Báo cáo khoa học: Differential susceptibility of Plasmodium falciparum versus yeast and mammalian enolases to dissociation into active monomers doc

Differential susceptibility of Plasmodium falciparum
versus yeast and mammalian enolases to dissociation into
active monomers
Ipsita Pal-Bhowmick, Sadagopan Krishnan and Gotam K. Jarori
Department of Biological Sciences, Tata Institute of Fundamental Research, Colaba, Mumbai, India

Keywords
enolase; monomers; Plasmodium
falciparum; rabbit muscle; yeast
Correspondence
G. K. Jarori, Department of Biological
Sciences, TIFR, Homi Bhabha Road, Colaba,
Mumbai 400 005, India
Fax: +91 22 22804610
Tel: +91 22 22782000
E-mail:
(Received 7 December 2006, revised 16
January 2007, accepted 12 February 2007)
doi:10.1111/j.1742-4658.2007.05738.x

In the past, several unsuccessful attempts have been made to dissociate
homodimeric enolases into their active monomeric forms. The main objective
of these studies had been to understand whether intersubunit interactions
are essential for the catalytic and structural stability of enolases. Further
motivation to investigate the properties of monomeric enolase has arisen
from several recent reports on the involvement of enolase in diverse nonglycolytic (moonlighting) functions, where it may occur in monomeric
form. Here, we report successful dissociation of dimeric enolases from Plasmodium falciparum, yeast and rabbit muscle into active and isolatable
monomers. Dimeric enolases could be dissociated into monomers by high
concentrations ( 250 mm) of imidazole and ⁄ or hydrogen ions. Two forms
were separated using Superdex-75 gel filtration chromatography. A detailed
comparison of the kinetic and structural properties of monomeric and

dimeric forms of recombinant P. falciparum enolase showed differences in
specific activity, salt-induced inhibition and inactivation, thermal stability,
etc. Furthermore, we found that enolases from the three species differ in
their dimer dissociation profiles. Specifically, on challenge with imidazole,
Mg(II) protected the enolases of yeast and rabbit muscle but not of
P. falciparum from dissociation. The observed differential stability of the
P. falciparum enolase dimer interface with respect to mammalian enolases
could be exploited to selectively dissociate the dimeric parasite enzyme into
its catalytically inefficient, thermally unstable monomeric form. Thus enolase could be a novel therapeutic target for malaria.

Enolase (EC 4.2.1.11) is a glycolytic enzyme that catalyzes the interconversion between 2-phospho-d-glycerate (PGA) and phosphoenolpyruvate. Enolases from
most organisms exist as homodimers of subunit mass
40–50 kDa [1], the exceptions being homo-octameric
enolases from thermophilic bacteria [2–4]. A variety of
physical data indicate that each dimer contains two
active sites. The active site in each subunit is completely independent [5,6]. As the active site is fully
contained in each subunit, attempts to dissociate the

dimeric form into an active monomeric form have been
made using genetic [7], chemical and physical methods,
but without much success. Although dissociation of
the dimer could be achieved, the monomers formed
were found to be inactive [8–11]. The formation of active monomers was inferred under conditions of high
temperature (40–45 °C) and low protein concentration
(in the nanomolar range) [12,13]. However, at such
low concentrations, neither the kinetic nor the structural characterization of the monomer could be carried

Abbreviations
Mim, imidazole-generated monomer; Mnt, native monomer; MpH, pH-generated monomer; 2-PGA, 2-phospho-D-glycerate; RMen, rabbit
muscle enolase; r-Pfen, recombinant Plasmodium falciparum enolase; Yen, yeast enolase.

1932

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I. Pal-Bhowmick et al.

out. Hydrostatic pressure, which is viewed as a gentle
and reversible perturbation, has also been employed in
attempts to obtain active monomers from dimeric enolase [8,11,14–17]. Although the dissociation of enolase
dimers into active monomers was inferred in some of
the above studies, it has never been unequivocally
demonstrated. As none of these attempts had resulted
in the isolation of active monomers and their characterization, it was not clear whether the monomeric
form is intrinsically inactive or the means of dissociation selectively inactivated it. Suggestions have also
been made that intersubunit interactions may be essential for completion of the catalytic cycle. In recent
years, enolase has also been shown to participate in a
variety of nonglycolytic (moonlighting) biological functions [18]. The oligomeric state of enolase recruited for
the moonlighting functions is not known.
Our interest in the enolase from the malarial parasite (Plasmodium falciparum) originates from the fact
that the intraerythrocytic stages of the parasite lack
active mitochondria and hence rely solely on glycolysis
for their energy needs [19,20]. The level of glycolytic
flux in parasite-infected cells is  50–100-fold greater
than that in uninfected red blood cells, and the activity
of many glycolytic enzymes is upregulated, enolase
being one of them [21–23]. P. falciparum enolase (Pfen)
could be a potential drug target, as there is only one
gene for this enzyme, and it shows greater resemblance

to plant enolases than to mammalian enolases. In
order to examine such possibilities and explore whether it has any moonlighting functions, we have
recently cloned the P. falciparum enolase gene, overexpressed it, and obtained pure protein [24]. We have
also raised polyclonal and monoclonal antibodies
against recombinant r-Pfen for subcellular localization
studies [25]. Our observations have shown a diverse
subcellular localization (enolase is associated with
plasma membrane, cytosol, cytoskeletal elements and
nucleus; unpublished results) for enolase, indicating
that it may be recruited for certain other nonglycolytic
functions.
One of the conventional approaches in rational drug
design has been to make active site-specific inhibitors
that can differentiate between host and parasite proteins and bring about selective inhibition of the parasite enzyme. The major limitation of this strategy is
that active sites are evolutionarily highly conserved,
and structural differences may be rather subtle or even
nonexistent. As there are numerous protein–protein
interactions that operate in living systems [26–28], and
most proteins exist as oligomers [29] with a predominance of homodimers [30], another approach could
be to target protein–protein interfaces for perturbing

Dissociation of enolase into active monomers

protein and cell functions [31]. In cases where oligomeric structure is essential for biological activity,
selective disruption of such an interface in a protein of
parasite origin can have therapeutic effects. Rationally
designed peptides that can compete for the interaction
between monomeric subunits (peptidomimetics) have
yielded encouraging results [32–35]. Attempts have also
been made to find nonpeptide small molecule inhibitors that effectively interfere with protein–protein

interactions [36–38]. Here, we have examined the possibility of dissociating malarial parasite enolase into
monomers using small molecules, and characterized
the properties of the monomeric state. We report the
successful dissociation of r-Pfen, and yeast and rabbit
muscle enolases, into isolatable active monomers. Thus
we demonstrate that the dimeric structure is not essential for catalysis. However, comparative kinetic and
structural studies on the monomeric and dimeric forms
of r-Pfen showed several interesting differences. The
monomeric form has low specific activity, is more thermolabile, and is more prone to lose activity in the
presence of salts as compared to the dimer. Our experiments have also identified conditions under which
selective dissociation of the parasite enolase may be
accomplished. Although the concentrations of dissociating ligand used in this study are rather high and
unrealistic for therapeutic applications, the possibility
remains that such differences may be exploitable for
targeting this enzyme for therapeutic purposes.

Results
Imidazole-induced dissociation of r-Pfen
into active monomers
Like most enolases, r-Pfen is a homodimer of two
50 kDa subunits [24]. Purified dimeric r-Pfen was dialyzed against increasing concentrations (0–250 mm) of
imidazole and then subjected to gel filtration chromatography on a Superdex-75 column. Figure 1A shows
gel filtration chromatograms obtained at several different concentrations of imidazole at pH 6.0. A dimer
peak was observed in the absence of imidazole. However, with increasing concentrations of imidazole, the
monomeric fraction increased, and at  250 mm
imidazole, > 95% of r-Pfen was dissociated. The effect
of pH on imidazole-induced dissociation was examined
by dialyzing the enzyme against 50 mm sodium phosphate at different pH values (6.0, 7.0 and 8.0) with
increasing concentrations of imidazole. The percentage
of monomer present in each of these samples was

determined from gel filtration chromatograms. The
results presented in Fig. 1B show that lower pH

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1933


Dissociation of enolase into active monomers

A

I. Pal-Bhowmick et al.

r-Pfen
100

[Imidazole]
250 mM

B

pH 6.0
pH 7.0

75
50

pH 8.0

25
0

80

50 mM

0

dimer

50

0 mM

0
40

50

60
Ve (ml)

70

100 150 200
[Imidazole](mM)

250
6

C

4
40
2
0

0

80

50

55

60

65

70

75

(----- )

2

50

Specific activity ( U/ mg )

0

150 mM

0

A 280 (mAU)

A 280 (mAU)

0
50

% monomer

monomer

75

80

Ve (ml)

Fig. 1. Imidazole-induced dissociation of r-Pfen. (A) Superdex-75 gel filtration chromatograms obtained at different concentrations of imidazole. Enzyme (0.5 mg) was dialyzed ( 14 h) against 50 mM sodium phosphate (pH 6.0) containing different amounts (0–250 mM) of imidazole. The column was pre-equilibrated with appropriate buffer, and chromatography was performed at room temperature (20 ± 1 °C). (B)
Effect of pH on imidazole-induced dissociation of r-Pfen. Each data point is an average of two chromatographic runs. (C) Gel filtration chromatogram of r-Pfen in 50 mM sodium phosphate containing 250 mM imidazole (pH 6.0). A280 nm and specific activity (r—r) for each fraction
are shown.

1934

Yen

A

A 280 (mAU)

2

dimer
(a)

0

dimer

10
(b)

0

monomer

3
(c)

0
40

B

50

60
Ve(ml)

70

80

RMen
dimer

2
0

A 280 (mAU)

favored the dissociation of r-Pfen. Figure 1C shows a
gel filtration chromatogram of a sample dialyzed
against 250 mm imidazole in 50 mm sodium phosphate
(pH 6.0). The enzyme activity of each fraction was
measured, and the specific activity was plotted along
with protein concentration (A280) as a function of elution volume. The results showed that both forms
(monomeric and dimeric) were catalytically active, with
the dimer having  3-fold greater specific activity than
the monomer. Experiments were also performed in
which the effects of adding NaCl to the dissociation
buffer were examined. The results showed that
the presence or absence of salt (300 mm NaCl) did

not have any effect on monomer–dimer equilibrium.
Imidazole-generated monomers of r-Pfen (Mim)
( 10 lm), when extensively dialyzed against imidazole-free buffer, could reassociate to form active
dimers. To the best of our knowledge, this is the first
demonstration of obtaining active monomers of
enolase that could be separated from dimers.
The ability of imidazole to dissociate enolases from
other species was also examined. Imidazole at 250 mm
and pH 6.0 failed to dissociate yeast enolase (Yen).
However, inclusion of 300 mm NaCl along with
250 mm imidazole at pH 6.0 resulted in almost complete dissociation of Yen (Fig. 2A). In the case of rabbit muscle enolase (RMen), 250 mm imidazole did not
dissociate the enzyme. As commercial preparations of
RMen contain Mg(II), we included 5 mm EDTA along
with imidazole. This resulted in partial dissociation of

(a)

35
0
15

dimer
(b)
dimer

monomer

(c)
0
40

50

60
Ve(ml)

70

80

Fig. 2. Imidazole-induced dissociation of Yen and RMen. Samples
were in 50 mM sodium phosphate (pH 6.0), and 0.2–0.6 mg of
protein was used for each chromatographic run on a Superdex-75
column. (A) Yen: (a) no imidazole; (b) 250 mM imidazole; and
(c) 250 mM imidazole + 300 mM NaCl. (B) RMen: (a) no imidazole;
(b) 250 mM imidazole; and (c) 250 mM imidazole + 5 mM EDTA.

FEBS Journal 274 (2007) 1932–1945 ª 2007 The Authors Journal compilation ª 2007 FEBS


I. Pal-Bhowmick et al.

Dissociation of enolase into active monomers

RMen (Fig. 2B). However, further inclusion of
300 mm NaCl did not lead to complete dissociation of
RMen. In this respect, RMen differs from Yen.
As Mg(II) is an essential cofactor for stabilization of
the active conformation and catalysis, extensive kinetic
and direct metal ion-binding studies have been performed in the past. These studies have shown that each

enolase subunit has three binding sites for the divalent
cation, namely a conformational site (site I), a catalytic
site (site II) and an inhibitory site (site III) [39–42]. As
binding of divalent cation induces large conformational changes in enolase, we examined the effect of
Mg(II) on imidazole-induced dissociation of r-Pfen,
Yen and RMen in the presence of different compounds. The results are summarized in Table 1. Inclusion of EDTA [to chelate residual Mg(II) in the
protein sample] and NaCl in the dissociation buffer
did not have any effect on the dimeric state. However,
the presence of Mg(II) affected the imidazole-induced
(or imidazole + NaCl-induced) dissociation of RMen
and Yen. It is interesting to note that in the presence
of 1.5 mm MgCl2, imidazole could dissociate r-Pfen
but had no effect on Yen and RMen.

Table 1. Effect of Mg(II) on monomer–dimer equilibrium in enolase.
Enolase, 10 lM (0.5 mgỈmL)1), was dialyzed against 50 mM
sodium phosphate (pH 6.0) containing different compounds as indicated below. Concentrations of these compounds when used
were: [imidazole] ¼ 250 mM; [EDTA] ¼ 5 mM; [NaCl] ¼ 300 mM;
[MgCl2] ¼ 1.5 mM. The Superdex-75 column was pre-equilibrated
with the same buffer as used for dialysis.
Dimeric–monomeric states of enolasesa

Enolase

P. falciparum
(r-Pfen)

Yeast
(Yen)

Rabbit muscle
(RMen)

+
+
+
+
+

Dimer
Dimer
Dimer
Monomer
Monomer

Dimer
Dimer
Dimer
Dimer
Dimer

+ Imidazole + NaCl

Monomer

Monomer

+ MgCl2 +
imidazole + NaCl
+ MgCl2 +

imidazole + EDTA
+ MgCl2 +
imidazole +
EDTA + NaCl

Monomer

Dimer

Dimer
Dimer
Dimer
Dimer
Partial
monomer
Partial
monomer
Dimer

Monomer

Dimer

Monomer

Monomer

a

MgCl2

EDTA
EDTA + NaCl
Imidazole
Imidazole + EDTA

Partial
monomer
Partial
monomer

When the amount of dimer or monomer is ‡ 90%, it is stated as
‘dimer’ or ‘monomer’. When the amount is ‡ 40%, it is stated as
‘partial monomer’.

pH-induced dissociation of enolases
The effect of pH on the oligomeric state of enolases
was examined by performing gel filtration chromatography on the enzyme, dialyzed against 50 mm sodium
phosphate of the desired pH. Figure 3A presents the
chromatographic profiles of r-Pfen in the pH range
4.5–8.0. Figure 3B shows that 50% dissociation of
r-Pfen occurs around pH 5.5. Monomeric r-Pfen generated by low pH (MpH) was also found to be enzymatically active (see below). Low pH could also dissociate
Yen and RMen. The half-dissociation point for Yen
was around pH 5 (Fig. 3C,D), whereas for RMen it
was around pH 5.5 (Fig. 3E,F).
Activity and reassociation of MpH and Mim
Measurements of enzyme activity in monomeric and
dimeric fractions showed that Mim had  3-fold less
specific activity than dimers (Fig. 1C). As the assay
solution did not contain imidazole, it is possible that
the observed activity of Mim may have arisen

from reassociation to dimers. Similarly, MpH could
also reassociate at the assay solution pH of 7.4. Such
reassociation of monomers into dimers during an
enzyme assay would result in an increase in the slope
of the reaction progress curve (as the dimer is  3-fold
more active than monomers). From these considerations, we can state that during an enzyme assay using
MpH or Mim: (a) if there is reassociation of monomers
into dimer on the timescale of the enzyme assay, the
reaction progress curve will exhibit a time-dependent
increase in slope; and (b) as the amount of dimer
formed increases as the square of the monomer concentration, a plot of monomer concentration versus
activity would have an upward curvature if there was
a significant amount of dimer formed during the
enzyme assay.
Figure 4A shows the reaction progress curves at
three different concentrations of MpH (0.04, 0.17 and
0.43 lm). The observed increase in slope with time suggests reassociation of MpH into dimers under our assay
conditions. A similar experiment performed using
Mim (Fig. 4B) did not show any change in slope with
time. We measured the concentration dependence of
specific activity for MpH at time  0.0 min as well as
at time > 1 min (Fig. 4C). The first slope reflects the
monomer activity, whereas the slope determined
after > 1 min reflects the dimer activity. With the
increasing concentration of MpH, initially we observed
a very low specific activity ([MpH] > 0.5 lm). However,
at higher protein concentrations, the rate of formation
of dimer increased rapidly, and only a single slope,

FEBS Journal 274 (2007) 1932–1945 ª 2007 The Authors Journal compilation ª 2007 FEBS

1935


Dissociation of enolase into active monomers

I. Pal-Bhowmick et al.

A 280 (mAU)

A 280 (mAU)

A 280 (mAU)

A

B

D

F

Fig. 3. pH-induced dissociation of enolases from different organisms. Each enzyme (0.2–0.6 mg) was dialyzed against 50 mM sodium phosphate (pH 4.5–8.0) containing 150 mM NaCl and subjected to gel filtration chromatography. (A) Gel filtration profile of r-Pfen at different pH
values. (B) pH versus % monomer (data are from two different experiments) for r-Pfen. The r-Pfen used here was prepared by eluting the
protein from Ni–nitrilotriacetic acid resin at low pH. (C) Gel filtration chromatograms for Yen. (D) pH versus% monomer for Yen. (E) Gel filtration chromatograms for RMen. (F) pH versus % monomer for RMen.

characteristic of dimer-specific activity, could be
observed (Fig. 4C). As in the low-concentration range
of MpH the observed activity is rather small, it is likely
that either MpH is inactive and the measured activity is

a reflection of the formation of tiny amounts of active
dimer, or the MpH has intrinsically very low activity.
Irrespective of these two possibilities, the observed second slope reflects the activity for the dimer formed
during the assay. As expected, the specific activity
computed using the second slope did not exhibit the
monomer concentration-dependent variation (Fig. 4C).
These results indicate that MpH rapidly reassociates to
form dimer. However, similar measurements of concentration dependence of activity for Mim gave a linear
increase in activity, suggesting that r-Pfen Mim did not
associate rapidly (with respect to assay timescale) to
form more active dimers (Fig. 4D). A replot of these

1936

data as specific activity versus Mim concentration
(Fig. 4D) showed that the observed specific activity
was low ( 1.8–1.9 unitsỈmg)1, a characteristic of Mim)
and was concentration invariant on enzyme assay timescales. These observations support the view that Mim
associates slowly to form dimers, whereas MpH associates rapidly. Thus MpH and Mim differ in their ability
to reassociate, indicating that they represent two different conformational states of the monomeric form of
the enolase. A schematic representation of the dissociation and association of r-Pfen is presented in
Scheme 1, where Mnt is the native monomer, which is
assumed to be the only monomeric form that can
dimerize. As Mim mostly remained in the monomeric
state under our assay conditions, kinetic characterization of the monomeric forms of enolase was performed
using Mim only.

FEBS Journal 274 (2007) 1932–1945 ª 2007 The Authors Journal compilation ª 2007 FEBS

I. Pal-Bhowmick et al.

A

Dissociation of enolase into active monomers

B

pH monomer (MpH ):
reaction progress curves

1.354

1.464

(µM)

1.34

Imidazole monomer (M im ):
reaction progress curves
(µM)

0.04

1.45
1.44

0.13

1.32

A 240 nm

0.17
1.30

A 240 nm

1.43
1.42
1.41

1.28

0.33

1.40
1.39

0.43

1.255

0.6

1.379

0.00

0.2

0.4

0.6

0.8

1.0

1.2

0.00

1.4 1.50

0.2

0.4

0.6

D

pH monomer (MpH ):
concentration dependence of activity

1.2

1.4 1.50

Imidazole monomer (Mim ):
concentration dependence of activity
0.40

)

2.5

1.8

1.5

1.0

0.5

1.6
0.30
1.4
0.25
1.2
0.20
1.0
0.15
0.8

Activity (units) (o

2.0

o)

0.35

Specific Activity (units·mg–1) (

Specific Activity (units·mg–1)

1.0

Time (minutes)

Time (minutes)

C

0.8

0.10
0.6
0.05

0.0
0.0

0.1

0.2

0.3

0.4

0.2

0.5

[Monomer] (µM)

0.3

0.4

0.5

0.6

0.7

[Monomer] (µM)

Fig. 4. Reaction progress curves for the monomeric r-Pfen-catalyzed reaction. Conversion of phosphoenolpyruvate to 2-PGA was monitored
at 240 nm. (A) MpH. (B) Mim. Note the time-dependent change in progress curve slopes in (A). (C) Variation in specific activity as a function
of [MpH], calculated from first (time  0.0 min) (d) and second (time > 1 min) (j) slopes of reaction progress curves. Note the change in
specific activity computed from the time  0.0 min slope, reflecting the rapid formation of dimer at higher concentrations of MpH. (D) Variation in activity (and specific activity) as a function of [Mim]. Activity showed a linear increase (o), with specific activity remaining constant
(d), suggesting that Mim did not associate to form high-activity dimer on the enzyme assay timescale.

MpH from Yen and RMen also showed nonlinear
reaction progress curves, very similar to those observed

for r-Pfen (data not shown). In enzyme assays, Mim
from Yen gave nonlinear reaction progress curves displaying an increase in slope with time. This is likely to
be due to Mg(II)-induced rapid dimerization of yeast
Mim. As Mg(II) does not affect the dimerization of rPfen monomers, such a change of slope was not
observed for Mim prepared from r-Pfen (Fig. 4B).
Observed low activities of monomeric (Mim and MpH)
enolases from all three species would imply that quaternary interactions between two subunits stabilize catalytically more active conformations of each subunit.
Comparison of kinetic properties of monomeric
(Mim) and dimeric forms of r-Pfen
As Mim did not reassociate rapidly into dimers, this
form of the enzyme was used for comparative kinetic

studies. Figure 5A shows the variation of enzyme activity as a function of phosphoenolpyruvate concentration.
Data were fitted to the Michaelis–Menten equation
using sigmaplot software. The nonlinear fit of data
gave Vmax ẳ 13.5 0.5 Uặmg)1, and Km (phosphoenolpyruvate) ¼ 0.28 ± 0.03 mm for dimers, and
Vmax ¼ 3.7 0.3 Uặmg)1, and Km (phosphoenolpyruvate) ẳ 0.38 0.07 mm for monomers, respectively.
Thus disruption of subunit–subunit interactions resulted
in a significant decline in enzyme activity ( 3-fold), but
did not have much effect on Km. We also compared the
thermal stability of monomeric (Mim) and dimeric
forms. Equal amounts of protein were incubated at
three different temperatures (4 °C, 37 °C and 50 °C),
and activity was assayed at different time intervals. The
dimeric form was stable for a prolonged (£ 250 min)
duration at 37 °C, and showed  20–25% inactivation
at 50 °C. In comparison, the monomeric form was
 80% inactivated at 37 °C (£ 250 min) and completely

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Dissociation of enolase into active monomers

I. Pal-Bhowmick et al.

A

Imidazole
monomer

+H+ (slow)

MpH

Mnt
-imidazole
(slow)

Native
monomer

-H+ (fast)

pH
monomer

Scheme 1. Schematic representation of dissociation of the dimeric

(D) form of enolase to the imidazole-induced and pH-induced monomeric (Mim or MpH) forms. The dimer is shown to dissociate into
the native form (Mnt), which has an enzyme activity similar to that
of the dimer. Lowering of the pH or addition of imidazole (or both)
stabilize different conformations of monomers. Mim and MpH differ
in activity and stability from the dimer and Mnt. MpH is rapidly converted into Mnt on raising of the pH, whereas Mim is slow to convert to Mnt, a form that is competent to form dimers.

inactivated at 50 °C in £ 150 min (Fig. 5B). Thus,
subunit–subunit interface interactions confer higher
thermal stability to the protein.
As enolases from different organisms are known to
differ in their response to different salts [43], we examined the effect of various salts on the catalytic activity of
the monomeric and dimeric forms. To assess the effect
of a salt, we assayed the enzyme with assay mixtures containing different concentrations of a salt.
Figure 6A–C shows the effects of NaCl, KCl and KBr
on the activity of the dimeric and monomeric (Mim)
forms of r-Pfen. NaCl inhibited both forms of the
enzyme, but the inhibition was stronger for the monomeric form (Fig. 6A). In the case of KCl and KBr, the
dimeric form was mildly activated ( 10–20%), whereas
the monomeric form was strongly inhibited (Fig. 6B,C).
For assessing the activating ⁄ inactivating effect of the
salts (NaCl, KCl and KBr), the enzyme was incubated in
buffer containing several different concentrations of salt
for 24 h at 20 ± 1 °C, and then assayed in the absence
of the salt. The results are presented in Fig. 6D–F. At
median concentrations ( 300 mm), NaCl and KCl had
an activating effect on the dimeric form of the enzyme,
whereas all three salts had a concentration-dependent
inactivating effect on the monomeric form.
Comparison of CD and fluorescence spectra
of monomeric and dimeric enolase

The effect of the loss of subunit–subunit interface
interactions on the secondary and ⁄ or tertiary structure
1938

8
6
4
2
0
0.0

0.2

0.4

0.6

0.8

1.0

[PEP] (m M)

B
1.4
1.2

Fractional activity

Mim

+ imidazole
(slow)

Activity (Units. mg -1)

10

Native
dimer (D)

1.0
0.8
0.6
0.4
0.2
0.0
0

50

100

150

200

250

300

Time (min)
Fig. 5. (A) Comparison of monomer and dimer activity with varying
concentrations of phosphoenolpyruvate. For activity measurements,
50 lL of 2 lM r-Pfen (monomeric or dimeric) was added to 450 lL
of assay mixture. As monomeric protein was in 250 mM imidazole,
the final concentration of imidazole in the assay mixture was
25 mM. The presence of 25 mM imidazole in the assay mixture had
no effect on dimer activity. Data were fitted to the Michaelis–
Menten equation. The best-fit parameters are Km (phosphoenolpyruvate) ¼ 0.38 ± 0.07 mM and Vmax (specic activity) ẳ
3.7 0.3 Uặmg)1 for monomer (j), and Km (phosphoenolpyruvate) ¼ 0.28 ± 0.03 mM and Vmax ¼ 13.5 ± 0.5 mg)1 for dimer
(h). (B) Temperature dependence of stability of dimeric (open symbols) and monomeric (Mim) (filled symbols) forms of r-Pfen. Enzyme
was incubated at 4 °C (s or d), 37 °C (n or m) and 50 °C (h or
j). The activity was assayed at different time intervals.

of enolase was probed by recording CD and fluorescence spectra of monomeric and dimeric forms of the
protein. As monomer preparations made by imidazole
treatment contain about 250 mm imidazole, which
interferes with fluorescence and CD spectra, it was not
possible to record meaningful spectra for Mim. Instead,
we recorded CD and fluorescence spectra for r-Pfen
and Yen MpH, and compared them with the spectra
from dimers (Fig. 7). CD spectra of monomeric and
dimeric forms (r-Pfen and Yen) were very similar
(Fig. 7A,C). Analysis of r-Pfen CD spectra using the

FEBS Journal 274 (2007) 1932–1945 ª 2007 The Authors Journal compilation ª 2007 FEBS


I. Pal-Bhowmick et al.

Dissociation of enolase into active monomers

140

140

A

120

140

B

100

100

80

80

80

60

60

60

40

40

40

20
140

20
140

C

120

100

% Activity

120

20
140

D

F

E

120

120

120

100

100

100

80

80

80

60

60

60

40

40

40
0

100 200 300 400 500 600
[NaCl] (mM)

0

100 200 300 400 500 600
[KCl] (mM)

0

100 200 300 400 500 600
[KBr] (mM)

Fig. 6. Effect of NaCl, KCl and KBr on the activity of monomeric (Mim) (j) and dimeric (h) r-Pfen. Upper panel (a, b, c): Enzyme samples
were assayed in the presence of different concentrations of salts. Lower panel (d, e, f): Enzyme samples (200 lL of 2 lM r-Pfen) containing
different concentrations of salts were incubated for 24 h at 20 °C, and assayed in the absence of the salt. For each assay, 50 lL of the
enzyme was added to 450 lL of the assay mixture. Activity is expressed as percentage of control where no salt was added to the enzyme
sample.

cdnn program for secondary structure analysis [44]
gave helix 36.6%, beta 14.1%, turn 19.3% and random 29.9% for the dimeric form, and helix 31.5%,
beta 18.3%, turn 18.4% and random 31.8% for the
monomeric form. Thus, it appears that dissociation of
dimers to form monomers lead to a slight decrease in
helical content with a concomitant increase in the beta
sheet content.
Fluorescence emission spectra of the monomic,

dimeric and denatured states of r-Pfen and Yen are
shown in Fig. 7B,D. For both of these enzymes, dissociation of enolase into monomers led to a decrease in
emission intensity. In the case of r-Pfen, the emission
maximum was blue-shifted upon dissociation (Fig. 7B;
compare traces a and b), whereas for Yen, a slight red
shift was observed (Fig. 7D). Such a red shift may
arise because the dimer interface of Yen has Trp56
(which becomes solvent-exposed upon dissociation),
whereas the analogous position in Pfen is occupied by
Tyr59.

Discussion
Alignment of enolase sequences from several different
organisms have shown that it is a highly conserved
protein [18]. Furthermore, the comparison of known
three-dimensional structures showed complete posi-

tional conservation of active site residues across species
[5,6,41,45–49]. The enolase polypeptide chain folds into
two domains, with the small domain having a mixture
of a-helices and b-sheets, and the large domain having
an a ⁄ b-barrel structure (Fig. 8D). The binding of substrate brings these two domains together to constitute
an active site. As most enolases are homodimeric, the
intersubunit interface is formed by contacts between
the small domain of one subunit and the large domain
of the other subunit. If the subunit–subunit interface is
large, the dissociated monomers are generally unstable,
due to exposure of large hydrophobic surfaces to the
solvent [50]. The intersubunit interface in enolases is
rather small ( 11–13% buried surface) and quite

hydrophilic, suggesting that it may be possible to dissociate the dimer into active monomers.
In the past, several attempts have been made to dissociate the dimeric enolase into active monomers
[7–9,11,14–17]. However, the active monomeric form
could never be obtained in isolation from the dimeric
form. Failure to obtain active monomers of enolase
strengthened the belief that enolases are active only in
the dimeric state [51] and that the quaternary structure
of enolase is necessary for catalytic activity. Thus, any
attempt to dissociate the dimer is accompanied by
changes in the tertiary and secondary structures that
are detrimental to enzyme activity [8]. However, the

FEBS Journal 274 (2007) 1932–1945 ª 2007 The Authors Journal compilation ª 2007 FEBS

1939


Dissociation of enolase into active monomers

I. Pal-Bhowmick et al.

4000

4000
2000

A

r-Pfen
Mean Residue Ellipticity

2
-1
(degrees cm dmol )

Mean Residue Ellipticity
(degrees cm2 dmol -1)

0
-2000
-4000
-6000
-8000

b

-10000

a

-12000

C Yen

2000
0
-2000
-4000
-6000
-8000

b

-10000
-12000

-14000

a

-14000
200

210

220

230

240

250

200

260

210

Wavelength (nm)

240

250

260

3.5e+8

B r-Pfen

D Yen

a

5e+8

b

4e+8
3e+8

c

2e+8

a

3.0e+8

Fluorescence Intensity

Fluorescence intensity

230

Wavelength (nm)

7e+8
6e+8

220

b

2.5e+8

c

2.0e+8
1.5e+8
1.0e+8

1e+8

5.0e+7

0

0.0

320

340

360

380

320

340

360

380

Wavelength (nm)

Wavelength (nm)

Fig. 7. Comparison of CD and fluorescence spectra of monomeric (MpH) and dimeric forms of r-Pfen and Yen. (A) CD spectra of (a) dimer
and (b) monomer of r-Pfen. (B) Fluorescence spectra of (a) dimer, (b) monomer and (c) urea-denatured r-Pfen. (C) Yen CD spectra: (a) dimer,
(b) monomer. (D) Fluorescence spectra of (a) dimer, (b) monomer and (c) urea-denatured Yen. Fluorescence emission intensities were normalized for protein concentration.

results presented here unequivocally establish that both
H+ and imidazole are able to dissociate enolase dimers
into active monomers that could be isolated from the
dimeric form (Figs 1–3). We observed that at any
given concentration, the ability of imidazole to dissociate the dimer was enhanced by low pH (Fig. 1B). In
the absence of imidazole, a change in pH (in the range

6–8) did not have any significant effect on the dissociation of the enzyme (Fig. 1B), suggesting that the
increased dissociation induced by imidazole at low pH
is mostly due to pH-dependent ionization of imidazole.
Imidazole has a pKa of 6.9, and hence at pH 6.0 it will
exist predominantly as an imidazolium ion. Thus, the
data presented in Fig. 1B are commensurate with the
idea that it is the imidazolium ion that is effective in
dissociating the r-Pfen dimer into monomers (Mim).
The pH-dependent dissociation of various enolases
indicate that it is the protonation of group(s) at the
intersubunit interface with a pKa  5–5.5 that is
responsible for the dissociation (Fig. 3).
An examination of the enolase intersubunit interface
showed that it is stabilized by two salt bridges, several
1940

hydrogen bonds, and p–cation (Tyr or Trp-Arg) and
hydrophobic interactions. A relatively low contribution
of the salt bridge in stabilizing the dimer interface was
evident from the fact that replacement of an interface
glutamate residue with a leucine (E414L) in RMen
did not result into the dissociation of dimer. DGo for
dissociation for the mutant enzyme decreased from
49.7 kJỈmol)1 to 42.3 kJỈmol)1 [7]. One may ask the following question: which are the subunit interface interactions that imidazolium and ⁄ or hydrogen ions can
possibly disrupt? An examination of the intersubunit
interface suggests the following two possibilities. (a)
There is a hydrogen bond between His191 NE2 and the
carbonyl oxygen of Gly15 (or Arg14) where His191
NE2 acts as a proton donor (Fig. 8A,B). Imidazole,
being an analog of histidine, can compete for this

hydrogen bond. The importance of this interaction in
stabilizing the dimer is evident from the observed changes in the intersubunit hydrogen bond pattern on
Mg(II) binding. In one Mg(II)-bound form of Yen
(Protein Data Bank: 2ONE, 1EBH), a hydrogen bond
forms between His191 and Arg14 (Fig. 8B) with a bond

FEBS Journal 274 (2007) 1932–1945 ª 2007 The Authors Journal compilation ª 2007 FEBS


I. Pal-Bhowmick et al.

Dissociation of enolase into active monomers

A

B

C

D

Fig. 8. Intersubunit interface hydrogen bond involving His191 in enolase crystal structures. (A) Yen with two Mg(II) ions bound per subunit
(Protein Data Bank: 1EBG) showing hydrogen bond between His191 and Gly15. (B) Yen with one Mg(II) ion bound per subunit (Protein Data
Bank: 2ONE) has a hydrogen bond between His191 and Arg14. (C) Neuronal enolase in which two subunits are asymmetrically bound to
one and two Mg(II) ions exhibits both types of hydrogen bond. The subunit bound to one Mg(II) ion has a hydrogen bond between His189
and Arg14, and the other subunit, which is bound to two Mg(II) ions (Protein Data Bank: 1TE6), has a His189–Gly15 hydrogen bond. (D) Ribbon diagram of dimeric Yen (Protein Data Bank: 1EBG), in which yellow and blue represent two subunits. Interface residues (Trp56 and
Arg184) involved in p–cation interaction are shown in a stick representation.

length (as measured between two non-hydrogen atoms
˚

in the hydrogen bond) of 3.2–3.3 A. In two Mg(II)bound forms (Protein Data Bank: 1ONE, 1EBG), the
hydrogen bond is between His191 and Gly15 (Fig. 8A)
˚
with a bond length of 2.9–3.0 A. Thus, the loss of
Mg(II) may favor weaker hydrogen bonds and dissociation, whereas addition of Mg(II) may reverse the process, leading to subunit–subunit association. It is
interesting to note that in the neuronal enolase crystal
structure (Protein Data Bank: 1TE6), where one subunit is bound to one Mg(II) ion and the other subunit
has two Mg(II) ions, both types of hydrogen bond

(His–Gly and His–Arg) are observed (Fig. 8C). Our
observations that Mg(II) favors dimer formation are in
agreement with earlier reports [15], and suggest that
interactions with His191 are critical for dimer stability.
(b) The imidazolium cation can compete with cationic
side chains if a p–cation interaction is present at the
dimer interface. Recent analysis of dimeric interfaces
has shown that p–cation interactions can make significant contributions to the binding energy for protein–
protein complex formation, and it is suggested that
such interactions be included in the list of criteria for
characterizing protein interfaces [52]. We examined the

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1941


Dissociation of enolase into active monomers

I. Pal-Bhowmick et al.

enolase interface using PP server, which identified
Trp56 in Yen (Tyr56 in RMen, Tyr59 in Pfen) as an
interface residue. The program capture (Cation-p
Trends Using Realistic Electrostatics) [53] was used to
identify interactions between the cationic group of
lysine or arginine and the aromatic rings of phenylalanine, tyrosine and tryptophan. Such a search on Yen led
to the identification of a Trp–Arg (Fig. 8D) interaction
with a calculated electrostatic energy of Ees ẳ
) 16.3 kJặmol)1 and a Van der Waals energy of Evdw ẳ
) 7.5 kJặmol)1. In the modeled structures of RMen and
r-Pfen, similar interactions occur with Tyr56 and Tyr59,
respectively. The energy of interaction computed for
Tyr and Arg in the modeled structures of RMen (Ees ẳ
) 4.4 kJặmol)1 and Evdw ẳ ) 4.2 kJặmol)1) and r-Pfen
(Ees ẳ ) 8.8 kJặmol)1 and Evdw ẳ ) 3.8 kJặmol)1) were
found to be much smaller than that for Yen. Thus, this
particular interaction appeared to be much stronger in
Yen than in r-Pfen and RMen. This could be a possible
reason for the inability of imidazole to dissociate Yen,
unless 300 mm NaCl is included in the buffer (Table 1),
whereas RMen and r-Pfen become dissociated in the
absence of salt. An aromatic ring can have two possible
modes of interaction with imidazole [54], namely p–p
and NH–p interactions, whereas the imidazolium cation
has an additional possibility of a p–cation interaction
[55]. The observed greater effectiveness of the imidazolium ion in dimer dissociation may arise from its ability
to undergo p–cation and stronger NH–p interactions
with an aromatic residue (Trp or Tyr) at the interface.
The enolase interface has two salt bridges (Glu20–
Arg414 and Arg8–Glu417 in Yen; Glu20–Arg411 and

Arg8–Glu414 in RMen; Glu23–Arg415 and Arg11–
Glu425 in r-Pfen) involving two glutamate residues.
pH-induced dissociation of the enzyme may arise due to
protonation of the side chains of these glutamate
residues at low pH (pKa  4.5–5.5).
Although r-Pfen is highly homologous to mammalian enolases, we have observed an interesting difference between these two proteins; namely, imidazole
can dissociate r-Pfen in presence of Mg(II), whereas
RMen could not be dissociated. Thus, there is a possibility of selective dissociation of r-Pfen into a lowactivity, unstable monomeric form that can lead to
significant reduction in enolase activity. Such a partial
decrease in enolase activity combined with low stability
could severely hamper the glucose-metabolizing capacity of the parasite. As activity of glycolytic enzymes is
known to be essential for the growth of Plasmodium in
the intraerythrocytic stages [56], analogs of imidazole
(with higher pKa) that may be able to dissociate parasite enolase at physiologic pH and in the presence of
Mg(II) might prove to be parasite growth inhibitors.
1942

The comparison of various properties between the
dimeric and monomeric forms of enolase showed that
the monomeric form of enolase is catalytically inefficient (Kcat ⁄ Km is  5-fold less than that of the dimer)
and thermally unstable. It is highly unlikely that such
a catalytically inefficient form of enolase is utilized for
high-flux glycolytic function in the parasite. However,
in recent years, enolase has also been recognized as a
multifaceted protein with a variety of nonglycolytic
novel biological functions [18,57]. It is likely that,
when executing some of these moonlighting functions,
enolase may interact with other proteins. In such cases,
the monomeric form of enolase may be more suitable,
as it has a freely accessible subunit-interacting surface.

One such case is the 48 kDa s-crystallin protein in the
eye lens of the lamprey [58], which has been shown to
be a monomeric form of a-enolase [59]. As enolases
undergo several post-translational modifications inside
cells, it is likely that some of these may stabilize the
monomeric form of enolase to enable it to participate
in diverse physiologic functions.

Materials and methods
Materials
Hexahistidine-tagged r-Pfen was purified as described previously [24]. Yen was purchased from Sigma Chemical Co.
(St Louis, MO, USA). RMen and all other enzymes were
purchased from Boehringer-Ingelheim GmbH (Ingelheim,
Germany). Phosphoenolpyruvate was purchased from Sigma. Tris, imidazole and magnesium chloride were obtained
from USB (Amersham-Buchler, Braunschweig, Germany).
All other reagents used in this study were of analytical
grade and were used as received.

Gel filtration chromatography
Gel filtration chromatography was performed with a Superdex-75 prep grade column (Hiload HR 16 ⁄ 60, using
an AKTA FPLC system supplied by Amersham-Pharmacia
Biotech (Kwai Chung, Hong Kong). Before every run, the
column was pre-equilibrated with two column volumes of
desired buffer. The typical flow rate was 1.5 mLỈmin)1, and
the fraction volume was 2 mL. Blue dextran (2000 kDa),
b-amylase (200 kDa), alcohol dehydrogenase (150 kDa),
BSA (67 kDa), ovalbumin (43 kDa) and chymotrypsinogen
(25 kDa) were used as molecular weight calibration markers.

Enzyme activity assay

Enolase activity was measured by a continuous spectrophotometric assay by monitoring the formation of 2-PGA

FEBS Journal 274 (2007) 1932–1945 ª 2007 The Authors Journal compilation ª 2007 FEBS


I. Pal-Bhowmick et al.

from phosphoenolpyruvate at 240 nm using a Perkin-Elmer
(Waltham, MA, USA) lambda 40 spectrophotometer (T ¼
20 ± 1 °C). The assay mixture contained 1.5 mm phosphoenolpyruvate and 1.5 mm MgCl2 in 50 mm Tris ⁄ HCl
(pH 7.4). A unit of enzyme was defined as the amount of
enzyme that converts 1 lmol of phosphoenolpyruvate into
2-PGA in 1 min at 20 °C. Appropriate correction was
applied for differences in absorbance of phosphoenolpyruvate at 240 nm with varying pH and Mg(II) concentration
[60]. Kinetic data were fitted to the Michaelis–Menten
equation using sigmaplot.

Fluorescence and CD spectra
The fluorescence measurements were carried out by using a
SPEX Fluorolog FL1T11 spectrofluorometer (Edison, NJ,
USA) equipped with a thermostated cell holder. All fluorescence intensities were corrected for variable background
emissions and lamp fluctuations (signal ⁄ reference: s ⁄ r).
Fluorescence emission intensities were normalized for protein concentration (2–10 lm range). The excitation wavelength used was 295 nm, and the emission was measured in
the range 310–390 nm. The scan speed was 1 nmỈs)1, and
two scans were averaged for each spectrum.
CD studies were carried out using a Jasco J-810 spectropolarimeter (Tokyo, Japan). Far-UV CD studies were carried out using a 1 mm path-length cuvette with 2–10 lm
protein concentration at constant temperature of
20 ± 1 °C. Far-UV CD spectra were measured over the
200–270 nm range, and each spectrum was taken by averaging 10 scans at 20 nmỈs)1 scan speed. The data are presented as mean residue ellipticities and were calculated using
the formula:

ẵh ẳ hobs =10 ClÞ
where hobs is the observed ellipticity in mdeg, l is the path
length in cm, and C is the concentration of peptide bonds
in molỈL)1.

Acknowledgements
We would like to thank Mr Hardeep K. Vora for help
with some of the experiments. I. Pal-Bhowmick would
like to thank the Kanwal Rekhi Career Endowment
Fund for partial support. We gratefully acknowledge
Dr Dinkar Sahal for reading the manuscript and making helpful suggestions.

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