Biology 2021, 10, 337. https://doi.org/10.3390/biology10040337 www.mdpi.com/journal/biology
Article
Biochemical Characterization of the Amylase Activity from the
New Haloarchaeal Strain Haloarcula sp. HS Isolated in the
Odiel Marshlands
Patricia Gómez‐Villegas
1
, Javier Vigara
1
, Luis Romero
2
, Cecilia Gotor
2
, Sara Raposo
3
, Brígida Gonçalves
3
and Rosa Léon
1,*
1 Laboratory of Biochemistry, Department of Chemistry, Marine International Campus of Excellence
(CEIMAR), University of Huelva, Avenida de las Fuerzas Armadas s/n, 21071 Huelva, Spain;
[email protected] (P.G.‐V.); [email protected] (J.V.)
2 Instituto de Bioquímica Vegetal y Fotosíntesis, Consejo Superior de Investigaciones Científicas and
Universidad de Sevilla, Avenida Américo Vespucio 49, 41092 Seville, Spain; [email protected] (L.R.);
[email protected] (C.G.)
3 CIMA—Centre for Marine and Environmental Research, FCT, Campus de Gambelas,
Universidade do Algarve, 8005‐139 Faro, Portugal; [email protected] (S.R.); [email protected] (B.G.)
* Correspondence: [email protected]; Tel.: +34‐959‐219‐951
Simple Summary: Amylases are a group of enzymes that degrade starch into simple sugars. These
proteins are produced by a wide variety of organisms and are supposed to be one of the most valu‐
able industrial enzymes. However, the extreme conditions required for many industrial operations
limit the applicability of most
amylases found in nature. In this context, halophilic archaea entail an
excellent source of novel proteins that tolerate harsh conditions, as they live in environments with
high salt concentration and temperature. In this work, a screening of haloarchaea, isolated from
Odiel salterns in the southwest of Spain, was carried out
to select a new strain with a high amylase
activity. This microorganism was identified as Haloarcula sp. HS and showed amylase activities in
both, the cellular and the extracellular extracts. Both amylase activities were poly‐extremotolerant,
as their optimal yields were achieved at 60 °C and 25% NaCl. Additionally, the study
of the protein
sequences from Haloarcula sp. HS allowed the identification of three different amylases, which con‐
served the typical structure of the alpha‐amylase family. Finally, the applicability of the extracellu‐
lar amylase to treat bakery wastes under high salinity conditions was demonstrated.
Abstract: Alpha‐amylases are a large
family of α,1‐4‐endo‐glycosyl hydrolases distributed in all
kingdoms of life. The need for poly‐extremotolerant amylases encouraged their search in extreme
environments, where archaea become ideal candidates to provide new enzymes that are able to
work in the harsh conditions demanded in many industrial applications. In this study,
a collection
of haloarchaea isolated from Odiel saltern ponds in the southwest of Spain was screened for their
amylase activity. The strain that exhibited the highest activity was selected and identified as Halo‐
arcula sp. HS. We demonstrated the existence in both, cellular and extracellular extracts of the new
strain,
of functional α‐amylase activities, which showed to be moderately thermotolerant (optimum
around 60 °C), extremely halotolerant (optimum over 25% NaCl), and calcium‐dependent. The tryp‐
tic digestion followed by HPLC‐MS/MS analysis of the partially purified cellular and extracellular
extracts allowed to identify the sequence of three alpha‐amylases, which despite
sharing a low se‐
quence identity, exhibited high three‐dimensional structure homology, conserving the typical do‐
mains and most of the key consensus residues of α‐amylases. Moreover, we proved the potential of
the extracellular α‐amylase from Haloarcula sp. HS to treat bakery wastes under high salinity con‐
ditions.
Keywords: amylase;
extremozymes; haloarchaea; enzymatic characterization; proteomics
Citation: Gómez‐Villegas, P.; Vigara,
J.; Romero, L.; Gotor, C.; Raposo, S.;
Gonçalves, B.; Léon, R. Biochemical
Characterization of the Amylase
Activity from the New Haloarchaeal
Strain Haloarcula sp. HS Isolated in
the Odiel Marshlands. Biology 2021,
10, 337. https://doi.org/10.3390/
biology10040337
Academic Editor: Xuehong Zhang
Received: 22 February 2021
Accepted: 15 April 2021
Published: 16 April 2021
Publisher’s Note: MDPI stays neu‐
tral with regard to jurisdictional
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tional affiliations.
Copyright: © 2021 by the authors. Li‐
censee MDPI, Basel, Switzerland.
This article is an
open access article
distributed under the terms and con‐
ditions of the Creative Commons At‐
tribution (CC BY) license (http://crea‐
tivecommons.org/licenses/by/4.0/).
Biology 2021, 10, 337 2 of 24
1. Introduction
Haloarchaea are the main representatives of extreme halophiles, which can thrive in
media with salt concentrations ranging from 20 to 30% [1,2]. These singular microorgan‐
isms are characterized by the accumulation of large amounts of KCl in the cytoplasm, to
maintain the osmotic balance with the medium, in
contrast to moderate or facultative hal‐
ophiles, which usually store compatible solutes for the same purpose [3]. Therefore, intra‐
and extracellular proteins from haloarchaea are specially adapted to work properly at
high salt concentrations. These proteins possess exceptional features that make them dis‐
tinguishable from non‐halophilic proteins; they have a
unique amino acid composition
with acidic surfaces and low overall hydrophobicity, to prevent aggregation and, at the
same time, retain flexibility in such high salinity [4]. Most haloarchaeal enzymes are con‐
sidered poly‐extremotolerant, as they work appropriately under more than one extreme
condition, usually elevated temperatures, in addition to
high salinity. For this reason, en‐
zymes from these halophilic microorganisms can be of interest in many harsh industrial
and biotechnological processes [5,6].
Amylases are a diverse group of hydrolase or transferase enzymes that degrade large
alpha‐linked polysaccharides, such as starch and related oligosaccharides, and are one of
the most
required enzymes in industrial operations. They stand for about 30% of the
world enzyme market, and this value is expected to grow in the following years, due to
the global increase in the demand for bakery and sugar‐derived products, biofuels, deter‐
gents, breweries, animal feeds, pharmaceuticals, paper, and textiles
[7,8]. At present, the
best market for amylases is in the production of maltose and glucose syrups from corn,
which are used as sweeteners for soft drinks [9,10]. They are also required for the enhance‐
ment of dough for baking, for clarification of fruit juices and beers, and in the pretreatment
of animal feed to improve the digestibility of fiber [11]. In addition, amylases are widely
used in textile and paper industries to remove the starch employed for the desizing pro‐
cess and the coating treatment, respectively. Moreover, several hydrolytic enzymes, in‐
cluding amylases, are usually added to detergents because they
permit the use of milder
conditions in laundry and automatic dishwashing, making them eco‐friendlier. Other in‐
teresting fields of application of amylases are biomedicine and pharmacy, to treat diges‐
tive disorders or as reporter genes in molecular biology [12]. Furthermore, amylases are
widely used for the conversion of starch‐rich
agronomic and food wastes into fermentable
sugars, which are required as feedstocks for the production of fuels and chemicals with
high demand and market value [13].
Amylases are ubiquitous ancient enzymes found in plants, animals, and microorgan‐
isms. Among them, bacteria of the genus Bacillus or fungi belonging to the
Aspergillus ge‐
nus are the most preferred source of amylases for large‐scale production [14]. Hydrolytic
amylases can be classified into two broad categories—endoamylases, which hydrolyze the
interior of the starch molecule; and exo‐amylases, which successively degrade starch from
the non‐reducing ends [9]. Most endoamylases belong to the α‐amylase
family (EC 3.2.1.1)
and cleave internal α,1‐4 glycosidic bonds between glucose units, producing oligosaccha‐
rides with varying lengths and α‐limit dextrins. Additionally, α‐amylases are typically
divided into two groups, according to the hydrolysis products and the degree of starch
hydrolysis; saccharifying α‐amylases that produce free sugars, and liquefying α‐amylases
that break down the starch polymer without producing free sugars [5]. The mechanism of
action and the catalytic properties of these amylases are well‐known and can be correlated
with their structural characteristics, as was detailed in several reviews [7,15,16].
The ability of haloarchaea to produce and excrete hydrolytic enzymes, including
am‐
ylases to degrade extracellular polysaccharides, as many other microorganisms do, was
previously described [6]. However, the application of archaeal amylases, which could be
beneficial to many industrial operations that require extreme conditions, remains scarcely
studied when compared to those from other microorganisms. Intracellular or cell‐associ‐
ated amylases from
haloarchaea are particularly understudied, although they are an im‐
portant haloarchaeal trait and can represent an interesting source of halotolerant enzymes.
Biology 2021, 10, 337 3 of 24
The saltern ponds of the Odiel Marshlands are an interesting saline ecosystem, which
harbors a rich diversity of prokaryotic and eukaryotic microorganisms. Our previous
studies showed that at very high salinity (33%), the most abundant archaea species belong
to the genera Halorubrum and Haloquadratum [17]. Metagenomic microbial profiling by
high‐
throughput 16S rRNA sequencing revealed the existence of various strains that be‐
longed to the Haloarcula genus. Although the abundance of these representatives was
quite low, with less than 0.2% of the total sequenced reads, our data suggest that some of
the haloarchaea of this group are able to produce bioactive
compounds [18] and excrete
hydrolytic haloenzymes, including proteases, amylases, or lipases [17].
In this study, a collection of haloarchaea isolated from the saltern ponds of the Odiel
Marshlands was screened for their amylolytic activity, and the one that exhibited the high‐
est activity was selected and identified on the basis
of its 16S rRNA coding gene. The ex‐
tracellular and cellular starch‐degrading activities of the selected archaea were character‐
ized, revealing different optimal parameters and modes of action. To get a further insight
into the identity of these starch‐degrading enzymes, the proteome composition of the par‐
tially purified cell
‐free supernatant and the cellular extracts was analyzed by tryptic di‐
gestion, followed by nano‐liquid chromatography coupled to an electrospray ionization
tandem mass spectrometry system. This study allowed the identification of three amylase
sequences (two were exclusively cell‐associated and one was also found in the extracellu‐
lar medium)
with high homology to amylases of other haloarchaeal species, and the typ‐
ical alpha‐amylase conserved regions. Furthermore, the potential applicability of the am‐
ylase enzymes of this new haloarchaea on the treatment of bakery waste was assessed and
compared with a commercial amylase.
2. Materials and Methods
2.1. Screening and
Selection of Amylase Producing Haloarchaea
The screening of amylase‐producing haloarchaea was performed by detection of the
extracellular amylase activity of the isolates on starch agar plates, with 20% NaCl. The
plates were flooded with commercial Lugol’s iodine solution, 0.5% I
2 and 1% KI (w/v)
(Chem Lab, Zedelgem, Belgium), every three days, to check the formation of degradation
halos around the colonies. The isolate that presented the highest ratio of halo zone with
respect to colony diameter was chosen for further studies. Screenings were done in tripli‐
cates.
2.2. Identification
of the Selected Microorganism
Genomic DNA of the isolated amylase‐producing strain was purified using the
GeneJET Genomic Purification kit (Thermo Fisher Scientific, Waltham, MA, USA), follow‐
ing the manufacturer’s instructions. The quantification and the purity assessment of the
genomic DNA obtained was done on a Nanodrop Spectrophotometer ND‐1000 (Thermo
Fisher Scientific). The full length of the 16S rRNA encoding gene was amplified with the
archaeal specific primers 21F (5′‐TTCCGGTTGATCCTGCCGGA ‐3′) and 1492R (5′‐
GGTTACCTTGTTACGACTT ‐3′). Polymerase chain reactions (PCR) were performed in a
total volume of 25 μL, using an Eppendorf thermo‐cycler. The reaction mixture
con‐
tained—1 μL of genomic DNA, 0.2 U REDTaq
®DNA polymerase from Sigma Aldrich (St.
Louis, MO, USA), and 2.5 μL of its specific 10× buffer that contained 10 pM of each primer,
0.2 mM dNTPs, and 2.5 mM MgCl
2. The thermal profile was set to 0.5 min at 96 °C, 0.5
min at 55 °C, and 1 min at 72 °C for 30 cycles, followed by 10 min of final extension. The
PCR products were analyzed by electrophoresis, on a 1% agarose gel to check their qual‐
ity, and
sent to Stabvida (Lisbon, Portugal) for Sanger sequencing. The 1.4 kb 16S rRNA
gene sequences obtained were compared to those available at the GenBank and the Euro‐
pean Molecular Biology Laboratory (EMBL) databases, using the Basic Local Alignment
Search Tool (BLAST) at the National Center for Biotechnology Information (NCBI) [19].
Biology 2021, 10, 337 4 of 24
2.3. Culture Conditions for Enzyme Production
All cultures were incubated at 37 °C with a shaking rate of 100 rpm, with either stand‐
ard rich medium or minimal medium. The standard rich medium for archaea growth con‐
tained per liter—156 g NaCl, 13 g MgCl
2∙6H2O, 20 g MgSO 4∙7H2O, 1 g CaCl 2∙6H2O, 4 g KCl,
0.2 g NaHCO
3, 0.5 g NaBr, and 5 g yeast extract, with a pH value of 7, measured before
autoclaving. For the minimal medium, yeast extract was substituted for 1% (w/v) of am‐
monium acetate. The amylase activity was induced by the addition of starch (3 g L
−1) to
either the rich or the minimal medium.
2.4. Partial Purification of Cell‐Associated and Extracellular Amylases
Haloarcula sp. HS cells were first cultured in the rich medium, containing yeast extract
and starch. When the culture reached the end of the exponential phase (OD
580 N 3), cells
were harvested through centrifugation, washed, and transferred to the minimal medium,
where the biomass was cultivated until the extracellular starch was completely exhausted,
about 3 days after the transference. Starch content was periodically measured every 24 h,
by mixing 1 mL of culture medium with 5 μL
of commercial Lugol’s iodine solution and
by reading the absorbance at 580 nm. Then, the biomass was harvested by centrifugation
for 20 min at 12,000 rpm and 4 °C. The supernatant was 100‐fold concentrated by an ul‐
trafiltration process in an Amicon
®
system with a 10 kDa cut‐off membrane and used as
the source of the extracellular amylase. The specific amylase activity in the medium su‐
pernatant was 8 U mg
−1 and it was increased to 350 U mg
−1 in the concentrated superna‐
tant, with a purification factor of 43.75. On its part, the cell pellet was disrupted by soni‐
cation in phosphate buffer (50 mM, 20% NaCl, pH 7) and centrifuged again to remove the
cell debris and unbroken cells. The obtained cell extract was loaded onto a
DEAE Sepha‐
cel
TM
column equilibrated with the same phosphate buffer. The absorbed proteins were
eluted using a linear gradient of NaCl from 0 to 500 mM and a final washing with NaCl 1
M, with a flow rate of 15 mL h
−1. All fractions that presented amylase activity were col‐
lected and used as the cellular amylase source. In this case, the specific activity was in‐
creased from 20 U mg
−1 in the crude cell extract to 120 U mg
−1 in the partially purified
preparation, with a purification factor of 6. Determination of the protein content in all the
obtained extracts was performed according to the Bradford method [20], using bovine
serum albumin (BSA) as standard.
2.5. Amylase Activity Assay
Unless otherwise indicated, the amylase activity was measured following the
degra‐
dation of soluble starch by the standard iodine assay, based on the decrease of the absorb‐
ance at 580 nm of the iodine–starch complex produced by starch hydrolysis. The standard
reaction mixture contained 50 μL of enzyme solution, 100 μL of 1% (w/v) potato starch
solution in 20% NaCl,
and 100 μL of phosphate buffer (50 mM, pH 7, 20% NaCl). The
reaction mixture was incubated at 50 °C for 30 min, previously set as the best time to
conserve the linearity of the activity. The reaction was stopped by cooling on ice and 100
μL were employed to
reveal the remaining starch, by mixing 5 μL of four times diluted
commercial Lugol’s iodine solution with the sample. Thereafter, 1 mL of distilled water
was added to the sample before reading the absorbance at 580 nm. A standard curve was
prepared with soluble starch. One unit of amylase activity
was defined as the amount of
enzyme degrading one microgram of starch per minute from soluble starch, under the
assay conditions. To study the substrate specificity, potato starch was substituted by the
indicated compounds (carboxymethyl cellulose, sucrose, and lactose) at a concentration
of 1% (w/v), and incubated in the same
conditions.
To analyze the starch hydrolysis products, aliquots were withdrawn from the incu‐
bation mixture at the initial reaction time and after 2 h of incubation at 50 °C. The reaction
mixture contained 300 μL of the corresponding amylase extract and 600 μL of 1% (w/v)
potato starch solution
in 20% NaCl (w/v). The hydrolysis products were examined by a
Biology 2021, 10, 337 5 of 24
high‐performance liquid chromatographic (HPLC) system (Merck‐Hitachi LaChrom
Elite), equipped with a refractive index detector (Merck‐Hitachi L‐2490) and an Aminex
®
HPX‐87H Column (Bio‐Rad, Hercules, CA, USA), using an isocratic elution method with
5 mM H
2SO4 at 50 °C, and a flow rate of 0.6 mL min
−1. Glucose, maltose, and dextrin stand‐
ards were obtained from Merck, Sigma‐Aldrich (St. Louis, MO, USA).
2.6. Native Electrophoresis and Zymogram
The presence of amylase activity in the concentrated supernatant and the cell extract
was revealed by in situ staining of a native PAGE containing 0.2% of soluble starch in
the
separating gel. A volume of 15 μL of the sample was mixed with 5 μL of loading dye and
electrophoretically separated into two parallel gels of acrylamide, 10% supplemented
with starch 0.2% (w/v) and run at 130 V. After electrophoresis, one of the gels was incu‐
bated in
phosphate buffer (50 mM, pH 7, 20% NaCl) at 50 °C and 50 rpm for 1 h. Subse‐
quently, the gel was stained with commercial Lugol´s reagent and the appearance of clear
bands revealed the amylase activity. Meanwhile, the other gel was stained with 0.1% (w/v)
Coomasie Brillant Blue R
‐250 in 45% (v/v) ethanol‐10% (v/v) acetic acid, and faded with
25% (v/v) ethanol‐10% (v/v) acetic acid. Molecular markers (NativeMark
TM
Unstained Pro‐
tein Ladder, Thermo Fisher Scientific, Waltham, MA, USA) were used as a reference for
the molecular weight of proteins. Molecular mass estimation of the proteins was calcu‐
lated by plotting the log (MW) as a function of Rf (migration distance of the protein/mi‐
gration distance of the dye
front).
2.7. Effect of NaCl, Temperature, pH, Metals, and Detergents on the Amylase Activities of the
New Isolated Strain Haloarcula sp. HS
The effect of salt concentration was evaluated until a maximum of 32% NaCl with
intervals of 4% salinity increase. The desired NaCl concentration was obtained by adding
the required
NaCl to the phosphate buffer (50 mM, pH 7) and to the 1% (w/v) starch solu‐
tion. The influence of temperature on cell‐associated and extracellular amylase activities
was studied in phosphate buffer (50 mM, pH 7, 20% NaCl), over the range of 30–80 °C,
with temperature increments of
10 °C. For pH studies, amylase activity was measured at
50 °C and 20% of salt, in the following buffers—50 mM acetate for pH 2 and 3; 50 mM
MES for pH from 4 to 6; and 50 mM Tris‐HCl for pH from 7 to 11. All the assays were
done at least in triplicates, and the results were presented as a percentage of relative ac‐
tivity.
To test the influence of different metals on cell‐associated and extracellular amylase
activities MgSO
4, CaCl2, CuCl2, FeCl2, FeCl3, or EDTA (ethylenediaminetetraacetic acid),
were added to the reaction mixture, in a final concentration of 10 mM. Similarly, the effect
of various surfactants were studied, including Tween20 (Polyoxyethylene (20) sorbitan
monolaurate), Tween80 (Polyoxyethylene (80) sorbitan monooleate), Triton‐X100 (2‐[4‐(2,
4, 4‐trimethylpentan‐2‐yl) phenoxy] ethanol), CHAPS (3
‐[(3‐Cholamidopropyl) dime‐
thylammonio]‐1‐propanesulfonate), SB‐12 (N‐Dodecyl‐N,N‐dimethylammonio‐3‐pro‐
pane sulfonate), and SDS (sodium dodecyl sulfate), in a final concentration of 0.5% (w/v).
Amylase residual activity was measured as previously detailed for the standard assay and
expressed as a percentage, with respect to a
control sample incubated in the absence of
additives. All the determinations were conducted in triplicates.
2.8. Proteomic Analysis
For the proteomic analysis, the concentrated supernatant and the partially purified
cell extract fractions with starch degrading activity were dialyzed for 48 h, against a solu‐
tion of 1% NaHCO
3 and 0.01% EDTA in milli‐Q water, to eliminate excess salt. The pro‐
teins were first precipitated with TCA/acetone and resuspended in ammonium bicar‐
bonate‐trifluoroethanol (50%). After that, the proteins were treated with dithiothreitol, 10
mM, and methyl ethanethiosulfonate, 10 mM. Prior to trypsin digestion, the samples were
Biology 2021, 10, 337 6 of 24
diluted with ammonium bicarbonate 25 mM, until the concentration of trifluoroethanol
was under 5%. The digestion with trypsin was done overnight at 37 °C. Subsequently, the
samples were analyzed by LC‐MS/MS in a triple quadrupole‐TOF system (5600 plus, AB‐
Sciex), equipped with a nano‐electrospray ion source, coupled
to a nano‐HPLC (Eksigent).
The Analyst TF 1.7 software was used for equipment controlling and data acquisition.
Peptide mass tolerance was set to 25 ppm and 0.05 Da, for fragment masses, and only 1 or
2 missed cleavages were allowed. The peptide and protein identifications were performed
using the Protein
Pilot software (version 5.0.1, SCIEX), with the Paragon algorithm. The
search was conducted against the Uniprotproteome_Haloarcula_hispanica database
11_24_2020. The false discovery rate (FDR) was set to 0.01 for both peptides and proteins.
Protein comparison was performed with the Basic Local Alignment Search Tool for pro‐
teins (BLASTp) of the NCBI
(National Center for Biotechnology Information). The ob‐
tained sequences were analyzed using the CLC Workbench software (version 8, Qiagen).
2.9. Identification of Amylase Coding Genes Based on Protein Sequences
With the aim of completing the full protein sequences of the amylases identified, the
sequences of their encoding genes were amplified by
PCR, using sets of primers specifi‐
cally designed on the basis of the sequences of peptides obtained in the proteomic analy‐
sis. Concretely, six pairs of primers were employed to cover almost the full length of the
DNA sequences of the three amylases found, obtaining two overlapping sequences for
each
amylase gene (Table 1).
Table 1. Sequences of the primers employed for the amylase encoding genes amplification.
Primers Forward (5′‐3′) Reverse (5′‐3′)
AMY_HS1
ACCGGCAGTAAGCAGGCGTCTC GGCGGCGTCCCAGCGAATACC
GGCTCGTCGGGCTGAAGGACC CCCTCTCGCTCGTAGACGTACAGGTC
AMY_HS2
CGTCGGCGAATCGGTCGAACT GTCGCGTTTCCGGTTCCACTGTC
GGAACGCGACAGTGGAACCGGA CGAAGTGCAGAACGACCACGAGCG
AMY_HS3
GGAGACGGCCCGGTCGAACA CGCGTCGAAGGGCGATTC
GCCGGCGATAGCGACGAAT TCGTACGGGATTCGGAGGAGG
Primer sets used for PCR amplification of the three amylase coding genes found in Haloarcula sp. HS. For each gene
(AMY_HS1, AMY_HS2, and AMY_HS3), two pairs of primers were designed on the basis of the sequences of the peptides
obtained by proteomics.
Polymerase chain reactions were performed in a total volume of 25 μL, using an Ep‐
pendorf thermo‐cycler. The reaction mixtures contained—1 μL of genomic DNA, 0.2 U
REDTaq
®
DNA polymerase from Sigma Aldrich (St. Louis, MO, USA), and 2.5 μL of its
specific 10× buffer that contained 10 pM of each primer, 0.2 mM dNTPs, and 2.5 mM
MgCl
2. The thermal profile was set to 0.5 min at 96 °C, 0.5 min at 62 °C, and 1 min at 72 °C
for 30 cycles, followed by 10 min of final extension. The PCR products were analyzed by
electrophoresis on a 1% agarose gel and sent to Stabvida (Lisbon, Portugal)
for Sanger
sequencing. The sequences obtained were translated to protein and both, DNA and pro‐
tein sequences, were compared to those available at National Center for Biotechnology
Information (NCBI) databases, using the Basic Local Alignment Search Tool (BLAST). Fi‐
nally, different alignments were conducted in the CLC Workbench software (version
8,
Qiagen), the predicted structural models were built using the Phyre2 [21] and NetSurfP
[22] online web servers, and three‐dimensional (3D) molecular graphics were analyzed in
the UCSF Chimera version 1.15 [23] (University of California, Oakland, CA, USA). Physi‐
cochemical characteristics of the proteins were obtained using the ProtParam tool
(ExPASy) [24].
Biology 2021, 10, 337 7 of 24
2.10. Starch Hydrolysis from Bakery Waste
Bread from bakery waste was chosen for the present experiment. Bread crumbs were
dried on a stove at 70 °C and milled in a porcelain mortar to obtain a fine powder. Starch
was recovered by mashing dried crumbs in distilled water, in saltwater at
20% NaCl (w/v),
and in saturated saline solution (33% NaCl). The ability of the extracellular amylase of
Haloarcula sp. HS to degrade the starch from bread was comparatively tested against a
commercial α‐amylase (Megazyme cat. no. E‐BSTAA). Starch and enzyme solutions were
mixed in a proportion of 1:1 (
v/v) in a final volume of 1 mL. The hydrolysis of the starch
was performed for 15 min at 60 °C in 50 mM acetate buffer pH 5. The amount of remaining
starch was measured by the iodine‐starch method. A control, containing starch recovered
from bread without the enzyme
solution, was incubated in the same conditions. All assays
were conducted in triplicates.
3. Results
3.1. Selection of Amylase‐Producing Haloarchaea Isolated from Odiel Salterns Ponds
An in vitro screening was carried out to select the best amylase‐producing strain,
among a collection of archaea previously isolated from the saltern ponds
of the Odiel
Marshlands (SW, Spain) with a salinity of 33%. Amylase activity of each isolate was
screened for 9 days on starch‐agar plates, as detailed in Material and Methods (Figure 1A).
Eight colonies showed a considerable amylase activity, and that with the largest halo was
selected and identified,
by amplification and sequencing of its 16S rRNA full‐length cod‐
ing gene (Supplementary Material, Figure S1), followed by the comparison of the obtained
sequence with the NCBI database using the BLASTn tool. The results showed that the
selected strain was closely related to the Haloarcula genus, showing 98% homology with
different species of this taxonomic group. Therefore, the new strain isolated was named
Haloarcula sp. HS.
Molecular phylogenetic analysis was performed using the Molecular Evolutionary
Genetics Analysis (MEGA X) [25], on a series of reference haloarchaeal species and on the
new isolated strain, Haloarcula sp. HS (Figure 1B). The hyperthermophilic
archaea Meth‐
anococcus vulcanus was used as an outgroup and the bootstrap was set at 1000 replicates.
The 16S rRNA encoding sequence of the isolate clustered with the corresponding genes
of the representatives of the Haloarcula genus, especially close to the species Haloarcula
hispanica.
3.2. Optimization of a Two‐Stage Culture
Strategy to Induce the Production of Amylase
In the studied archaea, significant levels of amylase activity were only found when
the biomass was grown under inductive conditions in the presence of starch. The culture
conditions that induced the production of amylases are widely studied for bacteria and
hyperthermophilic archaea, as
reviewed by Mehta and Satyanarayana [5], but more lim‐
ited information exists on the production of amylase in haloarchaea [26,27]. Most authors
agree that amylase production is growth‐associated and is strongly induced by starch.
To establish the best culture conditions for the production and excretion of amylase
by Haloarcula sp.
HS, the haloarchaea was cultured in a (i) rich medium, which contained
yeast extract and a (ii) minimal medium, in which the yeast extract was substituted by
ammonium acetate. In both cases, starch (3 g L
−1
) was added to the culture medium, as
detailed in Materials and Methods. The optical density of the cultures, protein secretion
into the media, and hydrolysis of extracellular starch were followed in both, rich and min‐
imal medium cultures. As shown in Figure 2, cell growth and protein secretion were
higher
when the microorganism was grown in the rich media, which contained yeast ex‐
tract. In this medium, the studied archaea reached the stationary phase of growth in about
4 days and excreted 3 mg L
−1 of proteins into the culture medium. The archaea cultured in
the minimal medium exhibited very slow growth and excreted much fewer proteins to
the culture medium, about 1 mg L
−1
, after 30 h of culture. However, starch degradation
Biology 2021, 10, 337 8 of 24
activity was much higher for the archaea cultured in the minimal medium, which despite
a much lower biomass, showed an initial starch degradation rate 5.5 times higher than
that of the rich medium. Although starch was completely hydrolyzed in both media, there
was a 24 h lag phase before starch
degradation started in the medium with yeast extract,
probably due to the presence of more easily assimilable carbon sources in this medium.
Figure 1. (A) In vitro selective screening for amylase‐releasing haloarchaea. Semi‐quantitative estimation of amylase ac‐
tivity from four haloarchaeal strains isolated from the Odiel Marshlands, grown on starch agar plates (top) and revealed
with Lugol´s iodine solution (down), as detailed in the Material and Methods section.
(B) Molecular phylogenetic analysis
by the maximum likelihood method. The tree represents a comparison among the complete 16S rRNA coding gene se‐
quences, including a series of reference haloarchaeal species and the new isolated strain, Haloarcula sp. HS. Multiple align‐
ments were generated by MUSCLE (MUltiple Sequence Comparison by
Log‐Expectation) and the tree was constructed
with MEGA X. The numbers at the nodes indicate the bootstrap values calculated for 1000 replicates. Arrows point to the
new strain Haloarcula sp. HS.
Biology 2021, 10, 337 9 of 24
Figure 2. Time course evolution of Haloarcula sp. HS cultures in rich and minimal media. Optical density (A), secretion of
proteins (B), and starch hydrolysis (C) were measured during the time of culture in rich (■) and minimal (
♦) broths. All
data are expressed as the mean ± SD of at least triplicate experiments.
For this reason, a two‐step culture was set up to get both, a high biomass and amylase
productivity. Cells were first grown in a rich medium in order to obtain a large amount
of biomass, and when the culture reached the end of the exponential phase of growth, at
the
third day of culture, the cells were transferred to the fresh minimal medium to induce
the production of amylase, and was cultured for another 4 days. Through this two‐step
approach, high starch consumption activity and a high protein excretion were achieved,
reaching an extracellular protein concentration of 10 mg
L
−1 and undetected levels of ex‐
tracellular starch, on the 7th day of culture (Figure 3).
Figure 3. Time course evolution of Haloarcula sp. HS in the two‐step culture. Optical density, se‐
creted proteins, and starch concentration were measured along the full cultivation time. The red
arrow indicates the moment of transference of the cells from the rich to the fresh minimum medium.
All data
are expressed as the mean ± SD of at least triplicate experiments.
3.3. Extracellular and Cell‐Associated Amylase Activities in Haloarcula sp. HS
To identify the enzyme responsible for the amylase activity and characterize its prop‐
erties, the haloarchaeal strain Haloarcula sp. HS was grown in a two‐stage culture with
starch (3 g L
−1), as previously described (Figure 3). Cell‐associated proteins and the con‐
0
1
2
3
0246810
O.D. at 580 mn
Days of culture
Minimal medium Rich medium
0
1
2
3
4
0 20406080
[Extracellular proteins] (mg L
−1
)
Time of culture (h)
Minimal medium Rich medium
0
1
2
3
4
0 20406080
[Starch] (mg L
−1
)
Time of culture (h)
Minimal medium Rich medium
BAC
Biology 2021, 10, 337 10 of 24
centrated extracellular proteins excreted into the cultured medium were electrophoreti‐
cally separated in a polyacrylamide gel containing starch. After electrophoresis, the gel
was split lengthwise with a razor blade. One half was stained with Coomassie blue and
the other with Lugol´s iodine solution to detect both proteins and amylase activity,
re‐
spectively. The zymogram analysis proved the presence of amylase activity in both sam‐
ples, the culture medium, and the crude extract. A unique band with amylase activity was
observed in the culture medium, after a 100‐fold concentration step through ultrafiltration
with a 10 kDa cut‐off membrane, as indicated
in Material and Methods. However, in the
crude extract, two bands with starch hydrolyzing activity were observed, indicating the
presence of several cell‐associated enzymes with amylase activity (Figure 4 and Figure
S2). The crude extract was partially purified through ion‐exchange chromatography in
DEAE Sephacel
TM, as indicated in the Materials and Methods section. All fractions with
amylase activity were pooled and used as the source of cellular amylase. The electropho‐
retic analysis of the purified extracts showed a unique band with amylase activity (Figure
4A). The size of the observed bands was between 20 and
27 kDa, however, the protein
mobility was strongly affected by the starch added to the polyacrylamide gel and these
apparent sizes observed were not representative.
Figure 4. Native‐PAGE and zymogram of cell‐associated (A) and extracellular (B) proteins obtained
from Haloarcula sp. HS. Lanes 1, 3, and 5—samples on native‐PAGE followed by Coomassie Blue
staining. Lanes 2, 4, and 6—samples on native PAGE followed by Lugol´s solution staining. Lane
M—molecular mass
marker in kDa, lanes 1 and 2—crude extract, lanes 3 and 4—partially purified
crude extract, and lanes 5 and 6—concentrated supernatant. The whole gels for each staining are
available in Supplementary Material, Figure S2.
3.4. Characterization of Extracellular and Cell‐Associated Amylase Activities
A series of in vitro experiments were carried out with the extracellular and cellular
amylase‐enriched extracts to characterize the hydrolysis products, the substrate specific‐
ity, and the optimal kinetic parameters of the amylase activity of these extracts. Both, ex‐
tracellular and
cellular, enzymatic preparations were incubated with 1 mg of starch in the
standard conditions, described in Material and Methods, excepting that the incubation
time was fixed at 2 h. The products of starch hydrolysis were identified by HPLC and an
IR detector, as detailed in Materials and Methods. A parallel
reaction with a commercial
Biology 2021, 10, 337 11 of 24
α‐amylase purchased from Megazyme (cat. no. E‐BSTAA) was done in the same condi‐
tions for comparison. The results (Table 2) revealed that starch degradation was almost
complete in all cases, being especially efficient in the case of the extracellular amylase ex‐
tract, with a remaining starch of only 1.7%.
The main product obtained with the three
enzymatic sources was maltose, which represents between 73.8% and 86.1% of the total
carbohydrate content in the reaction mixtures. In addition, the enzymatic preparation
from the cellular extract was also able to catalyze the liberation of glucose (6.6%). On the
other hand, the
extracellular extract and the commercial reference amylase catalyzed the
liberation of dextrins, which supposed 18.5% and 20.8% of the total carbohydrate content
in the reaction mixture, respectively, in addition to maltose. No glucose was found as the
end product in these reactions (Table 2).
Table 2. Percentage of the different products obtained from starch hydrolysis.
Dextrins (%) Maltose (%) Glucose (%) Starch (%)
Cell‐associated amylase ND 86.1 ± 3.6 6.6 ± 0.8 7.3 ± 2.9
Extracellular amylase 18.5 ± 1.1 79.7 ± 1.3 ND 1.7 ± 0.2
Commercial α‐amylase 20.8 ± 3.1 73.8 ± 3.4 ND 5.4 ± 0.3
Comparison of the products obtained from the hydrolysis of starch by cell‐associated or extracellular partially purified
extracts of Haloarcula sp. HS and a commercial α‐amylase. The percentage (%) of dextrins, maltose, and glucose produced
and the remaining starch are indicated as the mean of three replicates with the corresponding standard
deviation. ND,
not detected.
With respect to the substrate specificity, neither extracellular nor cellular Haloarcula
sp. HS extracts were able to hydrolyze other glucose polysaccharides, such as carboxyme‐
thyl cellulose, or disaccharides, such as sucrose or lactose, which not contain alpha‐1,4‐
linked glucose.
The most characteristic feature of halophilic enzymes is their ability to
operate under
very high salinities. As it is shown in Figure 5, the extracellular enzymatic preparation
presented the optimal activity at 28% salt and retained less than half of its activity when
the salt content was under 20% (Figure 5A), showing that it is more salt‐dependent than
the cellular
one, which instead showed the maximum activity at 16% salinity and retained
more than 60% of its activity at all salinities studied (Figure 5B).
The influence of pH in both amylase activities was studied in different buffers, as
detailed in Materials and Methods. The optimal activity was found at pH
5 for the extra‐
cellular amylase, and at pH 7 for the cellular amylase‐enriched extract, as shown in Figure
5C,D, respectively. It should be noted that the cell‐associated activity was stable under a
wide range of pH values, retaining more than 50% activity at pH values between 2 and
11, and more than 80% activity at pH values comprising pH 5 to 9. Contrarily, the extra‐
cellular amylase activity appeared to be more susceptible to extreme pH, losing more than
50% of its activity both at low and high pH values.
The effect of the temperature on the amylase
activities showed, once again, that the
extracellular amylase activity had a higher dependence on the physicochemical parame‐
ters of the assay than the cell‐associated one. The extracellular amylase activity showed
an optimal temperature of 60 °C, losing more than 65% of its activity below 50 °C or above
60 °C
(Figure 5E). The cellular amylase‐enriched extract, on the other hand, conserved a
high activity over a wide range of temperatures, retaining more than 75% of activity from
30 to 80 °C (Figure 5F). This weak temperature dependence was due to the fact that a mix
of three different cell‐
associated enzymes could contribute to the amylase activity, as later
shown by the proteomic analysis of the cellular enzymatic preparation.
Biology 2021, 10, 337 12 of 24
Figure 5. Effect of salt, pH, and temperature on amylase activities. Relative amylase activities in
different salt concentrations (%, w/v), pH values, and temperatures are shown for the extracellular
(A,C,E) and cellular (B,D,F) extracts. Relative activity was defined as the percentage of maximum
activity
for each case. The 100% activity corresponded to 70 ± 6.4 U mL
−1
(350 U mg
−1
) for the extra‐
cellular amylase extract and 60 ± 5.6 U mL
−1
(120 U mg
−1
) for the mix of cell‐associated amylases.
Mean and standard deviations are shown.
0
20
40
60
80
100
0 5 10 15 20 25 30 35Relative amylase activity (%)
NaCl (%)
0
20
40
60
80
100
0 5 10 15 20 25 30 35
Relative amylase activity (%)
NaCl (%)
A B
0
20
40
60
80
100
23456789Relative amylase activity (%)
pH
0
20
40
60
80
100
234567891011Relative amylase activity (%)
pH
C D
0
20
40
60
80
100
30 40 50 60 70 80
Relative amylase activity (%)
Temperature (°C)
0
20
40
60
80
100
30 40 50 60 70 80Relative amylase activity (%)
Temperature (°C)
EXTRACELLULAR AMYLASE ACTIVITY INTRACELLULAR AMYLASE ACTIVITY
E F
Biology 2021, 10, 337 13 of 24
3.5. Effects of Metals and Surfactants on the Amylase Enzymatic Activities
Many microbial α‐amylases are reported to be calcium‐dependent metalloenzymes
[9]. Therefore, the effect of EDTA, Ca
2+
, and other metallic ions including Mg
2+
, Cu
2+
, Fe
2+
,
and, Fe
3+ was tested. The results revealed that the addition of Ca
2+, Fe
2+, or Mg
2+ causes a
slight increase in the amylolytic activity of both extracellular and cellular enzymatic ex‐
tracts (Figure 6). The possible existence of divalent metals in the partially purified enzy‐
matic preparations makes it difficult to obtain accurate conclusions on the effects of these
metals on the amylase activities of Haloarcula
sp. HS. However, the strong inhibition ob‐
served in the presence of the metal chelating agent EDTA confirmed the divalent cation
dependence of both, extracellular and cell‐associated amylase activities, which decreased
to 39% and 33%, respectively, in the presence of EDTA.
On the other hand, the presence of Cu
2+ and Fe
3+ in the reaction mixture, at a concen‐
tration of 10 mM, caused a drastic reduction in both, extracellular and cell‐associated am‐
ylase activities (Figure 6). The cellular extract only retained 28% of its amylase activity in
Fe
3+; and similarly, the extracellular amylase conserved 26% of its activity. Under the pres‐
ence of the cupric ion, the activity of the extracellular amylase dropped to 8%, while the
activity of the cell‐associated amylase decreased to 33%. This fact suggests the involve‐
ment of tiol/carboxyl groups, typically inhibited by
Cu
2+, in the function of the enzymes
[28].
Figure 6. Effect of metal ions on the amylase activities. Extracellular and cell‐associated relative am‐
ylase activities under the presence of different metal ions and EDTA (10 mM) are represented. Rel‐
ative activity was defined as the percentage of maximal activity with respect to control, with no
additives. The control
activity was 70 ± 6.4 U mL
−1
(350 U mg
−1
) for the extracellular amylase and 60
± 5.6 U mL
−1
(120 U mg
−1
) for the cellular amylase. Mean and standard deviations are shown.
The in vitro effect of anionic (SDS), cationic (SB‐12), zwitterionic (CHAPS), and no
ionic (Tween 20, Tween 80, and Triton X‐100) detergents on the amylase activities were
assayed. The obtained results revealed that both activities were very stable in different
detergents, retaining more than 80% activity in all, with
the exception of SDS, which
caused a decrease by almost half in the amylase activities of both, extracellular and cellular
amylase enriched extracts.
3.6. Identification of Amylases in Cellular and Extracellular Concentrated Extracts of
Haloarcula sp. HS by a Proteomic Approach
A proteomic study of both, extracellular and cellular partially
purified extracts with
amylase activity, was carried out to identify the sequence of the proteins responsible for
0
20
40
60
80
100
120
140
Control MgSO4 CaCl2 CuCl2 FeCl2 FeCl3 EDTA
Relative amylase activity (%)
Cell‐associated amylase activityExtracellular amylase activity
Control MgSO
4CaCl
2CuCl
2 FeCl
2 FeCl
3EDTA
Biology 2021, 10, 337 14 of 24
these starch‐degrading activities in Haloarcula sp. HS. Both samples were submitted to
tryptic digestion and analyzed by LC‐MS/MS in a triple quadrupole‐TOF system, as de‐
scribed in Materials and Methods.
The results revealed that the main extracellular protein secreted to the culture media
was α‐amylase. The rest
of the proteins identified in the extracellular fraction were mem‐
brane‐ligated proteins, probably from broken cell remains. This extracellular amylase
(AMY_HS1) was identified by 42 unique peptides, presenting 73.56% identity and 60%
query cover with the α‐amylase of Haloarcula hispanica N601 (UniProt: V5TMJ3_HALHI).
For its part, in the cellular fraction,
three different amylases were found. One of them
corresponded to the same α‐amylase (UniProt: V5TMJ3_HALHI) detected in the extracel‐
lular fraction, which in this case was identified according to 13 unique peptides and
showed 81.48% identity and 36% query cover. The other two proteins, denoted as
AMY_HS2 and AMY_HS3, showed
high homology with two different α‐amylases of Halo‐
arcula hispanica N601 (UniProt codes: V5TRA6_HALHI and V5TQD3_HALHI, respec‐
tively), both determined according to 15 unique peptides. AMY_HS2 presented 45.25%
identity and 65% query cover with V5TRA6_HALHI, while AMY_HS3 showed 75.76%
identity and 31% query cover with V5TQD3_HALHI.
Due to the low percentage
of protein covering achieved, the sequences of amylase
coding genes were amplified by PCR, as detailed in Material and Methods, with primers
designed to target the peptide sequences identified by proteomics for each enzyme.
Through this approach, practically the full length of each protein sequence was com‐
pleted.
The alignment
of the three obtained protein sequences revealed that the extracellular
amylase only presented a 17% identity with the cell‐associated amylases, which in turn
showed a 38% identity between them. Nonetheless, as it is shown in their predicted three‐
dimensional (Figure 7) and secondary structures (Supplementary Material, Figure S3), the
three
amylase sequences from Haloarcula sp. HS conserve the typical structural domains
of the GH‐13 family, according to the Carbohydrate‐Active enZYmes (CAZY) database;
including the catalytic (β/α)
8‐barrel (TIM‐barrel) located in the domain A, a small domain
B located in the loop between the β
3‐strand and the α 3‐helix of the barrel, and the domain
C showing an antiparallel β‐sandwich structure in the C‐terminal end of the protein. In
addition to this typical conformation, the two cellular amylases show an N‐terminal do‐
main, which was reported in some maltogenic amylases and seems to be involved in
in‐
creasing the binding of the enzyme to raw starch. This N domain forms a large groove,
the N–C groove, which might be responsible for thermostabilization via oligomerization
and substrate affinity modifications in some microbial maltogenic amylases [29]. For its
part, the extracellular protein, AMY_HS1, presents the conserved TAT (Twin‐Arginine
‐
Translocation) motif and its corresponding processing site (Supplementary material, Fig‐
ure S4).
Regarding the physicochemical characteristics, the three alpha‐amylases of Haloar‐
cula sp. HS had a low isoelectric point, negative net charge, and low hydrophobicity (Table
3) as other haloarchaeal enzymes.
Table 3. Physicochemical properties of the amylase enzymes from Haloarcula sp. HS.
Name N MW (kDa) IP Z GRAVY Aliph. Index
AMY_HS1 393 43.70 4.27 −35.577 − 0.518 72.72
AMY_HS2 639 70.16 4.43 −66.227 − 0.382 73.43
AMY_HS3 549 60.02 4.37 −59.604 − 0.463 71.89
Main physicochemical characteristics of the amylase enzymes found in the Haloarcula sp. HS
strain. N, number of nucleotides; MW, molecular weight; IP, theoretical isoelectric point; Z, net
charge; GRAVY (Grand average of hydropathicity), mean of the hydropathy index of each amino
acid residue. The aliphatic index stands for the relative volume
occupied by the aliphatic side
chains.
Biology 2021, 10, 337 15 of 24
Figure 7. Three‐dimensional predicted structures of the three amylase sequences identified in Halo‐
arcula sp. HS; (A) mature extracellular amylase, AMY_HS1; (B) cellular amylase, AMY_HS2; (C)
cellular amylase, AMY_HS3. Phyre2 software and the Chimera program were employed for 3D
structure visualization. Helixes are represented by blue ribbons
and strands by red arrows.
Moreover, although the three protein sequences had a low percentage of sequence
homology, they conserved the catalytic triad of aspartate, glutamate, and aspartate in the
active site, along with other conserved residues that were described to be indispensable
for the structure of the enzyme [30]. These conserved amino acids are shown
in Figure 8.
One of the first consensus amino acids found is aspartic acid, which is essential for active
site integrity. This aspartic residue is in the position Asp92 for the mature AMY_HS1 pro‐
tein after TAT processing, and occupies the position 359 and 303 in AMY_HS2 and
Biology 2021, 10, 337 16 of 24
AMY_HS3, respectively (Asp92/359/303). Following the same notation, the rest of the con‐
served amino acids are distributed as follows—Asn96/363/307, which coordinates the con‐
served calcium ion between the A and B domains [31]; and His93/360/304, which stabilizes
the interaction between the C‐terminal of β3 and the rest of TIM
barrel through hydrogen
bonding to Asn61/328/272 and the backbone oxygen of Tyr57/324/268. The first catalytic
residue is Asp177/446/379, located in β4, which is preceded by Arg175/444/377, both of
which are indispensable amino acids for the catalytic activity. Lys and His are usually
present in this region in the position Asp+3 and
+4, binding the reducing end of the glu‐
cose chain in the substrate‐binding cleft [32], however, these residues were only found in
the extracellular amylase. The second catalytic residue is the proton donor
Glu205/475/408, which lies in the fifth L‐strand of the TIM‐barrel. The following conserved
residues
protect the active site from the solvent and contain the last catalytic amino acid
Asp268/539/471, postulated to be involved in substrate binding, substrate distortion, and
in elevating the pKa of Glu205/475/408. This residue is usually accompanied in α‐amyl‐
ases by His, Asn, Val, and Phe in the positions −1, −2, −4,
and −5, respectively, as it occurs
in AMY_HS1, while in AMY_HS2 and AMY_HS3, Phe is substituted by Tyr, and Val is
changed by Ala in AMY_HS3. Finally, the other two conserved residues were found in
Gly287/564/496, followed by Pro289/566/498 [33–36].
Figure 8. Alignment of the three amylase sequences from Haloarcula sp. HS. The mature extracellular protein is named
AMY_HS1, while the cell‐associated amylases are denoted as AMY_HS2 and AMY_HS3. Purple stars highlight the cata‐
lytic triad (Asp‐Glu‐Asp), blue star denotes the canonical calcium‐binding site and the black
stars point other essential
residues for enzyme structure. Identical residues in the three sequences are shaded in white, residues that coincide in two
of the sequences or do not coincide at all are shaded in pink and red, respectively.
Finally, a multispecies study was carried out to compare the degree of conservation
among the amylases reported and those from different haloarchaeal members. The pro‐
tein sequence of the extracellular amylase (AMY_HS1) was aligned and compared to var‐
ious extracellular amylase sequences available in the NCBI database, selecting different
representatives of
the order Halobacteriales (Supplementary Material, Figure S4). Among
these amylase sequences, different percentages of identity were found with the extracel‐
Biology 2021, 10, 337 17 of 24
lular amylases from Haloarcula hispanica N601 (90%), Halomicroarcula salina (73%), Halapri‐
cum salinum (57%), and Haloterrigena turkmenika (40%). Likewise, the two cell‐associated
amylases (AMY_HS2 and AMY_HS3) from Haloarcula sp. HS showed the following per‐
centages that identity with the amylase sequences from Haloarcula hispanica N601 (90 and
80%, respectively),
Halomicroarcula salina (70 and 65%, respectively), Haloferax mediterranei
(59, and 50%, respectively), Halogeometricum rufum (57% with AMY_HS2), and Haloge‐
ometricum limi (49% with AMY_HS3) (Supplementary Material, Figures S5 and S6). In all
alignments, it could be appreciated that the catalytic regions were conserved among the
different haloarchaeal species (Supplementary Material, Figures
S4–S6).
3.7. Hydrolysis of Bakery Waste
Bakery residues, including dough, flour dust, burnt or rejected bread, and biscuits,
can be exploited for the production of fermentable sugars. Among them, bread was cho‐
sen as the substrate for this assay, as it is one of the most abundant food waste products
worldwide. In addition, many of the discarded by‐products during the bread manufac‐
turing process are fundamentally constituted by starch, such as substandard bread and
the bread crust removed to make special types of sandwich bread [37].
The extracellular amylase activity was selected for this assay, as it works better in
high NaCl concentrations than the cell‐associated amylase. This was compared to a com‐
mercial thermostable α‐amylase. Sodium chloride was found to have a complex effect on
the gelatinization and rheological properties of starch. Some studies pointed out that the
enthalpy for the gelatinization process decreased at high salt concentrations,
an effect of
great importance for the production and properties of several cereal‐based products, as
well as for the manufacture of modified starches [38].
The results proved that a high percentage of the initial starch (75–85%) was hydro‐
lyzed by both enzymes, with maximum activities above 100 U mL
−1. However, the con‐
centration of salt was determined in their maximum activities. The extracellular amylase
hydrolyzed around 75% of the starch under 20% salt or even under salt saturation, with
maximum activities of 107 and 105 U mL
−1, respectively; while when no salt was added,
the degradation rate decreased to 22%, with an activity of 24.7 U mL
−1. Conversely, the
commercial α‐amylase presented the highest activity (101.8 U mL
−1
) when no salt was
added to the mixture, hydrolyzing 85% of the starch; and its activity dropped drastically
to 6–7 U mL
−1
, under elevated salt concentration, degrading only 5–6% of the starch.
The obtained hydrolysate was very rich in simple sugars that could now be used for
the bioproduction of many high‐value molecules like glycerol, hydrogen, ethanol, or lactic
acid, among others. These molecules are required for different purposes, including renew
‐
able energy sources, fuel additives, and food preservers [13]. To our knowledge, this study
supposes the first attempt to use a halophilic amylase to degrade starch from bread into
simple sugars. The applicability of the amylase from a haloarchaea was tested using starch
from agricultural waste [39], however, the starch
content of this residue was considerably
lower, and also the starch extraction method was costlier.
4. Discussion
Alpha‐amylases are a large family of endo‐glycosyl hydrolases that cleave the inter‐
nal α,1‐4 glycosidic bonds between the glucose units in polysaccharides, such as amylose
and amylopectin. They are common in
all kingdoms of life and their general properties,
three‐dimensional structures, and mechanisms of action are extensively reviewed [40–43].
Alpha‐amylases are particularly spread among microbial species, such as bacteria
and yeasts. These amylases are often extracellular enzymes that allow microorganisms to
use environmental polysaccharides for their nutrition. Thermostable α‐amylases
were
produced and commercially exploited from yeast and bacterial species, such as Bacillus
subtilis, B. licheniformis, or B. amyloliquefacines [14]. Some archaea, including halophilic ar‐
chaea, were found to produce halotolerant α‐amylases, which in addition were highly or
Biology 2021, 10, 337 18 of 24
moderately thermotolerant. The extracellular α‐amylases of Haloarcula sp. S‐1 [44], N. am‐
ylolyticus [45], H. mediterranei [26], H. xinjiangense [46], Haloferax sp. HA10 [47], H.turk‐
menica [39], or Halococcus GUVSC8 [48] are some examples. Although much less studied,
the intracellular α‐amylases of some haloarchaea, such as Haloarcula japonica [49]
were also
characterized.
The new strain isolated from the Odiel Marshlands and selected by its high ability to
degrade starch in iodine–starch agar plate assays was foumd to be closely related to the
genus Haloarcula, as shown in the phylogenetic tree (Figure 1). The high homology of its
16S rRNA
with that of Haloarcula hispanica (98%) or Haloarcula japonica (97%) confirmed it.
We found that the new strain exhibited a high amylolytic activity when cultured in the
presence of starch; this activity was higher in a minimal medium with ammonium acetate
(Figure 2).
In agreement with our observations, several reports
indicate that amylase production
in haloarchaea is induced by starch, as described for Halorubrum [46], Haloferax [47] Halo‐
arcula [50], and some halophilic bacteria [51]. Other culture conditions and nutrients that
were reported to influence the induction of alpha‐amylases are nitrogen, metal ions, or
phosphate [5]. Pérez‐Pomares et al.
[26], for example, reported low amylase excretion
when using a minimal medium containing ammonium acetate as carbon and nitrogen
source, plus starch in Haloferax mediterranei.
Our observations indicate that the conditions that induce the amylase activity are not
the best for growth (Figure 2). Therefore, a two‐step culture was
set up, in which a large
amount of biomass was obtained in a rich medium, followed by transfer to a minimal
medium with starch, to induce the production of amylase activity (Figure 3). This two‐
stage method allowed the improvement of amylase production, and at the same time fa‐
cilitated the
recovery of the extracellular enzyme. Since the minimal medium had no yeast
extract, there are no foreign proteins that could interfere with the amylase secreted into
the culture medium.
Partially purified extracellular and cellular‐amylase‐enriched extracts were obtained
from Haloarcula sp. HS, through ultrafiltration and anion exchange chromatography, re‐
spectively, and were electrophoretically separated in native conditions. In situ staining of
the obtained acrylamide gel allowed the identification of bands with starch degrading
ability, one band in the extracellular enzymatic preparation, and two bands in the cellular
extract, with apparent relative molecular masses between 21.6 and 30.4 kDa (Figure 4).
The zymogram indicates that the new isolated strain presents amylase activities in both,
the supernatant and the crude cell extract, suggesting that there is more than one cell‐
associated amylase that does not coincide with the extracellular one. The proteomic anal‐
ysis of the extracts and the subsequent amplification of
the whole gene sequences that
encode for these amylases allowed us to confirm this hypothesis (Figure 8), and also indi‐
cated that the real masses of the amylases were much higher than the apparent molecular
masses shown in the electrophoresis gel. These differences could be due to the fact that
the
electrophoresis was carried out in native conditions and in the presence of starch,
which could modify their electrophoretic mobilities, besides the fact that halophilic pro‐
teins usually show altered electrophoretic properties [52].
The apparent molecular masses reported for the amylases of other haloarchaea are
slightly higher than the molecular weight
observed for the amylases of Haloarcula sp. HS.
For example, the intracellular amylase of Haloarcula japonica presented a molecular mass
of 83 kDa [49]; the extracellular amylases of Haloterrigena tukmenica and Haloferax sp. HA10
showed a molecular weight of 66 kDa [39,47], in Haloferax mediterranei, the weight of the
enzyme
was around 50–58 kDa [26], 60 kDa in Halorubrum xinjiangense [46], and 74 kDa in
Natronococcus sp. Ah‐36 [45], while in the Haloarcula species, the molecular mass varied
from 43 to 70 kDa [27,44]. The physicochemical parameters of the new alpha‐amylases,
with low isoelectric point and negative net charge
(Table 3) also meet the usual character‐
istic of haloarchaeal enzymes, as reported by other authors. Yan and Wu [43] analyzed the
sequences of 88 α‐amylases from archaea and observed that amylases from haloarchaea
Biology 2021, 10, 337 19 of 24
have a highly negatively charged surface, and a higher percentage of acidic residues as a
mechanism of adaptation to high salinity. Other authors describe similar features for H.
hispanica, which has an extracellular amylase with an isoelectric point of 4.2 and a low
level of aromatic and hydrophobic residues [27].
In haloarchaea, most studies about am‐
ylases focused on extracellular enzymes, given that sometimes no activity was found in
the crude cell extract, as was observed in Haloferax mediterranei and Halorubrum xin‐
jiangense [26,46] or because, as in the case of Haloterrigena turkmenica, the amylase activity
was quite higher in
the supernatant than in the cell extract [39]. With respect to the Halo‐
arcula genus, Hutcheon et al. confirmed the overexpression of an extracellular amylase
(AmyH) in the mutant strain Haloarcula hispanica B3, which was secreted in a folded con‐
formation via the TAT (Twin‐Arginine‐Translocation) pathway, indicating that
it was ac‐
tive in the cytoplasm before the secretion to the media [27]. Additionally, Onodera et al.
reported the overexpression of an intracellular amylase (malA), which was not secreted
to the media in Haloarcula japonica [49]. Based on the above mentioned, it seems that there
were diverse amylases with probably
different modes of action, which to our knowledge,
are not yet deeply elucidated.
Maltose is the main end‐product released from the starch hydrolysis by the extracel‐
lular and cellular partially purified amylase extracts of Haloarcula sp. HS (Table 2). This is
the main product of maltogenic α‐amylases, like most α‐
amylases from haloarchaea, e.g.,
intracellular α‐amylase from Haloarcula japonica [49] or the extracellular α‐amylase from
Haloterrigena tukmenica [39]. A small proportion of glucose was found in the assay cata‐
lyzed by the cell extract. However, it is difficult to predict if it is directly due to the action
of the cellular
amylases in Haloarcula sp. HS or due to the contribution of additional cell‐
associated enzymes.
With regards to the optimal enzymatic parameters, both cell‐associated and extracel‐
lular starch‐degrading activities exhibited their maximum activities around 60 °C. The
low‐temperature dependence of the cell‐associated amylase, which only loses
around 25%
of its activity in the temperature range 20–80 °C, is noteworthy. However, it should be
considered that this activity might be, as shown in this study, the result of the action of
three different amylase enzymes. This fact contributes to broadening the range of optimal
temperatures for the cell
‐associated amylase activity. High retention of enzyme activity
over a wide range of temperatures was reported for other partially purified amylases,
such as that from Haloferax sp. HA10 [47], which showed the highest amylase activity at
55 °C.
The optimal pH values were 7 for the cellular amylase activity, and 5 for the extra‐
cellular activity. Additionally, extracellular and cell‐associated, amylase activities were
extremely halophilic, showing their maximum activities at 25% NaCl. Therefore, it is note‐
worthy that the cell‐associated amylase activity seemed to be
more tolerant to changes in
salinity, pH, and temperature than the extracellular one (Figure 5). This was probably due
to the presence of three different amylases in the cell extract, as was later confirmed in the
proteomic analysis. In addition, the high salt and temperature tolerance could be of inter‐
est for many industrial applications in which these extreme conditions are needed.
A comparison with the optimal parameters reported for amylases of other related
haloarchaea is summarized in Table 4. Most extracellular α‐amylases from haloarchaea
showed their best activity at temperatures from 45 to 70 °C, pH from 6.5 to 8.7,
and in salt
concentrations from 1 to 5 M. The extracellular amylase found in Haloarcula sp. HS is one
of the most halophilic and acidophilic α‐amylase described within the haloarchaea group
(Table 4).
Additionally, extracellular and cell‐associated amylase, activities from Haloarcula sp.
HS, exhibit a strong inhibition in the
presence of the metal chelating agent EDTA (Figure
6). In addition, the analysis of the amylase sequences obtained allowed the identification
of the canonical Ca‐binding residue in the three amino acidic sequences, as shown in Fig‐
ure 8, indicating that they must be calcium‐dependent. Dependence of calcium is a
com‐
mon feature within haloarchaeal amylases, as was reported for Haloferax mediterranei,
Biology 2021, 10, 337 20 of 24
Haloarcula hispanica, Haloarcula sp. S‐1, Halorubrum xijiangense, and [26,27,44,46]. However,
some haloarchaeal amylases showed to be resilient to EDTA, indicating no dependence
on Ca
2+
, as revealed by the studies on Haloterrigena turkmenica and Haloarcula japonica
[39,49].
Furthermore, both amylase activities showed high stability in most tested surfac‐
tants, excepting the anionic detergent SDS. There are few reports on the stability of amyl‐
ase from haloarchaea on surfactants. In this context, detergent‐stable amylases were
re‐
cently found in Haloterrigena tukmenica, Halorubrum xijiangense, and Haloferax sp. HA10
[39,46,47]. Additionally, a surfactant‐stable amylase was characterized from the halophilic
bacteria Nesterenkonia sp. F [53]. Therefore, to the best of our knowledge, the two deter‐
gent‐stable amylase activities described in this work entailed the first
report on this spe‐
cific feature of the amylase activity from the Haloarcula genus.
Table 4. Optimal parameters reported for α‐amylase activity in haloarchaea.
Microorganism Enzyme NaCl (M) pH Tª (°C) Ref.
Haloarcula sp. HS Cellular α‐amylase 2.6 7 50 This study
H. japonica Intracellular α‐amylase 2.6 6.5 45 [49]
Haloarcula sp. HS Extracellular α‐amylase 5 5 60 This study
Halococcus GUVSC8 Extracellular α‐amylase 2 6 45 [48]
Haloarcula sp. S‐1 Extracellular α‐amylase 4.3 7 50 [44]
H. hispanica B3 Extracellular α‐amylase 4–5 6.5 50 [27]
H. hispanica 2TK2 Extracellular α‐amylase 5 6.9 52 [50]
H. xinjiangense Extracellular α‐amylase 4 8.5 70 [46]
Haloferax sp. HA10 Extracellular α‐amylase 1 6 55 [47]
H. mediterranei Extracellular α‐amylase 3 7–8 50–60 [26]
H.turkmenica Extracellular α‐amylase 2 8.5 55 [39]
N.amylolyticus Extracellular α‐amylase 2.5 8.7 55 [45]
Analyzing the sequences of the peptides generated by the tryptic digestion of the
partially purified extracellular and cell extracts, it was possible to identify three different
amylases in Haloarcula sp. HS. One (AMY_HS1) was found both in cells and in the culture
medium, while the other two amylases (AMY_HS2 and AMY_HS3)
were exclusively
found in the cell extract.
Despite the low percentage of sequence identity that the three amylases of Haloarcula
sp. HS shared (Figure 8), the three enzymes exhibited a high three‐dimensional structure
homology, with the three typical domains of alpha‐amylases of the glycosyl hydrolase
GH‐13 family
(Figure 7) and many of the key conserved residues (Figure 8). For example,
the three amino acids (Asp‐Glu‐Asp), which constituted the active site of alpha‐amylases,
were identified in α‐amylases of haloarchaeal strains, such as Halogeometricum borinquense
[30] and Haloterrigena turkmenica [39]. These residues were conserved in all alpha
‐amyl‐
ases of the GH‐13 family of many different origins, which were compiled in the Carbohy‐
drate‐Active enZYmes (CAZy) database [54].
The calcium‐binding domain located between the 3rd beta‐strand and the 3rd alpha‐
helix contained an asparagine amino acid found in the three amylases of Haloarcula
sp. HS
(Figure 8), as was described for other haloarchaeal amylases [30] and many other α‐amyl‐
ases, which were calcium‐dependent metalloenzymes [55]. In some cases, more than one
Ca‐binding domain was described, as in H. orenii [56]. Additional conserved amino acids
reported in alpha‐amylases, which help to stabilize
the structure or the binding of the
substrate [5], were found in the three amylases of Haloarcula sp. HS (Figure 8).
Most available archaeal amylase sequences were from the thermophilic archaea and
could work at very high temperatures, which was of great industrial interest. The amyl‐
ases identified from the
Odiel Marshlands were medium or highly thermotolerant, in ad‐
dition to being extremely halophilic. The α‐amylases from hyperthermophilic archaea
were closely related to plant amylases [57]. However, as new potential α‐amylases from
Biology 2021, 10, 337 21 of 24
the halophilic archaea were identified, evident differences were observed with the se‐
quences of their known hyperthermophilic counterparts. Unfortunately, most of those
halophilic amylolytic enzymes were only putative proteins from genome sequencing pro‐
jects [58], or their complete sequences were not available [26,44,46], making it difficult to
establish accurate phylogenetic
relationships. In addition, an enormous diversity was ob‐
served among the amylases characterized and sequenced from a member of the Halobac‐
teriaceae family, which showed similarities with marine bacteria, fungal, or even animal
sources [59]. Therefore, more insightful biochemical characterization studies are needed
to reveal the exact features of these amylolytic
enzymes from haloarchaea.
Although several copies of alpha‐amylases appear in the sequenced genomes of halo‐
archaea, most studies are focused on the extracellular ones. The role of the extracellular
amylase in haloarchaea is well‐established, as it allows the conversion of starch, produced
by the marine plankton, into simple
sugars that could be incorporated into the cell and
used as a carbon source [60]. Nonetheless, the function of the intracellular amylases is less
understood, not only in haloarchaea but also in other heterotrophic microorganisms [61].
Most intracellular amylases of haloarchaea were assigned by sequence homology without
a functional characterization, with
few exceptions, like the intracellular α‐amylase from
H. japonica, whose activity is well‐studied [49]. Further insight is necessary to complete
the characterization of haloarchaeal amylases, to understand their role in archaeal metab‐
olism, and to evaluate their biotechnological applications.
5. Conclusions
The detailed biochemical characterization of the cell‐associated
and the extracellular
amylase activities from the new isolated strain Haloarcula sp. HS revealed that both are
active at high salinity conditions and at considerably high temperatures. These features,
joined to their stability under a wide range of surfactants, make them suitable for indus‐
trial applications. The proteomic analysis showed
that three different cell‐associated en‐
zymes, one of which was also found in the extracellular medium, might be responsible for
the amylase activities. The three proteins conserve the consensus domains and residues
of the α‐amylase family. Further studies aim to decipher the function of a hypothetical
ancestral gene and to
increase our understanding of the biochemical behavior of these
polyextremophilic enzymes. Developing new techniques for high‐scale production in the
industry is also needed.
Supplementary Materials: The following are available online at www.mdpi.com/2079‐
7737/10/4/337/s1. Figure S1: Full length of the 16S rRNA encoding gene from Haloarcula sp. HS. Fig‐
ure S2: Complete original polyacrylamide gels. Figure S3: Predicted structures of the three amylase
sequences identified in Haloarcula sp. HS. Figure S4: Multiple alignments of the
amino acid sequence
of the extracellular amylase identified in Haloarcula sp. HS (AMY_HS1). Figure S5: Multiple align‐
ments of the amino acid sequence of the cell‐associated amylase from Haloarcula sp. HS (AMY_HS2).
Figure S6: Multiple alignments of the amino acid sequence of the cell‐associated amylase from Halo‐
arcula
sp. HS (AMY_HS3).
Author Contributions: Conceptualization, J.V., R.L., and P.G.‐V.; Funding acquisition, J.V., R.L. and
P.G.‐V.; Investigation, P.G.‐V. and J.V.; Methodology, P.G.‐V., B.G. and S.R.; Supervision, J.V. and
R.L.; Software: P.G.‐V. and L.R. Data curation, P.G.‐V., J.V., L.R. and C.G.; Writing—original draft,
P.G.‐V. and
R.L.; Writing—review & editing, P.G.‐V., J.V. and R.L. All authors have read and agreed
to the published version of the manuscript.
Funding: This research was funded by the Operative FEDER Program‐Andalucía 2014–2020, the
University of Huelva, the Spanish Agencia Estatal de Investigación (grants PID2019‐109785GB‐I00
and PID2019‐110438RB‐C22 ‐AEI/FEDER) and the SUBV. COOP.ALENTEJO‐ALGARVE‐ANDA‐
LUCIA 2019. P.G.‐V. and C.G. acknowledge financial support from the University of Huelva (EPIT
2016‐17) and the Junta de Andalucía (grant P18‐RT‐3154), respectively.
Institutional Review Board Statement: This study did not involve humans or animals.
Informed Consent Statement: Not applicable.
Biology 2021, 10, 337 22 of 24
Data Availability Statement: All DNA and protein sequences of the studied enzymes are included
as Supplementary Materials; other information is available upon request.
Acknowledgments: Technical support of Rocío Rodríguez from the IBVF‐CSIC in the proteomic
analysis is acknowledged.
Conflicts of Interest: The authors declare no conflict of interest. The funders
had no role in the
design of the study; in the collection, analyses, or interpretation of data; in the writing of the manu‐
script, or in the decision to publish the results.
References
1. Kanekar: P.P.; Kelkar, A.S.; Dhakephalkar, P.K. Halophiles—Taxonomy, Diversity, Physiology and Applications. In Microor‐
ganisms in Environmental Management; Metzler, J.B., Ed.; Springer: Dordrecht, The Netherlands, 2012; Volume 9789400722, pp.
1–34.
2. Ventosa, A.; Márquez, M.C.; Sánchez‐Porro, C.; De La Haba, R.R. Taxonomy of Halophilic Archaea and Bacteria. In Advances
in
Understanding the Biology of Halophilic Microorganisms; Vreeland, R.H., Ed.; Springer: Dordrecht, The Netherlands, 2012; pp. 59–
80.
3. Oren, A. Life at high salt concentrations, intracellular KCl concentrations, and acidic proteomes. Front. Microbiol. 2013, 4, 315,
doi:10.3389/fmicb.2013.00315.
4. Mevarech, M.; Frolow, F.; Gloss, L.M. Halophilic enzymes: Proteins with a
grain of salt. Biophys. Chem. 2000, 86, 155–164,
doi:10.1016/s0301‐4622(00)00126‐5.
5. Mehta, D.; Satyanarayana, T. Bacterial and Archaeal α‐Amylases: Diversity and Amelioration of the Desirable Characteristics
for Industrial Applications. Front. Microbiol. 2016, 7, 1129, doi:10.3389/fmicb.2016.01129.
6. Cabrera, M. Ángeles; Blamey, J.M. Biotechnological applications of archaeal enzymes from extreme
environments. Biol. Res.
2018, 51, 1–15, doi:10.1186/s40659‐018‐0186‐3.
7. Vaidya, S.; Srivastava, P.; Rathore, P.; Pandey, A. Amylases: A prospective enzyme in the field of biotechnology. J. Appl. Biosci.
2015, 41, 1–18.
8. Kumar, V.; Sangwan, P.; Singh, D.; Gill, P.K. Global scenario of industrial enzyme
market. In Industrial Enzymes: Trends, Scope
and Relevance; Nova Science Publishers: New York, NY, USA, 2014; pp. 176–196, doi:10.13140/2.1.3599.0083.
9. Gupta, R.; Gigras, P.; Mohapatra, H.; Goswami, V.K.; Chauhan, B. Microbial α‐amylases: A biotechnological perspective. Process.
Biochem. 2003, 38, 1599–1616, doi:10.1016/s0032‐9592(03)00053‐0.
10. Fernandes, P. Enzymatic processing in
the food industry. In Reference Module in Food Science; Elsevier BV: Amsterdam, The
Netherlands, 2018.
11. Gopinath, S.C.B.; Anbu, P.; Arshad, M.K.M.; Lakshmipriya, T.; Voon, C.H.; Hashim, U.; Chinni, S.V. Biotechnological processes
in microbial amylase production. BioMed Res. Int. 2017, 2017, 272193, doi:10.1155/2017/1272193.
12. Mobini‐Dehkordi, M.; Javan, F.A.
Application of alpha‐amylase in biotechnology. J. Biol. Today World 2012, 1, 39–50,
doi:10.15412/j.jbtw.01010104.
13. Kumar, V.; Longhurst, P. Recycling of food waste into chemical building blocks. Curr. Opin. Green Sustain. Chem. 2018, 13, 118–
122, doi:10.1016/j.cogsc.2018.05.012.
14. van der Maarel, M.J.; van der Veen, B.; Uitdehaag, J.C.; Leemhuis,
H.; Dijkhuizen, L. Properties and applications of starch‐
converting enzymes of the α‐amylase family. J. Biotechnol. 2002, 94, 137–155, doi:10.1016/s0168‐1656(01)00407‐2.
15. Taniguchi, H.; Honnda, Y. Amylases. In Encyclopedia of Microbiology; Elsevier BV: Amsterdam, The Netherlands, 2009; pp. 159–
173.
16. John, J. Amylases‐bioprocess and potential applications:
A review. Int. J. Bioinform. Biol. Sci. 2017, 5, 41, doi:10.5958/2321‐
7111.2017.00006.3.
17. Gómez‐Villegas, P.; Vigara, J.; León, R. Characterization of the microbial population inhabiting a solar saltern pond of the odiel
marshlands (SW Spain). Mar. Drugs 2018, 16, 332, doi:10.3390/md16090332.
18. Gómez‐Villegas, P.; Vigara, J.; Vila,
M.; Varela, J.; Barreira, L.; Léon, R. Antioxidant, Antimicrobial, and Bioactive Potential of
Two New Haloarchaeal Strains Isolated from Odiel Salterns (Southwest Spain). Biology 2020, 9, 298, doi:10.3390/biology9090298.
19. Altschul, S.F.; Gish, W.; Miller, W.; Myers, E.W.; Lipman, D.J. Basic local alignment search tool. J. Mol. Biol. 1990, 215
, 403–410,
doi:10.1016/s0022‐2836(05)80360‐2.
20. Bradford, M.M. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of
protein‐Dye binding. Anal. Biochem. 1976, 72, 248–254, doi:10.1016/0003‐2697(76)90527‐3.
21. Kelley, L.A.; Mezulis, S.; Yates, C.M.; Wass, M.N.; Sternberg, M.J.E. The Phyre2 web
portal for protein modeling, prediction and
analysis. Nat. Protoc. 2015, 10, 845–858, doi:10.1038/nprot.2015.053.
22. Petersen, B.; Petersen, T.N.; Andersen, P.; Nielsen, M.; Lundegaard, C. A generic method for assignment of reliability scores
applied to solvent accessibility predictions. BMC Struct. Biol. 2009, 9, 51, doi:10.1186/1472‐6807‐9‐51.
23.
Pettersen, E.F.; Goddard, T.D.; Huang, C.C.; Couch, G.S.; Greenblatt, D.M.; Meng, E.C.; Ferrin, T.E. UCSF Chimera—A visuali‐
zation system for exploratory research and analysis. J. Comput. Chem. 2004, 25, 1605–1612, doi:10.1002/jcc.20084.
Biology 2021, 10, 337 23 of 24
24. Gasteiger, E.; Hoogland, C.; Gattiker, A.; Duvaud, S.; Wilkins, M.R.; Appel, R.D.; Bairoch, A. Protein Identification and Analysis
Tools on the ExPASy Server. In The Proteomics Protocols Handbook; Walker, J.M., Ed.; Humana Press: New York, NY, USA, 2005;
pp. 571–607.
25. Kumar, S.; Stecher, G.; Li, M.; Knyaz, C.;
Tamura, K. MEGA X: Molecular evolutionary genetics analysis across computing
platforms. Mol. Biol. Evol. 2018, 35, 1547–1549, doi:10.1093/molbev/msy096.
26. Pérez‐Pomares, F.; Bautista, V.; Ferrer, J.; Pire, C.; Marhuenda‐Egea, F.C.; Bonete, M.‐J.; Marhuenda‐Egea, F. α‐Amylase activity
from the halophilic archaeon Haloferax mediterranei. Extremophiles 2003, 7,
299–306, doi:10.1007/s00792‐003‐0327‐6.
27. Hutcheon, G.W.; Vasisht, N.; Bolhuis, A. Characterisation of a highly stable α‐amylase from the halophilic archaeon Haloarcula
hispanica. Extremophiles 2005, 9, 487–495, doi:10.1007/s00792‐005‐0471‐2.
28. Mehta, D.; Satyanarayana, T. Biochemical and molecular characterization of recombinant acidic and thermostable raw‐starch
hydrolysing α‐amylase
from an extreme thermophile Geobacillus thermoleovorans. J. Mol. Catal. B Enzym. 2013, 85–86, 229–238,
doi:10.1016/j.molcatb.2012.08.017.
29. Tan, T.‐C.; Mijts, B.N.; Swaminathan, K.; Patel, B.K.; Divne, C. Crystal Structure of the Polyextremophilic α‐Amylase AmyB
from Halothermothrix orenii: Details of a Productive Enzyme–Substrate Complex and an N Domain with a
Role in Binding
Raw Starch. J. Mol. Biol. 2008, 378, 852–870, doi:10.1016/j.jmb.2008.02.041.
30. Verma, D.K.; Vasudeva, G.; Sidhu, C.; Pinnaka, A.K.; Prasad, S.E.; Thakur, K.G. Biochemical and Taxonomic Characterization
of Novel Haloarchaeal Strains and Purification of the Recombinant Halotolerant α‐Amylase Discovered in the Isolate. Front.
Microbiol. 2020, 11,
2082, doi:10.3389/fmicb.2020.02082.
31. Boel, E.; Brady, L.; Brzozowski, A.M.; Derewenda, Z.; Dodson, G.G.; Jensen, V.J.; Petersen, S.B.; Swift, H.; Thim, L.; Woldike,
H.F. Calcium binding in alpha.‐amylases: An X‐ray diffraction study at 2.1‐.ANG. Resolution of two enzymes from Aspergillus.
Biochemistry 1990, 29, 6244–6249, doi:10.1021/bi00478a019.
32. Svensson, B.
Protein engineering in the α‐amylase family: Catalytic mechanism, substrate specificity, and stability. Plant Mol.
Biol. 1994, 25, 141–157, doi:10.1007/bf00023233.
33. Strokopytov, B.; Penninga, D.; Rozeboom, H.J.; Kalk, K.H.; Dijkhuizen, L.; Dijkstra, B.W. X‐ray Structure of Cyclodextrin Gly‐
cosyltransferase Complexed with Acarbose. Implications for the Catalytic Mechanism of Glycosidases.
Biochemistry 1995, 34,
2234–2240, doi:10.1021/bi00007a018.
34. Machovič, M.; Janeček, Š. The invariant residues in the α‐amylase family: Just the catalytic triad. Biol. Sect. Cell. Mol. Biol. 2003,
58, 1127–1132.
35. Zona, R.; Chang‐Pi‐Hin, F.; O’Donohue, M.J.; Janeček, Štefan Bioinformatics of the glycoside hydrolase family
57 and identifi‐
cation of catalytic residues in amylopullulanase from Thermococcus hydrothermalis. JBIC J. Biol. Inorg. Chem. 2004, 271, 2863–
2872, doi:10.1111/j.1432‐1033.2004.04144.x.
36. Roth, C.; Moroz, O.V.; Turkenburg, J.P.; Blagova, E.; Waterman, J.; Ariza, A.; Ming, L.; Tianqi, S.; Andersen, C.; Davies, G.J.; et
al. Structural and Functional Characterization
of Three Novel Fungal Amylases with Enhanced Stability and pH Tolerance. Int.
J. Mol. Sci. 2019, 20, 4902, doi:10.3390/ijms20194902.
37. Oda, Y.; Park, B.‐S.; Moon, K.‐H.; Tonomura, K. Recycling of bakery wastes using an amylolytic lactic acid bacterium. Bioresour.
Technol. 1997, 60, 101–106, doi:10.1016/s0960‐8524(97)00008‐4.
38. Chiotelli, E.; Pilosio, G.; Le Meste, M. Effect of sodium chloride on the gelatinization of starch: A multimeasurement study.
Biopolymers 2001, 63, 41–58, doi:10.1002/bip.1061.
39. Santorelli, M.; Maurelli, L.; Pocsfalvi, G.; Fiume, I.; Squillaci, G.; La Cara, F.; Del Monaco, G.; Morana, A. Isolation and charac‐
terisation of a
novel alpha‐amylase from the extreme haloarchaeon Haloterrigena turkmenica. Int. J. Biol. Macromol. 2016, 92, 174–
184, doi:10.1016/j.ijbiomac.2016.07.001.
40. Bertoldo, C.; Antranikian, G. Starch‐hydrolyzing enzymes from thermophilic archaea and bacteria. Curr. Opin. Chem. Biol. 2002,
6, 151–160, doi:10.1016/s1367‐5931(02)00311‐3.
41. Naidu, M.A.; Saranraj, P. Bacterial amylase:
A review. Int. J. Pharm. Biol. Arch. 2013, 4, 274–287.
42. Saranraj, P.; Stella, D. Fungal amylase‐a review. Int. J. Microbiol. Res. 2013, 4, 203–211, doi:10.5829/idosi.ijmr.2013.4.2.75170.
43. Yan, S.; Wu, G. Analysis on evolutionary relationship of amylases from archaea, bacteria and eukaryota. World J. Microbiol.
Biotechnol. 2016,
32, 24, doi:10.1007/s11274‐015‐1979‐y.
44. Fukushima, T.; Mizuki, T.; Echigo, A.; Inoue, A.; Usami, R. Organic solvent tolerance of halophilic α‐amylase from a Haloar‐
chaeon, Haloarcula sp. strain S‐1. Extremophiles 2004, 9, 85–89, doi:10.1007/s00792‐004‐0423‐2.
45. Kobayashi, T.; Kanai, H.; Hayashi, T.; Akiba, T.;
Akaboshi, R.; Horikoshi, K. Haloalkaliphilic maltotriose‐forming alpha‐amylase
from the archaebacterium Natronococcus sp. strain Ah‐36. J. Bacteriol. 1992, 174, 3439–3444, doi:10.1128/jb.174.11.3439‐3444.1992.
46. Moshfegh, M.; Shahverdi, A.R.; Zarrini, G.; Faramarzi, M.A. Biochemical characterization of an extracellular polyextremophilic
α‐amylase from the halophilic archaeon Halorubrum xinjiangense. Extremophiles 2013,
17, 677–687, doi:10.1007/s00792‐013‐0551‐
7.
47. Bajpai, B.; Chaudhary, M. Production and Characterization of α‐Amylase from an Extremely Halophilic Archaeon, Haloferax sp.
HA10. Food Technol. Biotechnol. 2015, 53, 11–17, doi:10.17113/ftb.53.01.15.3824.
48. Salgaonkar, B.B.; Sawant, D.T.; Harinarayanan, S.; Bragança, J.M. Alpha‐amylase Production by Extremely Halophilic Archae‐
onHalococcusStrain GUVSC8.
Starch Stärke 2018, 71, 1800018, doi:10.1002/star.201800018.
Biology 2021, 10, 337 24 of 24
49. Onodera, M.; Yatsunami, R.; Tsukimura, W.; Fukui, T.; Nakasone, K.; Takashina, T.; Nakamura, S. Gene Analysis, Expression,
and Characterization of an Intracellular α‐Amylase from the Extremely Halophilic Archaeon Haloarcula japonica. Biosci. Biotech‐
nol. Biochem. 2013, 77, 281–288, doi:10.1271/bbb.120693.
50. Erdagi, A.N.; Attar, A.; Basaran‐Elalmis, Y.; Yücel, S.;
Birbir, M. Production of α‐Amylase from Haloarcula hispanica 2TK2
Strain: Optimization of the Parameters That Effecting Activity. Adv. Sci. Lett. 2013, 19, 3551–3555, doi:10.1166/asl.2013.5229.
51. Sanchez‐Porro, C.; Martin, S.; Mellado, E.; Ventosa, A. Diversity of moderately halophilic bacteria producing extracellular hy‐
drolytic enzymes. J. Appl. Microbiol. 2003, 94
, 295–300, doi:10.1046/j.1365‐2672.2003.01834.x.
52. Lichi, T.; Ring, G.; Eichler, J. Membrane binding of SRP pathway components in the halophilic archaea Haloferax volcanii. JBIC
J. Biol. Inorg. Chem. 2004, 271, 1382–1390, doi:10.1111/j.1432‐1033.2004.04050.x.
53. Shafiei, M.; Ziaee, A.‐A.; Amoozegar, M.A. Purification and biochemical characterization of a novel SDS and
surfactant stable,
raw starch digesting, and halophilic α‐amylase from a moderately halophilic bacterium, Nesterenkonia sp. strain F. Process. Bio‐
chem. 2010, 45, 694–699, doi:10.1016/j.procbio.2010.01.003.
54. Sarian, F.D.; Janeček, Štefan; Pijning, T.; Ihsanawati; Nurachman, Z.; Radjasa, O.K.; Dijkhuizen, L.; Natalia, D.; Van Der Maarel,
M.J.E.C. A new group of
glycoside hydrolase family 13 α‐amylases with an aberrant catalytic triad. Sci. Rep. 2017, 7, srep44230,
doi:10.1038/srep44230.
55. Linden, A.; Mayans, O.; Meyer‐Klaucke, W.; Antranikian, G.; Wilmanns, M. Differential Regulation of a Hyperthermophilic α‐
Amylase with a Novel (Ca,Zn) Two‐metal Center by Zinc. J. Biol. Chem. 2003, 278, 9875–9884,
doi:10.1074/jbc.m211339200.
56. Sivakumar, N.; Li, N.; Tang, J.W.; Patel, B.K.; Swaminathan, K. Crystal structure of AmyA lacks acidic surface and provide
insights into protein stability at poly‐extreme condition. FEBS Lett. 2006, 580, 2646–2652, doi:10.1016/j.febslet.2006.04.017.
57. Janeček, Š.; Lévêque, E.; Belarbi, A.; Haye, B. Close evolutionary relatedness of α‐
amylases from archaea and plants. J. Mol. Evol.
1999, 48, 421–426, doi:10.1007/pl00006486.
58. Zorgani, M.A.; Patron, K.; Desvaux, M. New insight in the structural features of haloadaptation in α‐amylases from halophilic
Archaea following homology modeling strategy: Folded and stable conformation maintained through low hydrophobicity and
highly negative charged surface. J.
Comput. Mol. Des. 2014, 28, 721–734, doi:10.1007/s10822‐014‐9754‐y.
59. Janeček, Š. α‐amylases from Archaea: Sequences, structures and evolution. Biotechnol. Extrem. 2016, 1, 505–524.
60. Ştefan Andrei, A.; Banciu, H.L.; Oren, A. Living with salt: Metabolic and phylogenetic diversity of archaea inhabiting saline
ecosystems. FEMS Microbiol.
Lett. 2012, 330, 1–9, doi:10.1111/j.1574‐6968.2012.02526.x.
61. El‐Fallal, A.; Abou, M.; El‐Sayed, A.; Omar, A.E.‐S.A.N. Starch and microbial α‐amylases: From concepts to biotechnological
applications. Carbohydr. Compr. Stud. Glycobiol. Glycotechnol. 2012, doi:10.5772/51571.