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Cloning, expressionand characterizationofan extracellular enolasefromLeuconostocmesenteroidesJin-Ha Lee1, Hee-Kyoung Kang1, Young-Hwan Moon2, Dong Lyun Cho3, Doman Kim1,4,Jun-Yong Choe5, R. Honzatko6 & John F. Robyt6
1Engineering Research Institute, Chonnam National University, Gwang-Ju, South Korea; 2Department of Material and Chemical and Biochemical
Engineering, Chonnam National University, Gwang-Ju, South Korea; 3Faculty of Applied Chemical Engineering, Chonnam National University, Gwang-
Ju, South Korea; 4School of Biological Sciences and Technology, Chonnam National University, Gwang-Ju, South Korea and 5California Institute of
Technology, Division of Chemistry, Pasadena, CA, USA; and 6Department of Biochemistry, Biophysics and Molecular Biology, Iowa State University,
Ames, IA, USA
Correspondence: Doman Kim, School of
Biological Sciences and Technology, Chonnam
National University, Gwang-Ju, 500-757,
South Korea. Tel.: 182 62 530 1844;
fax: 182 62 530 0874;
e-mail: [email protected]
Received 4 January 2006; revised 28 March
2006; accepted 7 April 2006.
First published online 3 May 2006.
doi:10.1111/j.1574-6968.2006.00274.x
Editor: Dieter Jahn
Keywords
enolase; Leuconostoc mesenteroides ; cloning;
expression; 2-phospho-D-glucose;
phosphoenolpyruvate.
Abstract
Enolase on the surface of streptococci putatively facilitates pathogenic invasion of
the host organisms. The related Leuconostoc mesenteroides 512FMCM is nonpatho-
genic, but it too has an extracellular enolase. Purified isolates of extracellular
dextransucrase from cultures of L. mesenteroides contain minute amounts of
enolase, which separate as small crystals. Expression of L. mesenteroides enolase in
Escherichia coli provides a protein (calculated subunit mass of 47 546 Da) catalyzing
the conversion of 2-phsopho-D-glycerate to phosphoenolpyruvate. The pH opti-
mum is 6.8, with Km and kcat values of 2.61 mM and 27.5 s�1, respectively. At
phosphate concentrations of 1 mM and below, fluoride is a noncompetitive
inhibitor with respect to 2-phospho-D-glycerate, but in the presence of 20 mM
phosphate, fluoride becomes a competitive inhibitor. Recombinant enolase sig-
nificantly inhibits the activity of purified dextransucrase, and does not bind human
plasminogen. Results here suggest that in some organisms enolase may participate
in protein interactions that have no direct relevance to pathogenic invasion.
Introduction
Enolase (2-phospho-D-glycerate hydrolyase, EC 4.2.1.11) cat-
alyzes the dehydration of 2-phospho-D-glycerate (2PGA) to
phosphoenolpyruvate (PEP) in glycolysis, and the reverse
reaction in gluconeogenesis (Wold & Ballou, 1957; Wold,
1971). All characterized enolases require divalent metal ca-
tions for activity (Wold & Ballou, 1957; Holt & Wold, 1961;
Westhead & McLain, 1964; Wold, 1971; Wang & Himoe, 1974;
Faller et al., 1977; Pietkiewicz & Kustrzeba-Wojcicka, 1983;
Brewer, 1985). The natural cofactor is probably Mg21, which
confers the highest activity (Wold & Ballou, 1957; Faller et al.,
1977; Pietkiewicz & Kustrzeba-Wojcicka, 1983). Enolases are
homodimers in all eukaryotes examined thus far, with native
molecular weights of 80–100 kDa (Wold & Ballou, 1957; Holt
& Wold, 1961; Westhead & McLain, 1964; Wold, 1971; Wang
& Himoe, 1974). Enolases from prokaryotes and archaeons
either are dimers or octamers, the latter with native masses in
the range 350�400 kDa (Kaufmann & Bartholmes, 1992;
Peak et al., 1994; Schurig et al., 1995; Brown et al., 1998a).
Subunit molecular weights of most enolases are c. 45 kDa
(Wold & Ballou, 1957; Holt & Wold, 1961; Westhead &
McLain, 1964; Wold, 1971; Wang & Himoe, 1974; Faller
et al., 1977; Pietkiewicz & Kustrzeba-Wojcicka, 1983; Brewer,
1985; Kaufmann & Bartholmes, 1992; Peak et al., 1994;
Schurig et al., 1995; Brown et al., 1998a). Enolase from
Streptococcus rattus is evidently an exception to the general
trend, being a dimer of native mass 49 kDa (Huther et al.,
1990). Prokaryotic a-enolases are highly conserved proteins
that may function extracellularly in patho-physiological pro-
cesses (Pancholi, 2001). Surface-displayed a-enolase is a major
plasmin(ogen)-binding protein of Streptococcus pneumoniae
(Bergmann et al., 2001). Despite the absence of a signal
sequence and typical motifs required for membrane anchor-
ing, the attachment of enolase (and other glycolytic enzymes)
to cell surfaces of streptococci and pneumococci, and the
attendant effects on plasmin(ogen) acquisition are well estab-
lished (Pancholi & Fischetti, 1998; Bergmann et al., 2001). a-
Enolase (designated Eno) reassociates with the surface of
pneumococci, as observed by immunoelectron microscopy,
with a concomitant increase in plasmin(ogen)-binding capa-
city (Bergmann et al., 2001).
FEMS Microbiol Lett 259 (2006) 240–248c� 2006 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved
In the present study, enolase appears as an impurity in
extracellular preparations of dextransucrase from non-
pathogenic Leuconostoc mesenteroides, an organism used in
the production of clinical dextran and dextran found in
fermentation foods (Kim & Kim, 1999; Kim et al., 2003).
Although present in vanishingly small quantities, enolase
crystallizes from nearly pure preparations of dextransucrase.
To the best of our knowledge, this is the first report of an
extracellular enolase produced by Lactobacillaceae or any
other nonpathogenic prokaryote. In addition, small quan-
tities of recombinant enolase significantly inhibit dextran-
sucrase, suggesting a functional relationship between surface
enolase and dextransucrase in L. mesenteroides.
Materials and methods
Materials
2PGA was purchased from Sigma Chemical Co. (P-0257).
All other chemicals were of reagent grade and commercially
available. Leuconostoc mesenteroides B-512FMCM exhibits a
500-fold increase in extracellular dextransucrase levels rela-
tive to the commercial strain (B-512F) from which it
originated by vacuum UV irradiation and selection (Kim
et al., 1997). Production and secretion of dextransucrase by
strain B-512FMCM is constitutive, whereas dextransucrase
production by strain B-512F is exclusively sucrose inducible.
Restriction endonucleases, alkaline phosphatase and T4
DNA ligase came from Boehringer Mannheim (Mannheim,
Germany), Kosco (Seongnam, Korea) and Takara (Shiga,
Japan), respectively.
Conditions of bacterial culture
Leuconostoc mesenteroides B-512FMCM was grown in LW
medium [0.5% (w/v) yeast extract, 0.5% (w/v) KH2PO4,
0.02% (w/v) MgSO4 � 7H2O, 0.001% (w/v) NaCl, 0.001%
(w/v) FeSO4 � 7H2O, 0.001% (w/v) MnSO4 �H2O, 0.013%
(w/v) CaCl2 � 2H2O] containing 2% glucose at 28 1C without
aeration (Kim et al., 1997; Kim & Kim, 1999).
Preparation of dextransucrase
Isolation of secreted dextransucrase from cultures of L.
mesenteroides followed the procedure of Kitaoka & Robyt
(1998): cells were removed from the medium by centrifuga-
tion (15 000 g, 10 min, 4 1C), and the dextransucrase was
concentrated and dialyzed by passing the bacterium-free
culture through a polysulfone ultrafiltration hollow fiber
cartridge [H5P100-43 (100 kDa cut-off), Amicon, Inc.,
Beverly, MA]. Dialyzed and concentrated dextransucrase
was loaded onto a Toso Haas DEAE 5pw HPLC column,
equilibrated with 100 mM KPi, pH 6.5, and then eluted by a
gradient 0–1 M in NaCl. The eluent was monitored by UV
absorbance at 280 nm. Fractions containing dextransucrase
were verified by the addition of 5 mL of the eluent to 5 mL of
200 mM sucrose, followed shortly thereafter (10 min reac-
tion) by the addition of 30 mL of 100% ethanol. Copious
precipitation of dextran confirmed the presence of dextran-
sucrase. Protein purity was determined by sodium dodecyl
sulfate-polyacrylamide gel electrophoresis (SDS-PAGE)
(Laemmli, 1997), followed by staining with Coomassie
Brilliant Blue R250 (Laemmli, 1997).
Crystallization of enolase from dextransucrasepreparations
Crystals of extracellular enolase were grown from prepara-
tions of purified dextransucrase by the method of hanging
drops by the combination of 2 mL volumes of protein and
precipitant solutions. The protein solution contained dex-
transucrase (10 mg mL�1), ammonium sulfate (100 mM)
and sodium acetate (50 mM) at pH 5.4. The precipitant
solution contained 24% polyethylene glycol 3000, ammo-
nium sulfate (100 mM), and sodium acetate (50 mM), pH
5.4. Wells contained 500 mL of precipitant solution.
N-terminal amino-acid sequencing
Crystals from several droplets were washed thoroughly with
the precipitation buffer dissolved in de-ionized water and
submitted to the Iowa State University protein facility for N-
terminal sequencing by Edman degradation. Extracellular
proteins isolated from a L. mesenteroides culture were
separated by SDS-PAGE and then transferred electrophor-
etically to a polyvinylidene difluoride (PVDF) membrane
(Bio-Rad, Hercules, CA) (Otter et al., 1987).
DNA isolation and gene cloning
Genomic DNA from L. mesenteroides B-512FMCM was
prepared as described previously (Kim et al., 2000). Routine
DNA manipulations, including plasmid purification and
Escherichia coli transformation, followed protocols of Man-
iatis et al. (1989). Plasmid DNA was isolated from an
overnight culture of Escherichia coli using the alkaline lysis
method (Maniatis et al., 1989). Extraction of chromosomal
DNA and plasmid DNA from agarose gels employed an
AccuPreps Gel Purification kit (Bioneer Co., Daejeon,
Korea). Competent E. coli DH5a cells, prepared by the
procedure of Cohen et al. (1973), were transformed with
plasmid DNA using the CaCl2 method.
Two degenerate primers were designed from the
N-terminal amino-acid sequence, determined in this study,
and from conserved amino-acid sequences for other
bacterial enolases (GenBank accession numbers AF065394,
AAKO04742, AB029313, AJ401152 and AJ303085): E1F, 50-
ATCACTGATATTATCGCACGCGAAGTCCTT-30 and E1R,
FEMS Microbiol Lett 259 (2006) 240–248 c� 2006 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved
241Extracellular enolase from Leuconostoc mesenteroides
50-TGAAAGTGAACCAGTTTTGATACCAGC-30. These pri-
mers were used in the amplification of a 1173 bp fragment of
the complete gene. The PCR mixture contained 10 mM Tris/
HCl (pH 8.5), 50 mM KCl, 3 mM MgCl2, 2 mM of each the
deoxyribonucleotide triphosphates (dNTP), 0.25 mg of
genomic DNA from L. mesenteroides B-512FMCM, and
10 pmol of each of the two primers. After incubation for
5 min at 94 1C, 1 mL Taq DNA polymerase (Takara, Japan)
was added, followed by 25 cycles of denaturation (94 1C for
30 s), annealing (62 1C for 30 s) and elongation (72 1C for
30 s). The PCR fragment was ligated into pGEM-T Easy
vector (Promega) for DNA sequencing, followed by the
amplification of 50- and 30-regions by the thermal asym-
metric interlaced (TAIL) PCR method (Liu & Whittier,
1995). The nucleotide sequence of the initial clone facili-
tated the design of internal primers for the forward reaction
(E2R, 50-GCGTTTGCACCCAAGTTACCCTTGTTT-30) and
the backward reaction (E2F, 50-GTTGGTGATGACTT
CTTCGTTACTAACAC-30). Self-ligated L. mesenteroides
B-512FMCM chromosomal DNA, after partial digestion
with BamHI and PstI, was used as a template in PCR,
following the protocols described above. The amplified
fragment was inserted in pGEM-T Easy vector and se-
quenced. Finally, the whole enolase gene was amplified by
PCR using chromosomal DNA from L. mesenteroides B-
512FMCM and two oligonucleotide primers (E1F, as above,
and E3R, 50-TTACTTGTTTTCAATAACTTCG-30) derived
from the nucleotide sequences of the 50- and 30-termini. The
amplified gene was inserted into pGEM-T Easy vector and
cloned using E. coli DH5a.
Over expression and purification ofrecombinant enolase
PCR was performed to introduce a BamHI site at the
initiation codon and a KpnI site downstream from it. The
PCR fragment was ligated into the BamHI and KpnI sites of
the pRSETA vector (Invitrogen, The Netherlands) predi-
gested with BamHI and KpnI. The expression plasmid was
then transformed into E. coli BL21(DE3)pLysS (Invitrogen)
behind the T7 promoter. Escherichia coli BL21(DE3)pLysS
carrying enolA was grown in 50 mL LB medium containing
50mg mL�1 ampicillin and 1 mM isopropyl-a-D-thiogalac-
topyranoside (IPTG) at 28 1C for 6 h. The cells were
harvested by centrifugation (15 000 g, 10 min, 4 1C),
suspended in 200 mL 50 mM imidazole, pH 6.8, and
disrupted by sonication. One percent (v/v) Triton X-100
was added to release the enzyme from the cell membrane.
Cell debris was removed by centrifugation (15 000 g, 10 min,
4 1C). The N-terminal, 6-histidyl tagged (6xHis tag)
protein was purified by nickel-nitrilotriacetic acid-agarose
(Ni-NTA) affinity chromatography (Qiagen, Germany). The
molecular weight of recombinant enolase was determined,
using the Laemmli system (10% w/v) acrylamide gel
(Laemmli, 1997). Proteins were stained with Coomassie
Brilliant Blue R250.
Enolase kinetics
The concentration of recombinant His-tagged enolase was
determined by absorbance of UV radiation at 280 nm, using
an extinction coefficient of 0.9 mL mg�1 cm�1 determined
for rabbit muscle enolase (Kustrzeba-Wojcicka & Golczak,
2002). The activity of recombinant His-tagged enolase was
measured spectrophotometrically at 240 nm by the conver-
sion of 2PGA to phosphoenolpyruvate (Kustrzeba-Wojcicka
& Golczak, 2002). The temperature of assays was 30 1C.
Initial velocities came from the slopes of linear progress
curves of 1 min duration. The assay buffer was 50 mM
imidazole-HCl, pH 6.8, 400 mM KCl and 3 mM MgSO4 in
a total volume of 1.5 mL, unless noted otherwise. Under
these conditions and a substrate concentration of 3 mM, an
increase in absorbance of 0.2 corresponds to the conversion
of 0.226 mmol of substrate (Kustrzeba-Wojcicka & Golczak,
2002). One unit of activity was defined as the conversion of
1 mmol of substrate in 1 min under the described reaction
conditions in the presence of 3 mM 2PGA. In thermostabil-
ity studies, a 10 mL solution of enolase (34.7 U mg�1,
0.003 mg mL�1) was divided into two portions. One sample
was incubated at 50 1C in the assay buffer, with aliquots
drawn at time intervals of 30 min for 3 h. The second sample
was incubated at room temperature as a control. Variations
in enolase activity with respect to pH from 6.0 to 8.3 were
determined by assays using 3 mM 2PGA. The reaction was
initiated by the addition of 10 mL enolase (34.7 U mg�1,
0.003 mg mL�1) to the reaction mixture.
The kinetic constants, Km and kcat, were determined from
measurements of the initial reaction rates, using Linewea-
ver–Burk plots (Lineweaver & Burk, 1934). Concentrations
of 2PGA varied from 0.3 to 5 mM, and the reactions were
initiated by the addition of 10 mL enolase (34.7 U mg�1,
0.003 mg mL�1). The effects of fluoride ions on the kinetics
of enolase from L. mesenteroides B-512FMCM were studied
with and without orthophosphate. Initial reaction rates were
measured in a reaction mixture (1.5 mL) containing 10mL
enolase (34.7 U mg�1, 0.003 mg mL�1), 0.3–5 mM 2-PGA,
0–80 mM fluoride, at 0, 1 and 20 mM phosphate, and
incubated at 30 1C for 1 min. The effects of magnesium, zinc
and manganese divalent cations on enolase activity were
studied. The enzyme solution was dialyzed beforehand
against deionized water, because enzyme prepared in water
does not express any catalytic activity as reported by
Kustrzeba-Wojcicka et al. (1986) and Kustrzeba-Wojcicka
& Golczak (2002). Concentrations of divalent cations varied
from 25 to 1500 mM, containing with 3 mM 2PGA and 10mL
enolase (34.7 U mg�1, 0.003 mg mL�1).
FEMS Microbiol Lett 259 (2006) 240–248c� 2006 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved
242 J.-H. Lee et al.
Dextransucrase kinetics
The effect of recombinant His-tagged enolase (from
1.49� 10�3 to 14.89� 10�3 nmol per reaction, 1.78 U n-
mol�1) on dextransucrase (8.23� 10�2 nmol, 3.64 U n-
mol�1) was measured by the rate of fructose release from
sucrose (Kim & Kim, 1999). The highly purified dextransu-
crase (8.23� 10�2 nmol, 3.64 U nmol�1) used in these ex-
periments exhibited no detectable enolase activity. The
molecular weight of synthesized dextran was determined
using Bio Gel A-0.5 M (1.5� 100 cm), by the micro redu-
cing value and total carbohydrate analyses (Fox & Robyt,
1991). The degrees of dextranase hydrolysis of dextrans
produced with or without enolase were determined using
Penicillium dextranase (1.0 U; D-4668, Sigma Chemical Co.,
St Louis, MO). The dextranase was added to 1.0 mL,
containing 10 mg of the various dextrans, pH 5.5 (20 mM
citrate-phosphate buffer), at 37 1C and allowed to react
for 3 h. Aliquots (1–5 mL) were added to 20� 20 cm
Whatman K5F TLC plates, which were irrigated at 22 1C,
using two ascents (18 cm path length) of 4 : 10 : 3 volume
proportions of nitromethane–1-propanol–water. The
carbohydrates were visualized on the plate by rapidly
dipping the plate into a solution containing 0.3% (w/v)
N-(1-naphthyl) ethylenediamine and 5% (v/v) H2SO4
in MeOH, dried, and heated at 120 1C for 10 min (Kitaoka
& Robyt, 1998).
Detection of plasminogen binding
To detect plasminogen-binding activity, the recombinant
enolase was incubated overnight at 4 1C with 1%, human
plasminogen (Sigma) in 50 mM Tris-HCl, 110 mM NaCl,
pH 7.4. The enolase–plasminogen mixture was applied to
nickel–nitrilotriacetic acid-agarose (Ni–NTA) affinity col-
umn and checked whether plasminogen was bound to the
recombinant enolase using Chromogenic Assay Kit for
plasma plasminogen (American Diagnostica Inc.).
Results
Characterization of enolase crystals
Isolates of dextransucrase are at least 95% pure based on
SDS-PAGE (data not shown). Such preparations under
conditions of crystallization reproducibly produced tiny
bipyramidal crystals of c. 10–20 mm within 24 h. Protein
from washed crystals provided an unambiguous N-terminal
sequence (SLITDIIARE) that does not exist in the
known sequence of dextransucrase from L. mesenteroides
B-512FMCM, or for that matter, in any know sequence
of dextransucrase. A BLAST search for sequence homology,
using the first 10 residues tentatively identified the crystal-
line material as an enolase. Substantial 2PDG hydrolysis by
dissolved crystals, by the original preparation of dextransu-
crase and by the supernatant of new cultures of L. mesenter-
oides B-512FMCM corroborated the presence of enolase.
Crystals from different preparations of dextransucrase were
used in data collection at the APS-Structural Biology Center,
Argonne National Laboratory, Illinois, and National Syn-
chrotron Light Source, Brookhaven National Laboratory,
New York. All crystals belonged to space group I4 (unit cell
dimensions a = b = 145 A, and c = 101 A) and diffract to
2.4 A resolution. A preliminary structure clearly reveals an
enolase octamer, with two of its subunits defining the
asymmetric unit of the crystal (Jun-Yong Choe and Richard
B. Honzatko, unpublished results).
Cloning and nucleotide sequence of enolase
The 4.3 kb DNA fragment recovered from PCR protocols
and ligated into the BamHI and KpnI sites of the pRSETA
vector is called hereafter pENOLA. The nucleotide sequence
of subclones from pENOLA revealed one major open
reading frame of 1326 bp, coding for a polypeptide of
442 amino-acid residues (Fig. 1). The sequence has been
submitted to GenBank (accession identifier AB088633).
The 442 amino acids encoded by enolA correspond to a
molecular mass of 47 546 Da. The predicted amino-acid
sequence of L. mesenteroides B-512FMCM enolase was
compared with sequences in GenBank using the ClustalW
program (Thompson et al., 1994). ENOLA is similar in
sequence to various bacterial enolases: Lactococcus lactis
ssp. lactis (GenBank accession number, AAK04742)
(Bolotin et al., 2001), 67% identity and 80% homology;
Streptococcus pneumoniae (AJ303085) (Bergmann et al.,
2001), Staphylococcus aureus (AAC17130) (Kuroda et al.,
2001), 62% identity and 75% homology; and Bacillus
subtilis (NP391270), 62% identity and 75% homology
(Kunst et al., 1997).
Purification and characterization ofrecombinant enolase
Purified enolase migrated as a single band on SDS-PAGE,
with a molecular mass of 51.4 kDa (Fig. 2). The difference
between the observed and the calculated mass of 47 546 Da is
largely due to the His-tag. Specific activity of purified
recombinant enolase is 34.7 U mg�1. Enolase activity fell
30% and 36% after 0.5 and 3 h of incubation at 50 1C. The
pH of optimum enolase activity was 6.8 (Fig. 3). Above and
below pH 6.8, the catalytic activity remained constant at
30% maximal activity. Enolase displayed classical Michae-
lis–Menten kinetics. The Km and kcat for 2PGA were
2.61 mM (� 0.01) and 27.53 s�1 (� 1.21), respectively. The
inhibition type of enolase from L. mesenteroides B-
512FMCM by fluoride was noncompetitive with respect to
2PGA in the absence of phosphate (Fig. 4a) and in the
FEMS Microbiol Lett 259 (2006) 240–248 c� 2006 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved
243Extracellular enolase from Leuconostoc mesenteroides
presence of 1 mM phosphate (Fig. 4b), but competitive in
the presence of 20 mM phosphate (Fig. 4c). The Ki for
fluoride ions, calculated using the graphic method of Dixon
& Webb (1979), were 5.4, 4.6 and 9.2 mM in the 0, 1 and
20 mM phosphate, respectively. In the presence of Zn21,
Vmax was 4.11 (� 0.02) mmol min�1 (Table 1).
The addition of enolase to dextransucrase reduced
relative dextransucrase activity by up to 24.9% (Table 2
and Fig. 5) in the presence of 14.89� 10�3 nmol enolase/
8.23� 10�2 nmol dextransucrase (Table 2). As the ratio of
enolase to dextransucrase increased to greater than 0.06, the
relative dextransucrase activity remained much the same
(Fig. 5). The addition of enolase did not significantly change
the structure (branch formation) and polydispersity index
of dextran, based on dextranase hydrolysis and GPC (data
not shown).
Immobilized recombinant enolase did not retain plasmi-
nogen (data not shown), indicating the absence of a high-
affinity interaction between the recombinant enolase and
plasminogen.
Discussion
The sequence of the cloned gene enolA from L. mesenteroides
B-512FMCM is in Fig. 1. The N-terminal region of
Fig. 1. Nucleotide and deduced amino-acid sequences of enolase from
L. mesenteroides B-512FMCM. The N-terminal amino-acid sequence
from the purified L. mesenteroides B-512FMCM enolase crystal is boxed.
Conserved residues with other bacterial enolases (Lactococcus, Strepto-
coccus, Staphylococcus and Bacillus) are printed in boldface.
kDa
200
116
97.4
66.2
45
51.4 kDa
Fig. 2. SDS-PAGE and molecular weight determination of purified
recombinant enolase. The following molecular weight markers (Bio-
Rad) were used: myosin (200 kDa), b-galactosidase (116 kDa), phosphor-
ylase b (97.4 kDa), serum albumin (66.2 kDa), ovalbumin (45 kDa),
carbonic anhydrase (31 kDa), trypsin inhibitor (21.5 kDa), lysozyme
(14.4 kDa), and aprotinin (6.4 kDa).
0
0.1
0.2
0.3
0.4
5.5 6.5 7.5 8.5pH
2-P
GA
, µM
min
−1
Fig. 3. pH profile of enolase from L. mesenteroides B-512FMCM.
FEMS Microbiol Lett 259 (2006) 240–248c� 2006 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved
244 J.-H. Lee et al.
L. mesenteroides B-512FMCM enolase is 75�88% similar to
other bacterial enolases (Lactococcus, Streptococcus, Entero-
coccus and Bacillus). The calculated subunit molecular
weight the encoded protein is 47 546 Da, similar to
subunit molecular weight of the other enolases (Wold
& Ballou, 1957; Pietkiewicz & Kustrzeba-Wojcicka, 1983;
Peak et al., 1994): E. coli enolase 46 kDa (Wold &
Ballou, 1957), Pyrococcus furiousus enolase 45 kDa (Peak
et al., 1994), Saccharomyces cerevisiae enolase 46.7 kDa,
rabbit muscle enolase 41 kDa (Wold & Ballou, 1957) and
carp muscle enolase 49 kDa (Pietkiewicz & Kustrzeba-
Wojcicka, 1983).
Because enolase is a heat shock protein, its high thermo-
stability is not unusual. Similarly, highly thermostable yeast
enolase and Streptococcus rattus (Huther et al., 1990) enolase
have been characterized. Enolases from higher organisms
(carp, rabbit muscle and bovine brain), however, are less
resistant to thermal denaturation (Wold & Ballou, 1957;
Pietkiewicz & Kustrzeba-Wojcicka, 1983; Nazarian et al.,
1992; Kustrzeba-Wojcicka & Golczak, 2002).
0
1
2
3
4
5
−2 −1 0 1 2 3 41/s, mM
1/v
0
2
4
6
8
−1.25 0 1.25 2.5
1/s, mM
1/v
0
1
2
3
4
5
6
7
8
9
−3 −1.5 0 1.5 31/s, mM
1/v
(a)
(b)
(c)
Fig. 4. (a) Determination of the mode for fluoride ions inhibition with-
out phosphate ion. �, 0 mM fluoride ions (SD = � 0.209); ’, 20 mM
fluoride ions (SD = �0.793); m, 40 mM fluoride ions (SD = � 1.516).
(b) Determination of the mode for fluoride ions inhibition in the presence
of 1 mM phosphate ions. �, 0 mM fluoride ions (SD = � 0.256); ’,
20 mM fluoride ions (SD = � 0.702); ., 40 mM fluoride ions (SD =
� 1.622). (c) Determination of the mode for fluoride ions inhibition in
the presence of 20 mM phosphate ions. �, 0 mM fluoride ions (SD =
� 0.209); ’, 20 mM fluoride ions (SD = � 1.352); m, 40 mM fluoride
ions (SD = �1.645).
Table 1. Influence of magnesium, manganese and zinc ions on kinetics
parameters of enolase from L. mesenteroides B-512FMCM
Ion
Vmax (mmol conversion of
2-PGA per min of enzyme) Km (mM)
Mg21 2.91 (�0.02) 0.007 (� 0.001)
Mn21 3.14 (�0.02) 0.009 (� 0.002)
Zn21 4.11 (�0.02) 0.009 (� 0.001)
Parentheses show the standard deviation of Vmax and Km.
PGA, phospho-D-glycerate.
Table 2. Influences of enolase on dextransucrase activity on the average
molecular size and on resistance to dextranase hydrolysis of dextrans
produced
Enolase concentration
in digest
(� 10�3 nmol)�Average
MW (� 106)
Relative
dextransucrase
activity (%)
Unhydrolyzed
dextran
0 95.0 100 7.3
1.49 68.0 59.5 7.7
4.96 67.1 34.6 8.5
9.92 60.4 33.3 8.3
14.89 60.2 24.9 7.5
�Different amounts of the purified enolase (1.78 U nmol�1) were added
on dextransucrase (3.64 U nmol�1) digest.
The enolase activity in the purified dextransucrase was measured and
there was no detectable enolase activity found.
0
20
40
60
80
100
120
0 0.05 0.1 0.15 0.2Enolase/dextransucrase (nmol nmol−1)
Rel
ativ
ede
xtra
nsuc
rase
act
ivity
(%
)
Fig. 5. Influence of enolase on dextransucrase activity.
FEMS Microbiol Lett 259 (2006) 240–248 c� 2006 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved
245Extracellular enolase from Leuconostoc mesenteroides
The pH optima of the enolases from various organisms
are similar. The optimum pH of 6.8 for L. mesenteroides B-
512FMCM enolase is similar to the enolase optima of other
strains (Pietkiewicz & Kustrzeba-Wojcicka, 1983). The pH
activity profile for Leuconostoc enolase seems characteristic
insofar as its activity decreases sharply at pH values above
and below pH 6.8.
The Km value for the enolase from L. mesenteroides
B-512FMCM is 2.61 mM with 2-PGA as substrate, which is
greater than those of other reported enolases, e.g., the Km for
enolase from P. furiousus is 0.38 mM (Peak et al., 1994), carp
enolase Km is 0.31 mM (Pietkiewicz & Kustrzeba-Wojcicka,
1983) and yeast enolase Km is 0.12 mM (Westhead
& McLain, 1964). However, enolase from S. rattus has a Km
of 4.35 mM (Huther et al., 1990). In addition, the specific
activity of L. mesenteroides enolase (34.7 U mg�1) is low
relative to those from other bacterial sources for which
specific activities vary from 100 to 900 U mg�1 (Brown
et al., 1998b). The low specific activity is in part an artifact
due to the use of 3 mM 2PGA as the standard substrate
concentration in the definition of a unit of activity for
enolase. The Vmax value of Fig. 4a translates into a specific
activity (70 U mg�1) that differs only modestly from the
reported range in the literature.
Fluoride is an inhibitor of L. mesenteroides enolase, with
a Ki of 5.4, 4.6 and 9.2 mM in 0, 1 and 20 mM phosphate,
respectively. The Ki values are significantly higher
than inhibition constants for enolases from S. rattus
(Ki�0.85 mM Huther et al., 1990) and from carp muscle
(Ki�0.24 mM Pietkiewicz & Kustrzeba-Wojcicka, 1983) in
the absence of phosphate. Moreover, low concentrations of
phosphate greatly enhance fluoride inhibition of yeast enolase
(Maurer & Nowak, 1981; Nowak & Maurer, 1981). Pi, metal
cations and fluoride combine with the active site to form a
dead-end quaternary complex, which presumably blocks the
productive binding of the substrate in the yeast system.
Unlike the yeast system, fluoride inhibits L. mesenteroides
enolase with no difference in potency or kinetic mechanism
in the absence or presence of phosphate. In a true noncom-
petitive mechanism a fluoride ion must bind to the enzyme-
Mg21-2PGA complex, most likely displacing a water
molecule from the catalytic metal. Charge neutralization
of the catalytic Mg21 by fluoride should destabilize develop-
ing charge on the 3-OH group of 2PGA during the transition
state. The change in mechanism (noncompetitive to com-
petitive) at 20 mM Pi may simply reflect the depletion of
free Mg21 due to the formation of F�-Mg21-Pi complexes
in solution.
For most enolases the most effective metal activator is
Mg21 (Wang & Himoe, 1974; Pietkiewicz & Kustrzeba-
Wojcicka, 1983; Lee & Nowak, 1992a, b). Mg21 strongly
activates enolase from L. mesenteroides B-512FMCM, but
Mn21 and Zn21 are at least equally effective. Mn21 strongly
activates Candida albicans and Saccharomyces cerevisiae
enolases (Lee & Nowak, 1992a, b), but is a weaker activator
of the enolase from carp muscle. Zn21 induces higher
activity than Mn21 in carp enolase, but the converse is true
for the enolases of yeast and C. albicans (Pietkiewicz &
Kustrzeba-Wojcicka, 1983; Lee & Nowak, 1992a, b). In
much of the literature concerning enolases, the metal ion
that binds with highest affinity is called the ‘conformational’
or ‘structural’ metal ion, whereas the metal ion that binds
with lower affinity is the ‘catalytic’ metal ion (Faller et al.,
1977; Brewer, 1985). The activation effect by metal cations
studied here probably relates to metal binding at the
catalytic site of L. mesenteroides enolase.
Streptococcus pneumoniae can colonize the mucosal sur-
faces of the human respiratory tract without clinical symp-
toms (Austrian, 1996). Pneumococci and other Gram-
positive pathogens express specific cell surface components
called adhesins that mediate their adherence to host tissues,
thereby facilitating not only colonization but also invasion
(Cundell et al., 1995). Host plasmin(ogen) binds to cell
surface receptors of these pathogenic bacteria, and at least
two of these receptors are glycolytic enzymes, enolase and
glyceroaldehyde-3-phosphate dehydrogenase (Pancholi &
Fischetti, 1998; Winram & Lottenberg, 1998). Evidently, in
the case of surface enolase from Streptococcus pneumoniae,
a specific sequence (FYDKERKVYD) plays a critical role
in plasmin(ogen) binding (Bergmann et al., 2003). In
L. mesenteroides enolase, the corresponding sequence is
LYDAETKTYK. The absence of a binding interaction between
plasmin(ogen) and L. mesenteroides enolase is consistent with
the suggested critical role played by the central six residues of
the above sequence in the tight binding of synthetic peptides
to plasmin(ogen) (Bergmann et al., 2003).
As L. mesenteroides enolase does not bind plasmin(ogen),
consistent with the nonpathogenic properties of this organ-
ism, then what role does enolase play on the cell surface?
Optimum inhibition of dextransucrase at low molar ratios
of enolase is consistent with two general mechanisms: (i)
Enolase binds to single molecules of dextransucrase, catalyz-
ing a conformational change to a new conformational state
that slowly reverts to its original state after the dissociation
of enolase. (ii) Enolase binds to a dextransucrase multimer,
acting as an allosteric effector. A dextransucrase multimer of
more than a dozen subunits has no support from the
literature. The former mechanism, however, is analogous to
the role of a chaperone, and at least one isozyme of
eukaryotic enolase is a stressed-induced protein presumably
linked to protein folding activities (Aaronson et al., 1995).
Perhaps surface enolase in L. mesenteroides facilitates the
folding of dextransucrase, and dissociates from dextransu-
crase after folding is complete. Clearly protein–protein
interactions involving surface enolases may go well beyond
the recognition of plasmin(ogen).
FEMS Microbiol Lett 259 (2006) 240–248c� 2006 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved
246 J.-H. Lee et al.
Acknowledgements
This work was supported by a Korea Research Foundation
Grant (KRF-Y00-290) and National Institutes of Health
Research Grant NS 10546.
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