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.