Download Biosynthesis of Glucosyl Glycerol, a Compatible Solute, Using

Appl Biochem Biotechnol
DOI 10.1007/s12010-014-0889-z
Biosynthesis of Glucosyl Glycerol, a Compatible Solute,
Using Intermolecular Transglycosylation Activity
of Amylosucrase from Methylobacillus flagellatus KT
Jin-Woo Jeong & Dong-Ho Seo & Jong-Hyun Jung &
Ji-Hae Park & Nam-In Baek & Myo-Jeong Kim &
Cheon-Seok Park
Received: 9 January 2014 / Accepted: 24 March 2014
# Springer Science+Business Media New York 2014
Abstract A putative α-amylase gene (accession number, CP000284) of Methylobacillus
flagellatus KT ATCC51484 was cloned in Escherichia coli, and its gene product was
expressed and characterized. The purified recombinant enzyme (MFAS) displayed a typical
amylosucrase (ASase) activity by the demonstration of multiple activities of hydrolysis,
isomerization, and polymerization although it was designated as an α-amylase. The optimal
reaction temperature and pH for the sucrose hydrolysis activity of MFAS were determined
to be 45 °C and pH 8.5, respectively. MFAS has relatively high thermostable
characteristics compared with other ASases, as demonstrated by a half-life of
19.3 min at 50 °C. MFAS also showed polymerization activity using sucrose as a
sole substrate. Glycerol was transglycosylated by the intermolecular transglycosylation
activity of MFAS. Two major products were observed by thin-layer chromatography
and isolated by paper chromatography and recycling HPLC. Using 1H and 13C NMR,
their chemical structures were determined to be (2S)-1-O-α-D-glucosyl-glycerol or
(2R)-1-O-α-D-glucosyl-glycerol and 2-O-α-D-glucosyl-glycerol, in which a glucose
molecule is linked to glycerol via an α-glycosidic linkage.
Keywords Amylosucrase . Glucosyl glycerol . Glycerol . Methylobacillus flagellatus .
Transglycosylation
Jin-Woo Jeong and Dong-Ho Seo contributed equally.
J.<W. Jeong : D.<H. Seo : J.<H. Jung : J.<H. Park : N.<I. Baek : C.<S. Park (*)
Graduate School of Biotechnology and Institute of Life Science and Resources, Kyung Hee University,
Yongin 446-701, South Korea
e-mail: [email protected]
M.<J. Kim
Food Research Institute and School of Food and Life Science and Biohealth Products Research Center, Inje
University, Gimhae 621-749, South Korea
Appl Biochem Biotechnol
Introduction
Enzymes are highly selective biocatalysts that are responsible for a variety of cellular
biochemical reactions. Glycoside hydrolases (GH, EC 3.2.1) are a widespread group of
enzymes that cleave the glycosidic linkages in di-, oligo-, and polysaccharides [1]. GHs are
able to hydrolyze the glycosidic bonds of carbohydrate and non-carbohydrate moieties in
biological molecules. The GH13 family is the major GH family and acts on substrates
containing α-glycosidic linkages. Most of the known starch-modifying enzymes such as αamylases, pullulanases, α-1,6-glucosidases, maltogenic amylases, neopullulanases, and
cyclodextrinases (CDases) belong to this GH13 family [2, 3]. Amylosucrase (ASase, EC
2.4.1.4) is a versatile enzyme in the GH13 family that hydrolyzes sucrose to equal amounts of
glucose and fructose [4]. Simultaneously, it can transfer a released glucose to the four positions
of other glucose molecules or the non-reducing end of the resulting α-1-4 glucan molecules
resulting in the synthesis of an amylose-like glucan polymer [4, 5]. ASase also performs an
isomerization reaction to make sucrose isomers such as turanose and trehalulose [6]. Recently,
the intermolecular transglycosylation activity of ASase has gained increasing attention because
it uses a relatively cheap substrate, sucrose, as well as its broad range of acceptor
specificity [7, 8]. ASase can employ not only various glycones such as salicin [9] and
arbutin [10] but also numerous aglycone compounds including catechin [11] and
hydroquinone [12], as acceptor molecules.
With the development of advanced sequencing technology, data on numerous microbial
genomes are now available, and the presence of ASases and their homologues are being
revealed. However, to date, only five ASases from five different bacterial species, Neisseria
polysaccharea, Deinococcus radiodurans, Deinococcus geothermalis, Alteromonas macleodii,
and Arthrobacter chlorophenolicus, have been studied [4, 13–16]. A previous study reported
that the suh gene from Xanthomonas axonopodis pv. glycines, a causative agent of bacterial
pustule disease in soybeans, showed a very strong deduced amino acid homology with various
ASases [17]. However, it only displayed sucrose hydrolysis activity without any
glucosyltransferase or isomerization activities [18].
α-D-Glucosyl glycerol (GG) was formerly known as the main compatible solute in
moderately halotolerant cyanobacteria such as the Synechocystis sp. strain PCC 6803 [19].
This compound accumulates in cells to adjust the cellular osmotic potential to levels that allow
water uptake. In cyanobacteria, GG is synthesized by a two-step reaction in which the
enzymatic condensation of ADP-glucose and glycerol 3-phosphate by GG-phosphate synthase
(GgpS) is followed by the dephosphorylation of the intermediate by GG-phosphate phosphatase (GgpP) [20]. Interestingly, GG has been found in traditional Japanese foods such as sake,
miso, and mirin at concentrations of 0.1–0.5 % [21]. It was suggested that GG is formed from
glycerol by the transglycosylation activity of yeast α-glucosidase (EC 3.2.1.20) [22]. GG has
some industrially useful characteristics. It has approximately 0.55-fold the sweetness of
sucrose, but shows high thermostability, low heat-colorability, low Maillard reactivity, low
hygroscopicity, and high water-holding capacity. GG also exhibits some health benefits such as
non-cariogenicity and low digestibility [23].
In this study, a putative α-amylase gene (accession number, CP000284) was cloned from an
obligate methylotroph Methylobacillus flagellatus KT ATCC51484 and expressed in
Escherichia coli. The catalytic properties of the recombinant enzyme (hereafter referred
to as MFAS) were examined to confirm that it belongs to the ASase family. In
addition, the transglycosylation activity of MFAS was employed to synthesize GG.
The reaction products were purified to homogeneity, and their chemical structures
were determined.
Appl Biochem Biotechnol
Materials and Methods
Chemicals and Enzymes
Sucrose and glycerol were purchased from Duchefa Biochemistry (Haarlem, Netherlands) and
Sigma Chemical Co (St. Louis, MO, USA), respectively. Silica gel K5F thin-layer chromatography (TLC) plates (Whatman, Kent, UK) were used for sugar analysis. Restriction
endonucleases and other modifying enzymes, such as Pfu DNA polymerase and T4 DNA
ligase, were obtained from Stratagene Inc. (La Jolla, CA, USA) or Solgent (Daejeon, Korea).
Preparation of genomic DNA from M. flagellatus KT ATCC51484 was carried out using the
GeneAll TM total DNA purification kit (GeneAll Biotechnology, Seoul, Korea).
Oligonucleotides used for polymerase chain reaction (PCR) and DNA sequencing were
synthesized by Solgent Co. (Daejeon, Korea). The products of restriction enzyme digestion
and PCR amplifications were purified using the QIAquick Gel extraction kit (Qiagen, Hilden,
Germany). All other chemicals used in this study were of analytical reagent grade.
Bacterial Strains, Media, and Plasmids
Genomic DNA from M. flagellatus KT ATCC51484 was a generous gift from Dr. Mary E.
Lidstrom at the University of Washington (Seattle, WA, USA). Two E. coli strains were
employed for standard DNA manipulation or recombinant protein expression. E. coli DH10B
[F−Φ80lacZ M15 (lacZYA-argF) U169 recA1endA1 hsdR17(rk+, mk+) phoA supE44 thi-1
gyrA96 relA1λ−] was a host for typical DNA manipulation, and BL21 [F− ompT hsdSB (rB−,
mB−) gal dcm λ(DE3)plies T1R] was used as the expression vector. Selection of recombinant
cells was achieved on an Lysogeny broth (LB) agar plate with 100-μg/mL ampicillin, 0.5-mM
isopropyl-β-D-thiogalactopyranoside (IPTG), and 40-μg/mL 5-bromo-4-chloro-3-indolyl-β-Dgalactopyranoside (Xgal). The plasmids pGEM-T Easy vector (Promega Co., Madison, WI,
USA) and pET-21a(+) vector (Novagen, Darmstadt, Germany) were used for the cloning of
PCR products and construction of the expression vector, respectively.
E. coli strains containing recombinant DNA were cultured in 5 mL of LB medium in the
presence of ampicillin (100 μg/mL) at 37 °C overnight. Cells (1.5 mL) were collected by
centrifugation at 6,000×g for 5 min in a microcentrifuge, and the plasmid DNA was obtained
using the general alkaline lysis method. The final plasmid DNAs were dissolved and stored in
a 50-μL TE-RNase [DNase-free RNase A 20 μg/mL in 10-mM Tris-Cl and 1-mM EDTA, pH
8.0] until further use.
Construction of pETMFAS Expression Vector
The gene corresponding to the ASase homologue in M. flagellatus KT ATCC51484 was
amplified by PCR using the oligonucleotide primers MFAS-F (5’-CAT ATG TAC GAA CAA
GTC TCC CAC TCG-3’) and MFAS-R (5’-CTC GAG AAG CTG TAA CCA ATA AAA
GCC-3’). The PCR condition for mfas gene amplification was previously described [4, 16].
The amplified PCR product was purified using the QIAquick Gel extraction kit (Qiagen).
The PCR product was cloned into pGEM-T Easy vector (Promega), and the recombinant
plasmids were digested with Nde I and Xho I after confirming the absence of PCR-introduced
errors. The resulting fragments were inserted into pET-21a(+) vector treated with the same
enzymes to create pETMFAS, an expression vector for MFAS in which the mfas gene was
controlled by the T7 promoter. E. coli BL21(DE3) was transformed with pETMFAS, and the
resulting recombinant E. coli strain was used for the efficient production of MFAS.
Appl Biochem Biotechnol
Expression of pETMFAS and the Purification of the Recombinant MFAS
Recombinant E. coli BL21 harboring pETMFAS were grown in 1 L of LB medium supplemented with ampicillin (0.1 mg/mL) at 37 °C with agitation. The cells were harvested by
centrifugation at 4 °C after a 3-h induction of the mfas gene by the addition of IPTG to a final
concentration of 1 mM when the optical density at 600 nm (OD600) reached 0.5–0.6. The
collected pellets were resuspended in 5 mL of lysis buffer (50-mM NaH2PO4, 300-mM NaCl,
10-mM imidazole, pH 7.0) per gram (wet weight) of cells. Lysozyme (1 mg/mL) was added to
the resuspended cells, and the cell suspension was incubated for 30 min on ice before
disruption by sonication (Sonifier 450, Branson, Danbury, CT, USA; output four, six times
for 10 s, constant duty) in an ice bath. The cell lysate was centrifuged at 6,000×g for 10 min at
4 °C. The recombinant MFAS is then purified by affinity chromatography with nickelnitrilotriacetic acid (Ni-NTA) affinity column (Qiagen) as suggested in the manufacturer’s
protocol. Protein obtained from the eluted fraction was dialyzed to remove the imidazole, and
the protein concentration and enzyme activity were measured before further application.
Determination of Protein Concentration and Enzyme Activity
The bicinchoninic acid (BCA) assay was used to quantify protein concentration as described
by Smith et al. [24]. The BCA protein assay kit (Thermo Fisher Scientific, Agawam, MA,
USA) was used to prepare the working reagent solution, and bovine serum albumin (BSA) was
used as a standard.
Enzymatic activity of MFAS was measured by the 3,5-dinitrosalicylic acid (DNS) method
for detecting reducing sugar [4] or the D-glucose/D-fructose assay kit (Roche Diagnostics,
Darmstadt, Germany) [15] for measuring the fructose concentration. The reaction mixture
contained 100 μL of 25 % sucrose, 100 μL of double distilled water (DDW), and 250 μL of
100-mM Tris-HCl (pH 8.5). After the preincubation of the substrate solution at 45 °C for
5 min, the reaction was initiated by the addition of 50 μL of the enzyme solution. The reaction
was continued for 10 min before being terminated by the addition of 500 μL of DNS solution
and inactivation by boiling. The absorbance of reducing sugar at 545 nm was measured using a
spectrophotometer (UV1201, Shimadzu, Kyoto, Japan). For the D-glucose/D-fructose assay kit,
the reaction mixture was boiled for 5 min to inactivate the enzyme without adding DNS
solution. The enzyme reaction was used as a sample for the fructose assay. One unit of MFAS
hydrolysis activity was defined as the amount of enzyme that catalyzes the production of
1 μmole of fructose per minute in the assay conditions.
Determination of Optimal pH, Temperature, and Thermostability
The effect of pH on enzyme activity was investigated within the range of pH 4.0 to pH 10.0
(0.1-M sodium acetate buffer for pH 4 and 5; 0.1-M sodium citrate buffer for pH 5, 6, 6.5, and
7; 0.1-M Tris-HCl buffer for pH 7, 7.5, 8, 8.5, and 9; and 0.1-M glycine-NaOH for pH 9 and
10) at 45 °C. The effect of temperature on the hydrolysis activity of MFAS was studied
between 30 and 65 °C at pH 8.5. The thermostability of the purified MFAS was observed by
preincubation in the absence of substrate at 45, 50, and 55 °C. At various times, the residual
activity was measured under standard assay conditions.
Differential scanning fluorimetry (DSF) has been used to measure the melting temperature
of proteins [25]. The sample was prepared by mixing 10 μL of SYPRO orange (10×) with
10 μL of enzyme solution (0.1 mg/mL). The mixture of proteins and SYPRO orange was
incubated in a real-time polymerase chain reaction (RT-PCR) system using a temperature
Appl Biochem Biotechnol
gradient from 30 to 99 °C with 1 °C increments. All fluorescence measurements were carried
out using an Applied Biosystems 7500 Fast RT-PCR system (Applied Biosystems, Grand
Island, NY, USA). Sigma Plot 12.0 software (Systat Software Inc., Chicago, IL, USA) and
Microsoft Excel program (Redmond, WA, USA) were used for all graphical and statistical
analyses.
High-Performance Anion-Exchange Chromatography (HPAEC) Analysis
HPAEC analysis was carried out with analytical columns (CarboPac PA100 or CarboPac
MA1, Dionex Co., Sunnyvale, CA, USA) to detect carbohydrate in the reaction. Filtered
samples were applied and eluted with a linear gradient from 100 % buffer A (100 mM of
NaOH in water) to 60 % buffer B (500 mM of sodium acetate in buffer A) over
70 min. The flow rate of the mobile phase was maintained at 1.0 mL/min. Detection
was carried out using a Dionex ED50 module. The areas of the peaks were calculated
for quantitative analysis.
Gel Permeation Chromatography (GPC) Analysis
GPC (GE Healthcare, Little Chalfont, UK) analysis with a Superdex 200HR (GE Healthcare)
was used to confirm the molecular size of MFAS. To calculate the molecular weight of the
enzyme, standard proteins were loaded as controls, including carbonic anhydrase (29 KDa),
BSA (66 KDa), β-amylase (200 KDa), apoferritin (443 KDa), and thyroglobulin (669 KDa).
The concentration of injected protein was 1.0 mg/ml, and the flow rate was 0.4 mL/min.
Bioconversion of Glycerol to Glucosyl Glycerols (GGs)
GG biosynthesis reactions were carried out in 50-mM Tris-HCl buffer (pH 8.5) at 35 °C
instead of 45 °C because the transglycosylation activity of MFAS was similar to that of other
AS enzymes, which are known to have a higher transglycosylation activity at lower temperatures, in contrast to the hydrolysis activity [10].
The effect of different molar ratios of sucrose and glycerol on the production of GG was
examined. MFAS (1 unit) was added to 5 mL of substrate solution containing 100 mM of
sucrose as a glucosyl donor and 10, 20, 50, 100, 200, 500, or 1,000 mM of glycerol as an
acceptor (molar ratios of 1:0.1, 1:0.2, 1:0.5, 1:1, 1:2, 1:5, and 1:10, respectively) in 50-mM
Tris-HCl (pH 8.5). The reaction mixture was incubated at 35 °C for 12 h. The reaction was
stopped by heating in boiling water for 10 min, and the reaction products were analyzed by
TLC and HPAEC. In addition, a time course of the transglycosylation reaction was generated
over 24 h with sample collection every 3 h. The reaction was stopped by boiling for 10 min,
and the mixture was centrifuged at 15,300×g for 10 min to remove residual protein before
analysis.
Thin-Layer Chromatography (TLC) Analysis
TLC was used to detect and identify the transglycosylation products synthesized by MFAS.
One microliter aliquots of the reaction mixture were spotted onto Whatman K5F silica gel
plates that were activated at 110 °C for 30 min and developed with a solvent system of
isopropyl alcohol/ethyl acetate/water (3:1:1, v/v/v) in a TLC developing tank. Ascending
development was performed once at room temperature. The plate was allowed to air-dry in
a hood and was developed by soaking rapidly in 0.5 % (w/v) N-(1-naphthyl)-ethylenediamine
Appl Biochem Biotechnol
and 5 % (v/v) H2SO4 in methanol. The plate was dried and placed in a 110 °C oven for 10 min
to visualize the reaction spots.
Purification of Transglycosylated GGs
Paper chromatography was performed to purify transglycosylated GG products by multiple
descending techniques. Approximately 250 μL of the enzyme reaction product was loaded
onto Whatman 3-MM paper (23×55 cm). The paper was irrigated with a 3:1:1 (v/v/v) mixture
of isopropyl alcohol/ethyl acetate/water for 24 h in a glass paper chromatography developing
tank. After paper ascending development, the paper was allowed to air-dry in a hood. Spots on
the paper were detected using an AgNO3 reagent to verify the separation of purified carbohydrates. The paper was sectioned and eluted with deionized water. The separated compounds in
the elutant were freeze-dried (Il-shin Lab, Suwon, Korea) and collected for further analysis.
A recycling preparative HPLC system (LC-9104, JAI, Tokyo, Japan) was used for the
isolation of GGs. Samples (3 μL) were applied to a JAIGEL-W252 (2×50 cm, JAI) column
connected in tandem with a JAIGEL-W251 (2×50 cm, JAI) and guard column. The sample
was eluted with deionized water at a flow rate of 3 mL/min. The fractions corresponding to
GGs were collected and freeze-dried. The purity of each sample was determined using TLC
analysis.
NMR Analysis
1
H and 13C NMR spectra of the purified GGs were obtained with a Varian Inova AS 400 MHz
NMR spectrometer (Varian, Palo Alto, CA). The sample was dissolved in CD3OD at 24 °C
with tetramethylsilane (TMS) as the chemical shift reference.
Results and Discussion
Sequence Analysis, Cloning, and Expression of Putative mfas Gene
The putative mfas gene was designated an α-amylase gene (accession number, CP000284) in
NCBI Genbank. The gene consists of 1,953 nucleotides, which encode a predicted 650-amino
acid protein with a calculated molecular mass of 74,556 Da. The deduced amino acid analysis
revealed that the putative MFAS protein shared not only catalytic residues, but also five
consensus regions (CR I-V), with other ASases (Fig. 1 and Table 1), although the overall
protein showed a low level of amino acid sequence homology when compared with known
ASases (ACAS, AMAS, DGAS, DRAS, and NPAS). The topology of A, B, and B’ domains
creates an active pocket in ASases [5, 26, 27]. Interestingly, the amino acid homology of B and
B’ domains in MFAS with that of other ASases was higher than the overall sequence identity
(Table 1). Multiple amino acid sequence alignments revealed that the MFAS protein has
retained the key conserved residues required for ASase activity (Fig. 1). These include two
catalytic residues (Asp284 and Glu326) belonging to CR III and CR IV, respectively, and
Asp141 and Arg520 (located in CR I and CR V, respectively), which are able to form a salt
bridge that assists in the formation of an active pocket in ASases [26, 27]. Furthermore,
the residues of CR IV (395-HDDIG-399) that were proposed to play an important role
in the polymerization activity of ASase are present in the putative MFAS protein [28].
Therefore, we suspected that the putative mfas gene might encode an ASase rather
than an α-amylase.
Appl Biochem Biotechnol
Fig. 1 Amino acid sequence alignment of MFAS with previously identified ASases from various microorganisms. Fully conserved amino acid residues are blocked in black, and five conserved regions (CR), CR I, CR II,
CR III, CR IV, and CR V, are indicated by arrows. Triangles indicate amino acid residues required for the catalytic
active site. The solid line box indicates the B-domain of ASase, and the dotted line box indicates the B’-domain.
Arrows show residues that form a salt bridge, and the asterisk indicates oligosaccharide binding sites
The putative mfas gene was successfully amplified by PCR from M. flagellatus KT
genomic DNA and expressed in E. coli. An inducible expression vector pET21a was used
to construct pETMFAS for efficient expression in E. coli. When DGAS, DRAS, and NPAS
were previously expressed in a pGEX expression system, the expression levels of the
recombinant proteins were lower than in other systems such as pET and pHCE protein
expression systems [4, 12–14]. Recently, ACAS has been successfully expressed in the
constitutive and inducible expression systems pET21a and pHCXHD, respectively [16].
Although it was easier to achieve protein expression with the constitutive system (pHCAC
AS) than with the inducible system (pETACAS), the production level of pHCACAS was
lower than that of pETACAS [16]. As expected, MFAS was expressed in E. coli BL21(DE3)
as a dominant protein with a molecular mass of 74 kDa. The expressed recombinant MFAS
was efficiently purified by Ni-NTA affinity chromatography, and SDS-PAGE analysis showed
a single band at approximately 74 kDa, which matched the theoretical molecular mass of
MFAS.
Biochemical Properties of MFAS
The optimum pH of MFAS was investigated using various buffers, including sodium acetate
(pH 4.0 and 5.0), sodium citrate (pH 5.0, 6.0, 6.5, and 7.0), Tris-HCl (pH7.0, 7.5, 8.0, 8.5, and
9.0), and glycine-NaOH (pH9.0 and 10.0) (Fig. 2a). The pH optimum of MFAS was found to
Table 1 Alignment results for amylosucrase showing the percentage identity with Methylobacillus flagellatus
ASase (MFAS) for the whole protein, the B-domain, and the B’-domain
Enzyme
Overall amino acids (%)
B-domain amino acids
B’-domain amino acids
ACAS
37.48
53.52
51.52
AMAS
39.29
55.56
52.78
DGAS
DRAS
37.42
39.38
56.94
55.56
52.78
50.00
NPAS
37.26
52.11
43.28
ACAS=Arthrobacter chlophenoilcus, AMAS=Altermonas macleodii, DGAS=Deinococcus geothermalis,
DRAS=Deinococcus radiodurans, NPAS=Neisseria polysaccharea
Appl Biochem Biotechnol
Fig. 2 Biochemical properties of MFAS. a Optimum pH was determined in different buffers at 45 °C: filled
circle represents sodium acetate at pH 4.0–5.0, empty circle sodium citrate at pH 5.0–7.0, filled inverted triangle
Tris-HCl at pH 7.0–9.0, and empty triangle glycine-NaOH at pH 9.0–10.0. b Optimal temperature was
determined between 30 and 65 °C at pH 8. c Half-life of MFAS measured at various temperatures in 50-mM
Tris-HCl, pH 8.5: filled circle represents 45 °C, empty circle 50 °C, and filled inverted triangle 55 °C. d Melting
temperature was confirmed by differential scanning fluorimetry (DSF). A few proteins displayed a shoulder in
the peak of the DSF graph similar to MFAS due to the buffer conditions such as pH, salt concentration, and
presence or absence of ligand [39–41]
be 8.0–8.5. The pH profile of MFAS was similar to that of other ASases such as ACAS,
AMAS, and DGAS [4, 15, 16]. The optimum temperature of MFAS at pH 8.5 was determined
within the range of 30 to 60 °C (Fig. 2b). Highest enzymatic activity of MFAS was observed at
45 °C. The optimum temperature of MFAS was similar to that of DGAS, which has the highest
optimum temperature and thermostability among known ASases. Correlation between thermal
stability and enzyme activity for MFAS is shown in Fig. 2c. The half-life of MFAS was
calculated as 19.5 and 4.3 min at 50 and 55 °C, respectively, and the melting temperature of
MFAS was determined as 50.6 °C (Fig. 2d). These results indicate that the order of thermostability among known ASases is DGAS>MFAS≫AMAS >NPAS>ACAS >DRAS [4,
13–16]. Recently, the three-dimensional structure of DGAS was solved, revealing that it exists
as a dimeric form in solution [27]. The authors assumed that dimerization might contribute to
the enzyme stabilization of DGAS. To investigate the oligomeric state of MFAS in solution,
MFAS was subjected to gel permeation chromatography (GPC). GPC analysis showed that
MFAS displayed a peak at a molecular mass of approximately 137.9 kDa, implying that it also
exists in a dimeric conformation in solution.
The reaction products of MFAS were characterized using sucrose as a sole substrate and
compared with those of DGAS (Table 2 and Fig. 3). Although MFAS and DGAS are the two
most thermostable enzymes among known ASases, MFAS could not produce a significant
Appl Biochem Biotechnol
Table 2 Yields of three ASase reactions (hydrolysis, polymerization, and isomerization) for different enzymes
with sucrose as the sole substrate
Enzymes
Substrate
concentration (mM)
The ratio of reaction products (%)
Hydrolysis
(glucose and fructose)
References
Isomerization
Polymerization
(maltooligosaccharides (trehalulose
and insoluble glucan) and turanose)
MFAS
100
9.5
75.5
15.0
This study
ACAS
100
1.9
78.4
19.7
[16]
DGAS
100
7.0
71.0
22.0
[4]
DRAS
NPAS
100
100
9.6
5.4
56.9
80.1
33.5
14.5
[14]
[14]
amount of insoluble glucans from sucrose at a relatively high temperature (45 °C) [4]. Previous
studies showed that the polymerization activity of ASase decreased as temperature increased
[4, 10]. However, when sucrose was reacted with MFAS and DGAS at 40 °C for 48 h, MFAS
produced substantial amounts of turbid insoluble glucans. Moreover, the reaction mixture of
MFAS displayed more turbidity than that of DGAS, indicating that MFAS synthesized more
insoluble α-1,4-glucan from sucrose than DGAS (Fig. 3b). The comparison of the side-chain
distributions (DP>4) in the relative peak area of HPAEC chromatogram confirmed that MFAS
synthesized relatively longer glucans (DP>33) than DGAS. These results indicated that the
polymerization activity of MFAS was higher than that of DGAS at a high temperature. In
addition, the intermolecular transglycosylation activity of MFAS was examined with glycerol
as an acceptor molecule.
Bioconversion of Glycerol to GG by MFAS
GGs were enzymatically synthesized from glycerol through the intermolecular
transglycosylation activity of MFAS. In TLC analysis, few spots appeared in the reaction with
Fig. 3 High-performance anion-exchange chromatography (HPAEC) analysis of α-1,4 glucan synthesized by
MFAS. a HPAEC analysis of sucrose isomers and short maltooligosaccharides (shorter than maltohexaose), b
HPACE analysis of long maltooligosaccharides. G2 maltose, G3 maltotriose, G4 maltotetraose, G5
maltopentaose, G6 maltohexaose, G7 maltoheptaose, DP degree of polymerization. The image on the upper
right corner of b is the reaction mixtures of MFAS and DGAS. The reaction was performed for 48 h at 40 °C
with sucrose as a sole substrate. The sample was diluted 10-fold and filtered before injection
Appl Biochem Biotechnol
MFAS compared with the reaction mixture without glycerol (Fig. 4). Although the main spots
in TLC analysis with MFAS were maltooligosaccharides, as expected from a typical ASase
reaction with sucrose as the sole substrate, two spots that were not identical to any standard
molecule or maltooligosaccharide were clearly evident (GT1 and GT2 in Fig. 4a). In addition,
two new peaks other than glycerol, glucose, and fructose were observed in the HPAEC
analysis (Fig. 4a). These analytical experiments indicated that two unidentified glycerol
transglycosylation products (GT1 and GT2) were synthesized by MFAS. The amount of
GT2 produced was higher than that of GT1. Various enzymatic methods were previously
employed to produce GGs [23, 29–31]. Two β-GGs were synthesized by β-glucosidase from
Pyrococcus furiosus using 2-nitophenyl-β-D-glucopyranoside or cellobiose as a donor and
glycerol as an acceptor [31]. When the transglycosylation reaction of cyclodextrin
glucanotransferase (CGTase) was used to synthesize GGs using soluble starch as a donor
and glycerol as an acceptor, (2R/S)-1-O-α-D-glucosyl-glycerol and 2-O-α-D-glucosyl-glycerol
were isolated as the major and minor smallest transfer products, respectively [29]. 2-O-α-DGlucosyl-glycerol, (2R)-1-O-α-D-glucosyl-glycerol, and (2S)-1-O-α-D-glucosyl-glycerol were
synthesized by transglucosidase L “Amano” (Amano Enzyme Inc, Nagoya, Japan) with
maltose as a donor and glycerol as an acceptor. Among these three GGs, (2R)-1-O-α-Dglucosyl-glycerol was the main constituent [23]. 2-O-α-D-glucosyl-glycerol as a major product
was mainly synthesized through the transglycosylation reaction of sucrose phosphorylase with
sucrose or glucose-1-phosphate as a donor and glycerol as an acceptor [30]. Similar to these
reactions, two transglycosylation products were detected in the MFAS reaction and one major
product was predominantly synthesized, indicating that MFAS might have regioselectivity for
the transglycosylation reaction using glycerol as an acceptor.
Purification and Structural Determination of GGs
The GG products synthesized by MFAS were purified by paper chromatography and a
preparative recycling HPLC system. Two isolated GG products were each detected as a single
spot on TLC analysis, indicating that the purification was successful (Fig. 4b). Interestingly,
the isolated GT1 was detected as a double peak on HPAEC analysis whereas the isolated GT2
Fig. 4 HPAEC and TLC analyses of transglycosylation reaction by MFAS with glycerol as an acceptor and
sucrose as a donor (a) and purification of GGs synthesized by MFAS (b). a TLC analysis, Lane M standard
markers from G1 (glucose) to G7 (maltoheptaose); Lane 1 only sucrose reaction with MFAS; Lane 2 glycerol
transglycosylation reaction without MFAS; Lane 3 glycerol transglycosylation reaction with MFAS; GT1 and
GT2 glucosyl-glycerols; Gly glycerol. b TLC analysis, Lane M standard marker from G1 (glucose) to G7
(maltoheptaose); Lane 1 glycerol transglycosylation reaction with MFAS; Lane 2 purified GT1; Lane 3 purified
GT2. HPAEC analysis, solid lines represents purified GT1and dotted lines purified GT2
Appl Biochem Biotechnol
appeared as a single peak. Previous HPAEC analysis showed that stereoisomers appeared as a
double peak because they are not completely separated by HPAEC [32, 33]. This result led us
to assume that GT1 might be a mixture of GG stereoisomers.
1
H and 13C NMR analyses were used to determine the structure of isolated GT1 and GT2
(Table 3). In 1H and 13C NMR analyses, the signals of C1’, C1, C3, C4, and H1 in the GT1
compound were observed as double peaks, and the chemical shift of C1 in the glycerol
molecule changed greatly from 64 to 72 (8 ppm). The chemical shift of C1’ and C3 was
slightly changed. This results were comparable to 1H and 13C NMR spectra of a mixture of
(2R)-1-O-β-D-glucosyl-glycerol and (2S)-1-O-β-D-glucosyl-glycerol, suggesting GT1 was a
mixture of these two stereoisomers [34]. The bond between glucose and glycerol in GT1 was
determined to be an α-glycosidic linkage according to the coupling constant (J=3.6 Hz) in 1H
NMR analysis. These results indicated that GT1 (1-GGs) was a mixture of (2R)-1-O-α-Dglucosyl-glycerol (R-1-GG) and (2S)-1-O-α-D-glucosyl-glycerol (S-1-GG) (Fig. 5a).
GT2 was determined to be 2-O-α-D-glucosyl-glycerol (2-GG) by 1H and 13C NMR
analyses (Fig. 5a). The chemical shift of C2 in the glycerol molecule changed greatly from
72 to 81.5 (9.5 ppm) in GT2, confirming that the transferred glucose molecule was connected
to C2 in the glycerol molecule. The result of 13C NMR analysis was identical to 13C NMR
results of 2-O-β-D-glucosyl-glycerol synthesized by β-glucosidase [35]. In addition, 1H NMR
analysis revealed that the glucose molecule was transferred to C2 in the glycerol molecule with
an α-anomeric configuration based on the coupling constant (J=4.0 Hz) of the glucose
anomeric proton signal observed at 4.9 ppm.
Effect of Acceptor Concentration on Bioconversion of Glycerol to GGs
The production yield of GGs by MFAS was analyzed at various glycerol concentrations
with a fixed concentration of sucrose (100 mM) by HPAEC (Fig. 5b). As the glycerol
concentration increased, the production level of GGs increased. Interestingly, the production level of 2-GG was higher than that of 1-GGs under all reaction conditions. The
ratio of 2-GG over 1-GGs was calculated to be 4.5–5.3 when no less than 100 mM of
glycerol was used. The production yield of MFAS was 48 % (based on acceptor), which
Table 3
13
C (100 MHz) and 1H NMR (400 MHz) data for GGs synthesized by MFAS (units—ppm, in CD3OD)
1-GGs
13
C NMR
Glycerol
1
H NMR
13
C NMR
2-GG
1
H NMR
13
C NMR
1
63.8
72.3, 72.1
2
73.2
71.8
81.5
3
63.8
64.2, 64.1
62.3
α-D-Glucopyranose 1’
101.2
100.6, 100.3
2’
73.7
73.7
3’
75.3
75.2
4’
72.0
70.9, 70.1
5’
73.9
73.8
6’
62.7
62.6
Reference
[37, 38]
1
H NMR
62.6
4.80, 4,79
(d, J=3.6 Hz)
3.39
100.1
4.98
(d, J=4.0 Hz)
73.8
3.37
(dd, J=10.4, 4.0 Hz)
75.1
3.28
(dd, J=9.2, 9.2 Hz)
71.8
3.26
(dd, J=9.2, 9.2 Hz)
73.9
62.9
This study
This study
Appl Biochem Biotechnol
Fig. 5 Molecular structures of GGs synthesized by MFAS (a) and production of GGs in the transglycosylation
reaction with various concentrations of glycerol as the acceptor and 100 mM of sucrose as a donor (b). a
Structure of (2R)-1-O-α-D-glucosyl-glycerol and (2S)-1-O-α-D-glucosyl-glycerol and 2-O-α-D-glucosyl-glycerol. b Open bar represents the concentration of 2-O-α-D-glucosyl-glycerol whereas closed bar stands for the
concentration of (2R/S)-1-O-α-D-glucosyl-glycerol
was higher than those of other enzymes. The production yield of 19.9 % was observed
with CGTase reaction whereas 5.1 % with α-glucosidase [23, 29]. The most comparable
production yield (35 %) was obtained with sucrose phosphorylase [30]. When αglucosidase produced three GGs (R-1-GG, S-1-GG, and 2-GG) using maltose and
glycerol, the ratio of R-1-GG, S-1-GG, and 2-GG was 49:41:10 [23]. Likewise,
CGTase was shown to predominantly transfer glucose to the one (or three) position of
glycerol [29]. On the other hand, sucrose phosphorylase from Leuconostoc mesenteroides
synthesized mainly 2-GG using 800 mM of sucrose as a donor and 2,000 mM of glycerol
as an acceptor [30]. Similar to sucrose phosphorylase, MFAS predominantly synthesized
2-GG over 1-GGs. This result suggests that the active site of MFAS provided a regioselectivity
of glucosyl transfer using glycerol as an acceptor. It is quite interesting that previous studies
showed that both ASase and sucrose phosphorylase produced the same transglycosylation
products using hydroquinone and catechin as an acceptor [11, 12, 36]. This indicates that these
two enzymes share similar regioselectivity when they perform the intermolecular
transglycosylation reaction. Our results indicate that MFAS can be employed to make various
biological materials with high efficiency using inexpensive donor molecules.
Appl Biochem Biotechnol
Conclusions
The enzymatic properties of a putative amylosucrase (ASase) gene of M. flagellatus KT
ATCC51484 (mfas) were confirmed in this study. MFAS was relatively thermostable and
produced significant amounts of insoluble glucans at a high temperature (40 °C) compared
with other previously characterized ASases. The intermolecular transglycosylation activity was
employed in the production of GG, a compatible solute and enzyme stabilizer. Two major
products were observed in the MFAS reaction, and their chemical structures were identified as
(2S)-1-O-α-D-glucosyl-glycerol or (2R)-1-O-α-D-glucosyl-glycerol and 2-O-α-D-glucosylglycerol, in which a glucose molecule is linked to glycerol via an α-(1→4)-glycosidic linkage.
In conclusion, MFAS exhibited an intermolecular transglycosylation activity that might be
employed to synthesize a variety of functional materials from sucrose with high efficiency.
Acknowledgment This work was supported by a National Research Foundation of Korea (NRF) grant funded
by the Korean government (MEST) (No. 2013–031011) and by Business for Cooperative R&D between
Industry, Academy, and Research Institute funded Korea Small and Medium Business Administration in 2013
(Grant No. C0123542).
References
1. Davies, G., & Henrissat, B. (1995). Structure, 3, 853–859.
2. van der Maarel, M. J. E. C., van der Veen, B., Uitdehaag, J. C. M., Leemhuis, H., & Dijkhuizen, L. (2002).
Journal of Biotechnology, 94, 137–155.
3. Bertoldo, C., & Antranikian, G. (2002). Current Opinion in Chemical Biology, 6, 151–160.
4. Seo, D.H., Jung, J.H., Ha, S.J., Yoo, S.H., Kim, T.J., Cha, J., & Park, C.S. (2008). Carbohydrate-active
enzyme structure, function and application (Park, K.H., ed.), CRC press, Boca Raton, pp. 125–140.
5. Albenne, C., Skov, L. K., Mirza, O., Gajhede, M., Feller, G., D'Amico, S., et al. (2004). The Journal of
Biological Chemistry, 279, 726–734.
6. Wang, R., Bae, J.S., Kim, J.H., Kim, B.S., Yoon, S.H., Park, C.S., & Yoo, S.H. (2012). Food Chemistry. 132,
773–779
7. Park, H., Choi, K., Park, Y. D., Park, C. S., & Cha, J. (2011). Journal of Life Science, 21, 1631–1635.
8. Daudé, D., Champion, E., Morel, S., Guieysse, D., Remaud-Siméon, M., & André, I. (2013).
ChemCatChem, 5, 2288–2295.
9. Jung, J.H., Seo, D.H., Ha, S.J., Song, M.C., Cha, J., Yoo, S.H., Kim, T.J., Baek, N.I., Baik, M.Y., & Park,
C.S. (2009). Carbohydrate Research. 344, 1612–1619
10. Seo, D.H., Jung, J.H., Ha, S.J., Song, M.C., Cha, J., Yoo, S.H., Kim, T.J., Baek, N.I., & Park, C.S. (2009).
Journal of Molecular Catalysis B: Enzymatic. 60, 113–118
11. Cho, H. K., Kim, H. H., Seo, D. H., Jung, J. H., Park, J. H., Baek, N. I., et al. (2011). Enzyme Microbial
Technology, 49, 246–253.
12. Seo, D. H., Jung, J. H., Ha, S. J., Cho, H. K., Jung, D. H., Kim, T. J., et al. (2012). Applied Microbiology and
Biotechnology, 94, 1189–1197.
13. Potocki De Montalk, G., Remaud-Simeon, M., Willemot, R. M., Planchot, V., & Monsan, P. (1999). Journal
of Bacteriology, 181, 375–381.
14. Pizzut-Serin, S., Potocki-Véronèse, G., van der Veen, B. A., Albenne, C., Monsan, P., & Remaud-Simeon,
M. (2005). FEBS Letters, 579, 1405–1410.
15. Ha, S. J., Seo, D. H., Jung, J. H., Cha, J., Kim, T. J., Kim, Y. W., et al. (2009). Bioscience Biotechnology and
Biochemistry, 73, 1505–1512.
16. Seo, D. H., Choi, H. C., Kim, H. H., Yoo, S. H., & Park, C. S. (2012). Journal of Microbiology and
Biotechnology, 22, 1253–1257.
17. Kim, H. S., Park, H. J., Heu, S., & Jung, J. (2004). Journal of Bacteriology, 186, 411–418.
18. Kim, M. I., Kim, H. S., Jung, J., & Rhee, S. (2008). Journal of Molecular Biology, 380, 636–647.
19. Mikkat, S., Hagemann, M., & Schoor, A. (1996). Microbiology,142, 1725–1732.
20. Marin, K., Zuther, E., Kerstan, T., Kunert, A., & Hagemann, M. (1998). Journal of Bacteriology, 180, 4843–
4849.
Appl Biochem Biotechnol
21. Takenaka, F., Uchiyama, H., & Imamura, T. (2000). Bioscience Biotechnology and Biochemistry, 64, 378–
385.
22. Sawai, T., & Hehre, E. J. (1962). The Journal of Biological Chemistry, 237, 2047–2052.
23. Takenaka, F., & Uchiyama, H. (2000). Bioscience Biotechnology and Biochemistry, 64, 1821–1826.
24. Smith, P. K., Krohn, R. I., Hermanson, G. T., Mallia, A. K., Gartner, F. H., Provenzano, M. D., et al. (1985).
Analytical Biochemistry, 150, 76–85.
25. Niesen, F. H., Berglund, H., & Vedadi, M. (2007). Nature Protocols, 2, 2212–2221.
26. Skov, L. K., Mirza, O., Henriksen, A., De Montalk, G. P., Remaud-Simeon, M., Sarçabal, P., et al. (2001).
The Journal of Biological Chemistry, 276, 25273–25278.
27. Guérin, F., Barbe, S., Pizzut-Serin, S., Potocki-Véronèse, G., Guieysse, D., Guillet, V., et al. (2012). The
Journal of Biological Chemistry, 287, 6642–6654.
28. Schneider, J., Fricke, C., Overwin, H., Hofmann, B., & Hofer, B. (2009). Applied and Environmental
Microbiology, 75, 7453–7460.
29. Nakano, H., Kiso, T., Okamoto, K., Tomita, T., Bin Abdul Manan, M., & Kitahata, S. (2003). Journal of
Bioscience and Bioengineering, 95, 583–588.
30. Goedl, C., Sawangwan, T., Mueller, M., Schwarz, A., & Nidetzky, B. (2008). Angewandte Chemie,
International Edition, 47, 10086–10089.
31. Schwarz, A., Thomsen, M. S., & Nidetzky, B. (2009). Biotechnology and Bioengineering, 103, 865–872.
32. Hinz, S. W. A., Verhoef, R., Schols, H. A., Vincken, J. P., & Voragen, A. G. J. (2005). Carbohydrate
Research, 340, 2135–2143.
33. Masuda, T., Kawano, A., Kitahara, K., Nagashima, K., Aikawa, Y., & Arai, S. (2003). Journal of Nutritional
Science and Vitaminology. (Tokyo) 49, 64–68.
34. Wei, W., Qi, D., Zhao, H., Lu, Z., Lv, F., & Bie, X. (2013). Food Chemistry, 141, 3085–3092.
35. Suhr, R., Scheel, O., & Thiem, J. (1998). Journal of Carbohydrate Chemistry, 17, 937–968.
36. Goedl, C., Sawangwan, T., Wildberger, P., & Nidetzky, B. (2010). Biocatalysis and Biotransformation, 28,
10–21.
37. Seo, S., Tomita, Y., Tori, K., & Yoshimura, Y. (1978). Journal of the American Chemical Society, 100, 3331–
3339.
38. Cassel, S., Debaig, C., Benvegnu, T., Chaimbault, P., Lafosse, M., Plusquellec, D., et al. (2001). European
Journal of Organic Chemistry, 2001, 875–896.
39. He, F., Hogan, S., Latypov, R. F., Narhi, L. O., & Razinkov, V. I. (2010). Journal of Pharmaceutical
Sciences, 99, 1707–1720.
40. Goldberg, D. S., Bishop, S. M., Shah, A. U., & Sathish, H. A. (2011). Journal of Pharmaceutical Sciences,
100, 1306–1315.
41. Vedadi, M., Arrowsmith, C. H., Allali-Hassani, A., Senisterra, G., & Wasney, G. A. (2010). Journal of
Structural Biology, 172, 107–119.