchespermolecule(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 consistsofshortα(14)linkedchainswhilebranchingbyα(16)‐glycosidicbonds 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,itsstructureisoftendescribedbyaclustermodel(Fig.1.1)(4,8).Hizukuri(9) andKobayashietal.(10)haveshownthatanamylopectinmoleculecontainsseveral types of chains (A, B, and C) which differ in their chain length and are present in almostequalproportions.TheA‐chains(unbranched)aretheshortestchains,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 spanonlyonecluster,whileB2,B3,andB4chainsarelongchainsthatconnecttwo to four cluster chains in the amylopectin molecule (9). The C‐chain contains the reducingendgroupofthemolecule(11). Chapter 1 Page | 3 Fig1.1Schematicrepresentationoftheclustermodelofamylopectin.Fordefinitionof A, B1‐B2, and C, see text. Straight line, α(14) glycosidic bonds; dashed line, α(16) glycosidicbonds.AdaptedfromTesteretal.(12),withmodifications. The relative content of amylose and amylopectin varies widely, depending on the botanicalorigin(Table1.1).Theamylosecontentofregularstarchistypicallyfrom 10to35%ofthetotalstarch.High‐amylosestarchescontainapproximately35‐70% amylose,while‘waxy’(high‐amylopectin)starcheshavealmostnoamylose(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 amyloseintheamylopectin(14).Bothamyloseandamylopectinarepackedtogether in complexstructuresconsistingof crystallineandamorphous 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,smalleramountsofothercomponents,suchasproteinsandlipidsmay alsobepresentinamountsthatvarydependingonthebotanicaloriginandstarch isolationprocedure(3). Introduction Structure and properties of starch Nativestarchesaresemi‐crystallinestructures.Thedegreeofcrystallinityofnative Page | 4 starchvariesbetween15–45%dependingonthestarchsource(6,17).Observedby 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 crystallinelayersishighlyconservedat9nm(21–23)(Fig.1.1).Thestarchgranule organizationisverycomplexanddependsstronglyonthebotanicalorigin. A schematic representation of the granule’s architecture is given in Fig. 1.2. The outerbranchesoftheamylopectinmoleculeformdoublehelicesthatarearrangedin 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 predominantlylocatedintheamorphousgrowthringbasedontheirinvestigationof 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 diffractionpatternsforstarches,theA,B,andCpattern,thatexplainthedifference 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 fromeachotherbythedoublehelixpackingarrangementintheunitcellsaswellas 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 structureswithhexagonalchannelarrangementthatarefilledwithwatermolecules andcontainlongeraveragebranchchainlengths(26,27).Someoftherhizomeand 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 thechainlength(26,28,29),otherfactorsareknowntoinfluencethecrystallinity typeofstarches:temperature(30),concentrationofstarchsolution,salts(31),and organicmolecules(32). Page | 5 Amorphous Crystalline 9 nm Amorphous Crystalline Amorphous Granule diameter: 0.5 – 110 µm Growth ring diameter: 120 – 400 nm 9 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 crystallinelamellae.Crystallinelamellaeareseparatedbyamorphouslamellae,givingriseto semi‐crystallinezones.Crystalline‐amorphousrepeats,withaperiodicityof9nm,arecalled ‘growth rings’ and are organized into spherical blocklets to form granules. Adapted from Buleonetal.(6),withmodifications. 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), differenttypesofstems(sago),rootsandtubers(cassava,potato,sweetpotato),and even pollen (39, 40). Starches isolated from different botanical sources display a characteristic granule morphology (41). The diameter of starch granules varies dependingonthebiologicalorigin,andrangesfromsubmicron,suchassoybean,to more than 100 µm for edible canna and potato (Table 1.1). In addition, the shape Introduction rangesfromspheres,lenticulars(disks),irregulartubules,polygonals,toovals(42). Starchgranulesoccurindividuallyorasclusters,andnotwogranulesareidentical (5,43).Potatostarchgranuleshavebeenobservedtobebothovalandsphericalin Page | 6 shape(15).Cornstarchesaresphericalandpolygonalinshape(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 havealso beenfound(45).Thecharacteristicsattributedtoeachstarcharepresentedindetail inTable1.1. Fig1.3DoublehelixpackingarrangementintheA‐(left)andB‐type(right)ofstarch. MixturesofAandBaredesignatedC‐type.Eachcirclerepresentsaviewdownthez‐axisofa double helix. In the structure of A‐type starch, for each unit cell, four water molecules (diamondshape)arelocatedbetweenthehelices.AdaptedfromSarkoetal.(25)andBuleon etal.(46),withmodifications. 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) A 3–23(49) Spherical/ Polygonal Rice Cereal 1–8(50) Polygonal A 3–10(50) Wheat Cereal 18–30(51) Spherical A 22–26(51) Ediblecanna Root 28–38(17, Spherical B 10–101(41, 49) Potato Root 18–28(54, 49,52,53) Oval/ 55) Spherical Irregular Banana Fruit 16–40(36) Cassava Root 17–33 Lenticular/ B 15–110(15, 54,55) A,B,orC 20–60(36) AorC 7–23(47) Spherical Bean Legume 23–39(56, Oval C 57) Chickpea Legume 28–40(58) 8 – 55 (56, 57) Oval/ C 5–35(58) Spherical Smoothpea Legume 33–49(59) Oval C 2–40(59) Soybean Legume 12–16(37) C 0.7–4(37) Spherical/ Polygonal Sweetpotato 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. Leaf 4–34(62) Lentilcular n.d. 0.2–0.3(62) (tobacco) Leaves (Arabidopsis) n.d.,nodata;(number),reference Introduction A common feature of all starch granules is that they cannot be degraded directly, theymustbehydrolyzedfirstandtheenzymesinvolvedintheirdegradationprocess requirespecificentrypointsonthestructureofgranules,intheformofeitherpores, Page | 8 channels, or zones of attack (63, 64). Some granules as observed by Scanning ElectronMicroscopy(SEM)andAtomicForceMicroscopy(AFM)werefoundtohave manypores,othersafew,andsomenone(65).Poreswereobservedonthesurface ofsomecorn,sorghum,andmilletstarchgranules.Largegranulesofwheat,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,ediblecanna,orrice,whilethereismuchdisagreementastothe 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 processingwhenamylosemigratesfromthegranule’sinteriortotheouterlayerand surface(70).Otherporesareformedasaresultofmechanicaldamageorgranule’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 enzymeaccesstothegranuleinterior(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 photosyntheticallyactiveleaves,andthelong‐termstoragestarchinamyloplastsof 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,thestarchisdegradedtoprovideacontinuedsupplyofcarbohydrateforleaf Chapter 1 cell metabolism and also to provide substrate for sucrose synthesis to allow continuedexporttonon‐photosyntheticpartsoftheplants(74).Instorageorgans, 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 mostcultivars(76,77). In-vivo degradation of starch granules Hydrolysisofstarchtoglucoseisimportantforthemaximalutilizationofstarchto 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 degradationisbestunderstoodinleaves,wherethetransitorystarchisdegradedby respirationatnight(35,79),whilethepathwayinotherstarchcontainingtissueis 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 lightorinthedarkfor24‐48h(7,80–84).Leafstarchisconvertedtomaltodextrin 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 exportedtothecytosol,andlaterconvertedtosucrosethatisusedtoprovideenergy tothecellsortransportedtootherpartsoftheplantwhereitisconvertedtostarch forlongtermstorage(35).Inleaves,theamountofstarchaccumulatedduringthe entire photoperiod does not decrease, because during the night, the starch accumulatedduringthedayisdegradedatarelativelyconstantrate(78).Therateof degradationisregulatedinamannercomplementarytotherateofsynthesisduring theday(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 additionofphosphategroupsattheC‐6andC‐3positionsofasmallportionofthe 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 starchgranulesmaydisruptthepackingoftheamylopectinatthegranulessurface in such a way as to allow it to become degraded by exo‐amylolytic enzymes (87). α‐Amylase(EC3.2.1.1)hasalwaysbeenconsideredtobeinvolvedintheconversion oftransitorystarchtomaltodextrin,butrecentdataindicatethatα‐amylaseismost likely not required for transitory starch degradation in Arabidopsis (Arabidopsis thaliana)leaves,asevidenceforα‐amylaseinvolvementinleafstarchcatabolismis lacking(79,88).Currently,itisconsideredthatthehydrolysisofleafstarchgranules is catalyzed mainly by β‐amylase and debranching enzyme (isoamylase 3, EC 3.2.1.68).TheArabidopsisgenomecontainsninegenesencodingβ‐amylaseinwhich four of them are localized in the chloroplast (89, 90). Maltose and glucose accumulateduringstarchdegradation,andareexportedfromthechloroplasttothe cytosol.Eitherforsucrosesynthesis,glycolysis,ortheoxidativepentosephosphate pathway,conversionoftransportedmaltoseandglucoseoccurslaterinthecytosol (79). Duringgerminationofcereals,thesolubilisationofstarchisinitiatedbytheactionof 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 granulesindeadamyloplastcells.Inotherstarch‐storingorganssuchastubersand fruitsthereisnoclearevidenceforinitialattackonthestarchgranulebyα‐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 withα‐amylaseduringdegradationofbarleygrainandperhapspeaseedstarch.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),andα‐glucosidase.Theglucoseproducedintheendosperm 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 environmentalconditionsregulatethestarchdegradationprocesses.Asanexample changesinthelevelofgibberellins,agrowthhormone,promotesstarchdegradation Page | 11 bystimulatingtheproductionofα‐amylaseandotherhydrolyzingenzymesneeded for providing soluble sugars into the endosperm during germination and establishment of cereal seedling (95). The pathway of starch degradation is summarizedinFig.1.4. Starch granules Initial attack GWD and PWD α‐amylase Soluble branched glucans Debranching α‐amylase,β‐amylase, isoamylase,pullulanase Soluble linear glucans Degradation of linear chains α‐glucosidase 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,phosphoglucanwaterdikinase. The other typical storage starch is found in tubers. Starch degradation in tubers occursasaresultofstressduringsprouting(96,97).Inaddition,theconversionof starchintosugarscanalsobeinitiatedbycold‐inducedsweetening.Temporarycold sweetening process may be partially reversed since sugar levels decline by reconditioningofthetuberstohighertemperatureatsuchperiods(98,99).Aseries Introduction ofstarch‐degradingenzymeactivitieshavebeendetectedinpotatotubers,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 endospermhaveabundantsurfaceporesthataretheentrancesofchannelsleading to the interior of the granule (63, 66). During enzymatic degradation, pits in the cereal starch granulesbecome enlarged anddeeper. Internal material surrounding thechannelsiserodedandsubsequentlymoreofthegranularsurfaceisattacked.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 fromthatincerealendosperm(100). In order to maintain the cellular integrity and homeostasis, non‐photosynthetic organismshavetotakeuporganicmaterials,suchasstarch.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 starchdegradationissimilar,itdiffersinseveralaspectsfromthatofplants. Industrial starch conversion Among carbohydrate polymers, not only as the main source of food for human, starchisthemostimportantrawmaterialthatiswidelyusedinrangeofafoodand non‐food products. Starch is conventionally used in food as a thickener, colloidal stabilizer, gelling agent, encapsulation, coating agent, and water retention agent (106).Someofthenon‐foodapplicationsincludetheuseinpapermanufacturingfor surface sizing or in the textile industry as thickening and warp sizing agent (107– 109).Besidetheuseofstarchdirectlyasfoodsourceorinindustrialproducts,itcan 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 convertstarchintoglucoseandothersugarsisbasedeitheronacidhydrolysis(112) orenzymaticcatalyzedhydrolysis.Theadvantageofenzymatictreatmentisahigh Page | 13 yieldofsugarsandminimalformationofby‐products(113). Enzymatichydrolysisofstarchinvolvesthreesteps:gelatinizationofa15–35%(by weight)slurrystarchwithsteam(114–116),followedbyliquefactionbyα‐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 relativelyhighamountsofenergy.Toreducetheenergycostforcookingofstarchy materials,theno‐cookingconversionhasbeendevelopedresultinginareductionof energy consumption by approximately 50% (117). The no‐cooking concept for starch processing, although firstly reported in the 1940’s (118), has only been establishedatlargescaleduringthelastdecade. Starch-Degrading Enzymes Because of its complex structure, starch requires an appropriate combination of enzymesforitsconversiontooligosaccharidesandsmallersugars,suchasglucose and maltose. A number of enzymes that are known to be involved in starch degradationhavebeenidentifiedandtheyarebasicallyclassifiedintofourdistinct 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 regionofthestarchpolymerinarandomfashionandproducelinearandbranched oligosaccharidesintheα‐configuration.Exo‐actingamylaseshydrolyzethesubstrate 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)‐glycosidiclinkagesatthenon‐reducingends,producingmaltoseandβ‐limit Introduction dextrins.Glucoamylaseattacksα(14)linkages,releasingglucosemoleculesinthe β‐configuration;italsoattacksα(16)bondsatbranchingpointsintheamylopectin 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 pullulanasebasedontheirsubstratespecificityandproductformation;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 maltotrioseandlinearoligosaccharides,respectively.Incontrast,isoamylaseactson α(16)branchlinkagesofamylopectinandglycogen,butitisunabletohydrolyze 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)‐ glycosidiclinkagesontheterminalglucose(122). Glucosyltransferases (EC 2.4.1.‐) constitute another group of starch‐converting enzymesthathydrolyzetheα(14)‐glycosidiclinkagesinthedonormolecule 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,whilebranchingenzymes(EC2.4.1.18)cleaveα(14)‐glycosidiclinkages 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 amylomaltasescatalyzetransglycosylationtoformalinearchainproduct(123,124). Chapter 1 Amylolyticenzymesareabundantinmicroorganisms,plants,animals,andanumber ofrawstarch‐degradingenzymeshavebeenreportedthatareabletodegraderaw (native)starch(118,125).Theserawstarch‐degradingenzymesactonstarchinits 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 ofrawstarchdegradationhavebeenreportedtobeproducedbyavarietyofliving 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)representsenzymes that directly degraderawstarch belowthegelatinizationtemperatureofstarchandincludesdetaileddescriptionof theirfamiliesandbindingmodules.Themajorityoftheknownenzymeswithstarch‐ degrading activity belong to the α‐amylase family. They are known as glycosyl hydrolases(GH).ThesestarchdegradingenzymesarefoundinvariousGHfamilies (13,14,15,31,and119)(129).GHfamiliesaredescribedinmoredetailinthenext section. Page | 16 CGTase α-glucosidase/ glucoamylase glucose cyclodextrin amylomaltase α-amylase maltose β-amylase Debranching α-limit dextrin linear oligosaccharide Maltose and β-limit dextrin Isoamylase Pullulanase II 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)‐ glycosidiclinkages;○····○:α(16)‐glycosidiclinkages.AdaptedfromHorváthováetal.(119),withmodifications. 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 anotherα‐amylaseactionpattern,so‐calledpreferredattackaction(134).Likeinthe randomattackaction,onebondishydrolyzedpereffectiveencounter.However,this modeofactionpresumesthatbondsneartothechainendsandthebranchingpoints are less susceptible to hydrolysis (134, 135). The multi attack action is generally acceptedtoexplainthedifferencesinactionpatternofα‐amylases(136). Classification of α-amylases In 1991, Bernard Henrissat proposed a new classification system for glycoside hydrolases(GH)(137).Thisclassificationsystemisbaseduponaminoacidsequence andstructuralsimilaritiesratherthansubstratespecificityandcanbeusedtoreveal 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 subfamilieswithareliablepredictionofthesubstratespecificity(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 relatedoligo‐andpolysaccharides,includingenzymesspecificallyactingonα(11)‐ Page | 18 linkages(trehalose),arefoundwithinthefamilyGH13,currentlybyfarthelargest family of glycoside hydrolases, including more than 2000 representatives (140). ThisGH13familybelongstoclanGH‐HwhichcontainsalsofamiliesGH70andGH77, 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 arethought to sharea 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 betweenthesequencesandthepropertiesoftheenzymes.Fueledbytheimportance 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). ThefamilyGH13wassubdividedintosubfamiliesinordertoestablishrobustgroups 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 identityinsequence,structure,specificity,and/oreventaxonomyofthehostspecies (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 theN‐terminalendoftheprotein.Itconsistsofahighlyconserved(β/α)8‐ orTIM‐ barrelandcanberecognizedby4‐7conservedaminoacidregionscontainingthree Page | 19 activesiteresiduesinvolvedincatalysisandsubstratebinding,whicharebelieved torepresentacommonevolutionaryorigin(142,145).The(β/α)8 barrelconsistsof eightinnerparallelβ‐strandswhicharesurroundedbytheoutercylindercomposed of eight α‐helices so that the individual β‐strands and α‐helices alternate and are connectedbyloops(146).This(β/α)8 motifisrecognizedinatleastninefamiliesof glycosidehydrolases(147).Althoughthereisonly10%identitybetweentheamino 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,threeadditionalconservedregionswerelateridentifiedandwereshown tobeconnectedtoenzymespecificity(140,149).TheB‐domainisanirregularloop 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,itfoldsintoarandomcoilwithoutdefinedsecondarystructureelements (152). The active site is located in a cleft between domains A and B where a triad of catalyticresiduesperformscatalysis.Theglutamateresidueinvolved,corresponding toGlu230ofTAA,actsasgeneralacid/basecatalyst,performingprotondonationto 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 thecatalyticnucleophile,leadingtotheformationofacovalentintermediate,while the other aspartate residue (Asp297 in TAA) stabilizes the transition‐state (144, 150).TheC‐domain,whichformsaGreekkeymotif,isthedomainwiththelowest Introduction degreeofconservedsequenceintheGH13family.Ithasaβ‐sheetstructureandis linkedtotheA‐domainbyasimpleloop.Thetypeandsourceoftheα‐amylasewill affect the relative orientation between domain C and domain A (147, 153). The Page | 20 functionoftheC‐domainislargelyunknown,althoughvariousstudiesdemonstrate thatitisnecessaryforcatalyticactivityandsubstratebinding(154–156). B domain A domain C domain Fig 1.6 Ribbon representation of the overall structure of TAKA α‐amylase A, α‐amylasefromAspergillusoryzae(1TAA).DomainsA,B,andCarerepresentedasgreen, blue, and red, respectively. The triad of catalytic residues is marked as blue sticks. The structuremodelwasbuiltusingtheMacPyMOLprogram(157). Anewfamily,GH57,containingα‐amylaseswasestablishedin1996(158)afterno sequencesimilaritycouldbedetectedbetweentwosupposedα‐amylasesfromthe 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 familyGH57representsasecondandsmallerα‐amylasefamily(129).FamilyGH57 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 branchingenzyme(164),andα‐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 GH57memberssharea(β/α)7‐barrelcatalyticdomain,comprisedofsevenparallel β–strandsarrangedinabarrelencircledbysevenα‐helices.Structuralmodelsand 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.FiveconservedsequenceregionsproposedbyZonaetal.in2004can 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 recentlyitwasshownthatGH119isverycloselyrelatedinsequencetofamilyGH57 (129,142).ThefamilyGH119maythusbeconsideredasthethirdCAZyα‐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 catalyticdomainandcatalyticmachinerywithfamilyGH57(142). Introduction Physicochemical properties of α-amylases Many α‐amylases have been characterized in some detail. Table S1.2 presents a selectionofthemostextensivelystudiedα‐amylases.Mostofthemareextracellular 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 massesare generallyintherangeof45to60kDa(172–174),butsmallerorbiggerα‐amylases have also been reported. Ratanakhanokchai et al. (1992) reported that the α‐amylaseofChloroflexusaurantiacushasamolecularmassofabout210kDa(175), while the smallest α‐amylase so far is from Bacillus caldolyticus with a molecular massof10kDa(176). Mostoftheα‐amylasesarecalciummetalloenzymes,inwhichcalciumion(Ca2+)is requiredfortheiractivity,structuralintegrity,orstability(177,178).Theaffinityfor Ca2+ismuchstrongerthanthatforotherions,althoughcalciumcanbereplacedby otherdivalentmetalssuchasmagnesium,barium,andstrontiumwithoutmajorloss in activity (179, 180). The number of bound calcium atoms may vary from one to aboutten(181,182).TAAcontains10Ca2+ionsbutonlyoneatomofcalciumisvery tightly bound to the protein, while the other nine may be easily removed without anyeffectonthephysicalorbiologicalpropertiesoftheprotein(183).Mostofthe known α‐amylase structures share a common calcium‐binding site, located at the interfacebetweendomainsAandBandincloseproximitytotheactivesite(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) andeveninhibitionoftheenzymebyCa2+hasbeendocumented(189).Manymetal cations,especially heavy metals such as copper, mercury, silver, and lead, strongly inhibitα‐amylaseactivity(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,whileinmanyα‐amylasesahistidine(His)residueisinvolvedinsubstrate 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‐amylaseofBacillussubtilis(190,191),barley(192),humansaliva(193),aswellas 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.TheroleofthesecondAspresiduehasbeensuggestedtobeinvolvedin stabilizingtheoxocarbeniumion‐liketransitionstateandalsoinmaintainingtheGlu inthecorrectstateofprotonationforactivity(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). Membersoftheα‐amylasefamilyemploytheα‐retainingmechanism.Theprocessof hydrolysisofα‐glycosidiclinkagesbyaretainingmechanismisshowninFig.1.7.In theunboundstate,theun‐ionizedGlu230(TAAnumbering)ishydrogenbondedto thesidechaincarboxylofAsp297.Whenthesubstratebindstotheactivesiteofthe enzyme, the water molecules are displaced from the active site of the enzyme and thesubstratebreakstheoriginalnetworkofhydrogenbonds.Theglycosidicoxygen of the substrate interacts with Asp297 and points with its H1 toward the Asp206 (bottomsideofthecleft).Thenewhydrogenbondsandthehydrophobicinteraction betweenthesubstrateandthesubstrate‐bindingsitesareregeneratedandweaken the hydrogen bond between Glu230 and Asp297. Glu230 in the acid form easily protonatestheoxygenoftheO‐glycosidiclinkageandthereducing‐endfragmentis released from the cleft. Asp206, the catalytic nucleophile, located at the bottom of thecleftattackstheanomericcenter.Thetransitionstatepromotestheformationof theoxocarbeniumion‐likeintermediateandAsp206contributestothestabilityof the intermediate. The ionization of Glu230 produces electrostatic repulsive forces Introduction betweenGlu230andAsp297(197).Anorderedwatermoleculeispositionedatthe relativelyoutersideoftheclefttopromoteformationofahydrogenbondbetween Glu230 and Asp297 and leading to reduction of repulsive forces in the active site. Page | 24 Glu230actsasabasebyacceptingtheprotonfromthewatermoleculetoproduce an activated hydroxyl ion. The hydroxyl ion of the water then performs a nucleophilicattackontheoxocarboniumionintermediateleadingtothehydrolysis product.Theanomericconformationismaintained,becausethenucleophilicwater moleculeislocatedontheanomericsidewhichisclosetotheprotondonorinthe position between Glu230 and Asp297. Glu230 and Asp297 are located at the relativelyoutersidefromtheanomericcenter,whileAsp206ispositioneddeeperin thecoreofthe(β/α)8‐barrel(204). ThestructuralandbiochemicalinformationforGH57ismorelimited.Todate,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,includinganalysisofthe3Dcrystalstructureoftheenzyme.IntheGHfamily members that employ a retaining mechanism, the two catalytic residues are commonlyseparatedbyapproximately5.5Ådistance.Alargerthan5.5Ådistance usuallyindicatesthattheenzymehasaninvertingmechanism.Thecrystalstructure of free and ligand‐bound TLGT showed that the distance between Glu123 and Asp214residuesinbothstatesarewithintheaveragedistancecharacteristicofthe retainingmechanism(166,205,206).Furtherevidenceforaretainingmechanismof 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 througharetainingmechanismforcleavingtheα‐glycosidiclinkage. Chapter 1 Inconclusion,enzymesofGH13,57,and119meetthefollowingcriteria: (i) act on α‐O(14)‐, (16)‐ or (11)‐glycosidic linkages and hydrolyze these to produce α‐anomeric monosaccharides or oligosaccharides or formα‐glycosidiclinkagesbytransglycosylation,oracombinationofboth activities; (ii) containbetweenfourandsevenhighlyconservedregionsintheprimary structurethatcontaintheaminoacidsthatareessentialforthecatalytic reactionandsubstrate‐bindingsite; (iii) havea(β/α)8‐or(β/α)7‐barrelconformationforthecatalyticdomain; (iv) have the invariant Asp and Glu catalytic residue corresponding to the Asp206, Glu230, and Asp297 of TAA for family GH13 or equivalent to Glu123andAsp214ofTLGTofGH57andGH119members;and (v) employaretainingreactionmechanism. Page | 25 Introduction Asp206 Asp206 δ‐ δ+ Page | 26 δ‐ δ+ Glu230 Asp297 Asp297 Asp206 Asp297 Glu230 Glu230 Asp206 Asp297 Asp206 Glu230 Asp206 δ‐ δ+ δ‐ δ+ Asp297 Glu230 Asp297 Glu230 Fig 1.7 Scheme of retaining reaction mechanism of glycoside hydrolase of family GH13,asexplainedinthetext.AdaptedfromMacGregoretal.(145),withmodifications. Chapter 1 Starch binding domains Amylolytic enzymes are often multifunctional proteins composed of distinct domains,acatalyticdomainandoneormoredomainsinvolvedinsubstratebinding or multi‐enzyme complex formation. Various amylolytic enzymes, such as 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 CAZydatabase(http://www.cazy.org/,lastupdateon20October2015).Lessthan 5% of CBMs have been characterized experimentally and the sequence identities amongfamiliesarelessthan30%(211).CBMswithaffinityforinsolublerawstarch arecommonlyreferredtoasstarchbindingdomains(SBDs)andthelocationofSBD can be both, C‐ or N‐terminal or centrally positioned within the protein (147). All SBDsareapproximately100aminoacidsandshareseveralconservedaminoacids (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 arrangedintwoβ‐sheets(213).Basedonthestructures,sevenfoldtypeshavebeen observed in CBMs families: β‐sandwich, β‐trefoil, cysteine knot, unique, oligonucleotide/oligosaccharidebinding(OB)fold,heveinfold,andhevein‐likefold (214). To date, amino acid sequence‐based classification categorized SBDs into the 10 followingCBMfamilies:20,21,25,26,34,41,45,48,58,and69(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 representativeoftheotherfamilies(Fig.8).Despitethelowsequencesimilarities,all theseSBDCBMshaveβ‐sandwichfoldswithanimmunoglobulin‐liketopology(214, Page | 27 Introduction 219).Thissuggestsacommonevolutionaryancestrytowardsstarchrecognitionby allstarchbindingdomainfamilies(210). Page | 28 The CBM20 family Thefirstandbest‐characterizedSBDofstarchdegradingenzymesbelongstofamily CBM20 (210, 220). This family currently has 852 entries: 8 archaea, 616 bacteria, 220 eukaryota, and 8unclassified. So far,the tertiary structures of14 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).Thegranularrawstarch‐bindingfunctionofCBM20s hasbeendescribed,forexample,forα‐amylasefromBacillussp.strainTS‐23(222, 223), glucoamylase from Aspergillus niger (213), CGTase from Bacillus circulans strain 251 (220, 224), maltogenic α‐amylase from Geobacillus stearothermophilus (156),andβ‐amylasefromBacilluscereusvar.mycoides(225). TheCBM20domainsare90‐130aminoacidslargeandacomprehensiveanalysisof aminoacidsequencesshowedthattherearenoabsolutelyconservedresiduesinthe family(210,220).However,analysisofalargersetofCBM20ssuggestedthatcertain residuesappeartobeinvolvedspecificallyinsubstrateinteractions.Thestructure‐ 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 crystallographystructureoftheCBM20ofBacilluscirculansstrain251CGTase(224, 227). The CBM20 SBD has an open‐sided, distorted β‐barrel structure built by severalβ‐strands(213,220).Theoveralltopologyshowseight β‐strandsarranged intotwomajorβ‐sheets(Fig.1.8A)(228).Oneisafive‐strandedantiparallelβ‐sheet, while the second β‐sheet consists of one parallel β‐strand and one antiparallel Chapter 1 β‐strandpair.TheN‐andC‐terminiarelocatedatoppositeendsofthelongestaxis 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, andactsastheinitialrecognitionofthestarchmolecule.Itcomprisestwoconserved 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 providesurface‐siteandhydrophobicstackinginteractionstothesugarrings(226). Ontheotherhand,bindingsite2ismoreextended,flexible,andissuggestedasthe 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 aromaticrings,twotyrosine(Tyr527andTyr556,A.nigerSBDnumbering)andtwo tryptophan(Trp563andTrp615,A.nigerSBDnumbering)residues(226,229).The lattertryptophanisnotstrictlyconservedinCBM20(207).TheremovalofthisSBD from A. niger glucoamylase decreased the catalytic activity toward starch granules 100‐foldoftheoriginalactivity(231).Jugeetal.(2006)showedthatfusionofthis SBDtobarleyα‐amylaseisozyme1(AMY1)resultedinasix‐foldenhancementinthe affinityandactivityforstarchgranules(232). The CBM21 family Therearecurrently624entriesfortheCBM21family,ofwhich31arefrombacteria, 592arefromeukaryotes,and1isunclassified.ThebindingsitesofCBM21arevery 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 normallyfoundattheC‐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 hydrophobicenvironmenttointeractwiththesugarrings,andservesasthemajor 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 twobindingsiteswithhighaffinityforvariousmaltooligosaccharides(rangingfrom 61.9µMformaltotrioseto5.5µMformaltoheptaose)(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‐strandsinthiscase)(Fig.1.8C)(236).TheCBM25familywasestablishedbased 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 and930(Bacillussp.strain195numbering),andHis882,whereastheHisresidueis the only residue that is conserved at the structural and sequence level (Fig. 1.8C) (237). ThestructureofthebeststudiedCBM26,thatofthemaltohexaose‐formingamylase ofBacillushaloduransstrainC‐125wassolvedbyX‐raycrystallography(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 unusualorganizationoftheSBDrelatedtoamylasesfromfamilyGH13(239).Tyr25 andTrp36areresponsibleforstackinginteractionagainstthepyranoseringsofthe maltosesugarresidues.Tyr23contributestohydrogenbondformationwithoxygen 6ofthesugarstackingagainstTyr25(237).Thestarch‐bindingfunctionofCBM26 Chapter 1 hasalsobeendemonstratedinLactobacillusα‐amylasecontainingSBDsarrangedin an unusual manner, i.e. five identical modules in L. amylovorus and four in L.plantarum(240)andL.manihotivorans(241).Inapreviousreport,itwasshown thattheL.amylovorusα‐amylasewithnotandemSBDsisincapableofbindingand 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 amylasesandcyclodextrin(CD)‐hydrolyzingenzymeshaveN‐terminalregionsthat arecomposedof120‐140aminoacidswhichhavebeenclassifiedasCBM34(129). There are nearly 1450 records in the database of CBM34, of which 17 are from archaea,1427arefrombacteria,and5areunclassified. 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,TVAI(Fig.1.8E)andTVAII(243).Thismodulewasinitiallyreferredto as ‘domain N’. The presence of two glucose molecules bound to this module, supported by functional analysis that revealed the two maltooligosaccharides bindingsites,confirmingtheroleofdomain‘N’asaCBM(244).Thecrucialamino acidresidueinvolvedinrawstarchbindinginTVAI,Trp65,hasnocounterpartin TVAIIdespitethehighsequenceidentitybetweenthetworespectivedomains.TVAI hasbeenshowntohavehighhydrolyzingandbindingcapabilityagainststarchand pullulan,whereasthepreferredsubstrateofTVAIIismorelikelytobecyclodextrins (131,245). The CBM41 family Currently,theCBM41familyhasmorethan1000entriesintheCAZydatabase,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,themodulescompriseapproximately100residues(129).Thefunctionally characterized member is an α‐1,4‐glucan‐specific CBM41 from pullulanase of the marinebacteriumThermotogamaritima(TmPul13)(246).Thefamily41CBMfrom Page | 32 T.maritimaPul13hasonlyonemaincarbohydratebindingsiteontheprotein(247) andtheaffinitiesforβ‐cyclodextrin,pullulanandamylopectinwereintherangeof 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 contributedasinglewater‐mediatedhydrogenbondtothethirdglucoseinthechain (247).Thethirdresidue,Trp73,ofCBM41isconservedonlyatthestructurallevel andhasnoequivalentintheCBM20structure(248). Recently, the N‐terminal domain of the Klebsiella pneumoniae pullulanase (KpCBM41) was identified as a CBM41, using structural comparison. KpCBM41 sharesthesametopologyandbindingsiteasTmCBM41;however,onlytwoofthree bindingsitearomaticresiduesareconserved(248). The CBM45 and CBM69 families No 3D structures have been described for any member of the CBM45 and CBM69 families.CBM45membersaremainlyattachedtoeukaryoticproteins.Theyare,for example, present as N‐terminal tandem domains in Solanum tuberosum GWD (StGWD)andArabidopsisthalianaα‐amylase(AtAMY3)andseparatedbyalinkerof approximately200and50aminoacids,respectively(249).Adetailedbioinformatics and binding analysis of the CBM45 SBD of StGWD and AtAMY3, identified two conservedtryptophanresiduesasthesubstrate‐bindingsite(249,250).Trp139and Trp192 (StGWD numbering), appear to be essential for starch binding (249). An interestingobservationconcerningtheStGWDandAtAMY3SBDisthattheyhavea Chapter 1 lower‐affinitytowardscyclodextrinandstarchgranulescomparedtotypicalSBDsof microbial amylolytic enzymes, suggesting that this SBD is important in the reversibleinteractionwithstarch(duringstarchsynthesisanddegradation)(249). However,structuralelementsthatareresponsibleforthelowaffinityofCBM45in starchmetabolismhavenotyetbeendefined. TheCBM69familyhasonlyafewmembersandisexclusivelyfoundinenzymesfrom marine bacteria (215). No structural and almost no functional information is availableforthisfamily(129).ASBDwasidentifiedinanα‐amylasefromamarine metagenomiclibrary(AmyP)anddeletionofthisSBDdidnotaffecttheabilityofthe α‐amylasetodegraderawstarch.However,itwasobservedthatthedeletionmutant hadsignificantlyloweractivityforsolublestarchthanthewildtypeAmyP(215). The CBM48 family The family CBM48 is often found associated with enzymes from the GH13 family (251),exceptinthenon‐amylolyticSEX4proteinsofplantsandgreenalgae,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,andbranchingenzymes(251).TheseCBMswereoriginally 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‐bindingdomains(GBDs)insteadofCBMs,becauseseveralmembersofthis familyhavearoleinglycogenmetabolism(251).Secondaryandtertiarystructure‐ 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 commonancestralform(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 characterizedwasthatoftheGH13maltogenicamylaseofStaphylothermusmarinus Page | 33 Introduction (SMMA) (253). The crystal structure of SMMA revealed that the CBM48 domain containsalongloopthatextendsintothecatalyticsiteandparticipatesinsubstrate bindingatthecatalyticsite.OnlyonebindingsitehasbeenidentifiedinthisCBM48 Page | 34 comprisingofthreearomaticresidues,Phe95,Phe96,andTyr99(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 CBM41ofT.maritimapullulanasePul13(PDB2J71)(254).Todate,theonlycrystal 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 CBMfamilies,comprisingnineβ‐strandsorganizedintotwoopposingfour‐stranded and five‐stranded antiparallel β‐sheets and two additional parallel β‐strands that formasmallβ‐sheet(Fig.1.8H). Other Binding Sites Thecarbohydratebindingdomainshavebeenshowntobeimportantforadsorption tostarchanddegradationofstarchgranules.However,manyα‐amylaseslackingof 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 growingnumberofstructuralstudiesonvariousGHshaverevealedthepresenceof other additional non‐catalytic carbohydrates binding modules, such as fibronectin‐ likedomainsorsurfacebindingsites.ThepresenceofaSBDmaynotbethemajor requirement for degradation of raw starch, but it is still not clear why some α‐amylasescontainSBDwhereasothersdonot(147). Chapter 1 A B C Page | 35 D E G H F Fig 1.8 The structures of the SBDs from individual CBM families. The tryptophan and tyrosineresiduesinvolvedinstackinginteractionswiththesubstrateinbindingsite1are 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)CBM25ofBacillushaloduransα‐amylase,PDBcode:2C3V;(D)CBM26 ofBacillushaloduransα‐amylase,PDBcode:2C3G(237);(E)CBM34ofThermoactinomyces 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 usingtheMacPyMOLprogram(157). Introduction Fibronectin III modules Fibronectin III modules (FnIII) are one of three types of common constituents of animalplasmaproteinsthatactasalinkerandmediateprotein‐proteininteractions Page | 36 (258).SeveralanimalFnIIIstructureshavebeenelucidatedbyX‐raycrystallography andNMRspectroscopy(259,260).Theycontainapproximately90residuesandfold intoanimmunoglobulin‐likeβ‐sandwichmotif.Thesecondarystructureconsistsof 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).VerylittleinformationisavailableonthefunctionofFnIII 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.saccharolyticumNTOU1didnotalterthefunctionoftheenzyme(268).Therefore, itwassuggestedthatFnIIImaynotberequiredforthecatalyticreaction,butitmay serveas a linker between the other domains (discussed in more detail in Chapter 4). Surface Binding Sites (SBSs) Theexistenceofcarbohydratesurfacebindingsites(SBSs),sometimesreferredtoas secondary binding sites, was first described more than three decades ago for glycogenphosphorylasefromplants(269,270).SBSsoccureitheronthesurfaceof 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. AlthoughSBSsaredifferentfromCBMs,theyseemtohaveanimportantfunctionfor 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 alternativetoCBMs(273,274).ApproximatelyhalfoftheknowncarbohydrateSBSs havebeenfoundwithinseveralGH13subfamilies(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). ThecrystalstructuresofAMY1andAMY2revealedtwoSBSs,originallynamed“the starchgranule‐bindingsurfacesite”and‘thepairofsugartongs’,andnowreferred toasSBS1andSBS2,respectively(255).Thecrystalstructureandkineticproperties 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 oligosaccharidebinding(255,277).AnalysisofatripleSBSmutationwithTrp278, Trp279,andTyr380replacedbyalaninerevealedcompletelossoftheaffinityofthe resulting AMY1 mutant to starch granules and only 0.2% hydrolytic activity remainingcomparedtothewild‐typetowardsstarchgranules(255,271,278).The Trp278,Trp279,andTyr380thusappeartobethefunctionalresiduesforsubstrate binding(Fig1.9). These findings confirmed that SBSs play a variety of crucial roles to possess enzymaticactivitytowardsstarch,ascompensatingforapossiblelackofCBMs.The exactmechanisminvolvedinrawstarchdegradationstillremainslargelyunknown and none of those three residues were conserved when comparing with other subfamilies. Further studies thus are required to elucidate the presence and functionsofSBS. Introduction Page | 38 Fig1.9Representationofthecrystalstructureofbarleyα‐amylaseisozyme1(AMY1) highlightingthesurfacebindingsites.Thepictureshowsthepresenceofthe‘sugartongs’ surfacebindingsiteindifferentsurfaceregionsoftheAMY1incomplexwithmaltoheptaose (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,schematicrepresentationsareshownoftheoligosaccharidesboundatthesecondary bindingsitesofAMY1.Thediagramsofprotein‐substrateinteractionsweregeneratedusing MacPymol(157)andChemDrawbasedonXavieretal.(279). Chapter 1 Outline of this thesis Thesusceptibilityofstarchgranulestodegradationbyα‐amylasesdependsontheir botanicaloriginandontheα‐amylasesource(280).Considerableprogresshasbeen made in elucidating the nature, type, and mechanism of α‐amylases. This thesis focuses on the enzymatic degradation of starch, especially granular starch, by different types of microbial enzymes derived from several different places and conditions.Itissuggestedthatsomeofthenewlydiscoveredaspectsplayakeyrole 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)enablesthehydrolysisofnativestarchgranuleswithouttheneedfor 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 evaluatedinChapter2.Thesusceptibilityofstarchgranulesofsixdifferentplants (arrowroot,corn,ediblecanna,potato,sago,andsweetpotato)toStargenTM002was investigated.ItwasfoundthatStargenTM002degradessagoandsweetpotatoatthe samerateascornstarch,whiletheotherstarchesweremuchlessdegraded.Sweet potatoandsagothereforearerecommendedasalternativerawmaterialsourcesfor glucoseproductionbythestarchindustryintropicalregions. Chapter3describestherawstarchdegradingα‐amylasefromM.aurumstrainB8.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 Page | 39 Introduction supernatant showed several protein activity bands. The potential raw starch degradation ability ofM. aurum B8.A wasinvestigated by monitoring the extent of hydrolysisofvarioustypesofrawstarchgranules.ItturnedoutthatM.aurumB8.A Page | 40 isabletodegraderawcassava,wheat,andpotatostarch. 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‐terminaltailofMaAmyAwerestudied.DeletionoftheCBM25sresultedinalossof 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. InChapter5,characterizationofanovelα‐amylasefromBacillusmegateriumNL3, BmaN1, isolated from Kakaban landlocked marine lake in Indonesia, is presented. Basedontheaminoacidsequenceidentityandaphylogeneticanalysis,itwasfound 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),andpullulan,aswellasrawstarch. TheresultsofthisstudyaresummarizedanddiscussedinChapter6.Aperspective offutureresearchonthetopicofstarchdegradationmechanismisalsoincludedin thischapter. Chapter 1 References 1. Hermansson, A.-M., and Svegmark, K. (1996) Developments in the understanding of starch functionality. Trends Food Sci. Technol. 7, 345–353 2. Shaik, S. S., Carciofi, M., Martens, H. J., Hebelstrup, K. H., and Blennow, A. Page | 41 (2014) Starch bioengineering affects cereal grain germination and seedling establishment. J Exp Bot. 65, 2257–2270 3. Vamadevan, V., and Bertoft, E. (2015) Structure-function relationships of starch components. Starch - Stärke. 67, 55–68 4. Gallant, D. J., Bouchet, B., and Baldwin, P. M. (1997) Microscopy of starch: Evidence of a new level of granule organization. Carbohydr. Polym. 32, 177–191 5. Tester, R. F., Karkalas, J., and Qi, X. (2004) Starch - Composition, fine structure and architecture. J. Cereal Sci. 39, 151–165 6. Buléon, A., Colonna, P., Planchot, V., and Ball, S. (1998) Starch granules: Structure and biosynthesis. Int. J. Biol. Macromol. 23, 85–112 7. Streb, S., and Zeeman, S. C. (2012) Starch metabolism in Arabidopsis. Arabidopsis Book. 10, e0160 8. Robin, J. P., Mercier, C., Charbonniere, R., and Guilbot, A. (1974) Lintnerized starches. Gel filtration and enzymatic studies of insoluble residues from prolonged acid treatment of potato starch. Cereal Chem. 51, 389–405 9. Hizukuri, S. (1986) Polymodal distribution of the chain lengths of amylopectins, and its significance. Carbohydr. Res. 147, 342–347 10. Kobayashi, S., Schwartz, S. J., and Lineback, D. R. (1986) Comparison of the structures of amylopectins from different wheat varieties. Cereal Chem. 63, 71–74 11. Jane, J. (2006) Current understanding on starch granule structures. J. Appl. Glycosci. 53, 205–213 12. Tester, R. F., Karkalas, J., and Qi, X. (2004) Starch—composition, fine structure and architecture. J. Cereal Sci. 39, 151–165 13. Hoover, R., Hughes, T., Chung, H. J., and Liu, Q. (2010) Composition, molecular structure, properties, and modification of pulse starches: A review. Food Res. Int. 43, 399–413 Introduction 14. Jane, J. (2007) Structure of starch granules. J. Appl. Glycosci. 54, 31–36 15. Hoover, R. (2001) Composition, molecular structure, and physicochemical properties of tuber and root starches: A review. Carbohydr. Polym. 45, 253–267 Page | 42 16. Craig, S. A. S., Maningat, C. C., Seib, P. A., and Hoseney, R. C. (1989) Starch paste clarity. Cereal Chem. 66, 173–182 17. Zobel, H. F. (1988) Molecules to granules: A comprehensive starch review. Starch - Stärke. 40, 44–50 18. Jenkins, P. J., Cameron, R. E., and Donald, A. M. (1993) A universal feature in the structure of starch granules from different botanical sources. Starch - Stärke. 45, 417–420 19. Pilling, E., and Smith, A. M. (2003) Growth ring formation in the starch granules of potato tubers. Plant Physiol. 132, 365–371 20. Wang, S., Blazek, J., Gilbert, E., and Copeland, L. (2012) New insights on the mechanism of acid degradation of pea starch. Carbohydr. Polym. 87, 1941–1949 21. Jenkins, P. J., and Donald, a. M. (1995) The influence of amylose on starch granule structure. Int. J. Biol. Macromol. 17, 315–321 22. Oostergetel, G. T., and van Bruggen, E. F. J. (1989) On the origin of a low angle spacing in starch. Starch - Stärke. 41, 331–335 23. Oates, C. G. (1997) Towards an understanding of starch granule structure and hydrolysis. Trends Food Sci. Technol. 8, 375–382 24. Wang, T. L., Bogracheva, T. Y., Hedley, C. L., Centre, J. I., and Nr, N. (1998) Starch : As simple as A , B , C ? J. Exp. Bot. 49, 481–502 25. Sarko, A., and Wu, H.-C. H. (1978) The crystal structures of A-, B- and Cpolymorphs of amylose and starch. Starch - Stärke. 30, 73–78 26. Hizukuri, S., Kaneko, T., and Takeda, Y. (1983) Measurement of the chain length of amylopectin and its relevance to the origin of crystalline polymorphism of starch granules. Biochim. Biophys. Acta - Gen. Subj. 760, 188–191 27. Hanashiro, I., Abe, J., and Hizukuri, S. (1996) A periodic distribution of the chain length of amylopectin as revealed by high-performance anion-exchange Chapter 1 chromatography. Carbohydr. Res. 283, 151–159 28. Gidley, M. J., and Bulpin, P. V (1987) Crystallisation of malto-oligosaccharides as models of the crystalline forms of starch: Minimum chain-length requirement for the formation of double helices. Carbohydr. Res. 161, 291–300 29. Hizukuri, S. (1985) Relationship between the distribution of the chain length of amylopectin and the crystalline structure of starch granules. Carbohydr. Res. 141, 295–306 30. Fredriksson, H., Silverio, J., Andersson, R., Eliasson, A.-C., and Åman, P. (1998) The influence of amylose and amylopectin characteristics on gelatinization and retrogradation properties of different starches. Carbohydr. Polym. 35, 119–134 31. Hizukuri, S., Fujii, M., and Nikuni, Z. (1960) The effect of inorganic ions on the crystallization of amylodextrin. Biochim. Biophys. Acta. 40, 346–348 32. Gidley, M. J. (1987) Factors affecting the crystalline type (A-C) of native starches and model compounds: A rationalisation of observed effects in terms of polymorphic structures. Carbohydr. Res. 161, 301–304 33. Spoehr, H. A., and Milner, H. W. (1935) Leaf starch: Its isolation and some of its properties. J. Biol. Chem. 111, 679–687 34. Chang, C. W. (1979) Isolation and purification of leaf starch components. Plant Physiol. 64, 833–836 35. Zeeman, S. C., Smith, S. M., and Smith, A. M. (2004) The breakdown of starch in leaves. New Phytol. 163, 247–261 36. Zhang, P., Whistler, R. L., BeMiller, J. N., and Hamaker, B. R. (2005) Banana starch: Production, physicochemical properties, and digestibility—A review. Carbohydr. Polym. 59, 443–458 37. Stevenson, D. G., Doorenbos, R. K., Jane, J., and Inglett, G. E. (2006) Structures and functional properties of starch from seeds of three soybean (Glycine max (L.) Merr.) varieties. Starch - Stärke. 58, 509–519 38. Ridout, M. J., Parker, M. L., Hedley, C. L., Bogracheva, T. Y., and Morris, V. J. (2003) Atomic force microscopy of pea starch granules: Granule architecture of wild-type parent, r and rb single mutants, and the rrb double mutant. Carbohydr. Res. 338, 2135–2147 Page | 43 Introduction 39. Badorrek, P., Dick, M., Emmert, L., Schaumann, F., Koch, W., Hecker, H., Murdoch, R., Hohlfeld, J. M., and Krug, N. (2012) Pollen starch granules in bronchial inflammation. Ann. Allergy. Asthma Immunol. 109, 208–214.e6 Page | 44 40. Baker, H. G., and Baker, I. (1979) Starch in angiosperm pollen grains and its evolutionary significance. Am. J. Bot. 66, 591–600 41. Jane, J.-L., Kasemsuwan, T., Leas, S., Zobel, H., and Robyt, J. F. (1994) Anthology of starch granule morphology by scanning electron microscopy. Starch - Stärke. 46, 121–129 42. BeMiller, J. N., and Whistler, R. L. (2009) Starch: Chemistry and Technology, 3rd Ed. (BeMiller, J. N., and Whistler, R. L. eds), Food Science and Technology, Elsevier Science 43. Pérez, S., and Bertoft, E. (2010) The molecular structures of starch components and their contribution to the architecture of starch granules: A comprehensive review. Starch - Stärke. 62, 389–420 44. Dhital, S., Shrestha, A. K., and Gidley, M. J. (2010) Relationship between granule size and in vitro digestibility of maize and potato starches. Carbohydr. Polym. 82, 480–488 45. Hoover, R., and Sosulski, F. W. (1991) Composition, structure, functionality, and chemical modification of legume starches: A review. Can. J. Physiol. Pharmacol. 69, 79–92 46. Buléon, A., Gallant, D. J., Bouchet, B., Mouille, G., D’Hulst, C., Kossmann, J., and Ball, S. (1997) Starches from A to C. Chlamydomonas reinhardtii as a model microbial system to investigate the biosynthesis of the plant amylopectin crystal. Plant Physiol. 115, 949–957 47. Peroni, F. H. G., Rocha, T. S., and Franco, C. M. L. (2006) Some structural and physicochemical characteristics of tuber and root starches. Food Sci. Technol. Int. 12, 505–513 48. Morrison, W. R. R., Milligan, T. P. P., and Azudin, M. N. N. (1984) A relationship between the amylose and lipid contents of starches from diploid cereals. J. Cereal Sci. 2, 257–271 Chapter 1 49. Soni, P. L., Sharma, H., Srivastava, H. C., and Gharia, M. M. (1990) Physicochemical properties of Canna edulis starch — comparison with maize starch. Starch - Stärke. 42, 460–464 50. Wani, A. A., Singh, P., Shah, M. A., Schweiggert-Weisz, U., Gul, K., and Wani, I. A. (2012) Rice starch diversity: Effects on structural, morphological, thermal, and Page | 45 physicochemical properties—A review. Compr. Rev. Food Sci. Food Saf. 11, 417– 436 51. Soulaka, A. B., and Morrison, W. R. (1985) The amylose and lipid contents, dimensions, and gelatinisation characteristics of some wheat starches and their Aand B-granule fractions. J. Sci. Food Agric. 36, 709–718 52. Santacruz, S. (2002) Three underutilised sources of starch from the Andean region in Ecuador Part I. Physico-chemical characterisation. Carbohydr. Polym. 49, 63–70 53. Thitipraphunkul, K., Uttapap, D., Piyachomkwan, K., and Takeda, Y. (2003) A comparative study of edible canna (Canna edulis) starch from different cultivars. Part I. Chemical composition and physicochemical properties. Carbohydr. Polym. 53, 317–324 54. Kim, Y. S., Wiesenborn, D. P., Orr, P. H., and Grant, L. A. (1995) Screening potato starch for novel properties using differential scanning calorimetry. J. Food Sci. 40, 1060–1065 55. Noda, T., Tsuda, S., Mori, M., Takigawa, S., Matsuura-Endo, C., Saito, K., Arachichige Mangalika, W. H., Hanaoka, A., Suzuki, Y., and Yamauchi, H. (2004) The effect of harvest dates on the starch properties of various potato cultivars. Food Chem. 86, 119–125 56. Ovando-Martínez, M., Bello-Pérez, L. A., Whitney, K., Osorio-Díaz, P., and Simsek, S. (2011) Starch characteristics of bean (Phaseolus vulgaris L.) grown in different localities. Carbohydr. Polym. 85, 54–64 57. Lai, C. C., and Varriano-Marston, E. (1979) Studies on the charcateristics of black bean starch. J. Food Sci. 44, 528–530 58. Hughes, T., Hoover, R., Liu, Q., Donner, E., Chibbar, R., and Jaiswal, S. (2009) Composition, morphology, molecular structure, and physicochemical properties of starches from newly released chickpea (Cicer arietinum L.) cultivars grown in Introduction Canada. Food Res. Int. 42, 627–635 59. Ratnayake, W. S., Hoover, R., and Warkentin, T. (2002) Pea starch: Composition, structure and properties — a review. Starch - Stärke. 54, 217–234 Page | 46 60. Ahmad, F. B., Williams, P. A., Doublier, J.-L., Durand, S., and Buleon, A. (1999) Physico-chemical characterisation of sago starch. Carbohydr. Polym. 38, 361–370 61. Matheson, N. K. (1996) The chemical structure of amylose and amylopectin fractions of starch from tobacco leaves during development and diurnallynocturnally. Carbohydr. Res. 282, 247–262 62. Zeeman, S. C., Tiessen, A., Pilling, E., Kato, K. L., Donald, A. M., and Smith, A. M. (2002) Starch synthesis in Arabidopsis. Granule synthesis, composition, and structure. Plant Physiol. 129, 516–529 63. Huber, K. C., and BeMiller, J. N. (2000) Channels of maize and sorghum starch granules. Carbohydr. Polym. 41, 269–276 64. Tetlow, I. J. (2010) Starch biosynthesis in developing seeds. Seed Sci. Res. 21, 5– 32 65. Baldwin, P. M., Davies, M. C., and Melia, C. D. (1997) Starch granule surface imaging using low-voltage scanning electron microscopy and atomic force microscopy. Int. J. Biol. Macromol. 21, 103–107 66. Huber, K. C., and BeMiller, J. N. (1997) Visualization of channels and cavities of corn and sorghum starch granules. Cereal Chem. J. 74, 537–541 67. Baldwin, P. M., Adler, J., Davies, M. C., and Melia, C. D. (1998) High resolution imaging of starch granule surfaces by atomic force microscopy. J. Cereal Sci. 27, 255–265 68. Juszczak, L., Fortuna, T., and Krok, F. (2003) Non-contact atomic force microscopy of starch granules surface. Part II. Selected cereal starches. Starch Stärke. 55, 8–16 69. Fannon, J., Gray, J., Gunawan, N., Huber, K., and BeMiller, J. (2004) Heterogeneity of starch granules and the effect of granule channelization on starch modification. Cellulose. 11, 247–254 Chapter 1 70. Baldwin, P. M., Adler, J., Davies, M. C., and Melia, C. D. (1995) Starch damage Part 1: Characterisation of granule damage in ball-milled potato starch study by SEM. Starch - Stärke. 47, 247–251 71. Niemann, C., and Whistler, R. L. (1992) Effect of acid hydrolysis and ball milling on porous corn starch. Starch - Stärke. 44, 409–414 72. BeMiller, J. N., and Huber, K. C. (2015) Physical modification of food starch functionalities. Annu Rev Food Sci. 9, 19–69 73. Mitsui, T., Itoh, K., Hori, H., and Ito, H. (2010) Biosynthesis and degradation of starch. Bull Facul Agric Niigata Univ. 62, 49–73 74. Zeeman, S. C., Kossmann, J., and Smith, A. M. (2010) Starch: Its metabolism, evolution, and biotechnological modification in plants. Annu. Rev. Plant Biol. 61, 209–234 75. Smith, A. M., and Stitt, M. (2007) Coordination of carbon supply and plant growth. Plant. Cell Environ. 30, 1126–1149 76. Stanley, D., Farnden, K. J. F., and MacRae, E. a (2005) Plant α-amylases: Functions and roles in carbohydrate metabolism. Biologia (Bratisl). 60, 65–71 77. Simão, R. A., Silva, A. P. F. B., Peroni, F. H. G., do Nascimento, J. R. O., Louro, R. P., Lajolo, F. M., and Cordenunsi, B. R. (2008) Mango starch degradation. I. A microscopic view of the granule during ripening. J. Agric. Food Chem. 56, 7410– 7415 78. Zeeman, S. C., Delatte, T., Messerli, G., Umhang, M., Stettler, M., Mettler, T., Streb, S., Reinhold, H., and Kötting, O. (2007) Starch breakdown: Recent discoveries suggest distinct pathways and novel mechanism. Funct. Plant Biol. 34, 465–473 79. Smith, A. M., Zeeman, S. C., and Smith, S. M. (2005) Starch degradation. Annu Rev Plant Biol. 56, 73–98 80. Fondy, B. R., Geiger, D. R., and Servaites, J. C. (1989) Photosynthesis, carbohydrate metabolism, and export in Beta vulgaris L. and Phaseolus vulgaris L. during square and sinusoidal light regimes. Plant Physiol. 89, 396–402 81. Servaites, J. C., Fondy, B. R., Li, B., and Geiger, D. R. (1989) Sources of carbon for export from spinach leaves throughout the day. Plant Physiol. 90, 1168–1174 Page | 47 Introduction 82. Kakefuda, G., and Preiss, J. (1997) Partial purification and characterization of a diurnally fluctuating novel endoamylase from Arabidopsis thaliana leaves. Plant Physiol. Biochem. 35, 907–913 Page | 48 83. Graf, A., Schlereth, A., Stitt, M., and Smith, A. M. (2010) Circadian control of carbohydrate availability for growth in Arabidopsis plants at night. Proc. Natl. Acad. Sci. 107, 9458–9463 84. Appenroth, K.-J., Keresztes, A., Krzysztofowicz, E., and Gabrys, H. (2011) Lightinduced degradation of starch granules in turions of Spirodela polyrhiza studied by electron microscopy. Plant Cell Physiol. 52, 384–391 85. Kötting, O., Pusch, K., Tiessen, A., Geigenberger, P., Steup, M., and Ritte, G. (2005) Identification of a novel enzyme required for starch metabolism in Arabidopsis leaves. The phosphoglucan, water dikinase. Plant Physiol. 137, 242– 252 86. Ritte, G., Heydenreich, M., Mahlow, S., Haebel, S., Kötting, O., and Steup, M. (2006) Phosphorylation of C6- and C3-positions of glucosyl residues in starch is catalysed by distinct dikinases. FEBS Lett. 580, 4872–6 87. Ritte, G., Scharf, A., Eckermann, N., Haebel, S., and Steup, M. (2004) Phosphorylation of transitory starch is increased during degradation. Plant Physiol. 135, 2068–2077 88. Yu, T.-S., Zeeman, S. C., Thorneycroft, D., Fulton, D. C., Dunstan, H., Lue, W.-L., Hegemann, B., Tung, S.-Y., Umemoto, T., Chapple, A., Tsai, D.-L., Wang, S.-M., Smith, A. M., Chen, J., and Smith, S. M. (2005) α-Amylase is not required for breakdown of transitory starch in Arabidopsis leaves. J. Biol. Chem. 280, 9773– 9779 89. Scheidig, A., Fröhlich, A., Schulze, S., Lloyd, J. R., and Kossmann, J. (2002) Downregulation of a chloroplast-targeted β-amylase leads to a starch-excess phenotype in leaves. Plant J. 30, 581–591 90. Kaplan, F., and Guy, C. L. (2005) RNA interference of Arabidopsis β-amylase8 prevents maltose accumulation upon cold shock and increases sensitivity of PSII photochemical efficiency to freezing stress. Plant J. 44, 730–743 Chapter 1 91. Beck, E., and Ziegler, P. (1989) Biosynthesis and degradation of starch in higher plants. Annu Rev Plant Physiol Plant Mol Biol. 40, 95–117 92. Minamikawa, T., Yamauchi, D., and Wada, S. (1992) Expression of α-amylase in Phaseolus vulgaris and Vigna mungo plants. Plant Cell Physiol. 33, 253–258 93. Bernhardt, D., Trutwig, A., and Barkhold, A. (1993) Synthesis of DNA and the development of amylase and phosphatase activities in cotyledons of germinating seeds of Vaccaria pyramidata. J. Exp. Bot. 44, 695–699 94. Sun, Z., and Henson, C. A. (1990) Degradation of native starch granules by barley α-glucosidases. Plant Physiol. 94, 320–327 95. Kaneko, M., Itoh, H., Ueguchi-Tanaka, M., Ashikari, M., and Matsuoka, M. (2002) The α-amylase induction in endosperm during rice seed germination is caused by gibberellin synthesized in epithelium. Plant Physiol. 128, 1264–1270 96. Biemelt, S., Hajirezaei, M., Hentschel, E., and Sonnewald, U. (2000) Comparative analysis of abscisic acid content and starch degradation during storage of tubers harvested from different potato varieties. Potato Res. 43, 371–382 97. Davies, H. V, and Ross, H. A. (1984) The pattern of starch and protein degradation in tubers. Potato Res. 27, 373–381 98. Ohad, I., Friedberg, I., Ne’Eman, Z., and Schramm, M. (1971) Biogenesis and degradation of starch: I. The fate of the amyloplast membranes during maturation and storage of potato tubers. Plant Physiol. 47, 465–477 99. Sowokinos, J. (2001) Biochemical and molecular control of cold-induced sweetening in potatoes. Am. J. Potato Res. 78, 221–236 100. Alvani, K., Qi, X., Tester, R. F., and Snape, C. E. (2011) Physico-chemical properties of potato starches. Food Chem. 125, 958–965 101. Ao, Z., Quezada-Calvillo, R., Sim, L., Nichols, B. L., Rose, D. R., Sterchi, E. E., and Hamaker, B. R. (2007) Evidence of native starch degradation with human small intestinal maltase-glucoamylase (recombinant). FEBS Lett. 581, 2381–2388 102. Butterworth, P. J., Warren, F. J., and Ellis, P. R. (2011) Human α-amylase and starch digestion: An interesting marriage. Starch - Stärke. 63, 395–405 103. Hindle, V. A., Vuuren van, A. M., Klop, A., Mathijssen-Kamman, A. A., Van Page | 49 Introduction Gelder, A. H., and Cone, J. W. (2005) Site and extent of starch degradation in the dairy cow – a comparison between in vivo, in situ and in vitro measurements. J. Anim. Physiol. Anim. Nutr. (Berl). 89, 158–165 Page | 50 104. Anderson, K. L. (1995) Biochemical analysis of starch degradation by Ruminobacter amylophilus 70. Appl. Environ. Microbiol. 61, 1488–1491 105. Ze, X., Duncan, S. H., Louis, P., and Flint, H. J. (2012) Ruminococcus bromii is a keystone species for the degradation of resistant starch in the human colon. ISME J. 6, 1535–1543 106. Singh, J., Kaur, L., and McCarthy, O. J. (2007) Factors influencing the physicochemical, morphological, thermal and rheological properties of some chemically modified starches for food applications—A review. Food Hydrocoll. 21, 1–22 107. Radley, J. A. (1954) Starch and its derivatives, Chapman and Hall Ltd., London 108. Radley, J. A. (1976) The textile industry. in Industrial Uses of Starch and its Derivatives SE - 4 (Radley, J. A. ed), pp. 149–197, Springer Netherlands, 10.1007/978-94-010-1329-1_4 109. Aspinall, G. O. (2014) The Polysaccharides Volume 2 of Molecular Biology, revised (Aspinall, G. O. ed), Molecular Biology, Academic Press, New York 110. Miyazaki, M., Van Hung, P., Maeda, T., and Morita, N. (2006) Recent advances in application of modified starches for breadmaking. Trends Food Sci. Technol. 17, 591–599 111. Shigechi, H., Koh, J., Fujita, Y., Bito, Y., Ueda, M., Satoh, E., Matsumoto, T., Fukuda, H., Kondo, A., Bito, Y., Ueda, M., Satoh, E., Fukuda, H., and Kondo, A. (2004) Direct production of ethanol from raw corn starch via fermentation by use of a novel surface-engineered yeast strain codisplaying glucoamylase and αamylase. Appl. Environ. Microbiol. 70, 5037–5040 112. Hoover, R. (2000) Acid-treated starches. Food Rev. Int. 16, 369–392 113. Honsch, W. M. (1957) Preliminary work for the manufacture of wheat starch glucose. Starch - Stärke. 9, 45–47 114. Lee, S. H., Shetty, J. K., and Strohm, B. A. (2015) Liquefaction and saccharification of granular starch at high concentration. 1, 1–11 Chapter 1 115. Myat, L., and 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, 288–295 116. Singh, H., and Soni, S. K. (2001) Production of starch-gel digesting amyloglucosidase by Aspergillus oryzae HS-3 in solid state fermentation. Process Page | 51 Biochem. 37, 453–459 117. Matsumoto, N., Fukushi, O., Miyanaga, M., Kakihara, K., Nakajima, E., and Yoshizumi, H. (1982) Industrialization of a noncooking system for alcoholic fermentation from grains. Agric. Biol. Chem. 46, 1549–1558 118. Balls, A. K., and Schwimmer, S. (1944) Digestion of raw starch. J. Biol. Chem. 156, 203–210 119. Horvathova, V., Janecek, S., and Sturdik, E. (2000) Amylolytic enzymes: Their specificities, origins and properties. Biologia-Bratislava. 55, 605–616 120. van der Maarel, M. J., van der Veen, B., Uitdehaag, J. C., Leemhuis, H., and Dijkhuizen, L. (2002) Properties and applications of starch-converting enzymes of the α-amylase family. J. Biotechnol. 94, 137–155 121. Duffner, F., Bertoldo, C., Andersen, J. T., Wagner, K., and Antranikian, G. (2000) A new thermoactive pullulanase from Desulfurococcus mucosus: Cloning, sequencing, purification, and characterization of the recombinant enzyme after expression in Bacillus subtilis. J. Bacteriol. 182, 6331–6338 122. Singh, R. S., Saini, G. K., and Kennedy, J. F. (2008) Pullulan: Microbial sources, production and applications. Carbohydr. Polym. 73, 515–531 123. Fujii, K., Minagawa, H., Terada, Y., Takaha, T., Kuriki, T., Shimada, J., and Kaneko, H. (2005) Use of random and saturation mutageneses to improve the properties of Thermus aquaticus amylomaltase for efficient production of cycloamyloses. Appl. Environ. Microbiol. 71, 5823–5827 124. Weiß, S. C., Skerra, A., and Schiefner, A. (2015) Structural basis for the interconversion of maltodextrins by MalQ, the amylomaltase of Escherichia coli. J. Biol. Chem. 290, 21352–21364 125. Sun, H., Zhao, P., Ge, X., Xia, Y., Hao, Z., Liu, J., and Peng, M. (2010) Recent advances in microbial raw starch degrading enzymes. Appl. Biochem. Biotechnol. Introduction 160, 988–1003 126. Uthumporn, U., Zaidul, I. S. M., and Karim, A. A. (2010) Hydrolysis of granular starch at sub-gelatinization temperature using a mixture of amylolytic enzymes. Page | 52 Food Bioprod. Process. 88, 47–54 127. Cinelli, B. a., Castilho, L. R., Freire, D. M. G., and Castro, A. M. (2015) A brief review on the emerging technology of ethanol production by cold hydrolysis of raw starch. Fuel. 150, 721–729 128. El-Fallal, A., Dobara, M. A., El-Sayed, A., and Omar, N. (2012) Starch and microbial α-amylases: From concepts to biotechnological applications. Carbohydrates–Comprehensive Stud. Glycobiol. Glycotechnol. InTech 129. Cantarel, B. L., Coutinho, P. M., Rancurel, C., Bernard, T., Lombard, V., and Henrissat, B. (2009) The Carbohydrate-Active EnZymes database (CAZy): An expert resource for Glycogenomics. Nucleic Acids Res. 37, D233–238 130. Sakano, Y., Hiraiwa, S., Fukushima, J., and Kobayashi, T. (1982) Enzymatic properties and action patterns of Thermoactinomyces vulgaris α-amylase. Agric. Biol. Chem. 46, 1121–1129 131. Kamitori, S., Abe, A., Ohtaki, A., Kaji, A., Tonozuka, T., and Sakano, Y. (2002) Crystal structures and structural comparison of Thermoactinomyces vulgaris R-47 α-amylase 1 (TVAI) at 1.6 A resolution and α-amylase 2 (TVAII) at 2.3 A resolution. J. Mol. Biol. 318, 443–453 132. Azhari, R., and Lotan, N. (1991) Enzymic hydrolysis of biopolymers via singlescission attack pathways: a unified kinetic model. J. Mater. Sci. Mater. Med. 2, 9– 18 133. Robyt, J., and French, D. (1963) Action pattern and specificity of an amylase from Bacillus subtilis. Arch. Biochem. Biophys. 100, 451–467 134. Thoma, J. A. (1976) Models for deploymerizing enzymes: criteria for discrimination of models. Carbohydr. Res. 48, 85–103 135. Banks, W., and Greenwood, C. T. (1977) Mathematical models for the action of αamylase on amylose. Carbohydr. Res. 57, 301–315 136. Svensson, B., and Søgaard, M. (1993) Mutational analysis of glycosylase function. Chapter 1 J. Biotechnol. 29, 1–37 137. Henrissat, B. (1991) A classification of glycosyl hydrolases based on amino acid sequence similarities. Biochem. J. 280, 309–316 138. Henrissat, B., and Davies, G. (1997) Structural and sequence-based classification of glycoside hydrolases. Curr. Opin. Struct. Biol. 7, 637–644 139. Stam, M. R., Danchin, E. G. J., Rancurel, C., Coutinho, P. M., and Henrissat, B. (2006) Dividing the large glycoside hydrolase family 13 into subfamilies: Towards improved functional annotations of α-amylase-related proteins. Protein Eng. Des. Sel. 19, 555–562 140. Janeček, Š., Svensson, B., and MacGregor, E. A. (2014) α-Amylase: An enzyme specificity found in various families of glycoside hydrolases. Cell. Mol. Life Sci. 71, 1149–1170 141. Janeček, Š. (2002) How many conserved sequence regions are there in the αamylase family? Biologia (Bratisl). 57, 29–41 142. Janeček, Š., and Kuchtová, A. (2012) In silico identification of catalytic residues and domain fold of the family GH119 sharing the catalytic machinery with the αamylase family GH57. FEBS Lett. 586, 3360–3366 143. Janeček, Š., Svensson, B., and MacGregor, E. A. (2007) A remote but significant sequence homology between glycoside hydrolase clan GH-H and family GH31. FEBS Lett. 581, 1261–1268 144. Matsuura, Y., Kusunoki, M., Harada, W., and Kakudo, M. (1984) Structure and possible catalytic residues of Taka-amylase A. J. Biochem. 95, 697–702 145. MacGregor, E. A., Janeček, Š., and Svensson, B. (2001) Relationship of sequence and structure to specificity in the α-amylase family of enzymes. Biochim. Biophys. Acta - Protein Struct. Mol. Enzymol. 1546, 1–20 146. Sterner, R., and Höcker, B. (2005) Catalytic versatility, stability, and evolution of the (β/α)8-barrel enzyme fold. Chem. Rev. 105, 4038–4055 147. Janeček, Š., Svensson, B., and MacGregor, E. A. (2003) Relation between domain evolution, specificity, and taxonomy of the α-amylase family members containing a C-terminal starch-binding domain. Eur. J. Biochem. 270, 635–645 Page | 53 Introduction 148. Nakajima, R., Imanaka, T., and Aiba, S. (1986) Comparison of amino acid sequences of eleven different α-amylases. Appl. Microbiol. Biotechnol. 23, 355– 360 Page | 54 149. Janecek, S. (1992) New conserved amino acid region of α-amylases in the third loop of their (β/α)8-barrel domains. Biochem. J. 288, 1069–1070 150. Kuriki, T., and Imanaka, T. (1999) The concept of the α-amylase family: Structural similarity and common catalytic mechanism. J. Biosci. Bioeng. 87, 557–565 151. Machius, M., Wiegand, G., and Huber, R. (1995) Crystal structure of calciumdepleted Bacillus licheniformis α-amylase at 2.2 Å resolution. J. Mol. Biol. 246, 545–559 152. Kadziola, A., Abe, J., Svensson, B., and Haser, R. (1994) Crystal and molecular structure of barley α-amylase. J. Mol. Biol. 239, 104–121 153. MacGregor, E. A. (1988) α-Amylase structure and activity. J. Protein Chem. 7, 399–415 154. Holm, L., Koivula, A. K., Lehtovaara, P. M., Hemminki, A., and Knowles, J. K. C. (1990) Random mutagenesis used to probe the structure and function of Bacillus stearothermophilus α-amylase. Protein Eng. 3, 181–191 155. Vihinen, M., Peltonen, T., Iitiä, A., Suominen, I., and Mäntsälä, P. (1994) Cterminal truncations of a thermostable Bacillus stearothermophilus α-amylase. Protein Eng. 7, 1255–1259 156. Dauter, Z., Dauter, M., Brzozowski, A. M., Christensen, S., Borchert, T. V, Beier, L., Wilson, K. S., and Davies, G. J. (1999) X-ray structure of novamyl, the fivedomain “maltogenic” α-amylase from Bacillus stearothermophilus: Maltose and acarbose complexes at 1.7 Å resolution. Biochemistry. 38, 8385–8392 157. Schrodinger LLC (2010) The PyMOL molecular graphics system, Version 1.3r1 158. Henrissat, B., and Bairoch, A. (1996) Updating the sequence-based classification of glycosyl hydrolases. Biochem. J. 316, 695–696 159. Nakajima, M., Imamura, H., Shoun, H., Horinouchi, S., and Wakagi, T. (2004) Transglycosylation activity of Dictyoglomus thermophilum amylase A. Biosci. Biotechnol. Biochem. 68, 2369–2373 Chapter 1 160. Jeon, E.-J., Jung, J.-H., Seo, D.-H., Jung, D.-H., Holden, J. F., and Park, C.-S. (2014) Bioinformatic and biochemical analysis of a novel maltose-forming αamylase of the GH57 family in the hyperthermophilic archaeon Thermococcus sp. CL1. Enzyme Microb. Technol. 60, 9–15 161. Imamura, H., Fushinobu, S., Jeon, B.-S., Wakagi, T., and Matsuzawa, H. (2001) Page | 55 Identification of the catalytic residue of Thermococcus litoralis 4-α- glucanotransferase through mechanism-based labeling. Biochemistry. 40, 12400– 12406 162. Zona, R., Chang-Pi-Hin, F., O’Donohue, M. J., and Janecek, S. (2004) Bioinformatics of the glycoside hydrolase family 57 and identification of catalytic residues in amylopullulanase from Thermococcus hydrothermalis. Eur. J. Biochem. 271, 2863–2872 163. Kang, S., Vieille, C., and Zeikus, J. G. (2005) Identification of Pyrococcus furiosus amylopullulanase catalytic residues. Appl. Microbiol. Biotechnol. 66, 408–413 164. Palomo, M., Pijning, T., Booiman, T., Dobruchowska, J. M., van der Vlist, J., Kralj, S., Planas, A., Loos, K., Kamerling, J. P., Dijkstra, B. W., van der Maarel, M. J. E. C., Dijkhuizen, L., and Leemhuis, H. (2011) Thermus thermophilus glycoside hydrolase family 57 branching enzyme: Crystal structure, mechanism of action, and products formed. J. Biol. Chem. 286, 3520–3530 165. Lieshout, J. F. T. van, Verhees, C. H., Ettema, T. J. G., Sar, S. van der, Imamura, H., Matsuzawa, H., Oost, J. van der, and Vos, W. M. de (2009) Identification and molecular characterization of a novel type of α-galactosidase from Pyrococcus furiosus. Biocatal. [online] Biotransformation. http://www.tandfonline.com/doi/abs/10.1080/10242420310001614342 (Accessed November 24, 2015) 166. Imamura, H., Fushinobu, S., Yamamoto, M., Kumasaka, T., Jeon, B.-S., Wakagi, T., and Matsuzawa, H. (2003) Crystal structures of 4-α-glucanotransferase from Thermococcus litoralis and its complex with an inhibitor. J. Biol. Chem. 278, 19378–19386 167. Watanabe, H., Nishimoto, T., Kubota, M., Chaen, H., and Fukuda, S. (2006) Cloning, sequencing, and expression of the genes encoding an isocyclomaltooligosaccharide glucanotransferase and an α-amylase from a Bacillus Introduction circulans strain. Biosci. Biotechnol. Biochem. 70, 2690–2702 168. Raha, M., Kawagishi, I., Müller, V., Kihara, M., and Macnab, R. M. (1992) Escherichia coli produces a cytoplasmic α-amylase, AmyA. J. Bacteriol. 174, Page | 56 6644–6652 169. Satoh, E., Uchimura, T., Kudo, T., and Komagata, K. (1997) Purification, characterization, and nucleotide sequence of an intracellular maltotriose-producing α-amylase from Streptococcus bovis 148. Appl. Environ. Microbiol. 63, 4941–4944 170. Lim, W. J., Park, S. R., An, C. L., Lee, J. Y., Hong, S. Y., Shin, E. C., Kim, E. J., Kim, J. O., Kim, H., and Yun, H. D. (2003) Cloning and characterization of a thermostable intracellular α-amylase gene from the hyperthermophilic bacterium Thermotoga maritima MSB8. Res. Microbiol. 154, 681–687 171. Onodera, M., Yatsunami, R., Tsukimura, W., Fukui, T., Nakasone, K., Takashina, T., and Nakamura, S. (2013) Gene analysis, expression, and characterization of an intracellular α-amylase from the extremely halophilicarchaeon Haloarcula japonica. Biosci. Biotechnol. Biochem. 77, 281–288 172. Vihinen, M., and Mantsiila, P. (1989) Microbial amylolytic enzyme. Crit. Rev. Biochem. Mol. Biol. 24, 329–418 173. Yamamoto, T. (1994) Enzyme chemistry and molecular biology of amylases and related enzymes (Yamamoto, T. ed), CRC Press 174. Gupta, R., Gigras, P., Mohapatra, H., Goswami, V. K., and Chauhan, B. (2003) Microbial α-amylases: A biotechnological perspective. Process Biochem. 38, 1599–1616 175. Ratanakhanokchai, K., Kaneko, J., Kamio, Y., and Izaki, K. (1992) Purification and properties of a maltotetraose- and maltotriose-producing amylase from Chloroflexus aurantiacus. Appl. Environ. Microbiol. 58, 2490–2494 176. Grootegoed, J. A., Lauwers, A. M., and Heinen, W. (1973) Separation and partial purification of extracellular amylase and protease from Bacillus caldolyticus. Arch. Mikrobiol. 90, 223–232 177. Vallee, B. L., Stein, E. A., Sumerwell, W. N., and Fischer, E. H. (1959) Metal content of α-amylases of various origins. J. Biol. Chem. 234, 2901–2905 Chapter 1 178. Kandra, L. (2003) α-Amylases of medical and industrial importance. J. Mol. Struct. Theochem. 666-667, 487–498 179. Oikawa, A., and Maeda, A. (1957) The role of calcium in Taka-amylase A. J. Biochem. 44, 745–752 180. Heinen, W., and Lauwers, A. M. (1976) Amylase activity and stability at high and low temperature depending on calcium and other divalent cations. in Enzymes and Proteins from Thermophilic Microorganisms Structure and Function SE - 7 (Zuber, H. ed), pp. 77–89, Experientia Supplementum, Birkhäuser Basel, 26, 77– 89 181. Janeček, Š., and Baláž, Š. (1992) α-Amylases and approaches leading to their enhanced stability. FEBS Lett. 304, 1–3 182. Declerck, N., Machius, M., Wiegand, G., Huber, R., and Gaillardin, C. (2000) Probing structural determinants specifying high thermostability in Bacillus licheniformis α-amylase. J. Mol. Biol. 301, 1041–1057 183. Oikawa, A. (1959) The role of calcium in Taka-amylase A: II. The exchange reaction of calcium. J. Biochem. 46, 463–473 184. Agarwal, R. P., and Henkin, R. I. (1987) Metal binding characteristics of human salivary and porcine pancreatic amylase. J. Biol. Chem. 262, 2568–2575 185. Buisson, G., Duée, E., Haser, R., and Payan, F. (1987) Three dimensional structure of porcine pancreatic α-amylase at 2.9 A resolution. Role of calcium in structure and activity. EMBO J. 6, 3909–3916 186. Hagihara, H., Hayashi, Y., Endo, K., Igarashi, K., Ozawa, T., Kawai, S., Ozaki, K., and Ito, S. (2001) Deduced amino-acid sequence of a calcium-free α-amylase from a strain of Bacillus. Eur. J. Biochem. 268, 3974–3982 187. Sajedi, R. H., Naderi-Manesh, H., Khajeh, K., Ahmadvand, R., Ranjbar, B., Asoodeh, A., and Moradian, F. (2005) A Ca-independent α-amylase that is active and stable at low pH from the Bacillus sp. KR-8104. Enzyme Microb. Technol. 36, 666–671 188. Mehta, D., and Satyanarayana, T. (2013) Biochemical and molecular characterization of recombinant acidic and thermostable raw-starch hydrolysing αamylase from an extreme thermophile Geobacillus thermoleovorans. J. Mol. Catal. Page | 57 Introduction B Enzym. 85-86, 229–238 189. Babu, K. R., and Satyanarayana, T. (1993) Extracellular calcium-inhibited αamylase of Bacillus coagulans B 49. Enzyme Microb. Technol. 15, 1066–1069 Page | 58 190. Takase, K., Matsumoto, T., Mizuno, H., and Yamane, K. (1992) Site-directed mutagenesis of active site residues in Bacillus subtilis α-amylase. Biochim. Biophys. Acta - Protein Struct. Mol. Enzymol. 1120, 281–288 191. Fujimoto, Z., Takase, K., Doui, N., Momma, M., Matsumoto, T., and Mizuno, H. (1998) Crystal structure of a catalytic-site mutant α-amylase from Bacillus subtilis complexed with maltopentaose. J. Mol. Biol. 277, 393–407 192. Kadziola, A., Søgaard, M., Svensson, B., and Haser, R. (1998) Molecular structure of a barley α-amylase-inhibitor complex: Implications for starch binding and catalysis. J. Mol. Biol. 278, 205–217 193. Ramasubbu, N., Paloth, V., Luo, Y., Brayer, G. D., and Levine, M. J. (1996) Structure of human salivary α-amylase at 1.6 Å resolution: Implications for its role in the oral cavity. Acta Crystallogr. Sect. D. 52, 435–446 194. Kuriki, T., Takata, H., Okada, S., and Imanaka, T. (1991) Analysis of the active center of Bacillus stearothermophilus neopullulanase. J. Bacteriol. 173, 6147– 6152 195. Fritzsche, H. B., Schwede, T., and Schulz, G. E. (2003) Covalent and threedimensional structure of the cyclodextrinase from Flavobacterium sp. no. 92. Eur. J. Biochem. 270, 2332–2341 196. Yang, S.-J., Lee, H.-S., Park, C.-S., Kim, Y.-R., Moon, T.-W., and Park, K.-H. (2004) Enzymatic analysis of an amylolytic enzyme from the hyperthermophilic archaeon Pyrococcus furiosus reveals its novel catalytic properties as both an αamylase and a cyclodextrin-hydrolyzing enzyme. Appl. Environ. Microbiol. 70, 5988–5995 197. Klein, C., Hollender, J., Bender, H., and Schulz, G. E. (1992) Catalytic center of cyclodextrin glycosyltransferase derived from x-ray structure analysis combined with site-directed mutagenesis. Biochemistry. 31, 8740–8746 198. Knegtel, R. M. A., Strokopytov, B., Penninga, D., Faber, O. G., Rozeboom, H. J., Chapter 1 Kalk, K. H., Dijkhuizen, L., and Dijkstra, B. W. (1995) Crystallographic studies of the interaction of cyclodextrin glycosyltransferase from Bacillus circulans strain 251 with natural substrates and products. J. Biol. Chem. 270, 29256–29264 199. Uitdehaag, J. C. M., Mosi, R., Kalk, K. H., van der Veen, B. A., Dijkhuizen, L., Withers, S. G., and Dijkstra, B. W. (1999) X-ray structures along the reaction Page | 59 pathway of cyclodextrin glycosyltransferase elucidate catalysis in the α-amylase family. Nat Struct Mol Biol. 6, 432–436 200. Mathupala, S. P., Lowe, S. E., Podkovyrov, S. M., and Zeikus, J. G. (1993) Sequencing of the amylopullulanase (apu) gene of Thermoanaerobacter ethanolicus 39E, and identification of the active site by site-directed mutagenesis. J. Biol. Chem. 268, 16332–16344 201. Takata, H., Takaha, T., Kuriki, T., Okada, S., Takagi, M., and Imanaka, T. (1994) Properties and active center of the thermostable branching enzyme from Bacillus stearothermophilus. Appl. Environ. Microbiol. 60, 3096–3104 202. Abad, M. C., Binderup, K., Rios-Steiner, J., Arni, R. K., Preiss, J., and Geiger, J. H. (2002) The X-ray crystallographic structure of Escherichia coli branching enzyme. J. Biol. Chem. 277, 42164–42170 203. Chiba, S. (1997) Molecular mechanism in α-glucosidase and glucoamylase. Biosci. Biotechnol. Biochem. 61, 1233–1239 204. Swift, H. J., Brady, L., Derewenda, Z. S., Dodson, E. J., Dodson, G. G., Turkenburg, J. P., and Wilkinson, A. J. (1991) Structure and molecular model refinement of Aspergillus oryzae (TAKA) α-amylase: An application of the simulated-annealing method. Acta Crystallogr. Sect. B. 47, 535–544 205. Zechel, D. L., and Withers, S. G. (2000) Glycosidase mechanisms: Anatomy of a finely tuned catalyst. Acc. Chem. Res. 33, 11–18 206. Mhlongo, N. N., Skelton, A. A., Kruger, G., Soliman, M. E. S., and Williams, I. H. (2014) A critical survey of average distances between catalytic carboxyl groups in glycoside hydrolases. Proteins. 82, 1747–1755 207. Janeček, Š., and Ševčı́k, J. (1999) The evolution of starch-binding domain. FEBS Lett. 456, 119–125 208. Machovič, M., and Janeček, Š. (2006) Starch-binding domains in the post-genome Introduction era. Cell. Mol. Life Sci. C. 63, 2710–2724 209. Bolam, D. N., Ciruela, A., McQueen-Mason, S., Simpson, P., Williamson, M. P., Rixon, J. E., Boraston, A., Hazlewood, G. P., and Gilbert, H. J. (1998) Page | 60 Pseudomonas cellulose-binding domains mediate their effects by increasing enzyme substrate proximity. Biochem. J. 331, 775–781 210. Christiansen, C., Abou Hachem, M., Janeček, Š., Viksø-Nielsen, A., Blennow, A., and Svensson, B. (2009) The carbohydrate-binding module family 20 – Diversity, structure, and function. FEBS J. 276, 5006–5029 211. Chou, W.-Y., Pai, T.-W., Jiang, T.-Y., Chou, W.-I., Tang, C.-Y., and Chang, M. D.-T. (2011) Hydrophilic aromatic residue and in silico structure for carbohydrate binding module. PLoS One. 6, e24814 (1–10) 212. Rodríguez-Sanoja, R., Oviedo, N., and Sánchez, S. (2005) Microbial starchbinding domain. Curr. Opin. Microbiol. 8, 260–267 213. Sorimachi, K., Jacks, A. J., Le Gal-Coëffet, M. F., Williamson, G., Archer, D. B., and Williamson, M. P. (1996) Solution structure of the granular starch binding domain of glucoamylase from Aspergillus niger by nuclear magnetic resonance spectroscopy. J. Mol. Biol. 259, 970–987 214. Hashimoto, H. (2006) Recent structural studies of carbohydrate-binding modules. Cell. Mol. Life Sci. C. 63, 2954–2967 215. Peng, H., Zheng, Y., Chen, M., Wang, Y., Xiao, Y., and Gao, Y. (2014) A starchbinding domain identified in α-amylase (AmyP) represents a new family of carbohydrate-binding modules that contribute to enzymatic hydrolysis of soluble starch. FEBS Lett. 588, 1161–1167 216. Parker, M. W. (2003) Protein structure from X-ray diffraction. J. Biol. Phys. 29, 341–362 217. Shin, J., Lee, W., and Lee, W. (2008) Structural proteomics by NMR spectroscopy. Expert Rev. Proteomics. 5, 589–601 218. Mikkelsen, R., Suszkiewicz, K., and Blennow, A. (2006) A novel type carbohydrate-binding module identified in α-glucan, water dikinases is specific for regulated plastidial starch metabolism. Biochemistry. 45, 4674–4682 Chapter 1 219. Boraston, A. B., Bolam, D. N., Gilbert, H. J., and Davies, G. J. (2004) Carbohydrate-binding modules: fine-tuning polysaccharide recognition. Biochem. J. 382, 769–781 220. Machovič, M., Svensson, B., Ann MacGregor, E., and Janeček, Š. (2005) A new clan of CBM families based on bioinformatics of starch-binding domains from Page | 61 families CBM20 and CBM21. FEBS J. 272, 5497–5513 221. Machovic, M., and Janecek, S. (2006) The evolution of putative starch-binding domains. FEBS Lett. 580, 6349–6356 222. Lo, H.-F., Lin, L.-L., Chiang, W.-Y., Chie, M.-C., Hsu, W.-H., and Chang, C.-T. (2002) Deletion analysis of the C-terminal region of the α-amylase of Bacillus sp. strain TS-23. Arch. Microbiol. 178, 115–123 223. Lin, L.-L., Lo, H.-F., Chi, M.-C., and Ku, K.-L. (2003) Functional expression of the raw starch-binding domain of Bacillus sp. strain TS-23 α-amylase in recombinant Escherichia coli. Starch - Stärke. 55, 197–202 224. Penninga, D., van der Veen, B. A., Knegtel, R. M. A., van Hijum, S. A. F. T., Rozeboom, H. J., Kalk, K. H., Dijkstra, B. W., and Dijkhuizen, L. (1996) The raw starch binding somain of cyclodextrin glycosyltransferase from Bacillus circulans strain 251. J. Biol. Chem. 271, 32777–32784 225. Mikami, B., Adachi, M., Kage, T., Sarikaya, E., Nanmori, T., Shinke, R., and Utsumi, S. (1999) Structure of raw starch-digesting Bacillus cereus β-amylase complexed with maltose. Biochemistry. 38, 7050–7061 226. Sorimachi, K., Gal-Coëffet, M.-F. Le, Williamson, G., Archer, D. B., and Williamson, M. P. (1997) Solution structure of the granular starch binding domain of Aspergillus niger glucoamylase bound to β-cyclodextrin. Structure. 5, 647–661 227. Lawson, C. L., van Montfort, R., Strokopytov, B., Rozeboom, H. J., Kalk, K. H., de Vries, G. E., Penninga, D., Dijkhuizen, L., and Dijkstra, B. W. (1994) Nucleotide sequence and X-ray structure of cyclodextrin glycosyltransferase from Bacillus circulans strain 251 in a maltose-dependent crystal form. J. Mol. Biol. 236, 590–600 228. Jacks, A. J., Sorimachi, K., Le Gal-Coëffet, M.-F., Williamson, G., Archer, D. B., and Williamson, M. P. (1995) 1H and 15N assignments and secondary structure of Introduction the starch-binding somain of glucoamylase from Aspergillus niger. Eur. J. Biochem. 233, 568–578 229. Goto, M., Semimaru, T., Furukawa, K., and Hayashida, S. (1994) Analysis of the Page | 62 raw starch-binding domain by mutation of a glucoamylase from Aspergillus awamori var. kawachi expressed in Saccharomyces cerevisiae. Appl. Environ. Microbiol. 60, 3926–3930 230. Lo, H.-F., Chiang, W.-Y., Chi, M.-C., Hu, H.-Y., and Lin, L.-L. (2004) Sitedirected mutagenesis of the conserved threonine, tryptophan, and lysine residues in the starch-binding domain of Bacillus sp. strain TS-23 α-amylase. Curr. Microbiol. 48, 280–284 231. Svensson, B., Svendsen, T. G., Svendsen, I., Sakai, T., and Ottesen, M. (1982) Characterization of two forms of glucoamylase from Aspergillus niger. Carlsb. Res. Commun. 47, 55–69 232. Juge, N., Nøhr, J., Le Gal-Coëffet, M.-F., Kramhøft, B., Furniss, C. S. M., Planchot, V., Archer, D. B., Williamson, G., and Svensson, B. (2006) The activity of barley α-amylase on starch granules is enhanced by fusion of a starch binding domain from Aspergillus niger glucoamylase. Biochim. Biophys. Acta. 1764, 275– 284 233. Liu, Y.-N., Lai, Y.-T., Chou, W.-I., Chang, M. D.-T., and Lyu, P.-C. (2007) Solution structure of family 21 carbohydrate-binding module from Rhizopus oryzae glucoamylase. Biochem. J. 403, 21–30 234. Tung, J.-Y., Chang, M. D.-T., Chou, W.-I., Liu, Y.-Y., Yeh, Y.-H., Chang, F.-Y., Lin, S.-C., Qiu, Z.-L., and Sun, Y.-J. (2008) Crystal structures of the starchbinding domain from Rhizopus oryzae glucoamylase reveal a polysaccharidebinding path. Biochem. J. 416, 27–36 235. Chou, W.-I., Pai, T.-W., Liu, S.-H., Hsiung, B.-K., and Chang, M. D.-T. (2006) The family 21 carbohydrate-binding module of glucoamylase from Rhizopus oryzae consists of two sites playing distinct roles in ligand binding. Biochem. J. 396, 469–477 236. Sumitani, J., Tottori, T., Kawaguchi, T., and Arai, M. (2000) New type of starchbinding domain: the direct repeat motif in the C-terminal region of Bacillus sp. no. Chapter 1 195 α-amylase contributes to starch binding and raw starch degrading. Biochem. J. 350, 477–484 237. Boraston, A. B., Healey, M., Klassen, J., Ficko-Blean, E., van Bueren, A. L., and Law, V. (2006) A structural and functional analysis of α-glucan recognition by family 25 and 26 carbohydrate-binding modules reveals a conserved mode of Page | 63 starch recognition. J. Biol. Chem. 281, 587–598 238. Majzlová, K., and Janeček, Š. (2014) Two structurally related starch-binding domain families CBM25 and CBM26. Biologia (Bratisl). 69, 1087–1096 239. Guillén, D., Santiago, M., Linares, L., Pérez, R., Morlon, J., Ruiz, B., Sánchez, S., and Rodríguez-Sanoja, R. (2007) α-Amylase starch binding domains: Cooperative effects of binding to starch granules of multiple tandemly arranged domains. Appl. Environ. Microbiol. 73, 3833–3837 240. Giraud, E., and Cuny, G. (1997) Molecular characterization of the α-amylase genes of Lactobacillus plantarum A6 and Lactobacillus amylovorus reveals an unusual 3′ end structure with direct tandem repeats and suggests a common evolutionary origin. Gene. 198, 149–157 241. Morlon-Guyot, J., Mucciolo-Roux, F., Sanoja, R. R., and Guyot, J.-P. (2001) Characterization of the L. manihotivorans α-amylase gene. DNA Seq. 12, 27–37 242. Rodriguez Sanoja, R., Morlon-Guyot, J., Jore, J., Pintado, J., Juge, N., and Guyot, J. P. (2000) Comparative characterization of complete and truncated forms of Lactobacillus amylovorus α-amylase and role of the C-terminal direct repeats in raw-starch binding. Appl. Environ. Microbiol. 66, 3350–3356 243. Kamitori, S., Kondo, S., Okuyama, K., Yokota, T., Shimura, Y., Tonozuka, T., and Sakano, Y. (1999) Crystal structure of Thermoactinomyces vulgaris R-47 alphaamylase II (TVAII) hydrolyzing cyclodextrins and pullulan at 2.6 A resolution. J. Mol. Biol. 287, 907–921 244. Abe, A., Tonozuka, T., Sakano, Y., and Kamitori, S. (2004) Complex structures of Thermoactinomyces vulgaris R-47 α-amylase 1 with malto-oligosaccharides demonstrate the role of domain N acting as a starch-binding domain. J. Mol. Biol. 335, 811–822 245. Abe, A., Yoshida, H., Tonozuka, T., Sakano, Y., and Kamitori, S. (2005) Introduction Complexes of Thermoactinomyces vulgaris R-47 α-amylase 1 and pullulan model oligossacharides provide new insight into the mechanism for recognizing substrates with alpha-(1,6) glycosidic linkages. FEBS J. 272, 6145–6153 Page | 64 246. Lammerts van Bueren, A., Finn, R., Ausió, J., and Boraston, A. B. (2004) αGlucan recognition by a new family of carbohydrate-binding modules found primarily in bacterial pathogens. Biochemistry. 43, 15633–15642 247. van Bueren, A. L., and Boraston, A. B. (2007) The structural basis of α-glucan recognition by a family 41 carbohydrate-binding module from Thermotoga maritima. J. Mol. Biol. 365, 555–560 248. Mikami, B., Iwamoto, H., Malle, D., Yoon, H.-J., Demirkan-Sarikaya, E., Mezaki, Y., and Katsuya, Y. (2006) Crystal structure of pullulanase: Evidence for parallel binding of oligosaccharides in the active site. J. Mol. Biol. 359, 690–707 249. Glaring, M. A., Baumann, M. J., Hachem, M. A., Nakai, H., Nakai, N., Santelia, D., Sigurskjold, B. W., Zeeman, S. C., Blennow, A., and Svensson, B. (2011) Starch-binding domains in the CBM45 family – low-affinity domains from glucan, water dikinase and α-amylase involved in plastidial starch metabolism. FEBS J. 278, 1175–1185 250. Chou, W.-Y., Chou, W.-I., Pai, T.-W., Lin, S.-C., Jiang, T.-Y., Tang, C.-Y., and Chang, M. D.-T. (2010) Feature-incorporated alignment based ligand-binding residue prediction for carbohydrate-binding modules. Bioinformatics. 26, 1022– 1028 251. Janeček, Š., Svensson, B., and MacGregor, E. A. (2011) Structural and evolutionary aspects of two families of non-catalytic domains present in starch and glycogen binding proteins from microbes, plants and animals. Enzyme Microb. Technol. 49, 429–440 252. Machovič, M., and Janeček, Š. (2008) Domain evolution in the GH13 pullulanase subfamily with focus on the carbohydrate-binding module family 48. Biologia (Bratisl). 63, 1057–1068 253. Jung, T.-Y., Li, D., Park, J.-T., Yoon, S.-M., Tran, P. L., Oh, B.-H., Janeček, Š., Park, S. G., Woo, E.-J., and Park, K.-H. (2012) Association of novel domain in active site of archaic hyperthermophilic maltogenic amylase from Staphylothermus Chapter 1 marinus. J. Biol. Chem. 287, 7979–7989 254. Koropatkin, N. M., and Smith, T. J. (2010) SusG: A unique cell-membraneassociated α-amylase from a prominent human gut symbiont targets complex starch molecules. Structure. 18, 200–215 255. Bozonnet, S., Jensen, M. T., Nielsen, M. M., Aghajari, N., Jensen, M. H., Kramhøft, B., Willemoës, M., Tranier, S., Haser, R., and Svensson, B. (2007) The “pair of sugar tongs” site on the non-catalytic domain C of barley α-amylase participates in substrate binding and activity. FEBS J. 274, 5055–5067 256. Puspasari, F., Radjasa, O. K., Noer, A. S., Nurachman, Z., Syah, Y. M., van der Maarel, M., Dijkhuizen, L., Janeček, Š., and Natalia, D. (2013) Raw starch– degrading α-amylase from Bacillus aquimaris MKSC 6.2: isolation and expression of the gene, bioinformatics and biochemical characterization of the recombinant enzyme. J. Appl. Microbiol. 114, 108–120 257. Ranjani, V., Janeček, S., Chai, K. P., Shahir, S., Abdul Rahman, R. N. Z. R., Chan, K.-G., and Goh, K. M. (2014) Protein engineering of selected residues from conserved sequence regions of a novel Anoxybacillus α-amylase. Sci. Rep. 4, 5850–5858 258. Campbell, I. D., and Spitzfaden, C. (1994) Building proteins with fibronectin type III modules. Structure. 2, 333–337 259. de Vos, A., Ultsch, M., and Kossiakoff, A. (1992) Human growth hormone and extracellular domain of its receptor: crystal structure of the complex. Science (80-. ). 255, 306–312 260. María de Pereda, J., Wiche, G., and Liddington, R. C. (1999) Crystal structure of a tandem pair of fibronectin type III domains from the cytoplasmic tail of integrin α6β4. EMBO J. 18, 4087–4095 261. Leahy, D. J., Aukhil, I., and Erickson, H. P. (1996) 2.0 Å crystal structure of a four-domain segment of human fibronectin encompassing the RGD loop and synergy region. Cell. 84, 155–164 262. Goll, C. M., Pastore, A., and Nilges, M. (1998) The three-dimensional structure of a type I module from titin: A prototype of intracellular fibronectin type III domains. Structure. 6, 1291–1302 Page | 65 Introduction 263. Little, E., Bork, P., and Doolittle, R. (1994) Tracing the spread of fibronectin type III domains in bacterial glycohydrolases. J. Mol. Evol. 39, 631–643 264. Valk, V., Eeuwema, W., Sarian, F. D., van der Kaaij, R. M., and Dijkhuizen, L. Page | 66 (2015) Degradation of granular starch by the bacterium Microbacterium aurum strain B8.A involves a modular α-amylase enzyme system with FNIII and CBM25 domains. Appl. Environ. Microbiol. 81, 6610–6620 265. Watanabe, T., Suzuki, K., Oyanagi, W., Ohnishi, K., and Tanaka, H. (1990) Gene cloning of chitinase A1 from Bacillus circulans WL-12 revealed its evolutionary relationship to Serratia chitinase and to the type III homology units of fibronectin. J. Biol. Chem. 265, 15659–15665 266. Watanabe, T., Ito, Y., Yamada, T., Hashimoto, M., Sekine, S., and Tanaka, H. (1994) The roles of the C-terminal domain and type III domains of chitinase A1 from Bacillus circulans WL-12 in chitin degradation. J. Bacteriol. 176, 4465–4472 267. Kataeva, I. A., Seidel, R. D., Shah, A., West, L. T., Li, X.-L., and Ljungdahl, L. G. (2002) The fibronectin type 3-like repeat from the Clostridium thermocellum cellobiohydrolase CbhA promotes hydrolysis of cellulose by modifying its surface. Appl. Environ. Microbiol. 68, 4292–300 268. Lin, F.-P., Ma, H.-Y., Lin, H.-J., Liu, S.-M., and Tzou, W.-S. (2011) Biochemical characterization of two truncated forms of amylopullulanase from Thermoanaerobacterium saccharolyticum NTOU1 to identify its enzymatically active region. Appl. Biochem. Biotechnol. 165, 1047–1056 269. Goldsmith, E. J., Fletterick, R. J., and Withers, S. G. (1987) The three-dimensional structure of acarbose bound to glycogen phosphorylase. J. Biol. Chem. 262, 1449– 1455 270. Mori, H., Tanizawa, K., and Fukui, T. (1993) Engineered plant phosphorylase showing extraordinarily high affinity for various α-glucan molecules. Protein Sci. 2, 1621–1629 271. Søgaard, M., Kadziola, A., Haser, R., and Svensson, B. (1993) Site-directed mutagenesis of histidine 93, aspartic acid 180, glutamic acid 205, histidine 290, and aspartic acid 291 at the active site and tryptophan 279 at the raw starch binding site in barley α-amylase 1. J. Biol. Chem. 268, 22480–22484 Chapter 1 272. Cuyvers, S., Dornez, E., Delcour, J. A., and Courtin, C. M. (2012) Occurrence and functional significance of secondary carbohydrate binding sites in glycoside hydrolases. Crit. Rev. Biotechnol. 32, 93–107 273. Cockburn, D., Wilkens, C., Ruzanski, C., Andersen, S., Willum Nielsen, J., Smith, A., Field, R., Willemoës, M., Abou Hachem, M., and Svensson, B. (2014) Analysis Page | 67 of surface binding sites (SBSs) in carbohydrate active enzymes with focus on glycoside hydrolase families 13 and 77 — A mini-review. Biologia (Bratisl). 69, 705–712 274. Cockburn, D., Nielsen, M. M., Christiansen, C., Andersen, J. M., Rannes, J. B., Blennow, A., and Svensson, B. (2015) Surface binding sites in amylase have distinct roles in recognition of starch structure motifs and degradation. Int. J. Biol. Macromol. 75, 338–345 275. Robert, X., Haser, R., Gottschalk, T. E., Ratajczak, F., Driguez, H., Svensson, B., and Aghajari, N. (2003) The structure of barley α-amylase isozyme 1 reveals a novel role of domain C in substrate recognition and binding. Structure. 11, 973– 984 276. Hostinová, E., Janecek, S., and Gasperík, J. (2010) Gene sequence, bioinformatics and enzymatic characterization of α-amylase from Saccharomycopsis fibuligera KZ. Protein J. 29, 355–364 277. Nielsen, J. W., Kramhøft, B., Bozonnet, S., Abou Hachem, M., Stipp, S. L. S., Svensson, B., and Willemoës, M. (2012) Degradation of the starch components amylopectin and amylose by barley α-amylase 1: role of surface binding site 2. Arch. Biochem. Biophys. 528, 1–6 278. Nielsen, M. M., Bozonnet, S., Seo, E.-S., Mótyán, J. A., Andersen, J. M., Dilokpimol, A., Abou Hachem, M., Gyémánt, G., Naested, H., Kandra, L., Sigurskjold, B. W., and Svensson, B. (2009) Two secondary carbohydrate binding sites on the surface of barley α-amylase 1 have distinct functions and display synergy in hydrolysis of starch granules. Biochemistry. 48, 7686–97 279. Robert, X., Haser, R., Mori, H., Svensson, B., and Aghajari, N. (2005) Oligosaccharide binding to barley α-amylase 1. J. Biol. Chem. 280, 32968–32978 280. Planchot, V., Colonna, P., Gallant, D. J. J., and Bouchet, B. (1995) Extensive degradation of native starch granules by α-amylase from Aspergillus fumigatus. J. Introduction Cereal Sci. 21, 163–171 281. Mørkeberg, R., Carlsen, M., and Nielsen, J. (1995) Induction and repression of αamylase production in batch and continuous cultures of Aspergillus oryzae. Page | 68 Microbiology. 141, 2449–2454 282. Hayashida, S. (1975) Selective submerged productions of three types of glucoamylases by a black-koji, mold. Agric. Biol. Chem. 39, 2093–2099 283. Hayashida, S., Kuroda, K., Ohta, K., Kuhara, S., Fukuda, K., and Sakaki, Y. (1989) Molecular cloning of the glucoamylase I gene of Aspergillus awamori var. kawachi for localization of the raw-starch-affinity site. Agric. Biol. Chem. 53, 923–929 284. Anindyawati, T., Melliawati, R., Ito, K., Iizuka, M., and Minamiura, N. (1998) Three different types of α-amylases from Aspergillus awamori KT-11: their purifications, properties, and specificities. Biosci. Biotechnol. Biochem. 62, 1351– 1357 285. Matsubara, T., Ben Ammar, Y., Anindyawati, T., Yamamoto, S., Ito, K., Iizuka, M., and Minamiura, N. (2004) Degradation of raw starch granules by α-amylase purified from culture of Aspergillus awamori KT-11. J. Biochem. Mol. Bol. 37, 422–428 286. Abe, J. I., Bergmann, F. W., Obata, K., and Hizukuri, S. (1988) Production of the raw-starch digesting amylase of Aspergillus sp. K-27. Appl. Microbiol. Biotechnol. 27, 447–450 287. Demirkan, E. S., Mikami, B., Adachi, M., Higasa, T., and Utsumi, S. (2005) αAmylase from B. amyloliquefaciens: Purification, characterization, raw starch degradation and expression in E. coli. Process Biochem. 40, 2629–2636 288. Puspasari, F., Nurachman, Z., Noer, A. S., Radjasa, O. K., van der Maarel, M. J. E. C., and Natalia, D. (2011) Characteristics of raw starch degrading α-amylase from Bacillus aquimaris MKSC 6.2 associated with soft coral Sinularia sp. Starch Stärke. 63, 461–467 289. Nanmori, T., Nagai, M., Shimizu, Y., Shinke, R., and Mikami, B. (1993) Cloning of the β-amylase gene from Bacillus cereus and characteristics of the primary structure of the enzyme. Appl. Environ. Microbiol. 59, 623–627 Chapter 1 290. Candotti, M., Pérez, A., Ferrer-Costa, C., Rueda, M., Meyer, T., Gelpí, J. L., and Orozco, M. (2013) Exploring early stages of the chemical unfolding of proteins at the proteome scale. PLoS Comput. Biol. 9, e1003393 (1–11) 291. Watanabe, H., Nishimoto, T., Mukai, K., Kubota, M., Chaen, H., and Fukuda, S. (2006) A novel glucanotransferase from a Bacillus circulans strain that produces a Page | 69 cyclomaltopentaose cyclized by an α-1,6-linkage. Biosci. Biotechnol. Biochem. 70, 1954–1960 292. Goel, A., and Nene, S. (1995) A novel cyclomaltodextrin glucanotransferase from Bacillus firmus that degrades raw starch. Biotechnol. Lett. 17, 411–416 293. Gawande, B. N., Goel, A., Patkar, A. Y., and Nene, S. N. (1999) Purification and properties of a novel raw starch degrading cyclomaltodextrin glucanotransferase from Bacillus firmus. Appl. Microbiol. Biotechnol. 51, 504–509 294. Lin, L.-L., Tsau, M.-R., and Chu, W.-S. (1994) General characteristics of thermostable amylopullulanases and amylases from the alkalophilic Bacillus sp. TS-23. Appl. Microbiol. Biotechnol. 42, 51–56 295. Nagasaka, Y., Kurosawa, K., Yokota, A., and Tomita, F. (1998) Purification and properties of the raw-starch-digesting glucoamylases from Corticium rolfsii. Appl. Microbiol. Biotechnol. 50, 323–330 296. Galdino, A. S., Ulhoa, C. J., Moraes, L. M. P., Prates, M. V, Bloch, C., and Torres, F. A. G. (2008) Cloning, molecular characterization and heterologous expression of AMY1, an α-amylase gene from Cryptococcus flavus. FEMS Microbiol. Lett. 280, 189–194 297. Dicko, M. H., Searle-van Leeuwen, M. J. F., Beldman, G., Ouedraogo, O. G., Hilhorst, R., and Traoré, A. S. (1999) Purification and characterization of βamylase from Curculigo pilosa. Appl. Microbiol. Biotechnol. 52, 802–805 298. Adachi, M., Mikami, B., Katsube, T., and Utsumi, S. (1998) Crystal structure of recombinant soybean β-amylase complexed with β-cyclodextrin. J. Biol. Chem. 273, 19859–19865 299. Sarikaya, E., Higasa, T., Adachi, M., and Mikami, B. (2000) Comparison of degradation abilities of α- and β-amylases on raw starch granules. Process Biochem. 35, 711–715 Introduction 300. Sim, L., Quezada-Calvillo, R., Sterchi, E. E., Nichols, B. L., and Rose, D. R. (2008) Human intestinal maltase-glucoamylase: Crystal structure of the N-terminal catalytic subunit and basis of inhibition and substrate specificity. J. Mol. Biol. 375, 782–792 Page | 70 301. MacGregor, a. W., and Ballance, D. L. (1980) Hydrolysis of large and small starch granules from normal and waxy barley cultivars by α-amylases from barley malt. Cereal Chem. 57, 397–402 302. Tibbot, B. K., Wong, D. W. S., and Robertson, G. H. (2002) Studies on the Cterminal region of barley α-amylase 1 with emphasis on raw starch-binding. Biologia (Bratisl). 303. Binder, F., Huber, O., and Böck, A. (1986) Cyclodextrin-glycosyltransferase from Klebsiella pneumoniae M5a1: Cloning, nucleotide sequence and expression. Gene. 47, 269–277 304. Gawande, B. ., and Patkar, A. . (2001) Purification and properties of a novel raw starch degrading-cyclodextrin glycosyltransferase from Klebsiella pneumoniae AS22. Enzyme Microb. Technol. 28, 735–743 305. Giraud, E., Champailler, A., and Raimbault, M. (1994) Degradation of raw starch by a wild amylolytic atrain of Lactobacillus plantarum. Appl. Environ. Microbiol. 60, 4319–4323 306. Lei, Y., Peng, H., Wang, Y., Liu, Y., Han, F., Xiao, Y., and Gao, Y. (2012) Preferential and rapid degradation of raw rice starch by an α-amylase of glycoside hydrolase subfamily GH13_37. Appl. Microbiol. Biotechnol. 94, 1577–1584 307. Witt, W., and Sauter, J. J. (1995) In vitro degradation of starch grains by phosphorylases and amylases from poplar wood. J. Plant Physiol. 146, 35–40 308. Witt, W., and Sauter, J. J. (1996) Purification and characterization of α-amylase from poplar leaves. Phytochemistry. 41, 365–372 309. Takahashi, T., Tsuchida, Y., and Irie, M. (1978) Purification and some properties of three forms of glucoamylase from a Rhizopus species. J. Biochem. 84, 1183– 1194 310. Takahashi, T., Kato, K., Ikegami, Y., and Irie, M. (1985) Different behavior Chapter 1 towards raw starch of three forms of glucoamylase from a Rhizopus sp. J. Biochem. 98, 663–671 311. Celińska, E., Białas, W., Borkowska, M., and Grajek, W. (2015) Cloning, expression, and purification of insect (Sitophilus oryzae) α-amylase, able to digest granular starch, in Yarrowia lipolytica host. Appl. Microbiol. Biotechnol. 99, 2727– Page | 71 2739 312. Matsui, Y., Okada, S., Uchimura, T., Kondo, A., and Satoh, E. (2007) Determination and analysis of the starch binding domain of Streptococcus bovis 148 raw-starch-hydrolyzing α-amylase. J. Appl. Glycosci. 54, 217–222 313. Hang, M. T., Furuyoshi, S., Yagi, T., and Yamamoto, S. (1996) Purification and characterization of raw starch-digesting α-amylase from Streptomyces thermocyaneoviolaceus IFO 14271. J. Appl. Glycosci. 43, 487–497 314. Li, C., Du, M., Cheng, B., Wang, L., Liu, X., Ma, C., Yang, C., and Xu, P. (2014) Close relationship of a novel Flavobacteriaceae α-amylase with archaeal αamylases and good potentials for industrial applications. Biotechnol. Biofuels. 7, 18–30 315. Kim, J. W., Flowers, L. O., Whiteley, M., and Peeples, T. L. (2001) Biochemical confirmation and characterization of the family-57-like α-amylase of Methanococcus jannaschii. Folia Microbiol. (Praha). 46, 467–473 316. Jørgensen, S., Vorgias, C. E., and 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 317. Koch, R., Spreinat, A., Lemke, K., and Antranikian, G. (1991) Purification and properties of a hyperthermoactive α-amylase from the archaeobacterium Pyrococcus woesei. Arch. Microbiol. 155, 572–578 318. Kwak, Y. S., Akiba, T., and Kudo, T. (1998) Purification and characterization of αamylase from hyperthermophilic archaeon Thermococcus profundus, which hydrolyzes both α-1,4 and α-1,6 glucosidic linkages. J. Ferment. Bioeng. 86, 363– 367 319. Buonocore, V., Caporale, C., De Rosa, M., and Gambacorta, A. (1976) Stable, Introduction inducible thermoacidophilic α-amylase from Bacillus acidocaldarius. J. Bacteriol. 128, 515–521 320. Heinen, U. J., and Heinen, W. (1972) Characteristics and properties of a caldoPage | 72 active bacterium producing extracellular enzymes and two related strains. Arch. Mikrobiol. 82, 1–23 321. Ivanova, V. N., Dobreva, E. P., and Emanuilova, E. I. (1993) Purification and characterization of a thermostable α-amylase from Bacillus licheniformis. J. Biotechnol. 28, 277–289 322. Brumm, P. J., Hebeda, R. E., and Teague, W. M. (1991) Purification and characterization of the commercialized, cloned Bacillus megaterium α-amylase. Part I: Purification and hydrolytic properties. Starch - Stärke. 43, 315–319 323. Pfueller, S. L., and Elliott, W. H. (1969) The extracellular α-amylase of Bacillus stearothermophilus. J. Biol. Chem. 244, 48–54 324. 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, 67–71 330. 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 LCMS. 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 Page | 73 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 αamylase 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 Aspergillus Type awamori Source var. glucoamylase n.d. Raw Starch Activity Binding GH (sub) Domain Family References corn CBM20 GH15 (282, 283) cassava, corn, CBM20 GH13 (284, 285) n.d. (280, 286) no SBD GH13_5 (287) no SBD GH13_36 (288) kawachi Aspergillus awamori KT-11 α-amylase, air in Bogor, glucoamylase Indonesia potato, rice, (act together) sago, sweet potato, wheat Aspergillus fumigatus α-amylase soil corn, pea, n.d. wheat Bacillus amyloliquefaciens α-amylase Turkish soil corn, potato, rice, sweet potato, wheat Bacillus aquimaris α-amylase soft coral cassava, corn, 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 | 75 Page | 76 Table S1.1 continued. List of raw-starch degrading enzymes from different species. Organism Bacillus sp. no. 195 Type α-amylase Source soil Raw Starch Activity Binding GH (sub) Domain Family CBM25 GH13_32 (236) corn CBM20 GH13 (223, 294) cassava, corn, CBM20 GH15 (220, 295) CBM20 GH13_1 (296) n.d. (297) corn, potato, rice, References sweet potato, wheat Bacillus sp. strain TS-23 α-amylase soil Corticium rolfsii glucoamylase tomato stem potato, rice, sago, sweet potato Cryptococcus flavus α-amylase culture collection cassava Curculigo pilosa (flower β-amylase tubers plant) corn, potato, rice, n.d. 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) barley no SBD GH13_6 (301, 302) cassava, corn, CBM20 GH13_2 (303, 304) glucoamylase Hordeum vulgare (barley) α-amylase corn, rice, wheat germinated barley Klebsiella pneumoniae AS- CGTase n.d. 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 Marine metagenomics library α-amylase Source Raw Starch Activity subsurface rice, wheat Binding GH (sub) Domain Family References CBM69 GH13_37 (215, 306) CBM25 GH13_32 (264) n.d. (307, 308) sediments in South China sea Microbacterium aurum B8.A α-amylase Populus x canadensis α-amylase potato plant corn, wheat, waste water potato leaf and wood corn, potato n.d. 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. (313) collection rice, sago, thermocyaneoviolaceus IFO14271 n.d. sweet potato, potato n.d., no data; CBM, carbohydrate-binding module; SBD, starch-binding domain; GH, glycoside hydrolase Page | 77 Page | 78 Table S1.2 Physicochemical properties of α‐amylases Molecular mass (kDa) Optimum pH Optimum temperature (oC) Metal requirement 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) 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) 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+ Chloroflexus aurantiacus 210 7.5 71 Ca2+ Source References Archaea Bacteria Bacillus stearothermophilus n.d., no data (294, 325) (175) Table S1.2 continued. Physicochemical properties of α‐amylases 5.6 Optimum temperature (oC) 45 Metal requirement no (326) 67 5.5 80 n.d. (327) 59 5.5 70 n.d. (327) Geobacillus thermoleovorans 59 5.0 80 no (188) Halomonas meridiana 49 7.0 37 Ca2+ (328, 329) 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) Thermotoga maritima MSB8 50 7.0 70 Ca2+ (170) Thermoactinomyces vulgaris 67 5.0 40 Ca2+ (130, 334) Source Clostridium acetobutylicum ATCC Molecular mass (kDa) 84 Optimum pH References 824 Dictyoglomus thermophilum (AmyB) Dictyoglomus thermophilum (AmyC) Lactobacillus plantarum thermocyaneoviolaceus IFO14271 n.d., no data Page | 79 Page | 80 Table S1.2 continued. Physicochemical properties of α‐amylases Source Vibrio sp. Molecular mass (kDa) 52.5 Optimum pH 6.5 Optimum temperature (oC) 60 Metal requirement Ca2+, Co2+, Mg2+, 2+ Mn , Fe References (335) 2+ 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) Cryptococcus flavus 67 5.5 50 no Lipomyces kononenkoae 34 5.5 40 Ca2+ (341) Pichia burtonii 51 5.0 40 Ca2+ (342) 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) 55.2 6.9 25 Ca2+ (346) Populus canadensis Moench (leaf) 44 6.5 30 Ca2+, Mg2+ Sitophilus oryzae (insect) 53 5.0 40 no Yeasts (339, 340) Eukaryotes (other) Human saliva n.d., no data (307, 308) (311)
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