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I. INTRODUCTION….
1
Enzymes are biocatalysts that accelerate the rate of biological reactions through
defined pathways without being used in the process. These are made up of one or
more polypeptides organized in specific 3-dimensional structures. Enzymes have high
substrate specificity, stereospecificity and regiospecificity, which are expressed
during catalysis. Enzymatic reactions occur within a narrow temperature and pH
ranges. Any change in the vital factors may result in the loss of structural integrity of
the enzyme, thereby leading to loss of enzymatic activity. Although enzymes have
been exploited for long, the potential of enzymes in fermentation process was
understood in the beginning of 19th century. In 1860, Louis Pasteur recognized the
importance of enzymes in fermentation, and later in 1897, German chemist Edward
Buchner showed that the cell free extracts of yeast cells (Zyfnmase) could ferment
sugars to alcohol and carbon dioxide. American biochemist Sumner was the first to
isolate and crystallize enzyme (urease) in 1926. Later during 1930-1936, many other
enzymes like pepsin, trypsin and chymostrypsin were successfully crystallized.
Enzymes from different sources have wider applications in industries such as food,
pharmaceutical, leather, detergent, textile, paper and pulp, waste management and
others. Most of the enzymes for commercial applications are obtained from
microorganisms, including bacteria, fungi and yeasts. Among all industrial enzymes,
hydrolytic enzymes account for 85%. The market size was approximately US$ 1.6
billion in 2002, and about 12% annual growth has been witnessed in last one decade,
and therefore, this growth is expected to reach US$3 billion by 2008 (Pandey and
Ramachandran 2005). According to Sanchez and Demain (2010), the enzyme market
was US$5.1 billion in 2009.
1.1. Starch as a substrate
Starch is the second major food reserve polysaccharide in nature after cellulose. More
than a billion tonnes of starch is produced annually (The Food and Agriculture
Organization of the United Nations [FAO]; http://www.fao.org/) [Morell and Myers,
2005]. Plants are unique in synthesizing this α-glucan that serves as important source
of nutrition for other living organisms. A large number of bacteria, fungi, and yeasts
produce extracellular enzymes that degrade these substances in different
environmental niches (Antranikian 1992). A variety of polysaccharide hydrolyzing
enzymes suited for various industrial applications have emerged in last few decades
leading to the screening of enzymes with novel properties. Starch is the most easily
Introduction
2
available source of carbon and energy on earth, and is synthesized by plants in the
presence of sunlight and water through photosynthesis. Starch is biosynthesized as
semi-crystalline granules with different polymorphic types and degree of crystallinity.
Starch is synthesized in plastids present in the leaves and accumulated there as
insoluble granules of higher and lower plants. The granule size varies from 2 to 100
µm with round, oval and irregular shapes (Table 1.1). It is also synthesized by
amyloplasts found in tubers, seeds and other reserve tissues and is used by the plant in
one stage of life cycle to another, and such starch is called reserve carbohydrate.
Starch is major component of most of the staple foods and is used in many food and
non-food industries. The starch is mainly utilized in textile, paper, pharmaceutical,
beverage, alcohol and candy manufacture.
The structural organization of starch is mainly composed of two high
molecular weight compounds amylose and amylopectin, and both of these contain α-
D glucose as a sole monomer. Amylose is a linear water insoluble polymer of glucose
subunits joined by α-1, 4 bonds (99%) with the molecular weight of ~1x105 to 1x106.
On the other hand, amylopectin is branched water soluble polysaccharide with short
α-1, 4 linked (~95%) linear chains of 10-60 glucose units and α-1, 6 linked (~5%) side
chains with 15-45 glucose units that forms the volume of starch molecule (Buleon et
al.1998; Tester et al. 2004). The ratio of amylose to amylopectin varies among
starches, but representative levels of amylose to amylopectin are 25-28% and 72-75%,
respectively. Differences in amylose to amylopectin ratios of starches considered to
result in variations in granular structure, physiological properties and quality of useful
end products. The crystalline character of amylopectin depends on the regularity of
branching. At the end of the polymeric chain, a latent aldehyde group is present,
which is known as the reducing end.
Small amounts of lipids, phosphates and proteins are present in starch granules.
Lipid is only present in cereal starches, which is positively associated with the
amylose content where it represents ~1.5% of the granule (Morrison, 1993; Tester and
Karkalas, 2002). Lipid is thought to be complexed with some of the amylose that
produces ‘lipid free’ and ‘lipid complexed’ amylose. The structure and function of
starch granules is known to be affected by the complexation of amylose with lipid
(Tester et al. 2004). Depending on the source, starch lipids comprise free fatty acids
and lysophospholipids. Waxy starches generally have negligible amount of lipids
Introduction
3
(Buleon et al. 1998). Among commercial starches, potato starch is unique in having a
high level of phosphate groups that are covalently linked to the C6 and C3 positions
of the glucose monomers. The high swelling power of the potato starch is due to the
presence of these phosphate groups coupled with the large size of granules.
The presence of low level of protein in the purified starch represents the traces of
biosynthetic enzymes involved in the synthesis of starch.
Table 1.1. Characteristics of native starch granules from common sources (http://members.home.nl/ajansma/zetmeel/infoe/chapter2.htm)
Starch Type Size of grain in µm Shape
Range Average Oval spherical
Potato Tuber 5 – 100 40 Oval spherical
Maize Grain 2 - 30 15 Round polygonal
Wheat Grain 1 - 45 25 Round lenticular
Tapioca Root 4 - 35 25 Oval truncate
Waxy maize Grain 3 - 26 15 Round polygonal
1.1.1. Biosynthesis of starch
Starch is an insoluble polymer of Glucose (Glc) residues synthesized inside plastids of
higher plants. The pathway of starch synthesis has been clarified in the past and
known for many plant species (Preiss 1988; Ball and Morell 2003; James et al. 2003;
Geigenberger et al. 2004; Stitt et al. 2010; Vriet et al. 2010; Zeeman et al. 2010). The
first step of starch synthesis involves the conversion of Glc-1-P and ATP to ADP-Glc
and inorganic pyrophosphate (PPi), catalyzed by ADP-Glc pyrophosphorylase
(AGPase). ADP-Glc acts as the glucosyl donor for different classes of starch
synthases (SS), which elongate the α-1,4-linked glucan chains of the two insoluble
starch polymers amylose and amylopectin. Five distinct SS classes are known in
plants: granule-bound SS, that are responsible for the synthesis of amylose; and
soluble SS I to IV, involved in amylopectin synthesis. Branch points in amylopectin
are incorporated by two classes of starch-branching enzymes (SBE I and II), which
varies in length of the glucan chains transferred and substrate specificities.
Interestingly, starch synthesis also involves two types of debranching enzymes, which
cleave branch points and might be involved in modifying the branched glucans into a
form capable of crystallization within the granule. Genetic studies give evidence that
Introduction
4
the different isoforms of SS, SBE, and debranching enzyme probably play significant
roles in determining the complex structure of starch. Coordination of these enzymes is
required to synthesize the crystalline matrix of the starch granule. Interestingly, SS III
and SS IV have recently been reported to be responsible for starch granule initiation
(Szydlowski et al. 2009).
In most tissues, AGPase is located mainly in the plastid. In leaves, in the
presence of light, Glc-1-P is synthesized from Calvin-Benson cycle intermediates via
plastidic phosphoglucose isomerase and phosphoglucomutase (PGM), in the presence
of ATP supplied by photophosphorylation at the thylakoid membrane (Fig. 1.1.A).
In nonphotosynthetic tissues, such as potato tubers (Fig. 1.1.B), incoming sucrose
(Suc) is mobilized by a number of cytosolic reactions to Glc-6-P, and imported into
the amyloplast in counter exchange with inorganic phosphate (Pi) by a Glc-6-P/Pi
translocator (Kammerer et al. 1998) and lastly converted to Glc-1-P via plastidial
PGM. The second substrate of AGPase, ATP, is provided by mitochondrial
respiration and imported into the plastid through the envelope ATP/ADP exchanger
(Tjaden et al. 1998). However, in cereal seed endosperm, AGPase is situated in the
cytosol, with a overall AGPase activity of about 85% to 95% (James et al. 2003).
ADPGlc synthesized in the cytosol is then imported into the plastid to support starch
synthesis.
1.1.2. Hydrolysis of starch
The starch is mainly consumed after processing for domestic or industrial purposes.
The native starch has limited use mainly as thickener and binder. The hydrolyzed
starch has applications in food, beverage, pharmaceutical, textile and detergent
industries. Till 19th century, acid hydrolysis using dilute HCl was carried out for
starch saccharification, since the understanding of the potential advantages of
biological catalysts was limited. The enzymatic starch processing is advantageous
over chemical starch hydrolysis as the latter has limitations like high temperature
and low pH requirement, low glucose yields, formation of unwanted color, bitter
tasting compounds, and the need for corrosion resistant vessels (Glazer and Nikaido,
1995; Jensen and Olsen, 1999). Today starch saccharification is completely
enzyme based.
Introduction
5
(A) Leaves (B) Heterotrophic tissues
Fig. 1.1. Schematic representation of the pathway of starch biosynthesis, its subcellular compartmentation, in photosynthetic leaves (A) and heterotrophic tissues (B) The reactions of the pathway of starch biosynthesis are catalyzed by the following enzymes: 1, phosphoglucoisomerase; 2, PGM; 3, AGPase; 4, SS; 5, SBE; 6, starch debranching enzyme; 7, inorganic pyrophosphatase; 8, Suc synthase; 9 UDP-Glc pyrophosphorylase; 10, fructokinase; 11, ATP/ADP translocator; 12, Glc- 6-P/Pi translocator; 13, cytosolic AGPase; and 14, ADP-Glc/ADP translocator. (Adapted from Geigenberger 2011)
Gelatinization and retrogradation of starch
Starch is generally subjected to hydrolysis by amylolytic enzymes after gelatinization.
Gelatinization is the process in which starch becomes soluble, binds water and forms
a gel. This process makes starch more easily digestible. The use of starch as a
thickening agent is based on this process. Gelatinization or heating starch suspension
above critical temperature involves tangential swelling of the amorphous regions of
the granule, disruption of the readily ordered structure and eventually opening of the
crystal structures as the polymer chain becomes increasingly hydrated. The swollen
granules are enriched in amylopectin. The linear amylose diffuses out of the swollen
granule during and after gelatinization, and makes up the continuous gel phase outside
the granules. Starch swells up by heating, and it absorbs water and becomes viscous
with the increase in temperature. Gelatinization enhances the chemical reactivity of
inert starch granules towards amylolytic enzymes and has been widely adopted in the
manufacture of starch syrups (Hermansson and Svegmark 1996; Oates 1997)
[Fig. 1.2].
When a starch gel is left for some time, the amylose molecules will lose water
and bind together. A similar process occurs when starch rich products, such as
potatoes, will be stored for a long time. This process of recrystallisation of starch is
called retrogradation (Fig. 1.2).
Introduction
6
Fig. 1.2 Gelatinisation and retrogradation of starch. A: native starch, B: Gelatinised starch, C: Retrogradated starch (Adapted from http://www.food-info.net/uk/carbs/starch.htm)
1.2. Starch hydrolyzing enzymes
Amylolytic enzymes (α-glucanases) hydrolyze the glycosidic linkages in various
α-glucans (Fig. 1.3). They belong to mainly 3 families of glycoside hydrolases (GHs)
[Henrissat 1991]: GH 13 (the α-amylase family) [MacGregor et al. 2001; Kuriki et al.
2006], GH 14 (β-amylases) [Pujadas et al. 1996], and GH 15 (glucoamylases)
[Coutinho and Reilly1997] and they differ from each other by their amino acid
sequences, reaction mechanisms, catalytic machineries and structural folds.
A characteristic feature of the enzymes from the α-amylase family is that they all
employ the α-retaining mechanism but vary broadly in their substrate and product
specificities. These differences can be attributed to the attachment of different
domains to the catalytic core or to extra sugar-binding subsites around the catalytic
site (Table 1.2).
Based on the mode of action, the amylolytic enzymes have been divided into
two broad categories: endoamylases and exoamylases (Fig. 1.3).
Endoamylases: The dextrinogenic or liquefying amylases act randomly on the
α-1, 4 linkages only. As a result of their action, linear and branched oligosaccharides
of various chain lengths (dextrins) are formed. α-Amylase and pullulanase are endo-
acting enzymes.
Exoamylases: The saccharifying and saccharogenic amylases hydrolyze
polysaccharides from the non-reducing end successively liberating short end products.
One type hydrolyzes each α-1,4-gycosidic bond from the non-reducing end to produce
Introduction
7
only glucose (glucoamylases), and another type cleaves every alternate bond to
produce maltose (β-amylases).
Cyclodextrin glycosyl transferase (CGTase) is found only in bacteria,
produces a series of α, β, γ cyclodextrins (rings made up of 6, 7 and 8 glucose units,
respectively, bound by α-1, 4 bonds) from starch, amylose and other polysaccharides.
The CGTases catalyze the coupling reaction by which the rings are opened and
transferred to co-substrates like glucose, maltose or sucrose. The disproportionate
reaction is also catalyzed by the enzyme resulting in the transfer of one or more
glucosyl units between linear oligosaccharides.
Another enzyme, α-glucosidase also known as maltase, is the final enzyme
involved in the breakdown of starch. α-Glucosidase hydrolyses α-1,4 and/or α-1,6
linkages of the saccharides formed by the action of other amylolytic enzymes and
liberate α-D glucose units from the non-reducing ends.
Isoamylase hydrolyzes α-1, 6 linkages in polysaccharides like amylopectin,
glycogen and branched dextrins in an exo-fashion. While pullulanase is a debranching
enzyme, which hydrolyzes α-1, 6 linkages of amylopectin and pullulan, but exhibits
low activity on glycogen.
There are 2 types of pullulanses Type I and Type II. Type I is a bacterial
enzyme and exclusively attacks the α-1, 6 linkages in pullulan in an endo fashion to
yield maltotriose. While Type II pullulanases are mostly found in anaerobic bacteria,
which cleave α-1, 4 and α-1, 6 linkages, and thus cause complete conversion of starch
to small sugars without the requirement of other enzymes.
α-Amylases are extracellular enzymes, which catalyze the hydrolysis of α-1,
4 glycosidic linkages of starch liberating linear and branched oligosaccharides of
varying chain lengths and also glucose, the end products have an α-conformation at
C1 (Antranikian 1992). These are categorized on the basis of end product formation
as maltose-forming (B. acidicola) [Sharma and Satyanarayana 2010], maltotetraose-
forming (Pseudomonas sp. IMD 353) [Fogarty et al. 1994], maltopentaose-
forming (B. cereus NY-14) [Yoshigi et al. 1985] and maltohexaose-forming
(B. stearothermophilus US100) [Ali et al. 2001]. The α-amylase catalyses hydrolysis
of (1-4)-α-D-glucosidic linkages in polysaccharides and successively removes
Introduction
8
α-maltose, maltotetraose, maltopentaose and maltohexaose residues from the non-
reducing ends of the chains in the saccharification of starch.
Cyclodextrin
Fig. 1.3. Schematic representation of the action of different amylolytic enzymes on starch
Based on the degree of hydrolysis of substrate, α-amylases are divided into 2
categories: liquefying and saccharifying. Liquefying α-amylases carry out the rapid
reduction in viscosity of starch pastes without producing free sugars. On the contrary,
saccharifying α-amylases produce free sugars but reduce the viscosity slowly as
compared to liquefying α-amylases. The search for α-amylases with the desired
kinetic properties for diverse applications is encouraged because these will improve
the industrial process in terms of economics and feasibility (Martin et al. 1991a).
Based on the activity at different pH, acidic, neutral and alkaline α-amylases
are also known. The pH optima of α-amylases vary in the range between 2 and 12.
Most of the α-amylases display activity in acidic and neutral range (Pandey et al.
2000). α-Amylase from B. subtilis AX20, B. licheniformis, Micromonospora
melanospora and Geobacillus thermoleovorans display highest activity at pH 6.0, 6.5,
7 and 8, respectively (Najafi et al. 2005; Robyt and Ackerman 1971; Malhotra et al.
2000). α-Amylases with pH optima in acidic range are described in Table. 1.4. Acidic,
neutral and alkaline α-amylases are suited for different industrial applications.
Introduction
9
1.3. Evolutionary relatedness
There is a close evolutionary similarity of the α-amylases from Archaea and plants
from the family GH13 (Fig. 1.4). Leveque et al. (2000) showed that archaeal
α-amylases were closely related to the liquefying α-amylases like B. licheniformis
enzyme and distantly related to saccharifying ones represented by the B. subtilis and
L. amylovorus enzymes indicating that archaeal α-amylases are liquefying in nature.
Fig.1.4. α-Amylase evolutionary tree. Only representative from eubacterial, eukaryal and archaeal enzymes are included. This tree is based on a sequence alignment starting at strand β2 and ending at strand β8 of the (α/β)8-barrel and including the entire B domain (i.e. loop β3→α3). The branch lengths are proportional to the divergence of the sequences of the individual α-amylases. The sum of the lengths of the branches linking any two α-amylases is a measure of the evolutionary distance between them. The α-amylase sources are abbreviated as follows: Aerhy, Aeromonas hydrophila; Altha, Alreromonas haloplanctis; Bacli, Bacillus licheniformis; Bacsu, Bacillus subtilis; Ecoli, Escherichia coli; Lacam, Lactobacillus amylovorus; Stral, Streptomyces albidoflavus; Thtma, Thermotoga maritima; Pyrfu, Pyrococcus furiosus; Pyrsp,Pyrococcus sp. Rt-3; Thchy, Thermococcus hydrothermalis; Thcpr, Thermococcus profundus; Aspor, Aspergillus oryzae; Crysp, Cryptococcus sp.; BarHIG, Barley (high pI isozyme); BarLOW, Barley (low pI isozyme); Drome, Drosophila melanogaster; Chicke, Chicken; HumanS, Human (saliva); PigP, Pig (pancreas); Shrimp, Shrimp (Adapted from Leveque et al. 2000)
Introduction
10
Table 1.2. Enzymes of the α-amylase family that act on glucose-containing substrates, their corresponding EC number, the domain organization as far as it has been described, and main substrates
Enzyme EC number Domains Main substrate
Amylosucrase 2.4.1.4 Sucrose
Sucrose phosphorylase 2.4.1.7 Sucrose
Glucan branching enzyme 2.4.1.18 A, B, F Starch, glycogen
Cyclodextrin glycosyltransferase 2.4.1.19 A, B, C, D, E Starch
Amylomaltase 2.4.1.25 A, B1, B2 Starch, glycogen
Maltopentaose-forming amylase 3.2.1.– A, B, I Starch
α-Amylase 3.2.1.1 A, B, C Starch
Oligo-1,6-glucosidase 3.2.1.10 A, B Amylopectin
α-Glucosidase 3.2.1.20 Starch
Amylopullulanase 3.2.1.41 or 3.2.1.1 A, B, H, G, 1 Pullulan
Cyclomaltodextrinase 3.2.1.54 A, B Cyclodextrins
Isopullulanase 3.2.1.57 Pullulan
Isoamylase 3.2.1.68 A, B, F, 7 Amylopectin
Maltotetraose-forming amylase 3.2.1.60 A, B, C, E Starch
Glucodextranase 3.2.1.70 Starch
Trehalose-6-phosphate hydrolase 3.2.1.93 Trehalose
Maltohexaose-forming amylase 3.2.1.98 Starch
Maltogenic amylase 3.2.1.133 A, B, C, D, E Starch
Neopullulanase 3.2.1.135 A, B, G Pullulan
Malto-oligosyl trehalase hydrolase 3.2.1.141 Trehalose
Malto-oligosyl trehalase synthase 5.4.99.15 Maltose
1.4. Acidstable α-amylases
The demand for high maltose-forming α-amylases is increasing as they have diverse
commercial applications (Fogarty et al. 1993). The α-amylases currently used in
starch processing are active at 95 ˚C and pH 6.8, and stabilized by Ca2+, and therefore,
the process cannot be performed at low pH (3.2-4.5), the pH of the native starch
(Sivaramakrishnan et al. 2006). In order to be compatible with the pH optima of the
enzyme used in liquefaction, the pH of the starch slurry is raised from its native pH
3.2-4.5 to 5.8-6.2, and further, Ca2+ is added to enhance the activity and/or stability of
enzyme. The next saccharification step again requires pH adjustment to pH 4.2-4.5.
Introduction
11
Both these steps (adjustment of pH and removal of salts) need to be omitted, as they
are time consuming and add to the cost of the products (Antranikian 1992). The stress
is, therefore, on extremozymes from extremophiles that are naturally endowed with
the properties required for specialized industrial applications (Satyanarayana et al.
2004) [Fig. 1.5].
IsomerizationGlucose isomerasepH 8.0 at 65 °C for 1 hr
Starch slurry Starch slurry
Ideal process
Maltose syrup
Gelatinization at 105 °C for 5 mins
Liquefaction (α-amylase) pH 6.5 at 95 °C for 2 h (Ca2+- 50ppm)
Dextrins
GlucoamylasepH 4.5 at 60 °C for 48-72 h
Glucose syrup
β amylasepH 5.5 at 55 °C
Fructose syrup
Conventional process
Saccharification
Maltose syrup
Fructose syrup
Liquefaction pH 4.5-5.0At 90-105 ºC, Ca2+- independentNovel enzymes
Dextrins
Glucose syrup
IsomerizationGlucose isomerasepH 4.5 at 80 °C
Fig. 1.5. Conventional and ideal starch processing
α-Amylases are widely distributed among plants, animals and microorganisms.
Among amylases derived from various sources, microbial enzymes are known to
fulfill industrial demands. The microbial sources and characteristics of α-amylases are
shown in Table 1.3. From the estimated 25000 enzymes, approximately 3000
enzymes known to date catalyze different metabolic reactions. Among these, only
fewer than hundred enzymes are used industrially because of their high specificity and
stability. The world market for industrial enzymes is estimated to be around US $ 3
billion dollars (Pandey and Ramachandran 2005), and even more is estimated from
the products obtained from these enzymes. Acid- stable extracellular enzymes are
required as they are having applications in the degradation of polymeric or oligomeric
carbon sources the pH of which lie between 3.2 and 4.5 (Futterer et al. 2004).
The promising properties of enzymes from thermoacidophiles are expected to be
active at low pH and high temperatures, and therefore, these can be used in starch and
Introduction
12
textile industrial processes and in fruit juice industry. The demand for enzymes from
extremophiles may increase in future since they are active under harsh industrial
process conditions.
1.5. Production of α-Amylase
The production of α-amylase in submerged and solid state fermentations has been
studied extensively. The growth and enzyme production by various microorganisms
are affected by a number of physical and chemical parameters like carbon, nitrogen
and phosphate sources, metal ions, temperature, pH, agitation, aeration, inoculum
age and size.
α-Amylases are mostly known to be inducible enzymes. They are induced in
the presence of starch, maltose and other carbon sources like lactose, trehalose, and
α-methyl-D-glycoside. Among different strains of Aspergillus, maltose is a common
inducer that causes increase in enzyme production. Maltose and starch are reported to
be the strong inducers in Aspergillus oryzae (NRC 401013) and A. oryzae (DSM
63303) [Eratt et al. 1984; Lachmund et al. 1993]. Catabolite repression has been
reported by glucose and other sugars. The function of glucose in α-amylase
production is, however, controversial. Xylose or fructose were classified as highly
repressive sugars, although they support good growth in Aspergillus nidulans
(Arst and Bailey 1977). Among various carbon sources starch, fructose, glucose and
rice flour, supported high enzyme production (Ezeji et al. 2005, Prakash et al. 2009).
Carbon sources like glucose and maltose have been used for the production of
α-amylase, but the use of starch remains ubiquitous (Mamo et al. 1999; Sajedi et al.
2005; Liu and Xu 2008; Sharma and Satyanarayana 2011). Industrially important
enzymes have traditionally been produced in submerged fermentation, but recently
these enzymes are being produced by solid state fermentation. Hashemi et al. (2010)
reported the use of wheat bran for the economic production of α-amylase.
The combination of low molecular weight dextran with Tween-80 increased 27-fold
higher α-amylase production (Arnesen et al. 1998). The presence or absence of
various amino acids, and organic and inorganic nitrogen sources is correlated with the
synthesis of amylase in different microbes. For economical production of α-amylase,
soybean meal, casamino acids (Ueno et al. 1987), corn steep liquor (Shah et al. 1990),
and meat extract (Sajedi et al. 2005) have been employed.
Introduction
13
Besides carbon and nitrogen sources, phosphate is a vital requirement for
microbes as it regulates the synthesis of primary and secondary metabolites. Lower
and higher levels of phosphate in the medium significantly affect the growth and
enzyme production (Sharma and Satyanarayana 2011; Hillier et al. 1997; Zhang et al.
1983). Various metal ions like Ca2+, Fe2+, Mg2+, and K+ are added to the production
medium for α-amylase production (Sajedi et al. 2005; Liu and Xu 2008). The
presence of Co2+ in the production medium supported 13-fold higher biomass, but
reduction in the enzyme yield (McMohan et al. 1997).
Among the physical variables, pH of the growth medium acts as an inducer for
morphological change of the organism and indicator for the initiation and termination
of enzyme synthesis. Most of the Bacillus strains produce α-amylase in the production
medium with neutral pH. However, α-amylases produced at acidic pH are also known.
The pH change observed during the growth of the organisms also affects the stability
of the product in the medium. The growth of the organism and enzyme production is
influenced by the temperature. Most amylase studies have been done with mesophilic
fungi within temperature range 25-37 °C while in bacteria it is produced in wider
range of temperature. α-Amylase production has been reported from thermophilic
fungus Thermomonospora fusca (Busch and Stutzenberger 1997) and T. lanuginosus
(Mishra and Maheshwari 1996) and thermoacidophilic bacterium B. acidocaldarius
(Buonocore et al. 1976) and Alicyclobacillus sp. A4 (Bai et al. 2012). Agitation rates
are also known to influence enzyme production as it influences the mixing and
oxygen transfer rates. Agitation rate upto 300 rpm have been employed for the
production of amylases from various microorganisms. The production of amylases by
microbes is considerably affected by physical and chemical parameters of the medium
(Babu and Satyanarayana 1993a; Gigras et al. 2002). Traditionally ‘one-variable-at-a-
time’ approach has been used (Gokhale et al. 1991 and Pham et al. 1998), but it is
time consuming and does not permit understanding interactions among the process
parameters (Wenster-Botz 2000). The statistical Plackett and Burman design, on the
other hand, allows screening of critical culture variables (Sharma and Satyanarayana
2006; Kumar and Satyanarayana 2007), and response surface methodology (RSM)
provides information about the optimum levels of each variable, interactions among
them and their effects on the product yield (Rao and Satyanarayana 2003; Gu et al.
2005). The statistical approaches have been proved to be useful in optimizing medium
Introduction
14
components and cultural variables for maximizing enzyme titres in Bacillus acidicola
(Sharma and Satyanarayana 2011), Bacillus sp. KR 8104 (Hashemi et al. 2010),
Aspergillus awamori (Prakasham et al. 2007) and G. thermoleovorans (Rao and
Satyanarayana 2003, 2007).
1.6. α-Amylase production in fermentor
The effect of environmental conditions on the regulation of extracellular enzymes in
batch cultures is well documented (Amanullah et al. 1999). α-Amylase production and
biomass of B. flavothermus peaked twice and highest production was attained after 24
h in a 20 L fermentor (Kelly et al. 1997). A 70% enhancement in the production of
α-amylase was achieved when G. thermoleovorans was cultivated in a laboratory
fermentor (Rao and Satyanarayana 2003b). The production of α-amylase by B. subtilis
TN106 (pAT5) was enhanced by extending the batch cultivation with fed batch
operation (Baig et al. 1984; Lee and Parulekar 1993). In B. amyloliquefaciens, the
addition of limiting substrate and ammonium resulted in a 2-fold increase in amylase
production (Kole and Gerson 1989). Schwab et al. (2009) had also reported a high
yield of α-amylase in B. caldolyticus using exponential fed fermentation. The use of
fed-batch has an advantage over batch fermentation because the concentration of
limiting substrate is maintained at low level, and thus avoiding the repressing effect of
high substrate concentration and minimizing the accumulation of inhibitory
metabolites (Sharma and Satyanarayana 2011; Schwab et al. 2009; Huang et al.
2004). The reduction in fermentation period was observed in fermentor as compared
to shake flask and high enzyme titres were produced in lesser time in fermentor than
in the shake flasks (Iftikhar et al. 2010; Singh and Satyanarayana 2008).
1.7. Whole Cell Immobilization
A number of biological processes using various biocatalysts such as enzymes,
microorganisms, organelles, plant and animal cells have been investigated to exploit
the advantages of immobilized biosystems. The use of immobilized microbial cells
prevents the laborious and expensive steps involved in the extraction, isolation and
purification of the intracellular enzymes. In case of enzymes bound to subcellular
structures like membranes the stability of the desired enzyme is generally enhanced
by maintaining the natural environment during its operation (Brodelius and
Vandamme 1987). In addition, cofactor requirements and regeneration can be
Introduction
15
achieved in situ using whole cells. In many instances it have been observed that
bound cell systems are more tolerant to environmental changes like temperature, pH,
etc. and less susceptible to toxic substances (Brodelius and Vandamme 1987). The use
of whole cell immobilization allows fermentation on heterogeneous catalysis basis
and the developments in the design and operation of chemical reactions can be
applied to this system also (Brodelius and Vandamme 1987). The reduction in the
equipment size and cost can be achieved by replacing batch fermentation with
continuous column reactors. The continuous system reactor offer advantages like
better process control, reduced operational costs, minimization of fermentation time,
product uniformity, use of high dilution rates without cell washout, higher cell
concentration in reactor, use of high substrate concentration, less product inhibition
and continuous removal of toxic metabolites from the reactor (Chetham 1980;
Brodelius and Vandamme 1987; Corcoran 1985; Furusaki and Seki 1992). A large
number of cell materials or supports are available for whole cell immobilization and
they vary in the quantity and quality of reactive groups, which interact with the cell
surface. Inorganic and organic carriers for microbial systems are known, among them
organic carriers are more commonly being used for many applications as a variety of
reactive groups are present on their surfaces (Brodelius and Vandamme 1987). Three
major classes of organic support are polysaccharides (cellulose, agar/agarose,
carrageenan, alginate, dextran, xanthan gum etc.), proteins (collagen, gelatin, albumin
and fibrin), and synthetic polymers (polyacrylamide, polyurethane, epoxy resin,
polyester, polypropylene etc.). The immobilization techniques like entrapment,
adsorption and cross linking can be used for all three these types of carriers (alumina,
Zirconia, silica, glass, ceramics, sand etc.) and also grafted (supports grafted with
various coupling agents). Adsorption and covalent coupling are two techniques used
for inorganic carriers. Immobilization systems using living cells should be mild
enough to maintain cell viability and activity. Several attempts have been made to
produce enzymes using immobilized cells (Furusaki and Seki 1992). The cells of
Bacillus spp. have been immobilized in matrices such as k-carrageenan (Shinmyo et
al. 1982), alginate (Koshcheyenko et al. 1983), chitosan (Abdel-Naby et al. 2011),
agar (Jamuna and Ramakrishna 1992), agarose (Dobreva et al. 1996) and used for the
production of extracellular enzymes. PUF has been found to be better than other
commonly used matrices for immobilizing bacterial cells because of its high
permeability (Kapoor et al. 2000). The studies on bacteria of the genera Clostridium
Introduction
16
and Thermoanaerobacter immobilized on calcium alginate revealed the significance
of this method for continuous or fed-batch fermentation process to obtain high
enzyme yields. Polyurethane foam is preferable for immobilizing microorganisms as
it allows entrapment of high concentration of cells without a lag phase during their
repeated use (Ghosh and Nanda 1991). The bacterial cells adhere to the surface of
PUF and also partially infuse into the pores. The technique of whole cell
immobilization in PUF has been successfully used for a wide variety of microbial
cells (Beg et al. 2000). G. thermoleovorans and B. acidicola bacterial cells were
immobilized in PUF and were repeatedly used over 15 and 7 cycles, respectively,
with sustained α-amylase secretion (Rao and Satyanarayana 2009; Sharma and
Satyanarayana 2012).
1.8. α-Amylase assays
α-Amylase catalyses the hydrolysis of α-1, 4 glycosidic linkages in starch to produce
glucose, dextrins and limit dextrins. The reaction is examined by an increase in
reducing sugar levels or decrease in the iodine color of the treated substrate under
optimum conditions of pH and temperature. Many methods are available for the
determination of α-amylase activity (Priest 1977), which are based on the decrease in
the intensity of color of starch-iodine complex, increase in reducing sugars,
degradation of color complexed substrate and decrease in viscosity of the starch
suspension.
Dinitrosalicylic acid (DNSA) method is a routinely followed method that
estimates the liberation of reducing sugars by the action of amylase on starch and was
originally described by Bernfeld (1955). The major defect in this assay is slow loss in
the amount of color produced and destruction of glucose by various constituents of
the DNSA reagent. To overcome these limitations in the DNSA reagent, a modified
method for the estimation of reducing sugars is developed (Miller 1959). In the
modified reagent, Rochelle salt was excluded and 0.05% sodium sulfite was added to
prevent the oxidation of the reagent. Since then the modified method has been used
widely to measure reducing sugars without any further alterations in the procedure.
In another method, dextrinizing activity of α-amylases is determined by
soluble starch as a substrate. The reaction is terminated with dilute HCl, and adding
0.1 ml of iodine solution. The decrease in optical density at 620 nm is then measured
Introduction
17
against substrate control. Ten percent decline in absorbance is considered as one unit
of enzyme (Fuwa 1954; Babu and Satyanarayana 1994). Recently a modified
dextrinizing method was suggested for determining the dextrinizing activity
(Nguyen et al. 2002). The major limitation of this assay is interference of media
components such as tryptone, peptone, corn steep liquor and thiol compounds with
starch-iodine complex. The interference of these media components is protected by
the addition of copper sulfate and hydrogen peroxide (Manonmani and Kunhi 1999a).
Further, zinc sulfate was found to be the best for counteracting interference of various
metal ions. Several workers (Hansen 1984; Carlsen et al. 1994) have successfully
used the original assay procedure in combination with flow injection analysis (FIA).
The flow system comprised an injection valve, a peristaltic pump, a photometer with a
flow cell and 570 nm filter and a pen recorder. The samples were allowed to react
with starch in a coil before iodine is added. The absorbance is then read at 570 nm.
This method has various advantages as high sampling rates, fast response, flexibility
and apparatus being simple.
Starch forms a deep blue complex with iodine (Hollo and Szeitli 1968) and
with further hydrolysis of starch, it changes to reddish brown. This method also
determines the dextrinizing activity of α-amylase in terms of decrease in the iodine
color reaction.
Sandstedt Kneen and Blish (SKB) method (Sandstedt et al. 1939) is commonly
used for the assay of amylases used in baking industry. The potency of most of the
commercial amylases is described in terms of SKB units (25 SKB ~ 1000 IU of
saccharogenic activity). This method is used to express the diastatic power of the malt
and not for expressing α-amylase activity alone (Kulp 1993).
Indian pharmacopoeia method is used to estimate α-amylase in terms of grams
of starch digested by a given volume of enzyme. This method involves incubation of
the enzyme preparation in a range of dilutions in buffered starch substrate at 40 °C for
1 h. The solutions are then treated with iodine solution. The tube, without any blue
color, is used to calculate activity as grams of starch digested. This method is
mainly employed for estimating the α-amylase activity in cereals. Besides these,
other assay methods for α-amylase have recently been described briefly by
Gupta et al. (2003).
Introduction
18
A number of methods to differentiate acidic amylase (AA) from that of neutral
amylase (NA) are known. A conventional assay method used for distinguishing
between AA and NA activities is based on the acid-labile property of NA;
preincubation under acidic conditions can inactivate NA activity alone without
modifying AA activities (Suganuma et al. 1997). The remaining activity is that of AA,
when a specific substrate for endo-type amylase is used (Shirokane et al. 1996;
Nagamine et al. 2003).
A new method for distinguishing AA with that of NA (Suganuma et al. 1997)
was developed using 2-chloro-4-nitrophenyl-α-maltotrioside (CNP-α-G3) as a
substrate. Both enzymes, AA and NA, can degrade the substrate at pH 5.4 to release
the CNP group, which is directly observed at 405 nm without the addition of an
alkaline solution. The rate of CNP release is affected by an SCN ion. In the presence
of 500 mM KSCN, the NA reaction rate increases noticeably whereas the AA reaction
rate decreases.
Another alternate method also distinguishes AA from NA (Suganuma et al.
1997). The cleavage pattern of maltopentaose (G5) could be determined by the
analysis of anomer products using HPLC. The column (YMC AQ type) can separate
anomers of products bigger than maltose. The maltose peak can be detected on the
chromatogram, but the peak of its anomer cannot be separated. AA mainly produces
α-anomer of maltotriose (G3). This signifies that AA hydrolyzes G5 mainly at the
third glycoside bond from the non-reducing end. On the contrary, NA produced a
mixture of anomers of G3 that indicates second glycosidic bond cleavage.
1.9. Purification and kinetics of acid-stable α-amylases
Extraction of a protein from the biological environment requires a series of
purification steps, each step removing some of the impurities and making the product
closer to the final purified form. A number of strategies are used for purification of
α-amylases involving conventional as well as the modern fast purification techniques
as listed in Table 1.4. Initial processes include crude fractionation, clarification,
concentration of crude enzyme using processes such as centrifugation, ultrafiltration
(Bohdziewicz 1996) and salt precipitation (Babu and Satyanarayana 1993b). The
regularly used method for the concentration and purification of acid-stable α-amylases
is ammonium sulfate precipitation and ion exchange chromatography (Table 1.4).
Introduction
19
The concentrated protein is then further purified using high-resolution techniques
based on chromatographic and electrophoretic separations. It is designed to remove
aggregates, degradation products and to prepare a solution fit for the final formulation
of the purified enzyme. The commercial use of α-amylases does not require enzyme
in a purified form, but enzyme applications in pharmaceutical and clinical sectors
need high purity. The purified enzyme is also a requirement in studies of structure-
function relationships and biochemical properties. Some purified microbial
α-amylases and their characteristics are listed in Table 1.3.
1.10. Characterization of α-amylase
1.10.1. Substrate specificity
Substrate specificity of α-amylases varies from microorganism to microorganism.
α-Amylases exhibits highest specificity towards starch as compared to other
substrates like amylose, amylopectin, cyclodextrin, glycogen and maltotriose
(Antranikian 1992).
1.10.12. Temperature and pH optima and stability
The α-amylase displays activity in a broad pH range between 2.0 and 12.0
(Vihinen and Mantsala 1989). The pH optima of most of the α-amylases are in the
acidic and neutral range (Pandey et al. 2000). The acid-stable α-amylases are listed in
Table 1.3.
Thermostability of the enzyme is important characteristic and determines
primary structure of the protein. Temperature optima ranging between 45 ˚C and 115
˚C have been observed in α-amylases (Table 1.3)
A number of acid-stable α-amylases have been purified and characterized
from different microorganisms, which exhibited varying physico-chemical properties.
It has been observed that acid-stable α-amylases contain 30% less acidic and basic
amino acids as compared to neutral ones and this avoids the electrostatic repulsion of
charged groups at acidic pH and contributes to the acid stability of proteins
(Schwermann et al. 1994), as the nature of acid-stable amylases depend on isoelectric
point. The pH lower than isoelectric point signifies that the basic amino acids carry
large number of positive charges resulting in the expansion of protein structure that
affects the catalytic center of activity (Liping et al. 2002). Only 18% charged amino
Introduction
20
acids are present in A. niger α-amylase, suggesting the acid resistance of the enzyme
(Zeng et al. 2011).
Most of the known acid-stable amylases lack thermostability at elevated
temperatures, which is a major constraint for their application in starch industry.
Research is in progress to isolate extremophilic microorganisms producing enzymes
bestowed with the desired properties.
1.10.3. Molecular weight
Molecular weights of α-amylases vary between 10 and 210 kDa. Molecular weights of
microbial α-amylases range between 50 and 115 kDa (Table 1.3). The acidic
α-amylase (AA) from A. niger alone has a lower molecular weight than AAs from
other acid-producing molds. Most acid-stable α-amylases are high molecular weight
enzymes as reported in B. stearothermophilus US 100 (Ben Ali et al. 2001),
Lactobacillus manihotivorans (Aguilar et al. 2000), Bacillus sp. WN 11 (Gashaw and
Amare 1999), and Bacillus sp. KR8104 (Sajedi et al. 2005) B. acidicola (Sharma and
Satyanarayana 2012).
1.10.4. Inhibitors
Among inhibitors, heavy metal ions, sulfhydryl group reagents, N-bromosuccinimide
(NBS), p-hydroxy mercuribenzoic acid, iodoacetate, EDTA and EGTA are known to
inhibit α-amylases (Hamilton et al. 1999b). Many α-amylases are inhibited by
Hg2+ (Asoodeh et al. 2010), and this indicates the presence of carboxyl group in
enzyme molecule (Dey et al. 2002). Further, Hg2+ is known to oxidize indole ring and
to interact with aromatic ring present in tryptophan (Zhang et al. 2007; Liu et al.
2010). Inhibition of enzyme activity by NBS demonstrates the catalytic role of
tryptophan (Sharma and Satyanarayana 2012). Dithiothreitol and β-mercaptoethanol
are the reducing agents and suggests the role of –SH groups in the catalytic activity of
enzyme. There are reports where DTT has stimulated and inhibited the activities of α-
amylases (Ballschmiter et al. 2006; Rao and Satyanarayan 2007). In maltogenic α-
amylase from Bacillus sp. WPD616, there was no effect of DTT indicating that –SH
groups are not involved in the catalytic activity or these enzymes have no free and
accessible –SH groups (Liu et al. 2006).
The inhibition of α-amylase by PMSF suggests the role of seryl hydroxyl
group in enzyme catalysis. The inhibition of α-amylase by Woodward’s Reagent
Introduction
21
K (WRK) signifies the chemical modification of aspartic and glutamic acid residues
involved in the active site (Paoli et al. 1997). The inactivation of enzyme by WRK
also indicates the involvement of acidic amino acids in the active site of the enzyme
(Chauthaiwale et al. 1994; Komissarov et al. 1995).
1.10.5. Metal ion and stability of α-amylase
Various cations, substrate and other stabilizers influence thermostability of the
enzymes (Vihinen and Mantasala 1989). α-Amylase is a metal activated enzyme and
has high affinity for Ca2+. The Ca2+ alters the activity and thermal stability of most of
the α-amylases (Dong et al 1997; Khajeh et al. 2001) and it is known that
thermal stability is usually enhanced in the presence of Ca2+ (Laderman et al. 1993;
Neilsen et al. 2003). The number of bound Ca2+ varies from 1 to 10. Usually one
Ca2+ is sufficient to stabilize the enzyme, but the crystalline TAKA amylase A
contains ten Ca2+ but only one is tightly bound (Oikawa and Maeda 1957). Dialysis
against EDTA can remove Ca2+ and also Ca2+ free enzyme can be reactivated with the
addition of Ca2+. Although α-amylase is known to be Ca2+-dependent, there are
reports of Ca2+-independent acid-stable α-amylases that do not require Ca2+ for
stability and activity (Asoodeh et al. 2010; Sajedi et al. 2005; Hmidet et al.2008;
Gashaw and Amare 1999; Rao and Satyanarayana 2007; Sharma and Satyanarayana
2010). The Hg2+ completely inhibited α-amylase activity (Mamo and Gessesse 1999
and Asoodeh et al. 2010), indicating the presence of carboxyl groups in enzyme
molecule (Dey et al. 2002). Further, Hg2+ is also known to oxidize indole ring and to
interact with aromatic ring present in tryptophan (Zhang et al. 2007; Liu et al. 2010).
1.11. α-Amylase gene cloning and expression
Genetic engineering has been used broadly for cloning α-amylase gene from amylase
producing strains. Attempts have been made on cloning of α-amylase genes in
different microbes, mostly in E. coli and Saccharomyces cerevisae. α-Amylase was
one of the first proteins adopted for molecular biological studies because of many
reasons like existence of easy screening assay, availability of amylase negative strains,
knowledge of genetics, protein production and fermentation technology of α-amylase
in B. subtilis. Sajedi et al. (2007) reported α-amylase gene (1328 bp) from Bacillus sp.
KR-8104 designated as KRA encoding 440 amino acids without 20 amino acids of
N and C terminus. The gene encoding 460 aa extracellular α-amylase from
Intr
oduc
tion
22
Tab
le 1
.3. C
hara
cter
istic
s of a
cids
tabl
e α-
amyl
ases
Sour
ce
Mol
ecul
ar
pH
Te
mpe
ratu
re/s
tabi
lity
pI
Km
, Vm
ax, K
cat
R
efer
ence
w
eigh
t
B
acte
ria
Alic
yclo
baci
llus s
p. A
4
64
4.2
75/
75 (>
95%
for 1
h)
-
-
B
ai e
t al.
2012
A.
aci
doca
ldar
ius
1
60
3.
0
75
M
atzk
e et
al.
1997
Ba
cillu
s sp.
US
100
5.
6/4.
5-8.
0 82
/90-
95
-
Ali
et a
l. 19
99
Baci
llus s
p. W
N 1
1
Am
y 1-
76
5.5/
5.5-
9.0
(1h)
75-
80/8
0 (4
h)
- -
Mam
o &
Ges
sess
e 19
99
Am
y 2-
53
Baci
llus a
cido
cald
ariu
s 66
3
.5
70
Km
-0.1
6 m
g m
l-1
K
anno
198
6b
B. c
aldo
lytic
us
70
5
.5
70
/70
(60m
in)
Koc
h et
al.
1987
Ba
cillu
s circ
ulan
s
48
4.
9
48
K
m-1
1.66
, Vm
ax-6
8.97
U D
ey e
t al.2
002
Ba
cillu
s lic
heni
form
is
58
4-
9
90
-
Hm
idet
et a
l. 20
08
Baci
llus s
ubtil
is
53
5-
7
65-7
0 2.
6
Vm
ax-9
09 U
mg-1
N
agra
jan
et a
l. 20
06
Baci
llus s
p. Y
X1
56
5.0
40
-50
-
Liu
and
Xu
2008
Ba
cillu
s sp.
KR
8104
59
4.
0-6.
0
75
-80
-
Saje
di e
t al.
2005
Ba
cillu
s sp.
53
4.5
70
-
A
sood
eh e
t al.
2010
Ba
cillu
s ste
arot
herm
ophi
lus
5.6
80
-
K
hem
akhe
m e
t al.
2009
B.
stea
roth
erm
ophi
lus -
4.6-
5.1
55
-70
4.
82
-
Man
ning
& C
ampb
ell 1
961
Geo
baci
llus s
p. L
H8
52
5-7
80
-
K
haje
h et
al.
2009
La
ctob
acill
us m
anih
otiv
oran
s 135
5.5
55
-
A
guila
r et a
l. 20
00
L. k
onon
enko
ae
76
4.5-
5.0/
5.0-
7.0
70
3
.5
Km
- 0.8
g l-1
Pr
ieto
et a
l. 19
95
Kca
t- 62
2 s-1
Py
roco
ccus
furi
osus
48
5.6
115
Sav
chen
ko e
t al.
2002
Fu
ngi
Aspe
rgill
us a
wam
ori
54
4
.8-5
.0
5
0/40
(60
min
)
-
K
m-1
.0 m
g m
l-1
Bhe
lla &
Alto
saar
198
5 A.
ben
nebe
rgi
50
5
.5
5
0
-
-
A
laza
rd&
B
alde
nspe
rger
198
2 A.
che
valie
ri 68
5.5
4
0/60
(15
min
)
-
Km
-0.1
9 m
g m
l-1
Out
iola
198
2 A.
hen
nebe
rgi
50
5.
5
50/4
0 (1
5 m
in)
-
-
Ala
zard
& B
alde
nspe
rger
198
2 A.
flav
us
-
5.25
/5.0
-8.0
50
/55
(10
min
)
-
- Pe
revo
zche
nko
& T
sype
rovi
ch
Intr
oduc
tion
23
1972
A.
foet
idus
41
.5
5.
0
45/3
5 (6
0 m
in)
K
m-1
.14
(am
ylop
ectin
) Mic
hele
na &
Cas
tillo
198
4
(mg
ml-1
) 2.1
9 m
g m
l-1
(s
tarc
h) V
max
-313
(am
ylop
ectin
)
606
(sta
rch)
174
8 (a
myl
ose)
A.
ory
zae
52
4.0
5
0
4.0
K
m -0
.13
%
Yab
uki e
t al.
1977
A.
nig
er
-
5.0
5
0
-
-
Bhu
mib
ham
on 1
983
A.
nig
er
58
4.
0-5.
0/2.
2-7.
0
3.4
4 -
M
inod
a &
Yam
ada
1963
Fu
sari
um v
asin
fect
um
4.
4-5.
0
45-5
0/50
(30
min
)
-
-
N
aray
anan
&
Sh
anm
ugas
unda
ram
196
7 Pa
ecilo
myc
es sp
. 69
4.0
45
-
Ze
nin
& P
ark
1983
Th
erm
omyc
es la
nugi
nosu
s 61
4.0
80
0
.68
N
guye
n et
al.
2002
T.
lanu
gino
sus
42
5.6
65
/50
(> 7
h)
-
-
Mis
hra
& M
ahes
hwar
i 199
6 Tr
icho
derm
a vi
ride
5.0-
5.5/
4.0-
7.0
60 (1
0 m
in)
-
S
chel
lart
et a
l. 19
76
Yea
st
Cry
ptoc
occu
s fla
vus
84
.5
5.
5
50
0.
056
mg
ml-1
W
ande
rley
et a
l. 20
04
Sa
ccha
rom
yces
cer
evisi
ae
54.1
5.0
50
-
-
De
Mor
aes e
t al.
1999
Intr
oduc
tion
24
Tab
le 1
.4. S
trate
gies
use
d fo
r pur
ifica
tion
of b
acte
rial α
-am
ylas
es
Sour
ce
Purif
icat
ion
stra
tegy
Fo
ld p
urifi
catio
n
Ref
eren
ces
Alic
yclo
baci
llus s
p. A
4
Ultr
afilt
ratio
n w
ith 6
kD
a (M
otia
nmo,
Tia
njin
, Chi
na)
21.
8%
B
ai e
t al.
2012
HiT
rap
SP X
L co
lum
n (I
on e
xcha
nge)
r
ecov
ery
(Am
ersh
am P
harm
acia
, Upp
sala
, Sw
eden
) Ba
cillu
s circ
ulan
s GR
S 31
3
Org
anic
solv
ent f
ract
iona
tion
2.
54
D
ey e
t al.
2002
S
epha
dex
G-1
00, C
M-S
epha
dex
Ba
cillu
s sp.
B3
Aff
inity
chr
omat
ogra
phy
with
alg
inic
aci
d-C
ELB
EDS
Am
ritka
r et a
l. 20
04
Baci
llus s
p. K
R-8
104
Am
mon
ium
sulfa
te p
reci
pita
tion
Saje
di e
t al.
2005
D
EAE-
Seph
aros
e, p
heny
l Sep
haro
se
Baci
llus s
p. W
N11
60%
(NH
4SO
4), D
EAE
Seph
aros
e (p
H 5
.3)
A
my
I- 6
5/13
Mam
o &
Ges
sess
e 19
99
Seph
adex
G-7
5
Am
y II
- 40.
7/9.
5 Ba
cillu
s sp.
YX
-1
A
mm
oniu
m su
lfate
pre
cipi
tatio
n
34
Li
u an
d X
u 20
08
DEA
E Se
phar
ose
fast
flow
S
epha
dex
G-7
5 Ba
cillu
s lic
heni
form
is N
CIB
634
6
DEA
E-C
ellu
lose
DE5
2 (p
H 5
.3)
33/6
6
Mor
gan
& P
riest
1981
La
ctob
acill
us p
lant
arum
A
ffin
ity c
hrom
atog
raph
y, S
epha
rose
6B
(pH
5.5
)
Sa
noja
et a
l. 20
00
Intr
oduc
tion
25
Tab
le 1
.5. E
nzym
es b
elon
ging
to α
-am
ylas
e fa
mily
and
the
four
hig
hly
cons
erve
d re
gion
s. Th
ree
cata
lytic
site
s ar
e in
dica
ted
as b
old.
Num
berin
g of
th
e am
ino
acid
sequ
ence
s of t
he e
nzym
e st
arts
at t
he a
min
o-te
rmin
al a
min
o ac
id o
f eac
h m
atur
e en
zym
e.
E
nzym
e
Ori
gin
Reg
ion
I R
egio
n II
R
egio
n II
I R
egio
n IV
α-A
myl
ase
A. o
ryza
e
117D
VV
AN
H
202G
LRID
TVK
H 2
30E
VLD
29
2FV
ENH
D
CG
Tase
Ba
cillu
s mac
eran
s
13
5DFA
PNH
22
5GIR
FDA
VK
H 2
58E
WFL
32
4FID
NH
D
Pullu
lana
se
Kle
bsie
lla a
erog
enes
600D
VV
YN
H 6
71G
FRFD
LMG
Y 70
4EG
WD
827Y
VSK
HD
Is
oam
ylas
e
Ps
eudo
mon
as a
myl
oder
amos
a
292D
VV
YN
H 3
71G
FRFD
LASV
435
EPW
A
505F
IDV
HD
B
ranc
hing
enz
yme
E.
col
i
335D
WV
PGH
40l
ALR
VD
AV
AS
458E
EST
5
2lLP
LSH
D
Neo
pullu
lana
se
B.
stea
roth
erm
ophi
lus
24
2DA
VFN
H 32
4GW
RLD
VA
NE
357E
IWH
41
9LLG
SHD
α-
Am
ylas
e pu
llula
nase
C
lost
ridi
um th
erm
ohyd
rosu
lfulc
um
488D
GV
FNH
594G
WR
LDV
AN
E 62
7EN
WN
699
LLG
SHD
α-
Glu
cosi
dase
Sacc
haro
myc
es c
arls
berg
enes
is l0
6DLV
INH
2
10G
FRID
TAG
L 27
6EV
AH
34
4YIE
NH
D
Cyc
lode
xtrin
ase
Th
erm
oana
erob
acte
r eth
anol
icus
23
8DA
VFN
H 32
1GW
RLD
VA
NE
354E
VW
H
416L
IGSH
D
Olig
o-1,
6 gl
ucos
idas
e
Baci
llus c
ereu
s
98
DLV
VN
H
195G
FRM
DV
INF
255
EM
PG 32
4YW
NN
HD
D
extra
n gl
ucos
idas
e
Stre
ptoc
occu
s mut
ans
98
DLV
VN
H 1
90G
FRM
DV
IDM
236
ETW
G 30
8FW
NN
HD
A
myl
omal
tase
Stre
ptoc
occu
s pne
umon
iae
22
4DM
WA
ND
29
1IV
RID
HFR
G 3
32E
ELG
39
1YTG
THD
G
lyco
gen
debr
anch
ing
Hum
an
29
8DV
VY
NH
504
GV
RLD
NC
HS
534E
LFT
60
3MD
ITH
D
Enzy
me
Introduction
26
Pyrococcus furiosus was cloned in E. coli. The P. furiosus α-amylase was a
liquefying enzyme with a specific activity of 3,900 U mg at 98°C. It was optimally
active at pH 5.5 to 6.0 and 100 ºC with a half life of 13 h at 98 °C and did not require
Ca2+ for activity (Dong et al. 1997). Another α-amylase gene, Amy N, from
B. licheniformis NH1 was also cloned, sequenced and expressed in E. coli using
pDEST17 expression system. This recombinant α-amylase showed high
thermostability at 85 °C (60 min) as compared to wild type amylase (8 min) [Hmidet
et al. 2008]. The gene encoding acid-stable α-amylase from Aspergillus niger was
cloned in pPIC9K vector and expressed in Pichia pastoris with a very high production
of 2838 U ml-1. The 58 kDa recombinant α-amylase was optimally active at pH 4.0
and 70 ºC (Zeng et al. 2011). A 1920 bp gene encoding 640 amino acids of an acid-
stable α-amylase was cloned from Aspergillus kawachii IF04308. The amino acid
sequence from the N-terminus to the 479th residue showed 97% homology with the A.
niger acid-stable α-amylase. The amino acid sequence in the C-terminal region
between T-502 and T-538 was rich in threonine and serine also known as TS region is
essential for the digestion of raw starch (Kaneko et al. 1996). Four highly conserved
regions are reported in different enzymes of amylase family like α-amylase,
CGTase (Binder et al. 1986), isoamylase, pullulanase (Amemura et al 1988), α-
glucosidase, cyclodextrinase, amylomaltase, neopullulanase (Table 1.5). These
conserved regions contains all of the three catalytic residues and the substrate-
binding residues that bind glucosyl residues adjacent to the scissile linkage in
the substrates by the enzyme, according to the substrate-binding model of Taka-
amylase A, the α-amylase from Aspergillus oryzae, proposed by Matsuura et al.
(1984). Acidophilic protein contains three exchanges in residues that are uniformly
conserved among all members of the enzyme family. The α-amylase gene from
Alicyclobacillus acidocaldarius was expressed in Escherichia coli to find whether
these exchanges are responsible for the acidic pH optimum. The adaptation of protein
to the acidic environment was considered to be due to the reduction of density of the
both positive and negative charges on the surface of the protein; this effect avoids the
electrostatic repulsion of charged groups at acidic pH and contributes to the acid-
stability of proteins (Schwermann et al. 1994). The temperature and pH optima of the
enzyme produced in E. coli were similar to those of the native enzyme (Matzke et al.
1997). The α-amylase encoding gene of an acidophile B. acidicola with N and C
terminal truncation has been cloned recently in pET28a(+) and expressed in E. coli
Introduction
27
(Sharma and Satyanarayana 2012). The 62 kDa recombinant α-amylase was optimally
active at pH 4.0 and 60 ºC.
1.12. Structural conformation studies
Circular dichroism spectroscopy and X-ray crystallography are extensively used
techniques for acquiring information about protein structure and conformation.
The sensitivity of far-UV protein CD spectra to protein secondary structure is used in
one of the most successful applications of CD in determining secondary structure
composition of protein, and also the spectra of protein at different temperatures and
chemical environments is used to study the changes in protein folding. α-Amylases
have 3 domains. A central (α/β)8 TIM-barrel (Fig. 1.6), known as domain A forms the
core of the molecule and consist of three active site residues Asp231, Glu261 and
Asp328 [B. licheniformis α-amylase (BLA) numbering], while domains B and C are
situated at the opposite sides of this TIM-barrel. The amino acid residues in the active
site are strictly conserved but a few positional changes are seen when
B. stearothermophilus α-amylase (BSTA) was superimposed with BLA particularly in
the catalytic residues. This indicates the flexible nature of catalytic residues, playing
important role in catalytic reactions. The C-terminal part of the sequence is present in
domain C and it contains a Greek key motif. Domain B is a projection between the
third strand and the third helix of the TIM barrel and it forms an irregular β-like
structure which is possibly responsible for the differences in substrate specificity and
stability.
Fig. 1.6. Schematic representation of the (β/α)8 barrel (A) and 3D structure of the α-amylase of Aspergillus oryzae or Taka amylase (B), obtained from the Protein Database
Introduction
28
Carboxyl-terminal truncation has been observed in some glycosyl hydrolases
such as α- amylases from B. subtilis, Pseudomonas stutzeri, and α-amylase 1 in malt
[Nakada et al. 1990; Ohdan et al. 1999; Sogaard et al. 1991; Yamane et al. 1984],
while artificial truncation has been performed on various amylolytic enzymes from
Bacillus sp., B. subtilis, and B. stearothermophilus [Lin et al. 1997; Marco et L. 1996;
Vihinen et al. 1994] to study the function of C-terminal region of α-amylase.
The involvement of C-terminal, in translocation of enzyme across the outer membrane
of E. coli as reported in A. haloplanctis (Feller et al. 1998) and Bacillus KR8104
(Ali et al. 2012), binding to raw starch (Rodriguez et al. 2000), and thermal stability
(Vihinen at al. 1994; Marco et al. 1996; Rodriguez et al. 2000) has been
demonstrated. However, Ali et al. (2012) suggests that, the C-terminal truncation did
not affect the thermal stability, optimum pH and end products of starch hydrolysis.
The C-terminal carbohydrate binding domain (CBD) deletion mutant of AA
from Aspergillus kawachii was found to be active under acidic conditions
suggesting that the C-terminal CBD does not affect the acid-stability of the protein
(Suganuma et al. 2007).
Some amylases contain a carbohydrate-binding module (CBM) for the
hydrolysis of insoluble starch. A CBM is an ancillary module of 40 to 200 amino
acids with a discrete fold that possesses carbohydrate-binding activity and is usually
contiguous to a carbohydrate-active enzyme. It does not have catalytic activity; and it
helps in bringing the substrate to the active site in the catalytic domain and
consequently improving hydrolysis (23).
Approximately 10% of the amylolytic enzymes possess a separate domain for
binding to raw starch and it has been found in filamentous fungi, gram positive
bacteria, proteobacteria, actinobacteria and archaea. The starch binding function has
been reported from some glycoside hydrolases, α-amylases, cyclodextrin
glucanotransferases, and acarviose transferases from glycoside hydrolase family
GH13, β-amylases from GH14, and glucoamylases from GH15. Florencio et al.
(2000) and Morlon-Guyot et al. (2001) reported the presence of starch binding
domains (SBDs) in three α-amylases from Lactobacilli. The genes encoding the
α-amylases have been sequenced (Giraud and Cuny 1997; Guyot et al. 2001) and
amino acid sequence analysis of these enzymes showed more than 96% identity and a
structure comprising two discrete functional domains, the N-terminal or catalytic
Introduction
29
domain (GH13) and the C-terminal domain or SBD (CBM-26) formed by direct
tandem repeat units; four modules are reported in Lactobacillus plantarum and
L. manihotivorans α-amylases and five in the L. amylovorus enzyme.
The enhanced affinity in the SBD of the maltohexaose-forming amylase from
B. halodurans is due to the simultaneous interaction of the two tandem CBMs
(carbohydrate binding modules) present in the enzyme (one from family CBM25 and
the other from family CBM26). However, Santiago et al. (2007) reported the
involvement of five-tandem-module SBD not only as distinct modules but also as a
part of the whole amylase.
Three steps are involved in the catalytic mechanism for retaining glycosyl
hydrolases (Sinnott 1990; Davies and Henrissat 1995). Firstly, the protonation of the
glycosidic oxygen by the proton donor (Glu261) followed by a nucleophilic attack on
the C1 of the sugar residue in subsite-1 by Asp231 (Neilson et al. 1999). Once the
aglycon part of the substrate leaves, a water molecule is activated presumably by the
deprotonated Glu261. This water molecule hydrolyses the covalent bond between the
nucleophilic oxygen and the Cl of the sugar residue in subsite-1, thereby completing
the catalytic cycle (Neilson et al. 1999) [Fig. 1.7].
Fig. 1.7. The double displacement mechanism and the formation of a covalent intermediate by which retaining glycosylhydrolases act (Van der Maarel et al. 2002)
The 3-D structure and amino acid sequences of AA from A. niger was
explained by the Novo company group (Boel et al. 1990) that displayed 3-D structure
similar to TAKA amylase A (TAA). Several applications of α-amylases are
performed at different pH values which are different from those where α-amylases act
optimally, and therefore, there is compelling need to change the pH performance
profile of the α-amylases and related enzymes.
The 8-anilino-1-naphthalenesulfonic acid (ANS) binding and light scattering
experiments revealed that at acidic pH, unfolding of B. amyloliquefaciens α-amylase
Introduction
30
(BAA) was observed in such a way that its hydrophobic surface is exposed to a
greater extent in comparison with the native form. In addition, acrylamide quenching
of the intrinsic tryptophan residues in the protein molecules indicate that at pH 3.0,
the protein is in a partially unfolded conformation with more tryptophan residues
exposed to the solvent as compared to the native conformation in the neutral pH.
Ca2+ is required for the refolding of the molten globule state to the native form.
All known α-amylases contain Ca2+ which is situated at the interface between
domains A and B (Boel et al. 1990; Machius et al. 1995; Machius et al. 1998), which
is required for the activity and/or stability of the enzyme. It has also been suggested
that the function of conserved Ca2+-ion is structural (Larson et al. 1994; Machius et al.
1998) as it is distantly located from the active site to participate in the catalysis.
One or more than one Ca2+-ions are present in many structures, and Ca I is strictly
conserved in all distantly related α-amylases and connects domain A to B and helps in
the stabilization of active site structure and controls the formation of the extended
substrate binding site (Machius et al. 1998). The second Ca2+ (Ca II) is situated close
to CaI and in the presence of a sodium ion, Ca-Na-Ca arrangement was observed in
BLA (Machius et al. 1998), BAA (Brzozowski et al. 2000) and BStA
(Suvd et al. 2001). On comparing the metal-containing and metal-free crystal
structures of BLA, it was observed that the loss of metal ion causes numerous
conformational changes around the metal triad and the active site containing 21
residues. In the metal free form of BLA, the segment between residue 182 and 192
contains Asp183, a metal liganding residue, which is completely disordered, while on
the other hand, an ordered large loop like structure is formed upon metal binding.
Residues 178-182 undergo large conformational changes that contribute to the
stabilization of the metal-ligand area by the formation of ionic interaction between
Lys180 and Asp202, another metal liganding residue. Besides Ca2+, chloride ion has
been shown to enhance the catalytic efficiency of the enzyme. The deduced amino
acid sequence of Ca2+-independent α-amylase from Bacillus sp. KR-8104 (KRA)
revealed maximum sequence homology to BAA [85% identity and 90% similarity]
and BLA [81% identity and 88% similarity] α-amylases. The 3D structure of KRA
shows one amino acid substitution in comparison with BLA and BAA in the region
engaged in calcium binding sites, while at the interface of A and B domains and
around the metal triad and active area, many amino acid differences between BLA
Introduction
31
and KRA have been observed. The amino acid differences at the active site cleft and
around the catalytic residues resulted in the shifting of pH profile of KRA in the
acidic range. The shifting of pH activity profile towards acidic pH in the acidic
amylase from Bacillus KR-8104 (KRA) as compared to neutral ones from
B. licheniformis (BLA) may be because of some amino acid substitutions that affect
the putative active site leading to the formation of an extra hydrogen bond between
Glu261 and Arg229 (BLA numbering) [Alikhajeh et al. 2007] {Fig. 1.8}. The
presence of chloride ions at active sites is dominated in mammalian α-amylases
(Larson et al. 1994; Brayer et al. 1995; Ramasubbu et al. 1996).
Fig. 1.8. Depiction and its local hydrogen bonding networks of the real active site of α-amylase of B. licheniformis (BLA) [a], and putative active site of Bacillus KR-8104 (KRA) [b]. Hydrogen bond distances are shown on the picture in terms of Angstrom. Adapted from Alikhajeh et al. (2007)
1.13. Alteration of properties of α-amylases by directed evolution
Naturally occurring enzymes are wonderful biocatalysts with abundant potential
applications in industries and medicine. To be compatible with the specific
requirements for an application, the catalytic properties of the enzyme are required to
be tailored. Directed evolution mimics Darwinian evolution and has emerged as a
powerful tool for engineering enzymes with new or improved functions. It can be
used to modify various enzyme properties like activity, selectivity, substrate
specificity, stability and solubility (Rubin-Pitel and Zhao 2006). Various strategies are
used for directed evolution like error prone PCR, DNA shuffling, staggered extension
process, random priming recombination, heteroduplex recombination, random
Introduction
32
chimera genesis on transient templates, recombinant extension on truncated templates,
incremental truncation for the creation of hybrid enzymes, degenerate oligonucleotide
gene shuffling, random drift mutagenesis, sequence saturation mutagenesis and
nucleotide excision and exchange technology (Sen et al. 2007). The protein
engineering of the α-amylase was also tried to understand the determinants of pH
activity profile. Based on the structural studies, it is difficult to engineer a protein, as
there are many key factors that are responsible for the pH activity profile of
α-amylases. Site directed mutagenesis of α-amylase produced by Bacillus strain was
performed in order to understand the pH activity profile of the enzyme (Declerck et
al. 2000; Nielsen and Borchert 2000). Based on the mutagenic studies, it was
concluded that the modification of dynamics of the active site could be a substitute for
engineering pH activity profile of the protein. The chance of rational engineering of
the enzyme activity is expected to succeed in case the detailed description of enzyme
mobility and dynamics of the active site are available while designing the point
mutations (Neilsen et al. 2001).
The gene-targeted mutants of extremely thermoacidophilic archaea are a major
challenge, but some progress has been made in this area. The mutant of Sulfolobus
solfataricus 98/2 termed S. solfataricus PBL 2025 lacks about 50 genes including
lacS (Schelert et al. 2004). The inability of the mutant to grow on lactose based
minimal media provides a selectable marker (Albers and Driessen 2007). A deficient
mutant of S. solfataricus was used to study the function and regulation of α-amylase
(Worthington et al. 2003).
Presently, starch industry has grown to be the largest market of enzymes after
detergent industry. The properties of starch and α-glucan acting enzymes are altered
by directed evolution as the naturally occurring enzymes from hyperthermophilic
bacteria and archaea are unfit for the unfavorable industrial applications or have
restricted shelf lives. Richardson et al. (2002) found two robust chimeric α-amylases
using DNA shuffling by high throughput screening with superior properties suited for
industrial applications. Error prone PCR (epPCR) was used to improve the
performance of a maltogenic amylase (Novamyl) in baking (Jones et al. 2008).
Site directed mutagenesis and saturated mutagenesis have been employed for tailoring
the pH optimum of a number of enzymes like α-amylase from B. licheniformis
(Verhaert et al. 2002) and soyabean β-amylase (Hirata et al. 2004), but the catalytic
Introduction
33
rate of these enzymes was affected. In contrast, the directed evolution approach
ensures the selection of variants with enough activity at the desired pH. Liu et al.
(2008) reported the enhancement of acid-stability of α-amylase of B. licheniformis
CICC 10181 by directed evolution. The mutations at two crucial positions Leu134 and
Ser320 together affected the acid resistance of the enzyme.
α-Amylase catalyzes the enzyme substrate reaction at low pH by the
protonation of the nucleophile (D231) and at high pH by deprotonation of the hydrogen
donor (E261), and the correlation between activity of enzyme and pH is determined by
pKa values of these two active site groups (Kyte 1995). The pKa value of the amino
acid residue depends on the free energy difference between the neutral and the
charged states of the residue in the protein, and this difference in free energy is
influenced by the desolvation effects and by the charges and dipoles in the protein and
the substrate. The pKa value of the residue is lower when it is placed in a positive
environment (Nielsen et al. 2001). At most physiological pH values, arginine with a
guanidyl group is expected to attain a positive charge. Therefore, positioning of a
positive charge at a distance from the nucleophile (D231) was assumed to stabilize the
negative charge on this aspartate residue and to reduce its pKa, thereby stabilizing its
deprotonated form. This resulted in a shift of the acidic limb to more acidic values for
mutant L134R and improving the activity and stability of the enzyme at acidic pH.
In the interior of domain A, Ser320 is the first residue of β-strand 7.
As compared to Ser, which has a strong tendency with forming β-turn, Ala is a small
residue which usually exists β-sheet forming. Protein unfolding is a mutual process
and the most energy consuming step is the breakage of the polar contacts to
neighboring residues of the first residue in a α-helix or a β-strand. Therefore, Ala320
could be situated at important position for primary unfolding of the molecule. The Ser
to Ala substitution, replaces a polar residue interiorly by a more hydrophobic residue,
and thus, expected to stabilize the protein. Ser320 is engaged in a hydrogen bonding
network: Ser320 ↔ Asp285 and Ser320 ↔ Tyr358 (Fig. 1.9). On the other hand, Ala320
only gives a hydrogen bond to Tyr358 without Asp285. Asp285 is more solvent
accessible as the hydrogen bond provided with Ala320 is lost. The pKa of Asp285 was
increased slightly by the more solvent accessibility resulting in the change of the
electrostatic fields in the protein. At most physiological pH values, Asp285 near the
hydrogen donor (E261) is expected to bear a negative charge and this in turn stabilizes
Introduction
34
the protonated form of Glu261. The higher pKa of Asp285 was responsible for
destabilizing the protonated form of Glu261 that can lead to decrease of the pKa of
Glu261. Therefore, a shift in the basic limb of the pH activity profile for mutant S320A
toward acidity was expected which in turn was effective in increasing stability at low
pH. The combined effect of the double mutated L134R/S320A indicates that the amino
acids inserted at each site contribute independently to the overall stability of the
protein, as generally seen for stabilizing mutations in protein structures (Matsumura et
al. 1986; 1989; Pantoliano et al. 1989; Serrano et al. 1993). The changes in the
electrostatic field due to charged groups play a significant role in determining the
stability of BLA in strongly acidic environment.
Fig. 1.9. The position of the point mutation in wild type α-amylase gene from B. licheniformis CICC 10181. Domain A, blue; domain B, green; domain C, gray. The active site acids Asp231, Glu261, Asp328 are showed in red. The positions of the point mutation are shown in yellow (Adapted from Liu et al. 2008)
1.14. Industrial applications of α-amylases
Presently, amylases have the major world market share of enzymes (Aehle and Misset
1999). Many amylase preparations are available with various enzyme manufacturers
for specific use in different industries (Fig. 1.10). A detailed account on commercial
applications of α-amylases has been provided by Godfrey and West (1996). Bacterial
amylase is generally preferred over fungal amylase due to a number of characteristic
advantages that it offers (Hyun et al. 1985; Babu and Satyanarayana 1994;
Malhotra et al. 2000). Acid-stable α-amylases can be preferred as their application
minimizes contamination risk.
Introduction
35
Laundry and detergents
Paper Industry
Baking Industry
Sugar Industry
Brewing Industry
Textile Industry
Fig. 1.10. Applications of α-amylase
1.14.1. Starch liquefaction and saccharification
Starch is converted into high fructose corn syrups (HFCS) with help of biocatalysts.
Due to their high sweetening property, these are used in huge quantities in the
beverage industry as sweeteners for soft drinks (Guzman et al. 1995; Crabb
and Mitchinson 1997). The acid-stable, Ca2+-independent α-amylases are preferred
over the currently used enzyme in starch processing, as the latter is active at 95 ˚C and
pH 6.8, and stabilized by Ca2+, and therefore, the process cannot be performed at low
pH (3.2-4.5), the pH of the native starch (Sivaramakrishnan et al. 2006). The stress is,
therefore, on extremozymes from extremophiles that are naturally endowed with the
properties required for specialized industrial applications (Satyanarayana et al. 2004).
Amylolytic enzymes that produce specific malto-oligosaccharides in high yields from
starch have gained significant attention. Such enzymes are widely used in the food,
chemical and pharmaceutical industries (Nigam and Singh 1995). Although
maltogenic α-amylases that yield 53-80% maltose have been reported from
Actionobacteria (Kelly et al. 1993), their industrial potential is limited because of
their moderate thermostability and Ca2+ requirement. The use of Ca2+-independent
enzymes in starch hydrolysis eliminate the addition of Ca2+ in starch liquefaction and
its subsequent removal by ion exchangers from the product streams (Pandey et al.
2000; Van der Maarel et al. 2002).
Introduction
36
1.14.2. Baking Industry
For, decades, enzymes such as malt and microbial α-amylases have been widely used
in the baking industry (Hamer 1995; Si 1999, Kumar and Satyanarayana 2008). These
enzymes were used in bread and allied products to give high quality products having
better color and softer crumb. Many enzymes such as proteases, lipases, xylanases,
pullulanases, pentosanases, cellulases, glucose oxidase, lipoxygenase and others are
being used in the bread industry for different purposes, but none had been able to
replace α-amylase.
Bread is an important food item and a major constituent of balanced diet all
over the world. The shelf-life of bread is short and thus lead to major financial loss to
both customers and bakers. On storage, bread deteriorates as the crumb becomes dry
and firm, the firmness of bread crumb increases with the evaporation of water
from the surface of sliced bread. The complex physico-chemical changes (staling)
result in the loss of flavor and crispiness of bread. It is accepted that the
retrogradation/recrystalisation along with water migration are important factors for
the firming of the bread (Zobel and Kulp 1996; Gray and Bemiller 2003).
The importance of retrogradation of starch fraction in bread staling has, therefore,
been highlighted (Kulp and Ponte 1981; Gupta et al. 2003). To prevent the staling of
bread and other baked goods and to improve its texture and shelf-life, the dough is
supplemented with various additives (Pritchard 1992). Supplementation of various
enzymes as flour additives are used as dough conditioners and are considered to be
safer replacements of chemical ingredients. The bacterial maltogenic α-amylases with
intermediate thermostability are known to act as antistaling agents, thereby reducing
the crumb firmness during storage (Hebeda et al. 1991; Kumar and Satyanarayana
2008) by shortening the amylopectin chain length and production of malto-
oligosaccharides (DP 2-12) [Qi Si and Simmonsen 1994] and allowing the yeast to act
continuously during dough fermentation and early stages of baking. The
supplementation of α-amylases to the dough improves the crumb grain, volume,
texture, flavor and shelf-life of the bread (Van Dam and Hille 1992; Rao and
Satyanarayana 2007).
1.14.3. Bioethanol production
For thousands of years, ethanol has been produced for human consumption, and for at
least a thousand years it has been possible to make concentrated alcoholic drinks by
Introduction
37
means of distillation. In recent years, the attention has turned again to the production
of ethanol for chemical and fuel purposes by fermentation.
Traditionally, ethanol fermentation depends on sugar-rich substrates, mainly
sugarcane, as their carbohydrate is in fermentable form. However, sugarcane is an
expensive material and not continuously available as it is a seasonal crop (De Moraes
et al. 1995). Thus there are great economic benefits in expanding the substrate range
of ethanol-fermenting microorganisms so that the ethanol may be produced from
cheap substrates such as starchy crops (Coombs 1984) and cellulosic materials (Hung
and Chen 1989; Szczodrak and Targonski 1989).
Ethanol-fermenting microorganisms such as S. cerevisiae and Zymomonas
mobilis lack amylolytic enzymes and are unable to directly convert starch into
ethanol. Traditionally, starch is hydrolysed enzymatically into fermentable sugar via
liquefaction and saccharification processes prior to ethanol fermentation (Rao and
Satyanarayana 2006).
In the process currently employed on industrial-scale, ethanol production from
starchy materials involves enzyme hydrolysis. The liquefied starch is hydrolyzed to
glucose with a saccharifying enzyme glucoamylase, and glucose is fermented to
ethanol by the yeast. Acid-stable α-amylase can be used for hydrolyzing starch since
the pH of native starches is acidic.
1.14.4. Miscellaneous applications
Among starch-hydrolyzing enzymes (α-amylase, pullulanase, cyclodextrin
glucosyltransferase, and maltogenic amylase) used in various industrial applications,
α-amylase is the widely used enzyme. Besides their use in the saccharification or
liquefaction of starch, these enzymes are also used in the preparation of viscous and
stable starch solutions used for the warp sizing of textile fibers, the clarification of
haze formed in beer or fruit juices, and in the animal feeds for improving the
digestibility. The new area of application of α-amylases is in the fields of laundry,
textile desizing and dish-washing detergents. The present trend among consumers is
to use lower temperatures for doing the laundry or dishwashing. The removal of
starch from porcelain has become more problematic. Detergents supplemented with
α-amylases which are optimally active at moderate temperatures and alkaline pH can
solve this problem (Van der Maarel et al. 2002).
Introduction
38
α-Amylase is one of the most important industrial enzymes employed in the
starch processing industry for the production of starch hydrolysates. The pH of starch
is 3.2 – 4.5, and therefore, thermostable acidic and Ca2+-independent α-amylases suit
better in the conversion of starch to various sugar syrups. Acidic α-amylases are
known to b produced by bacteria, archaea and fungi. α-Amylases are also used in the
removal of starch in beer, fruit juices, and from textiles and porcelain. The maltogenic
amylase is used as an antistaling agent in order to prevent the retrogradation of starch
in bakery products. α-Amylases are now gaining importance in biopharmaceutical
applications too. Their application in food and starch based industries is the major
market, and the demand for α-amylases would be expected to rise in future too.
The potential applications of α-amylases are immense, especially in starch
saccharification process and baking. The commercial demands are so enormous that
even small improvements in production and catalytic efficiency could be beneficial.
Enzymes used today in starch processing have varying temperature and pH
requirements according their thermostability and physicochemical properties.
Performing starch liquefaction and saccharification in similar conditions of pH and
temperature would decrease the cost of glucose, maltose and fructose production.
Although, several α-amylases have been isolated, cloned and characterized, the
α-amylase story is still incomplete due to the unavailability of α-amylase possessing
activity at low pH, thermostability at high temperature (95 ºC) and Ca2+-independence.
Elucidating the three dimensional structures of these unique α-amylase would help in
understanding the physiological and biochemical basis of their adaptation to extreme
conditions, which could further be exploited for tailoring the enzyme to suit the
process parameters.
1.15. Objectives of the present investigation
In view of foregoing discussion, the present investigation was planned and carried out
with following objectives:
Selection of a bacterial isolate producing acid-stable, thermostable, Ca2+-independent α-amylase.
Production optimization, characterization and structure-function analysis of acidic α-amylase
Cloning and expression of α-amylase encoding gene from B. acidicola. Testing the applicability of enzyme in starch saccharification and baking.
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