NC State
BioResources
Lim, S. J., Oslan, S. H., and Oslan, S. N. (2020). "Purification and characterisation of thermostable α-amylases from microbial sources," BioRes. 15(1), Page numbers to be added.

Abstract

α-Amylases (E.C 3.2.1.1) hydrolyse starch into smaller moieties such as maltose and glucose by breaking α-1,4-glycosidic linkages. The application of α-amylases in various industries has made the large-scale productions of these enzymes crucial. Thermostable α-amylase that catalyses starch degradation at the temperatures higher than 50 °C is favourable in harsh industrial applications. Due to ease in genetic manipulation and bulk production, this enzyme is most preferably produced by microorganisms. Bacillus sp. and Escherichia coli are commonly used microbial expression hosts for α-amylases (30 to 205 kDa in molecular weight). These amylases can be purified using ultrafiltration, salt precipitation, dialysis, and column chromatography. Recently, affinity column chromatography has shown the most promising result where the recovery rate was 38 to 60% and purification up to 13.2-fold. Microbial thermostable α-amylases have the optimum temperature and pH ranging from 50 °C to 100 °C and 5.0 to 10.5, respectively. These enzymes have high specificity towards potato starch, wheat starch, amylose, and amylopectin. EDTA (1 mM) gave the highest inhibitory effect (79%), but Ca2+ (5 mM) was the most effective co-factor with 155%. This review provides insight regarding thermostable α-amylases obtained from microbial sources for industrial applications.


Download PDF

Full Article

Purification and Characterisation of Thermostable α-Amylases from Microbial Sources

Si J. Lim,a Siti Nur Hazwani-Oslan,d and Siti N. Oslan a,b,c,*

α-Amylases (E.C 3.2.1.1) hydrolyse starch into smaller moieties such as maltose and glucose by breaking α-1,4-glycosidic linkages. The application of α-amylases in various industries has made the large-scale productions of these enzymes crucial. Thermostable α-amylase that catalyses starch degradation at the temperatures higher than 50 °C is favourable in harsh industrial applications. Due to ease in genetic manipulation and bulk production, this enzyme is most preferably produced by microorganisms. Bacillus sp. and Escherichia coli are commonly used microbial expression hosts for α-amylases (30 to 205 kDa in molecular weight). These amylases can be purified using ultrafiltration, salt precipitation, dialysis, and column chromatography. Recently, affinity column chromatography has shown the most promising result where the recovery rate was 38 to 60% and purification up to 13.2-fold. Microbial thermostable α-amylases have the optimum temperature and pH ranging from 50 °C to 100 °C and 5.0 to 10.5, respectively. These enzymes have high specificity towards potato starch, wheat starch, amylose, and amylopectin. EDTA (1 mM) gave the highest inhibitory effect (79%), but Ca2+ (5 mM) was the most effective co-factor with 155%. This review provides insight regarding thermostable α-amylases obtained from microbial sources for industrial applications.

Keywords: Purification; Characterisation; Thermostable; α-Amylase; Microorganism

Contact information: a: Department of Biochemistry, Faculty of Biotechnology and Biomolecular Sciences, Universiti Putra Malaysia, 43400 UPM Serdang, Selangor; b: Enzyme and Microbial Technology Research Centre, Centre of Excellence, Universiti Putra Malaysia, 43400 UPM Serdang, Selangor; c: Institute of Bioscience, Universiti Putra Malaysia, 43400 UPM Serdang, Selangor, Malaysia; d: Bioprocessing and Biomanufacturing Research Centre, Faculty of Biotechnology and Biomolecular Sciences, Universiti Putra Malaysia, 43400 UPM Serdang, Selangor, Malaysia; currently at School of Biology, Faculty of Applied Sciences, Universiti Teknologi MARA, Cawangan Negeri Sembilan, Kampus Kuala Pilah, 72000 Kuala Pilah, Negeri Sembilan, Malaysia;

* Corresponding author: snurbayaoslan@upm.edu.my

INTRODUCTION

The International Union of Biochemistry (IUB) establishes the categorisation of enzymes into six different classes, based on the mechanism of enzyme action. They are E.C 1 oxidoreductases, E.C 2 transferases, E.C 3 hydrolases, E.C 4 lyases, E.C 5 isomerases, and E.C 6 ligases. Amylases are enzymes that hydrolyse the glycosidic linkages in starch, and are thus categorised in the class of E.C 3 hydrolases. Amylases can be categorised into endo- and exo-amylases as well as 3 classes including α-, β-, and γ-amylases, catalysing the hydrolysis of α-1,4 and α-1,6-glycosidic bonds in starch, yielding a variety of disaccharides and monosaccharides.

Microorganisms, especially bacteria have proven to have short generation time and are one of the main sources of α-amylase. Thermophilic, mesophilic, and extremophilic bacteria are good sources for thermostable α-amylases. These enzymes work optimally at extreme temperatures.

Saccharomyces cerevisiae (an edible yeast) as well as other fungi (Aspergillus oryzae) and bacteria (Bacillus licheniformis and Bacillus stearothermophilus) have been used to produce α-amylase especially in the food industry because of its “Generally Recognised as Safe” (GRAS) status honoured by the U.S. Food and Drug Administration (FDA) (Nevoigt 2008).

Many purification methods have also been established to purify α-amylases from microbial sources. The methods are ultrafiltration, salt precipitation, dialysis, and column chromatography. These methods give different yields and folds of purification. Characterisation of α-amylases from microbial sources, in terms of optimum temperature, optimum pH, thermostability, and pH stability has become important in determining their related applications as biocatalysts in many processes in industrial fields.

This review article provides an overview on microbial sources of thermostable α-amylases. Purification methods and characterisation of microbial extracellular thermostable α-amylases in terms of optimum temperature and pH, thermostability and pH stability, substrate specificity as well as effects of metal ions and inhibitors are also focused in this article. However, information on purification and characterisation of non-thermostable and non-microbial α-amylases are excluded.

AMYLASE AS BIOCATALYST

Amylases are biological catalysts or enzymes that catalyse the hydrolysis of starch; thus, they are categorised in the E.C 3 class of hydrolases. Amylases are classified into two groups, namely endo- and exo-amylases, depending on their mode of action. Endo-amylases randomly hydrolyse α-1,4-glycosidic linkages in the amylose or amylopectin of starch, yielding linear and branched oligosaccharides of different chain lengths. Exo-amylases only hydrolyse starch from the non-reducing end, forming short end products successively. Table 1 summarises the class, glycosidic bond specificity, mode of action, and products of amylases.

α-Amylase or glucan-1,4-α-glucanohydrolase (E.C 3.2.1.1) is a starch degrading, calcium metalloenzyme that hydrolyses starch into smaller moieties such as maltose and glucose (Singh et al. 2016). This endo-amylase catalyses the internal hydrolysis of α-ᴅ-1,4-glycosidic linkages in the starch to yield small molecular weight carbohydrate moieties of α-glucose, α-maltose, and α-limit dextrin (Singh and Guruprasad 2014). These hydrolysed products have their functional hydroxyl group (-OH) in the α-configuration; hence, this enzyme is named α-amylase.

β-Amylase (glucan-1,4-α-maltohydrolase; glycogenase; saccharogen amylase, E.C 3.2.1.2) is an exo-amylase that catalyses the hydrolysis of α-1,4-glycosidic linkages of starch, producing β-maltose and β-limit dextrin (Oktiarni et al. 2015). This exo-amylase is not synthesized by animal tissues but present in microorganisms contained in the digestive tract. γ-Amylase (glucan-1,4-α-glucosidase; amyloglucosidase; exo-1,4-α-glucosidase; glucohydrolase, E.C 3.2.1.3) can act as exo- or endo-amylase due to its ability to hydrolyse both α-1,4 and α-1,6-glycosidic linkages. However, γ-amylases have the optimum of pH 3 and are most efficient in acidic environments (Saini et al. 2017).

Table 1. Classification of Amylases (Singh et al. 2016)

THERMOSTABLE α-AMYLASE

Thermostable α-amylases are relatively stable at high temperature. Most studies focus on the purification and characterisation of thermostable α-amylase secreted from bacteria, but not from fungi and yeast. Thermophilic bacteria are the most commonly used as α-amylase producers as they can survive in high temperature and produce enzymes having optimum temperatures higher than 50 °C.

Thermostability is crucial in industrial applications, as most processes are optimally performed at elevated temperature, where thermostable enzymes are not deactivated by heating the mixture to a certain temperature over a period due to their high denaturing temperature, unlike the mesophilic enzymes. Thermostable enzymes can be stored at room temperature, thus lowering the costs (Straathof and Adlercreutz 2014). There are three steps in starch hydrolysis, which are gelatinization, liquefaction, and saccharification. The gelatinization of starch is industrially carried out at 110 °C; thus thermophilic and extremophilic α-amylases are preferred for their efficiency and economical value (Zhang et al. 2017).

A novel α-amylase has been discovered in the strain of Bacillus licheniformis B4-423, exhibiting the optimal activity at 100 °C and pH 5.0. The enzyme is stable over a wide pH range (4.0 to 10.0) and exhibits more than 90% activity from 20 °C to 80 °C (Wu et al. 2018). Because of these favourable properties, the thermostable enzyme has been applied in many production processes such as wine brewing and fermentation, baking and food processing, the pulp and paper industry, and detergent treatment systems. Table 2 shows the optimum temperature, thermostability, and potential industrial applications of microbial α-amylases.

Table 2. Microbial Thermostable α-Amylases and their Industrial Applications

MICROBIAL SOURCES OF THERMOSTABLE α-AMYLASE

α-Amylase can be extracted from many sources such as animals, plants, and microorganisms. It is preferred to be industrially extracted and purified from microorganisms, especially bacteria and fungi. Microbial α-amylase can be easily isolated and selected using substrate specificity, serial dilution, and extreme conditions such as temperature and extreme pH. The desired α-amylase properties for specific industrial applications can be designed and improved due to the advancement of genetic engineering and media optimization (Xie et al. 2014).

Gandhi et al. (2015) stated that the main reasons for selecting microorganisms as sources of enzymes are the physiologically and physicochemically controlled access of microorganisms, higher product yield than other sources, convenient and easy recovery in downstream processes, and cost benefits in processing. Moreover, having microorganisms as expression systems of α-amylase is beneficial because of inexpensive media, great adaptability, not affected by seasonal fluctuations, more stability, and catalytic variation compared with other sources (Borrelli and Trono 2015).

Fungus is a preferred source compared with other microbial sources because fungal α-amylases have more accepted GRAS status (Gupta et al. 2003). Espargaró et al. (2012) also stated that bacteria such as Escherichia coli forms inclusion bodies (IBs) containing infectious prion if it is used as expression host for yeast proteins. As a eukaryotic expression host, yeast has its post-translational modifications (PTMs) more similar to higher level eukaryotes than bacteria (Ahmad et al. 2014).

Although it is beneficial as a eukaryotic expression system, there has not been much research performed to purify and characterise α-amylase from yeast. Gandhi et al. (2015) expressed and characterised recombinant SR74 recombinant α-amylase in Komagataella phaffii GS115 with the SR74 α-amylase gene transformed from Geobacillus sp. SR74 using the vector of pPICZαB/SR74 α-amylase. A higher yield of α-amylase from K. phaffii GS115 was recorded than in E. coli transformed by Kassaye (2009) using pET-32b/α-amylase as a vector. However, the expression of SR74 α-amylase in K. phaffii GS115 under the regulation of alcohol oxidase (AOX) promoter required high methanol concentration (1% (v/v) every 24 h) to induce the expression for 120 h. Thus, Nasir (2019) has cloned the gene into pFLDα expression vector under the control of formaldehyde dehydrogenase (FLD1) promoter before transforming into a new yeast expression system, i.e.Meyerozyma guilliermondii strain SO (Oslan et al. 2012). Optimization was performed and highest production was found after 12 h of cultivation without any inducers.

In a study concerning marine yeast isolation and industrial applications conducted by Zaky et al. (2014), enzymes from marine yeast (Aureobasidium sp. and Pichia sp.) are expected to have high salt tolerance, thermostability, barophilicity, and cold adaptivity as the yeasts live in high salinity environment. M. guilliermondii has been used as the research model organism named “flavinogenic yeasts”, being capable of riboflavin over-synthesis during starvation for iron as well as the expression system of thermostable T1 lipase gene (Sibirny and Boretsky 2009; Oslan et al. 2015; Abu et al. 2017). Table 3 shows the sources of microbial α-amylase from different expression hosts and its mode of production.

Table 3. Sources of Microbial Thermostable α-Amylases

PURIFICATION OF MICROBIAL EXTRACELLULAR α-AMYLASE

Enzyme purification is crucial in obtaining a pure enzyme fraction from an impure enzyme crude extracted from available sources. Without enzyme purification, protein and enzyme activity cannot be characterised accurately due to the impurities in the crude extract, resulting in faulty information and data. The α-amylase gene must be overexpressed in the induction medium before purification is conducted. For every purification step performed, total protein content, total activity, specific enzyme activity, yield, and purification fold are calculated to indicate the effectiveness of the steps taken.

Ultrafiltration

Ultrafiltration is a widely used technique in concentrating and purifying proteins by their molecular weight (Mw). The most commonly used filtration membranes are of 10-kDa and 30-kDa molecular weight cut-off membranes. This technique is usually equipped before or after ammonium sulfate precipitation. Before being subjected to ammonium sulfate precipitation, the crude α-amylase expressed in Bacillus subtilis KIBGE HAS was filtrated twice against 100-kDa and 30-kDa molecular weight cut-off (MWCO) ultrafiltration membrane, whereby 3.4-fold purification and 20.61% yield recovery were obtained (Bano et al. 2011). While purifying α-amylase expressed in Anoxybacillus sp. YIM 342, the crude enzymes were subjected to an Amicon ultrafiltration cell with 3-kDa MWCO membrane. The yield of 82% and 1.33-fold purification were reported after ultrafiltration technique (Zhang et al. 2016).

An example of ultrafiltration after ammonium sulfate precipitation was performed by Baltas et al. (2016). The work involved purifying α-amylase expressed in a thermophilic Anoxybacillus thermarum A4 strain. After the precipitation of salt was suspended in MOPS buffer, the enzyme solution was washed and subjected to an Amicon ultrafiltration membrane with the MWCO of 30 kDa. A 75.2% yield recovery as well as 4.4-fold purification were reported with this ultrafiltration technique after performing salt precipitation (Baltas et al. 2016). Similarly, after performing salt precipitation, enzyme solution containing α-amylase expressed in Talaromyces pinophilus 1-95 was concentrated using a 10-kDa MWCO ultra-filtration membrane with 80.13% yield recovery and 1.77-fold purification being reported (Xian et al. 2015).

Salt Precipitation and Desalting

Salt precipitation is a technique to purify proteins from the crude enzymes by increasing the salt concentration gradually. The most common salt used in this method is ammonium sulfate, (NH3)2SO4. Precipitation is started by salting in, i.e., adding (NH3)2SO4 salt into the crude enzymes slowly in a conical flask on a magnetic stirrer until all salt has dissolved completely.

While adding salt into solution, the increase in water surface tension increases the hydrophobic interaction between proteins and water, resulting in the folding of protein to decrease the contact surface area of the proteins to the solvent. Finally, the proteins are precipitated. The saturation of (NH3)2SOused in precipitation is majorly dependent on the molecular weight of the proteins, where low molecular weight protein, e.g., IL-1β (17.5 kDa), requires higher salt concentration compared with IgG (150 kDa) with the addition of 40% to 45% saturation (NH3)2SO4 (Wingfield 2016).

Table 4. Salt Precipitation and Desalting of Microbial α-Amylases

While purifying a 44.0 kDa α-amylase from B. methylotrophicus P11-2, Xie et al. (2014) added solid (NH3)2SOwith 80% saturation under gentle stirring, and the suspension was centrifuged at 10,000 rpm for 30 min at 4 °C after incubation at 4 °C overnight. The percentage yield of α-amylase was 70.8% with a 2.3-fold purification and a specific activity of 57.6 U/mg. However, when Karim et al. (2018) were precipitating α-amylase expressed from A. flavus NSH9, the percentage yield of the enzyme was only 30.7% with 1.84 purification fold and a specific activity of 34.8 U/mg. It was interesting when Du et al. (2018) performed salt precipitation at 70% saturation on the crude enzyme containing α-amylase expressed from Bacillus amyloliquefaciens BH072, but the pellet was dissolved and dialysed against sterile deionized water overnight. Such desalting technique was still able to achieve the purification fold of 3.23 as well as a yield of 71.1% which was high on average. This might be caused by the purified α-amylase exhibited its optimal activity at pH 7 (neutral). Even though salt precipitation cannot lead to highly purified protein, this technique can eliminate some unwanted protein and concentrate the sample.

Referring to Table 4, the precipitation and purification α-amylases from different microbial expression hosts were performed at the salt concentrations ranging from 33 to 90%, but the most common concentration used was 80%. However, the results reflected that α-amylases produced by bacteria required higher salt concentration compared to fungi. This phenomenon might be due to higher solubility and stronger interaction between bacterial α-amylases with water molecules compared to fungal α-amylases.

The most commonly used desalting technique is dialysis, depending on the buffers used to dissolve the pellet. Dialysis is the step following salt precipitation. It removes the salt after the pellet from post-precipitating centrifugation has been resuspended in buffer or to undergo buffer exchange when expression medium has different pH with purification column’s pH. While Xian et al. (2015) were purifying α-amylase expressed from T. pinophilus 1-95, a 0.22 µm filter membrane (HiPrep 16/10 desalting column) was equipped to dialyse and filter out the eluted (NH3)2SOafter resuspending in 20 mM sodium phosphate buffer, pH 6.5. A higher yield of 80.1% was found compared with other fungal α-amylases desalted using dialysis, e.g., 30.69% for Aspergillus flavus NSH9 (Karim et al. 2018) and 36.95% for Aspergillus terreus NCFT 4269.10 (Sethi et al 2016a) (Table 4).

Column Chromatography

Ion-exchange chromatography

Ion-exchange column chromatography (IEX) is based on the ionic bonds between cations and anions. Duong-Ly and Gabelli (2014b) stated that IEX separates molecules by their surface charge, which deviates greatly between different proteins and enzymes. There are two distinct mechanisms in purification using IEX: competitive ionic binding and ion exclusion due to repulsion between similarly charged analyte ions and ions fixed on the column (Acikara 2013). To ensure a protein or an enzyme has a particular charge, it should be dissolved in buffers with pH lower or higher than its isoelectric point (pI). There are two phases involved in this chromatography namely mobile and stationary phases. The mobile phase is generally an aqueous buffer system that contained the crude enzyme. Nevertheless, the stationary phase is an inert organic matrix, which is chemically derived from ionisable functional groups that carries a displaceable oppositely charged ion (Cummins et al. 2010).

The common desorption (elution) method increases the concentration of a similarly charged species within the mobile phase, thus competing and eluting the enzyme of interest from the column. In the purification of α-amylase, the most commonly used ion-exchange column is DEAE Sepharose, with commercially available HiTrap DEAE Sepharose FF and HiTrap Q Sepharose FF. Referring to Table 5, all the resins used in IEX are anionic exchangers, indicating that all the tabulated α-amylases are negatively charged at the respective working pH from the buffers used. This could be explained based on the fact that the pH of buffers used are higher than the pI of these enzymes. Negatively charged α-amylases are able to bind to the positively charged resins and are eluted with different concentrations of chloride ion (Cl), which depends on its overall strength of negative charge. Other positively charged contaminants will flow out from the column without binding to the resins, while other negatively charged contaminants will be separated from the α-amylases depending on the elution strength, thus in different elution fractions.

Referring to Table 5, the most commonly equipped columns in IEX are DEAE-Sephadex A-50 (Wu et al. 2017; David et al. 2017) and Q-Sepharose (Chen et al. 2015; Sudan et al. 2018). While Sudan et al. (2018) were purifying α-amylase from Geobacillus bacterium K1C, the dialysed enzyme sample was loaded on a Q-Sepharose column pre-equilibrated with 20 mM Tris-HCl buffer, pH 8.0 followed by elution with step gradient of 1 M NaCl. Although the purification fold had the range from 2.55 (Karim et al. 2018) to 34.33 (Xian et al. 2015), the yields (11.73 to 42.91%) were lower compared to other types of column chromatography. Table 5 also shows that KCl and NaCl have been used frequently during elution to desorb the enzyme of interest from the column matrix (stationary phase).

Table 5. Ion-exchange Column Chromatography of Microbial α-Amylases

Size-exclusion chromatography

Size-exclusion chromatography (SEC) or gel-filtration chromatography are often used for enzyme purification. Proteins of varying sizes are separated by columns consisting of a matrix of beads, which contain sieves of a particular size. Larger molecules are eluted earlier than small compounds, as the beads have cross-linked polyacrylamide, agarose, and dextran, where smaller compounds enter the sieves in the matrix of the stationary phase (Duong-Ly and Gabelli 2014a). According to Giridhar et al. (2017), porosity, i.e., pore size, is an important parameter. Because SEC separates molecules according to their size in solution, the process occurs wholly within the pore volume, which should be as large as possible. Due to the porosity of SEC, larger components of the analyte will be sampled by larger pores and vice versa. Thus, the larger molecules elute from the column first and smaller components will elute later (Striegel 2017; Berg et al. 2002).

Referring to Table 6, the most frequently equipped SEC matrix is Sephadex G-100 (Chen et al. 2015; Baltas et al. 2016; Allala et al. 2019; El-Sayed et al. 2019). This matrix shown promising yields while purifying α-amylases produced by Anoxybacillus thermarum A4 strain (74.6%), Bacillus subtilis WB800 (41.7%), Escherichia coli BL21 (76.53%), and Tepidimonas fonticaldi strain HB23 (41%). However, the highest purification fold was achieved by Sudan et al. (2018) at 49-fold, although its yield was the lowest at only 5.2%. Besides Sephadex G-100Superdex 75 has been used while purifying α-amylases produced by Anoxybacillus sp. YIM 342 (Zhang et al. 2016) and Geobacillus K1C (Sudan et al. 2018).

Zhang et al. (2016) performed gel filtration chromatography to achieve 32-fold increase in specific activity and a yield of about 10.4%. A Hiprep QXL 26/60 column (Superdex 75) was loaded with concentrated enzyme sample in 50 mM Tris-HCl buffer (pH 7.5) and eluted using the same buffer using an AKTATM time at a flow rate of 1 mL/min with 3.0 mL per fraction. In recent research, Sephadex G-100 was loaded with enzyme solution before eluting with 0.1 M phosphate buffer (pH 7.5) while purifying recombinant α-amylase AmyLa from Laceyella sp. DS3 expressed in E. coli BL21 (El-Sayed et al. 2019). Sephadex G-100 has a molecular weight fractionation range of 1-100 kDa, thus, the AmyLa from Laceyella sp. DS3 was shown to have 51.5 kDa from SDS-PAGE was small enough to enter the pores of the resin and was within the intermediate period of elution (El-Sayed et al. 2019).

SEC is also known as gel-filtration chromatography where the column resin acts as a filter to remove salts from the samples loaded. This desalting technique is usually performed as the finishing or polishing step to remove excessive salt after ammonium sulfate precipitation and ion-exchange chromatography, as high salt concentration may affect the downstream characterisation and crystallisation processes. After purifying bacterial α-amylase (44 kDa) expressed in B. methylotrophicus strain P11-2 using anionic exchanger DEAE FFSuperdex 75 10/300GL was used as a filter to remove NaCl from the active fractions of IEX (Xie et al. 2013). A similar desalting procedure was performed by Abdulaal (2018) while purifying fungal α-amylase (30 kDa) expressed in Trichoderma pseudokoningii, where Sephacryl S-200 was equipped to filter off salt (0.2 M NaCl) from the active fraction of IEX (DEAE-Sepharose). To remove excessive ammonium sulfate salt from sample to be loaded into Q-SepharoseSephadex G-100 was used, while Chen et al. (2015) was purifying bacterial α-amylase expressed in B. subtilis WB800 because unnecessary salt may affect the binding efficiency of IEX.

Table 6. Size-exclusion Chromatography of Various Microbial α-Amylases

Affinity column chromatography

Affinity column chromatography purifies proteins according to their specific affinity towards a ligand. Such chromatography is also known as immobilization, which is normally called immobilized metal affinity chromatography (IMAC). When the analyte molecules in the crude enzymes interact with the solid resin of IMAC, which has a covalent linkage with a polydentate metal-chelating group binding to a metal ion, e.g., nickel (Ni2+), surface-exposed amino acid residues of the enzyme of interest will exchange with the water molecule in the metal coordination site, thus the enzyme is immobilized (Chang et al. 2017).

While purifying thermostable α-amylase from B. subtilis DR8806 but expressed in E. coli BL21 (DE3), Emtenani et al. (2015) loaded the clear supernatant containing intracellular α-amylase through Ni2+-NTA matrix for affinity binding, yielding 60% recovery. Likewise, Gandhi et al. (2015) used IMAC to purify the α-amylase expressed in fungus with polyhistidine tag on 5 mL HiTrap IMAC FF, fast flow column with AKTA purifier system, yielding 1.9-fold purification with 52.6% recovery. Table 7 summarises affinity chromatography used to purify various microbial α-amylases.

Table 7. Affinity Chromatography of Various Microbial α-Amylases

CHARACTERISATION OF MICROBIAL EXTRACELLULAR α-AMYLASE

α-Amylase can be characterised in many respects such as the effects of temperature and pH, thermostability, pH stability, substrate specificity, effects of metal ions and chelating reagents, inhibitors and activators, and kinetics constants. The determination of optimum temperature and pH as well as the stabilities are crucial especially in identifying the most suitable microorganisms to be used in specific industrial production processes. In every characterisation, DNS method (Gandhi et al. 2015) is used to quantify the enzyme activity.

Optimum Temperature and pH

Characterisation of α-amylase in terms of optimum temperature and pH enables industrial processes utilizing these α-amylases to be performed at the optimal rate, thus maximizing their yield. Referring to Table 8, α-amylase produced by Bacillus licheniformis B4-423 (Wu et al. 2017) showed highest activity at 100 °C compared to the lowest optimum temperature exhibited by that from Streptomyces fragilis DA7-7 (Nithya et al. 2017) in terms of bacterial α-amylases. However, both fungal thermostable α-amylases expressed in Aspergillus flavus NSH9 (Karim et al. 2018) and Trichoderma pseudokoningii (Abdulaal 2018) exhibited the lowest optimum temperature at 50 ˚C, while α-amylase produced by Komagataella phaffii (Wang et al. 2015) showed the highest optimum temperature at 90 °C. Bacterial α-amylases have a wide range of optimum pH from pH 5.0 (Wu et al. 2017; Emtenani et al. 2015) to 10.5 (Baltas et al. 2016), while fungal α-amylases were shown to have their optimum pH ranging from pH 5.0 (Karim et al. 2018; Sethi et al. 2016b; Xian et al. 2015) to pH 9.0 (Ali et al. 2014).

The difference of optimum temperature between bacterial and fungal α-amylases could be due to the characteristics of the bacteria and fungus, which are the expression hosts of the enzymes. Thermophilic bacteria generally have higher resistance toward high temperature compared to thermophilic fungi; thus, the α-amylases expression in thermophilic bacteria will probably exhibit higher optimum temperature compared to those expressed in thermophilic fungi.

Table 8. Optimum Temperature and pH of Extracellular α-Amylase from Microorganisms

Thermostability and pH Stability

Thermostability and pH stability are important factors in industrially applied α-amylase because most of the industrial processes are performed at elevated temperature and non-neutral pH. Most studies on thermostability of α-amylase used a range near to its optimal temperature. While characterising α-amylase expressed from Anoxybacillus sp. YIM342, a maximum activity was observed at 80.0 °C; thus, the range of temperature was set from 70 °C to 90 °C. α-Amylase expressed from strain YIM342 had its half-life after 30 min incubation at 80 °C, remaining >49% of its activity, thus suitable to be used in starch saccharification process.

In terms of pH stability, the enzyme was found to retain more than 80% of its activity after incubation at pH 8.0 and pH 9.0 for 210 min. 45% of original activity was still retained by α-amylase from strain YIM342 after being pre-incubated at pH 10.0 for 210 min (Zhang et al. 2016). α-Amylase expressed from Aspergillus flavus NSH9 was found to be thermally stable at 50 °C, with 87% residual activity after incubation for 60 min. It was also observed that α-amylase from strain NSH9 was able to retain almost 100% of its original activity after incubation at pH 6.0 and pH 7.0 for 24 h (Karim et al. 2018).

Although characterisation of α-amylase in term of thermostability is important, the stability of enzymes while they are stored at 30 C as well as refrigerated at 4 C is also significant to be determined. A study by El-Sherbiny and El-Chaghaby (2012) showed that recovery of α-amylase (expressed in Bacillus sp.) with glycerol as a carrier or stabilizer at the storage temperature of 4 C (114%) was higher than the sample stored at 30 C (103%). However, when there was only water as carrier without glycerol as the stabilizer, the α-amylase recovery at 4 C (117%) was significantly higher than the sample stored at 30˚C (30.7%). These results had shown that the significance and importance to have α-amylase shipped with glycerol as stabilizer at around 4 C as the ambient temperatures for each country can be varied at high levels of fluctuation (El-Sherbiny and El-Chaghaby 2012).

Substrate Specificity

The substrate specificity profile is crucial because it characterises and determines the kind of starch that is degraded most effectively and efficiently by α-amylase. In the recent research conducted by Allala et al. (2019), the α-amylase TfAmy48 from Tepidimonas fonticaldi strain HB23 had the highest relative activity towards soluble potato starch (100%) while the enzyme had no activity towards some of the starches such as native potato, maize, rice starches, CMC and α-cyclodextrin. However, Baltas et al. (2016) found that the partially purified α-amylase from Anoxybacillus thermarum A4 strain had its highest specificity towards amylose (113%) and subsequently to soluble potato starch (100%) and amylopectin (93%), while the enzyme showed no activity towards cellulose as well as β-cyclodextrin (Table 9).

Both profiles in Table 9 reflected the preference of α-amylases to catalyze the hydrolysis of α-ᴅ-1,4-glycosidic linkages present at higher percentage in amylopectin, amylose, as well as soluble starches. Having a spontaneous hydrolysis rate of approximately 2 × 10-15 s-1 at room temperature, the α-glycosidic bond is very stable (Wolfdenden et al. 1998). The α-retaining double displacement proposed by Koshland (1953) is the most generally accepted catalytic mechanisms of the α-amylase family.

Five conserved amino acid sequence regions can be identified in members of the α-amylase family, where the two most conserved catalytic residues are located at the active site (glutamic acid as acid or base catalyst, and an aspartate as nucleophile) (Van Der Maarel et al. 2002). The third conserved residue, which is the second aspartate, binds to second and third hydroxyl groups (OH-2 and OH-3) of the substrate via hydrogen bonds, distorting the substrate (Uitdehaag et al. 1999). The fourth conserved amino acid residues can be histidine, arginine, and tyrosine, playing roles in ensuring correct orientation of the substrate into the active site, proper orientation of the nucleophile, transition state stabilization, as well as the polarization of the electronic structure of the substrate (Nakamura et al. 1993; Lawson et al. 1994; Strokopytov et al. 1996; Uitdehaag et al. 1999). An additional fifth conserved region also contains an aspartate, which is a calcium ligand (Janecek 1992).

Apart from the difference in conserved amino acid sequences, domain organization in various enzymes in the α-amylase family also has an effect on its substrate specificity. α-Amylase (E.C. 3.2.1.1), having A-domain (a highly symmetrical fold of eight parallel β-strands arranged in a barrel encircled by eight α-helices), B-domain (protruding between β-sheet no. 3 and α-helix no. 3 and playing a role in substrate or Ca2+ binding), as well as C-domain (unknown function), is meant to have starch (amylose and amylopectin) as its main substrate (Van der Veen et al. 2002). Thus, both conserved amino acid sequence and domains of the enzymes may contribute to their specificity to the substrate even though they are all in the α-amylase family.

Fig. 1. The α-retaining double displacement method of α-amylase reaction mechanism (Van Der Maarel et al. 2002; Kumari et al. 2011)

Table 9. Substrate Specificity Profile of the Purified α-Amylases (Baltas et al. 2016; Allala et al. 2019)

Metal Ions and Inhibitors

Some metal ions of optimal concentration may act as cofactor in increasing the activity of α-amylase in degrading starch, while some reagents and inhibitors act to decrease its activity disregarding their concentration. Being a calcium metalloenzyme, α-amylase has elevated activity when calcium ion (Ca2+) or salt (CaCl2) is added in the reaction mixture; its activity increases by 8 ± 5%when 4 mM of Ca2+ is added to the reaction mixture containing α-amylase expressed from Bacillus licheniformis AT70 (Afrisham et al. 2016).

Allala et al. (2019) showed 55 ± 3.9% increased activity when 5 mM of Ca2+ was added to the reaction mixture with α-amylase purified from T. fonticaldi strain HB23. However, mercury ion (Hg2+) showed an inhibitory effect on amylolytic activity (15 ± 3%) of α-amylase from strain AT70, which might be due to the non-specific binding and aggregation of the enzyme (Afrisham et al. 2016; Sethi et al. 2016b).

Referring to Table 10, Agüloglu et al. (2014) found the highest inhibitory effect when 1 mM EDTA (21%) and 10 mM EDTA (13%) were added separately to reaction mixtures with α-amylase from Anoxybacillus flavithermus. This result also demonstrated that the chelating agent EDTA inactivated α-amylase, which is a metalloenzyme. When 10 mM EDTA was added to α-amylase from Anoxybacillus sp. AH1, the amylolytic activity dropped to 37%, with a 63% decrease in enzyme activity (Acer et al. 2016).

Table 10. α-Amylase Activity Remaining after Incubation for 30 min at 37 °C
(Agüloglu et al. 2014)

CONCLUSIONS

  1. Affinity chromatography has shown the highest purification fold (1.72 to 13.2-fold) and recovery (38 to 60%) while purifying thermostable α-amylases in comparison to other purification methods such as ultrafiltration, salt precipitation, dialysis, and other means of column chromatography. An established purification method for microbial thermostable α-amylase is critical in fulfilling the demand of well-decontaminated and non-toxic enzymes in the industries.
  2. Most studies have shown that microbial thermostable α-amylases have optimum temperature and pH values ranging from 50 °C to 100 °C and pH 5.0 to 10.5, respectively. Microbial thermostable α-amylase also shown to have high specificity towards soluble potato starch, wheat starch, amylose, and amylopectin. Both EDTA (1 mM) and mercury ion (Hg2+) have been proven to strongly inhibit α-amylase activity, while calcium ions (Ca2+) shown promising inducing effect (55%) on microbial α-amylase activity.
  3. Purification and characterisation of α-amylase have been focused on enzymes from microbial sources (bacteria and fungi) as well as exhibiting thermostability. Such trends should be expected to have research contributing to an established purification method with more than 90% yield recovery and very high purification fold so that thermostable α-amylases can be purified thoroughly from microbial sources in large scale to ensure their safety to be used in various industrial applications.

ACKNOWLEDGEMENTS

This review was supported by Putra-IPS grant GP-IPS/2017/9516700 from Universiti Putra Malaysia, which was awarded to the last author.

REFERENCES CITED

Abdulaal, W. H. (2018). “Purification and characterization of α-amylase from Trichoderma pseudokoningii,” BMC Biochemistry 19(4), 1-6. DOI: 10.1186/s12858-018-0094-8

Abu, M. L., Nooh H. M., Oslan, S. N., and Salleh, A. B. (2017). “Optimization of physical conditions for the production of thermostable T1 lipase in Pichia guilliermondii strain SO using response surface methodology,” BMC Biotechnology 17(78), 1-10. DOI: 10.1186/s12896-017-0397-7

Acer, Ö., Bekler, F. M., Pirinççioğlu, H., Güven, R. G., and Güven, K. (2016). “Purification and characterization of thermostable and detergent-stable α-amylase from Anoxybacillus sp. AH1,” Food Technology and Biotechnology 5(1), 70-77. DOI: 10.17113/ftb.54.01.16.4122.

Acikara, Ö. B. (2013). “Ion-exchange chromatography and its applications,” in: Column Chromatography, Martin, D. F., Martin, B. B. (ed.), Intech, London, UK. DOI: 10.5772/55744

Afrisham, S., Badoei-Dalfard, A., Namaki-Shoushtari, A., and Karami. Z. (2016). “Characterization of a thermostable, CaCl2-activated and raw-starch hydrolyzing α-amylase from Bacillus licheniformis AT70: Production under solid state fermentation by utilizing agricultural wastes,” Journal of Molecular Catalysis B: Enzymatic 132, 98-106. DOI: 10.1016/j.molcatb.2016.07.002

Agüloglu, S. G. F., Enez, B., Özdemir, S., and Bekler, F. M. (2014). “Purification and characterization of thermostable α-amylase from thermophilic Anoxybacillus flavithermus,” Carbohydrate Polymers 102, 144-150. DOI: 10.1016/j.carbpol.2013.10.048

Ahmad, M., Hirz, M., Pichler, H., and Schwab, H. (2014). “Protein expression in Pichia pastoris: Recent achievements and perspectives for heterologous protein production,” Applied Microbiology and Biotechnology 98(12), 5301-5317. DOI: 10.1007/s00253-014-5732-5

Ali, I., Ali, A., Anwar, M., Yanwisetpakdee, B., Prasongsuk, S., Lotrakul, P., and Punnapayak, H. (2014). “Purification and characterization of extracellular, polyextremophilic α-amylase obtained from halophilic Engyodontium album,” Iranian Journal of Biotechnology 12(4), 35-40. DOI: 10.15171/ijb.1155

Allala, F., Bouacem, K., Boucherba, N., Azzouz, Z., Mechri, S., Sahnoun, M., Benallaoua, S., Hacene, H., Jaouadi, B., and Bouanane-Darenfed A. (2019). “Purification, biochemical, and molecular characterization of a novel extracellular thermostable and alkaline α-amylase from Tepidimonas fonticaldi strain HB23,” International Journal of Biological Macromolecules 132, 558-574. DOI: 10.1016/j.ijbiomac.2019.03.201.

Baltas, N., Dincer, B., Ekinci, A. P., Kolayli, S., and Adiguzel, A. (2016). “Purification and characterization of extracellular α-amylase from a Thermophilic Anoxybacillus
thermarum
 A4 strain,” Brazilian Archives of Biology and Technology 59, 1-14. DOI: 10.1590/1678-4324-2016160346

Bano, S., Qader, S. A. U., Aman, A., Syed, M. N., and Azhar, A. (2011). “Purification and characterization of novel α-amylase from Bacillus subtilis KIBGE HAS,” AAPS PharmSciTech 12(1), 255-261. DOI: 10.1208/s12249-011-9586-1

Berg, J. M., Tymoczko, J. L., and Stryer, L. (2002). “The purification of proteins is an essential first step in understanding their function,” in: Biochemistry, 5th Ed., WH Freeman, New York.

Borrelli, G. M., and Trono, D. (2015). “Recombinant lipases and phospholipases and their use as biocatalysts for industrial applications,” International Journal of Molecular Sciences 16, 20774-20840. DOI: 10.3390/ijms160920774.

Chang, Y. Y., Li, H., and Sun, H. (2017). “Immobilized metal affinity chromatography (IMAC) for metalloproteomics and phosphoproteomics,” in: Inorganic and Organometallic Transition Metal Complexes with Biological Molecules and Living Cells, Elsevier, Amsterdam, Netherlands, pp. 329-353. DOI: 10.1016/B978-0-12-803814-7.00009-5

Chen, J., Chen, X., Dai, J., Xie, G., Yan, L., Lu, L., and Chen, J. (2015). “Cloning, enhanced expression and characterization of an α-amylase gene from a wild strain in B. subtilis WB800,” International Journal of Biological Macromolecules 80: 200-207. DOI: 10.1016/j.ijbiomac.2015.06.018

Cummins, P. M., Dowling, O., and O’Connor, B. F. (2010). “Ion-exchange chromatography: Basic principles and application to the partial purification of soluble mammalian prolyl oligopeptidase,” in: Protein Chromatography Methods and Protocols, pp. 215-228. DOI: 10.1007/978-1-60761-913-012

David, S., Femi, B., Gbenga, A., and Saanu, A. B. (2017). “Purification and characterization of α-amylase from Bacillus subtilis isolated from cassava processing sites,” Journal of Bioremediation & Biodegradation 8(6), 1-7. DOI: 10.4172/2155-6199.1000417

Deljou, A., and Arezi, I. (2016). “Production of thermostable extracellular α-amylase by a moderate thermophilic Bacillus licheniformis-AZ2 isolated from Qinarje hot spring
(Ardebil Prov. of Iran),” Periodicum Biologorum 118 (4). DOI: 10.18054/pb.v118i4.3737

Du, R., Song, Q., Zhang, Q., Zhao, F., Kim, R. C., Zhou, Z., and Han, Y. (2018). “Purification and characterization of novel thermostable and Ca-independent α-
amylase produced by Bacillus amyloliquefaciens BH072,” International Journal of Biological Macromolecules 115: 1151-1156. DOI: 10.1016/j.ijbiomac.2018.05.004

Duong-Ly, K. C., and Gabelli, S. B. (2014a). “Gel filtration chromatography (Size exclusion chromatography) of proteins,” In Methods in Enzymology, 105-114. DOI: 10.1016/B978-0-12-420119-4.00009-4

Duong-Ly, K. C., and Gabelli, S. B. (2014b). “Using ion exchange chromatography to purify a recombinantly expressed protein,” Methods in Enzymology 541 (January): 95-103. DOI: 10.1016/B978-0-12-420119-4.00008-2

El-Sayed, A. K.A., Abou-Dobara, M. I., El-Fallal, A. A., and Omar, N. F. (2019). “Heterologous expression, purification, immobilization and characterization of
recombinant α-amylase AmyLa from Laceyella sp. DS3,” International Journal of Biological Macromolecules 132, 1274-1281. DOI: 10.1016/j.ijbiomac.2019.04.010

El-Sherbiny, M., and El-Chaghaby, G. (2012). “Storage temperature and stabilizers in relation to the activity of commercial liquid feed enzymes: A case study from Egypt,” Journal of Agrobiology 28(2), 129-137. DOI: 10.2478/v10146-011-0014-7

Emtenani, S., Asoodeh, A., and Emtenani. S. (2015). “Gene cloning and characterization of a thermostable organic-tolerant α-amylase from Bacillus subtilis DR8806,” International Journal of Biological Macromolecules 72, 290-298. DOI: 10.1016/j.ijbiomac.2014.08.023

Espargaró, A., Villar-Piqué, A., Sabaté, R., and Ventura, S. (2012). “Yeast prions form infectious amyloid inclusion bodies in bacteria,” Microbial Cell Factories 11(1), 89-101. DOI: 10.1186/1475-2859-11-89

Gandhi, S., Salleh, A. B., Rahman, R. N. Z. R. A., Leow, T. C., and Oslan, S. N. (2015). “Expression and characterization of Geobacillus stearothermophilus SR74 recombinant α-amylase in Pichia pastoris,” BioMed Research International 2015, article ID 529059. DOI: 10.1155/2015/529059

Giridhar, G., Manepalli, R. K. N. R., and Apparao, G. (2017). “Size-exclusion chromatography,” in: Thermal and Rheological Measurement Techniques for Nanomaterials Characterization, 51-65. Elsevier. DOI: 10.1016/B978-0- 323-46139-9.00003-7

Gupta, R., Gigras, P., Mohapatra, H., Goswami, V. K., and Chauhan, B. (2003). “Microbial α-amylases: A biotechnological perspective,” Process Biochemistry 38, 1599-1616. DOI: 10.1016/S0032-9592(03)00053-0

Hammami, A., Fakhfakh, N., Abdelhedi, O., Nasri, M., and Bayoudh, A. (2018). “Proteolytic and amylolytic enzymes from a newly isolated Bacillus mojavensis SA: Characterization and applications as laundry detergent additive and in leather processing,” International Journal of Biological Macromolecules 108, 56-68. DOI: 10.1016/j.ijbiomac.2017.11.148

Janecek, S. (1992). “New conserved amino acid region of α-amylases in the third loop of their (β/α)8-barrel domains,” Biochemical Journal 288(3), 1069-1070. DOI:10.1042/bj2881069

Jiang, T., Cai, M., Huang, M., He, H., Lu, J., Zhou, X., and Zhang, Y. (2015). “Characterization of a thermostable raw-starch hydrolyzing α-amylase from deep-sea thermophile Geobacillus sp,” Protein Expression and Purification 114, 15-22. DOI: 10.1016/j.pep.2015.06.002

Karim, K. M. R., Husaini, A., Sing, N. N., Sinang, F. M., Roslan, H. A., and Hussain, H. (2018). “Purification of an α-amylase from Aspergillus flavus NSH9 and molecular
characterization of its nucleotide gene sequence,” 3 Biotech 8(4), 204. DOI: 10.1007/s13205-018-1225-z.

Kassaye, E. K. (2009). Molecular Cloning and Expression of a Thermostable α-Amylase from Geobacillus sp, Master’s Thesis, Universiti Putra Malaysia.
http://psasir.upm.edu.my/id/eprint/7576/.

Koshland, D. E. (1953). “Stereochemistry and the mechanism of enzymatic reactions,” Biological Reviews 28(4), 416-436. DOI: 10.1111/j.1469 185x.1953.tb01386.x

Kumari, A., Singh, K., and Kayastha, A. M. (2011). “α-Amylase: General properties, mechanism and biotechnological applications – A review,” Current Biotechnology 1(1), 98-107. DOI: 10.2174/2211551X11201010098

Lawson, C. L., van Montfort, R., Strokopytov, B., Rozeboom, H. J., Kalk, K. H., de Vries, G. E. et al. (1994). “Nucleotide sequence and X-ray structure of cyclodextrin glycosyltransferase from Bacillus circulans strain 251 in a maltose-dependent crystal form,” Journal of Molecular Biology 236(2), 590-600. DOI: 10.1006/jmbi.1994.1168

Nakamura, A., Haga, K., and Yamane, K. (1993). “Three histidine residues in the active center of cyclodextrin glucanotransferase from alkalophilic Bacillus sp. 1011: Effects of the replacement on pH dependence and transition-state stabilization,” Biochemistry 32(26), 6624-6631. DOI: 10.1021/bi00077a015

Nasir, N. S. M. (2019). Expression of a Thermostable α-Amylase using Formaldehyde Dehydrogenase Promoter (pFLD) in Meyerozyma guilliermondii Strain SO, Master’s Thesis, Universiti Putra Malaysia.

Nevoigt, E. (2008). “Progress in metabolic engineering of Saccharomyces cerevisiae,” Microbiology and Molecular Biology Reviews 72(3), 379-412. DOI: 10.1128/MMBR.00025-07

Nithya, K., Muthukumar, C., Kadaikunnan, S., Alharbi, N. S., Khaled, J. M. and Dhanasekaran, D. (2017). “Purification, characterization, and statistical optimization of a thermostable α-amylase from desert actinobacterium Streptomyces fragilis DA7- 7,” 3 Biotech 7(5). DOI: 10.1007/s13205-017-0981-5

Oktiarni, D., Lusiana, Simamora, F. Y., and Gaol, J. M. L. (2015). “Isolation, purification and characterization of β-amylase from Dioscorea hispida Dennst,” in: AIP Conference Proceedings 1677(1). DOI: 10.1063/1.4930749

Oslan, S. N., Salleh, A. B., Rahman, R. N. Z. R. A., Leow, T. C., Sukamat, H., and Basri, M. (2015). “A newly isolated yeast as an expression host for recombinant lipase,” Cellular and Molecular Biology Letters 20(2), 279-293. DOI: 10.1515/cmble-2015-0015

Oslan, S. N., Salleh, A. B., Rahman, R. N. Z. R. A., Leow, T. C. and Basri, M. (2012). “Locally isolated yeasts from Malaysia: Identification, phylogenetic study and characterization,” Acta biochimica Polonica 59(2), 225-229. DOI: 10.18388/abp.2012

Özdemir, S., Okumus, V., Ulutas, M. S., Dundar, A., Akarsubasic, A. T., and Dumontet, S. (2016). “Production and characterization of thermostable α-amylase from thermophilic Anoxybacillus flavithermus sp. nov. SO-19,” Starch/Staerke 68(11-12), 1244-53. DOI: 10.1002/star.201500071

Saini, R., Saini, H. S., and Dahiya, A. (2017). “Amylases: Characteristics and industrial applications,” Journal of Pharmacognosy and Phytochemistry 6(4), 1865-
1871.

Sethi, B. K., Jana, A., Nanda, P. K., DasMohapatra, P. K., Sahoo, S. L., and Patra, J. K. (2016a). “Production of α-amylase by Aspergillus terreus NCFT 4269.10 using pearl
millet and its structural characterization,” Frontiers in Plant Science 7(May): 639. DOI: 10.3389/fpls.2016.00639.

Sethi, B. K., Nanda, P. K., Sahoo, S., and Sena, S. (2016b). “Characterization of purified α-amylase produced by Aspergillus terreus NCFT 4269.10 using pearl millet as
substrate,” Cogent Food & Agriculture 2(1158902). DOI: 10.1080/23311932.2016.1158902.

Simair, A. A., Qureshi, A. S., Khushk, I., Ali, C. H., Lashari, S., Bhutto, M. A., Mangrio, enzyme from Bacillus sp. BCC 01-50 and potential applications,” BioMed Research
International
, 2017: 1-9. DOI: 10.1155/2017/9173040.

Singh, R, Kumar, M., Mittal, A., and Mehta, P. K. (2016). “Amylases: A Note on current application,” International Research Journal of Biological Sciences 5 (11): 27-32.
http://www.isca.in/IJBS/Archive/v5/i11/6.ISCA-IRJBS-2016-127.pdf.

Singh, S., and Guruprasad, L. (2014). “Structure and sequence-based analysis of α- amylase evolution,” Protein & Peptide Letters 21(9), 948-956. DOI: 10.2174/092986652109140715124139

Straathof, J. J. A., and Adlercreutz, P. (2014). “How to get the biocatalyst,” in: Applied Biocatalyst, 2nd Ed., pp. 181-182.

Striegel, A. M. (2017). “Size-exclusion chromatography,” in: S. Fanali, P. R. Haddad, C. Poole, & M.-L. Riekkola (Eds.), Liquid Chromatography: Fundamentals and Instrumentation, 2nd Ed., pp. 245–273. Elsevier. DOI: 10.1016/B978-0-12-805393-5.00010-5

Strokopytov, B., Knegtel, R. M. A., Penninga, D., Rozeboom, H. J., Kalk, K. H., Dijkhuizen, L., and Dijkstra, B. W. (1996). “Structure of cyclodextrin glycosyltransferase complexed with a maltononaose inhibitor at 2.6 Å resolution. Implications for product specificity,” Biochemistry 35(13), 4241-4249. DOI: 10.1021/bi952339h

Sudan, S. K., Kumar, N., Kaur, I., and Sahni, G. (2018). “Production, purification and Characterization of raw starch hydrolyzing thermostable acidic α-amylase from hot springs, India,” International Journal of Biological Macromolecules 117, 831-839. DOI: 10.1016/j.ijbiomac.2018.05.231

Uitdehaag, J. C. M., Mosi, R., Kalk, K. H., Van der Veen, B. A., Dijkhuizen, L., Withers, S. G., and Dijkstra, B. W. (1999). “X-ray structures along the reaction pathway of cyclodextrin glycosyltransferase elucidate catalysis in the α-amylase family,” Nature Structural Biology 6(5), 432-436. DOI: 10.1038/8235

Van Der Maarel, M. J. E. C., Van Der Veen, B., Uitdehaag, J. C. M., Leemhuis, H., and Dijkhuizen, L. (2002). “Properties and applications of starch-converting enzymes of the α-amylase family,” Journal of Biotechnology 94(2): 137–155. DOI: 10.1016/S0168-1656(01)00407-2

Wang, J. R., Li, Y.Y., Liu, D. N., Liu, J. S., Li, P., Chen, L. Z., and Xu, S. D. (2015). “Codon optimization significantly improves the expression level of α-amylase gene
from Bacillus licheniformis in Pichia pastoris,” BioMed Research International 2015, Article ID 248680, 9 pages. DOI: 10.1155/2015/248680

Wang, J., Li, Y., and Lu, F. (2018). “Molecular cloning and biochemical characterization of an α-amylase family from Aspergillus niger,” Electronic Journal of
Biotechnology
 32, 55-62. DOI: 10.1016/j.ejbt.2018.01.004

Wingfield, P. T. (2016). “Protein precipitation using ammonium sulfate,” In Current Protocols in Protein Science, P. T. Wingfield (ed.), John Wiley & Sons, Inc., Hoboken, NJ, USA, A.3F.1-A.3F.9. DOI: 10.1002/0471140864.psa03fs84

Wu, X., Wang, Y., Tong, B., Chen, X., and Chen, J. (2018). “Purification and biochemical characterization of a thermostable and acid-stable α-amylase from Bacillus licheniformis B4-423,” International Journal of Biological Macromolecules 109, 329-337. DOI: 10.1016/j.ijbiomac.2017.12.004

Xian, L., Wang, F., Luo, X., Feng, Y. L. and Feng, J. X. (2015). “Purification and characterization of a highly efficient calcium-independent α-amylase from Talaromyces pinophilus 1-95,” PLoS ONE 10(3), 1-18. DOI: 10.1371/journal.pone.0121531

Xie, F., Quan, S., Liu, D., Ma, H., Li, F., Zhou, F., and Chen, G. (2014). “Purification and characterization of a novel α-amylase from a newly isolated Bacillus methylotrophicus strain P11-2,” Process Biochemistry 49(1), 47-53. DOI: 10.1016/j.procbio.2013.09.025

Zaky, A. S., Tucker, G. A., Daw, Z. Y., and Du, C. (2014). “Marine yeast isolation and industrial application,” FEMS Yeast Research 14(6), 813-825. DOI: 10.1111/1567-1364.12158

Zhang, F., Yang, X., Geng, L., Zhang, Z., Yin, Y., and Li, W. (2016). “Purification and characterization of a novel and versatile α-amylase from thermophilic Anoxybacillus sp. YIM 342,” Starch/Staerke. 68, 446-453. DOI: 10.1002/star.201400056

Zhang, Q., Han, Y., and Xiao, H. (2017). “Microbial α-amylase: A biomolecular overview,” Process Biochemistry. DOI: 10.1016/j.procbio.2016.11.012

Article submitted: September 4, 2019; Peer review completed: October 19, 2019; Revisions accepted: November 15, 2019; Published: November 26, 2019.

DOI: 10.15376/biores.15.1.Lim