 |
INTRODUCTION |
Type II DNA topoisomerases catalyze the ATP-dependent
transport of one duplex DNA segment through a transient break in
another (for recent reviews, see Refs. 1-3). One essential function of this class of enzymes is to separate intertwined daughter chromosomes (4). Additionally, due to their ability to change DNA topology, these
enzymes exert a global influence on DNA metabolism (2). Clinically
these enzymes are of interest because they are targets of a diverse
group of anticancer and antibiotic drugs (5-7).
Type II topoisomerases catalyze two types of chemical reactions: ATP
hydrolysis and DNA breakage/religation. These are large homodimeric
enzymes with an ATPase domain at the amino terminus of each monomer.
Several types of experiments have indicated that when ATP binds, the
ATPase domains dimerize (8, 9). Hydrolysis is thought to occur by the
direct in-line attack of an activated water molecule on the
-phosphate of ATP (10). The steady-state ATPase rates of enzymes
purified from several organisms are DNA-stimulated (11-13).
Pre-steady-state ATPase experiments show a rapid burst in ATP
hydrolysis, but only when the topoisomerase is bound to DNA (14). These
results indicate that DNA binding stimulates the rate of ATP binding
and/or the rate of ATP hydrolysis. Analysis of pH rate profiles
suggests that binding of DNA primarily stimulates ATP binding (15).
Additionally, the pre-steady-state results, in conjunction with results
of inhibitor studies, show that although the topoisomerase binds two
ATPs, it hydrolyzes only one rapidly (15). Following the first
hydrolysis, the enzyme apparently releases the Pi and ADP
produced before hydrolyzing the second ATP.
The second chemical reaction, double strand DNA breakage, is catalyzed
by the nucleophilic attack from the hydroxyl groups of two active site
tyrosines, one in each monomer, on a staggered pair of phosphodiester
bonds in the DNA (reviewed in Ref. 2). This attack results in the
transient covalent attachment of each monomer to the 5'-end of one
strand of DNA. Religation of the DNA occurs by essentially the reverse
reaction in which the 3'-OH ends of DNA attack the pair of
phosphotyrosyl linkages. The equilibrium between DNA cleavage and
religation in a topoisomerase II-DNA complex normally lies strongly
toward religation (16). This equilibrium is somewhat perturbed by ATP
binding (17, 18) and is strongly perturbed by the presence of
anticancer drugs, such as etoposide, amsacrine, and mitoxantrone
(5-7).
These two chemical reactions, ATP hydrolysis and DNA
breakage/religation, can occur independently. However, in order to
understand how the enzyme can transport one segment of DNA (termed the
transported (T)1 segment)
through a protein-mediated gate in another segment (termed the gated
(G) segment) while using the energy available from ATP binding and
hydrolysis, it is essential to understand how these two reactions are
coordinated. In the present study, we begin to probe the interactions
between the ATPase and DNA breakage reactions. Without the hydroxyl
group of the active site tyrosine, the topoisomerase cannot cleave DNA
through covalent attachment. Without covalent attachment to the G
segment of DNA, the topoisomerase cannot undergo the conformational
changes normally associated with opening of the gate without first
breaking its noncovalent DNA binding interactions. To study how the
inability to cleave DNA and related disturbances in protein
conformational changes affect the ATPase reaction cycle, the active
site tyrosine, amino acid 782 in the S. cerevisiae
enzyme,2 has been mutated to
phenylalanine. The DNA-stimulated ATPase activity of this mutant is
shown to differ from the wild type activity. The difference in
steady-state ATPase rates for the mutant and wild type enzyme could be
due to differences in DNA binding, the rates of ATP
binding/dissociation, the rates of ATP hydrolysis/synthesis, or the
rates of ADP/Pi dissociation. To interpret how these
differences in ATPase rate may provide clues to the overall
topoisomerase II mechanism, it is necessary to understand precisely
which of these parameters is affected. DNA binding and pre-steady-state
ATPase experiments are described that help to define the differences
between the wild type and mutant enzymes. To ensure that this
perturbation in ATPase activity is solely due to a lack in DNA breakage
ability by this mutant enzyme, and not some unrelated consequence of
mutation, a second DNA cleavage mutant (19) is also studied.
 |
EXPERIMENTAL PROCEDURES |
Materials--
Standard reagents were purchased from the
following commercial resources: ATP, Amersham Pharmacia Biotech;
[
-32P]ATP (3000 Ci/mmol) and
[
-32P]dCTP (3000 Ci/mmol), NEN Life Science Products;
5'-adenylyl-
,
-imidodiphosphate, AMPPNP, and ultra-pure HEPES,
Boehringer Mannheim; NADH, phospho(enol)pyruvate (trisodium salt
hydrate), and pyruvate kinase (700 units/ml)/lactate dehydrogenase (100 units/ml) mixture from rabbit muscle, Sigma. All buffers were filtered
(0.45 µm). The sheared salmon sperm DNA used for ATPase assays was
prepared as described previously (14).
Expression and Purification of Saccharomyces cerevisiae
Topoisomerase II--
A new method of topoisomerase II purification
based on the IMPACT system (New England Biolabs) was used for these
studies. The expression plasmid used for the wild type enzyme, pJEL236, is essentially YEpTOP2-PGAL1 (20) with the insertion of the sequences
coding for the modified intein and chitin binding domains (intein/CBD)
from pCYB2 (New England Biolabs) just prior to the stop codon. This
vector expresses a fusion of yeast topoisomerase II with the intein/CBD
that allows for rapid purification. The constructs were made using
polymerase chain reactions that replaced the final aspartic acid codon
and stop codon of the TOP2 gene with a unique blunt cutting
Ecl136II restriction site that could be ligated to a SmaI site at the
5'-end of the intein/CBD coding sequences. The construct was designed
such that the only difference between wild type S. cerevisiae topoisomerase II and the final purified wild type
topoisomerase is that the final aspartic acid has been changed to
glycine. The purified protein with this one change has identical
biochemical characteristics to the fully wild type enzyme, and for the
purposes of these studies, it will be referred to as wild type
topoisomerase II.
The expression plasmid (pSKM1) for the mutant in which the active site
tyrosine has been changed to phenylalanine, Y782F, was made from
plasmid YEpTOP2-PGAL1-Y782F (a kind gift of Brian Davis, United States
Department of Agriculture). The topoisomerase II coding sequence was
verified by sequencing, which showed that the only alteration was a
change from TAT to TTT at codon 782. The intein/CBD coding sequences
were then inserted into this plasmid to make pSKM1 as described above
for the wild type expression plasmid. The expression plasmid for the
mutant R690A fused to the intein/CBD was derived from the plasmid
pSW201-R690A (a kind gift of Qiyong Liu and James C. Wang, Harvard
University). pSW201-R690A was cut with KpnI and
AvrII, and the fragment encoding the mutated portion of
topoisomerase II was swapped with the similar fragment from
YEpTOP2-PGAL1 to make pSKM2. The final expression plasmid (pSKM3) was
made by inserting the intein/CBD sequences as above. Both of the final
purified mutant proteins, Y782F and R690A, also have the final aspartic
acid replaced by glycine.
The enzyme-intein/CBD fusions were expressed to very high levels from
the plasmids pJEL236, pSKM1, and pSKM3 in the yeast strain BCY123 as
described previously for the wild type enzyme (21). The basic buffer
used throughout purification, Buffer I, contained 50 mM
HEPES-KOH, pH 7.5, 1 mM EDTA, 1 mM EGTA, and 10% glycerol. The cells (36 g) were washed and cracked as described previously (22). The cracked cells were diluted to 200 ml with load
buffer (Buffer I + 500 mM KCl, 1 mM
phenylmethylsulfonyl fluoride, 1 µg/ml leupeptin, 1 µg/ml
pepstatin, and 150 µg/ml benzamidine) and centrifuged for 30 min at
45,000 × g. The supernatant was loaded onto a 50-ml (5 cm in diameter) chitin bead (New England Biolabs) column by gravity.
The column was washed with 5 volumes of load buffer at 2 ml/min
followed by 10 volumes of wash buffer (Buffer I + 1 M KCl + 0.1% Triton X-100) at 3 ml/min. Cleavage of the intein/CBD from
topoisomerase II was induced by washing the column with four volumes of
Buffer I + 500 mM KCl + 30 mM dithiothreitol.
The released topoisomerase II was eluted 12 h later. The eluant
was diluted 2× in Buffer I prior to loading onto a phosphocellulose
column equilibrated with Buffer I + 150 mM KCl. The size of
the phosphocellulose column was determined by the amount of protein
eluted from the chitin column; 1 ml of phosphocellulose was used for
each 3 mg of protein. The column was washed with three volumes of
Buffer I + 300 mM KCl, and the topoisomerase II was eluted
with a minimal volume of Buffer I + 1 M KCl. The peak
fractions were combined, frozen in liquid nitrogen, and stored at
70 °C. For DNA binding studies, the protein was dialyzed
extensively in Buffer I + 250 mM KCl prior to freezing. This method provided 45 mg of highly purified wild type or 15 mg of
mutant topoisomerase II from 12 liters of cell growth. These large
quantities of protein were required because 15-20 mg of enzyme is
needed for each pre-steady-state ATPase assay.
Steady-state ATPase Assays--
A coupled assay using pyruvate
kinase, phospho(enol)pyruvate, lactate dehydrogenase, and NADH was used
as described previously (13). Reactions were performed at 30 °C in
ATPase reaction buffer (50 mM HEPES-KOH (pH 7.5), 150 mM KOAc, and 10 mM Mg(OAc)2) at 10 different ATP concentrations (25 µM to 1 mM).
Purified and sheared salmon sperm DNA was used at a ratio of 200 bp/enzyme dimer; this DNA was shown to stimulate the ATPase identically to supercoiled plasmid DNA (14). Topoisomerase II binds ATP with
positive cooperativity, as revealed by Michaelis-Menten plots with
slight sigmoidal character at the lowest ATP concentrations. However,
in the present analysis, this cooperativity was ignored and the data
were fit to the standard Michaelis-Menten equation as follows using
KaleidaGraph 3.0.
|
(Eq. 1)
|
DNA Binding Assays--
The binding affinities of wild type and
Y782F topoisomerase II for DNA were measured using a double-filter
method developed by Wong and Lohman (23). The DNA used was a
32P-labeled 46-mer duplex, similar in sequence to
oligonucleotides used previously for DNA binding and cleavage studies
with type II topoisomerases isolated from various organisms (24, 25). The substrate was made by annealing complementary 43-mer
oligonucleotides (5'-GGGTGAAATCTAACAATGCGCTCATCGTCATCCTCGGCACCGT
and 5'-GGGACGGTGCCGAGGATGACGATGAGCGCATTGTTAGATTTCA) such that the
duplex had a 3-guanine overhang on each 5'-end. Equal concentrations of
the oligonucleotides (final concentration, 41 µM) were
mixed in 10 mM Tris-HCl, pH 7.5, 1 mM EDTA, 20 mM LiCl2, heated to 96 °C, and allowed to
cool to room temperature over 4 h. The overhangs were filled in
with Klenow fragment, 100 µCi of [
-32P]dCTP (3000 Ci/mmol), and 200 µM each of dTTP, dGTP, and dATP. After
15 min, the reaction was chased with 200 µM unlabeled
dCTP for 1 min. The substrate was purified away from free nucleotides using a Chroma spin TE-10 column (CLONTECH).
Following phenol:chloroform extraction and ethanol precipitation, the
substrate was resuspended in 10 mM Tris-HCl, 1 mM EDTA, and its concentration was determined by absorbance
at 260 nM (
260 = 6.16 × 105 M
1 cm
1).
The double-filter method employed uses a nitrocellulose membrane to
trap labeled DNA bound to protein on top of a DEAE membrane that traps
free DNA (23). This method allows one to measure both protein-bound DNA
and free DNA counts, improving precision and accuracy over conventional
binding methods. We used the same membranes, prepared in the same
fashion, and a similarly altered dot blot apparatus, as described
previously (23). Topoisomerase II and the labeled DNA substrate were
mixed at room temperature such that the final DNA concentration was 2 nM and the final enzyme dimer concentration ranged from 0.5 to 400 nM. The reactions performed in the absence of
Mg2+ also contained 85 mM KCl, 25 mM HEPES-KOH (pH 7.5), 5-7% glycerol, and 250 µg/ml
bovine serum albumin. The reactions that contained 5 mM
MgCl2 were similar except that the KCl concentration was lowered to 45-60 mM to maintain a similar overall ionic
strength between experiments. Using the optimal ATPase reaction buffer including 150 mM KCl, the maximum percentage of DNA bound
was only 10-20% (not shown). The concentrations of KCl were therefore dropped to those listed above to achieve binding near 100% for accurate determination of Kd values. The slight
range in glycerol and KCl concentrations was inevitable due to the
different amounts of enzyme added to the binding reactions. The binding reactions were allowed to equilibrate for 10-30 min at room
temperature prior to filtration through the nitrocellulose and DEAE
membranes at 6 ml/min. Aliquots (25 µl) of each reaction were
filtered in triplicate and rinsed with 150 µl of cold binding buffer.
DNA counts on the dried membranes were determined at each spot by phosphorimage analysis (Molecular Dynamics PhosphorImager TM model 400 with ImageQuant (version 3.3) software). Control reactions lacking
topoisomerase were also filtered as described to determine the
nonspecific DNA counts trapped by the nitrocellulose. For each binding
reaction, the ratio of bound to total DNA was determined as indicated
below and plotted versus the topoisomerase II
concentration.
|
(Eq. 2)
|
The data were fit to the binding mechanism,
|
(Eq. 3)
|
where T represents the topoisomerase II dimer, D represents the
duplex DNA oligomer, and Kd represents the
equilibrium dissociation constant. The values for Kd
were determined by fitting the data to the equation describing this
mechanism,
|
(Eq. 4)
|
where C is a constant that approaches 1 as the
asymptote approaches 1, using KaleidaGraph 3.0.
Trapping of Circular DNA Analyzed by CsCl Density Gradient
Ultracentrifugation--
The technique used is essentially that
described previously (26). Reactions of 30 µl containing 150 nM topoisomerase II dimer, 250 nM relaxed or
linearized pBluescript plasmid DNA (3 kilobase pairs), 50 mM Tris-HCl, pH 7.5, 80 mM KOAc, and 8 mM Mg(OAc)2 were incubated for 10 min at
30 °C. AMPPNP was added to a final concentration of 1 mM, and incubation was continued an additional 30 min. To
an aliquot of 20 µl of each reaction, 130 µl of buffer (50 mM Tris-HCl, pH 7.5, 80 mM KOAc, and 8 mM Mg(OAc)2) and 334 µl of saturated CsCl
were added. These samples were spun at 40,000 rpm in an analytical
ultracentrifuge (XL-A ultracentrifuge, Beckman Instruments) with scans
at 260 and 280 nm taken at 36 and 40 h. Data plots were made using
Microcal Origin 3.78.
Pre-steady-state ATPase Assays--
Rapid chemical quench and
pulse-chase experiments were performed as described previously (14)
using the KinTek Model RQF-3 rapid quench apparatus (27). Briefly,
topoisomerase II bound to DNA was rapidly mixed with
[
-32P]ATP (0.01 µCi/µl) in ATPase reaction buffer.
The final topoisomerase II dimer concentration in the reactions ranged
from 5.5 to 7.5 µM (see legends to Figs. 3 and 4),
whereas the DNA concentration was kept at a constant ratio of 200:1
base pairs:enzyme dimer. The final labeled ATP concentration was
typically 350 µM, a concentration previously shown to
essentially saturate the ATP active sites while still providing a good
signal to noise ratio (14). The enzyme and substrate were allowed to
react for 20 ms to 1.5 s. For chemical quench experiments, the
reaction was rapidly quenched by addition of 250 mM EDTA in
100 mM Tris base (pH 10) at the indicated time points. SDS
(final concentration, 1%) was present in the sample collection tubes
to ensure that no residual enzyme activity persisted. For pulse-chase
experiments, 10.6 mM unlabeled ATP was mixed with the
reaction at the indicated time points. After sufficient time for
multiple enzyme turnovers, 1.5 s for the wild type enzyme and
3 s for the mutant enzymes, the reactions were chemically quenched
as indicated above. The concentrations of [
-32P]ADP
produced at each time point were determined in quintuplicate as
described previously (14). The data were all fit to a single exponential equation with a linear term called the burst equation, A(1
e
Bt) + Ct, using
KaleidaGraph 3.0. The value A is the burst amplitude, B is the burst rate constant, and C is the
steady-state term.
 |
RESULTS |
Steady-state ATPase Analysis--
As a first step to understanding
the relationship between ATP hydrolysis and DNA cleavage by
topoisomerase II, steady-state ATPase parameters were determined for
wild type and Y782F enzymes. This mutant cannot cleave DNA through
covalent attachment because it lacks the active site hydroxyl
nucleophiles that normally would attack a pair of staggered 5'
phosphates. The results of the steady-state ATPase assays are given in
Table I. Although the results shown are
from a single set of experiments, the same relative values have been
found a minimum of four times using four different sets of protein
preparations. In the absence of DNA, kcat and
kcat/Km values are identical for
the wild type and mutant enzymes. This is expected if the mutation only affects interactions with DNA. It has been previously shown for several
type II topoisomerases that DNA stimulates the steady-state ATPase
rates (11-13). The present study is in agreement and shows that when
either the wild type or mutant enzyme is bound to DNA, kcat and
kcat/Km values increase. Whereas kcat/Km values increase to the
same extent for the two proteins, the kcat value for
Y782F is reproducibly 4-fold lower than that for wild type. Results of
previous studies indicate that
kcat/Km for the mechanism of ATP hydrolysis by topoisomerase II essentially includes only rate constants
involved in ATP binding, whereas kcat reflects the
rate-determining steps (15). These present data are consistent with the
Y782F mutant enzyme-DNA complex binding ATP in a similar fashion to
wild type-DNA complex. However, why is the kcat of
ATP hydrolysis lower for a mutant that cannot covalently cleave DNA?
The following experiments were performed to address this question.
DNA Binding--
Because topoisomerase II is a DNA-stimulated
ATPase, any perturbation in DNA binding could affect the ATPase
activity of the enzyme. Therefore, to determine whether the decreased
kcat value for the Y782F protein is due to altered
noncovalent interactions with DNA, DNA binding studies were performed.
A double-filter binding method (23) was used to measure the affinity of
wild type and Y782F enzymes for a 46-bp duplex oligonucleotide
substrate. Wild type topoisomerase II requires a divalent cation
for covalent attachment to DNA. To ensure that only noncovalent
interactions were being measured in the first binding experiment,
reaction and wash buffers lacked Mg2+. The results of these
binding studies are shown in Fig.
1A. The data fit well to an
equation describing a single DNA molecule binding per enzyme dimer,
with Kd values of 29 nM for both wild
type and Y782F mutant enzymes.

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Fig. 1.
Affinity of wild type and Y782F
topoisomerase II for DNA as measured by double-filter nitrocellulose
binding. The ratios of bound DNA to total DNA were determined in
the absence (A) and presence (B) of 5 mM MgCl2 in reactions including varying
concentrations of wild type ( ) or Y782F ( ) topoisomerase II, as
described under "Experimental Procedures." The data were fit to Eq. 4, which resulted in identical Kd values for the
wild type and Y782F mutant of 29 and 11 nM in A
and B, respectively. The error bars represent the
deviation from mean of data collected in triplicate.
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The presence of Mg2+ not only allows the wild type enzyme
to covalently cleave DNA but may affect important noncovalent
interactions as well. Therefore, the binding experiments were
repeated in the presence of 5 mM MgCl2
(Fig. 1B). In the presence of Mg2+, both enzymes
bound DNA more tightly than in its absence, with Kd
values of 11 nM for both wild type and Y782F. Again, the
enzymes bound DNA with equal affinities, even though under these
conditions, the wild type can covalently attach to the DNA. These
results are in agreement with many previous results indicating that the
DNA cleavage-religation equilibrium for topoisomerase II strongly
favors religation (16).
Topologic Trapping of Circular DNA--
Topoisomerase II can
topologically trap circular DNA if it subsequently binds a
nonhydrolyzable ATP analog (18, 28, 29). If a high concentration of DNA
circles is used in such an experiment, the wild type enzyme can
catenate two circles; the enzyme remains directly topologically linked
to one circle, which is catenated to the other circle (28). Given that
the buoyant densities of free topoisomerase II and
Escherichia coli plasmid DNA are approximately 1.37 g/ml
(21) and 1.71 g/ml (30), respectively, complexes between the two will
have intermediate densities depending on the ratios of DNA to enzyme.
This property has been used previously to analyze topoisomerase
II-circular DNA complexes by equilibrium sedimentation in a CsCl
gradient (26, 31). The results of similar experiments performed with
wild type and Y782F topoisomerase II and a molar excess of 3 kilobase
pairs relaxed circular or linearized DNA are shown in Fig.
2. In each absorbance trace, free DNA is
seen as a large peak at the bottom of the density gradient. Two peaks
of intermediate density are seen for reactions with wild type enzyme
bound to circular DNA and AMPPNP (Fig. 2A). When the binding
reactions were analyzed by agarose gel electrophoresis, a band
migrating at the position expected for catenated circles was detected
for only the wild type reaction (not shown). The most reasonable
interpretation of these data is that the intermediate peak of higher
density represents the enzyme linked to a catenane it has produced. The
lower density peak indicates the enzyme linked to a single DNA circle.
Because the DNA:topoisomerase ratio is high in these experiments, no
peaks caused by multiple topoisomerase dimers per DNA circle are seen.
In contrast, the reaction with the Y782F mutant produced only the lower
density intermediate peak (Fig. 2D). Because this mutant
cannot perform a catenation reaction, it is not surprising that the
higher density intermediate is missing. However, these experiments do
confirm that a mutant unable to transport one DNA through another can
still only topologically trap one DNA circle (3). Even under conditions
where the wild type enzyme can catenate two circles, Y782F apparently
cannot trap both the G and T segments of DNA.

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Fig. 2.
AMPPNP-induced trapping of DNA circles by
wild type and Y782F topoisomerase II observed by analytical
ultracentrifugation. Absorbance traces of the CsCl density
gradients taken at 40 h and 260 nm are shown for each reaction.
The large peak seen at the right end of each
trace is from the absorbance of free DNA running at the bottom of each
gradient.
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Control reactions were performed to ensure that the intermediate peaks
that were seen result from the topologic trapping of circular DNA by
topoisomerase II forming an AMPPNP-induced clamp around the DNA. When
linear DNA was used instead of circular, no peaks of intermediate
density were seen (Fig. 2, B and E). Additionally, in the absence of AMPPNP, no intermediate peaks were
detected (Fig. 2, C and F).
Pre-steady-state ATPase Assays--
Because the difference in
kcat values for ATP hydrolysis by the wild type and
Y782F mutant enzymes is apparently not due to differences in DNA
affinities, pre-steady-state chemical quench and pulse-chase
experiments were used to further explore the differences. In the
pre-steady state, information about the rates of individual steps in
the reaction pathway can be obtained; this information is obscured in
steady-state kinetics. The ATPase activity of wild type topoisomerase
II has been previously studied extensively using these rapid quench
methods (14, 15). New results of chemical quench and pulse-chase time
courses for the wild type enzyme bound to DNA are shown in Fig.
3A. The results of these
present time courses are essentially the same as those reported
previously. The chemical quench time course shows a rapid burst of ATP
hydrolysis, followed by the steady-state rate of enzyme turnover. The
burst amplitude (6.0 ± 0.6 µM) approximately equals
the enzyme dimer concentration (6.7 µM), or half the
concentration of ATP active sites.

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Fig. 3.
Pre-steady-state ATPase analysis of wild type
and Y782F topoisomerase II. A, chemical quench ( )
and pulse-chase ( ) time courses were performed with wild type
topoisomerase II at final concentrations of 6.7 µM enzyme
dimer, 1.6 mM DNA bp, and 350 µM ATP. The
data were fit as described under "Experimental Procedures" with
values for the chemical quench reaction of A = 6.0 ± 0.6 µM, B = 25 ± 5 s 1, and C = 14 ± 1 µM
s 1, and for the pulse-chase reaction of A = 14 ± 1 µM, B = 50 ± 10 s 1, and C = 14 ± 1 µM
s 1. B, similar chemical quench ( ) and
pulse-chase ( ) time courses were performed with Y782F at final
concentrations of 7.5 µM enzyme dimer, 1.8 mM
DNA bp, and 350 µM ATP. The parameters obtained by
fitting the data are A = 8 ± 1 µM,
B = 6 ± 1 s 1, and C = 6 ± 1 µM s 1 and A = 15 ± 1 µM, B = 50 ± 10 s 1, and C = 6 ± 1 µM
s 1 for the chemical quench and pulse-chase reactions,
respectively.
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Fig. 3A also shows the results of a pulse-chase experiment
performed with the same enzyme-DNA sample. If instead of chemically quenching the ATPase reaction, a large excess of unlabeled ATP is added
at the indicated time points, the enzyme can either continue to
hydrolyze any labeled ATP that it had bound prior to the chase or
release the unhydrolyzed ATP. Any new ATP bound will be unlabeled and
therefore undetectable in the assay system. The reaction is allowed to
continue for several turnovers before it is chemically quenched. A
burst in ADP production is again seen in the pulse-chase results, but
this time the burst amplitude (14 ± 1 µM)
approximately equals the enzyme active site concentration (13.4 µM). This experiment shows that at 350 µM
ATP, the enzyme active sites are essentially saturated with ATP, and
once the two ATP are bound, they are hydrolyzed faster than they are
released. In conjunction with inhibitor studies, these results showing
a doubling of the burst amplitude from one half the ATP active site
concentration in chemical quench experiments to essentially equal the
ATP active site concentration in pulse-chase experiments have been used
to define the ATPase reaction mechanism for the DNA-bound topoisomerase
II (15). The simplest mechanism in agreement with all of these data is
that the enzyme binds two ATP, hydrolyzes one rapidly, releases the
products of the first hydrolysis, and then hydrolyzes the second ATP
and releases those products. The clearly defined burst in hydrolysis of
one ATP shows that a rate-determining step occurs after hydrolysis of
the first ATP and before hydrolysis of the second ATP. A reaction
mechanism describing this pathway is shown in Scheme 1, where
E2 represents dimeric topoisomerase II bound
to DNA, S represents ATP, P represents ADP·Pi, and E2S*
represents a state of topoisomerase II bound to a single ATP that
differs from E2S.
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Similar chemical quench and pulse-chase reaction time courses were
performed with the Y782F mutant enzyme (Fig. 3B). The results of the chemical quench experiment again show a burst in ATP
hydrolysis, with the burst amplitude (8 ± 1 µM)
approximating the dimeric enzyme concentration (7.5 µM).
As expected, the calculated steady-state rate is also slower for Y782F
(6 µM s
1) as compared with the wild type
(14 µM s
1). The same chemical quench
results were obtained when the ATP concentration was increased to 1 mM, confirming that the low value for the burst rate
constant is not a result of subsaturation of the ATP active sites at
350 µM ATP (data not shown). The pulse-chase results show
a burst amplitude (15 ± 1 µM) equal to the enzyme active site concentration (15 µM) and a burst rate
constant equal to that seen for the wild type enzyme (50 ± 10 s
1). A comparison of the chemical quench and pulse-chase
results indicates that Y782F binds two ATP with a high commitment for catalysis, just as the wild type enzyme. If Y782F had bound ATP at
slower rates or lower affinities than the wild type enzyme, the
pulse-chase burst rate constant and amplitude would have been reduced.
This is not seen. Additionally, the identical
kcat/Km values support the
conclusion that there is no significant decrease in DNA-stimulated ATP
binding by Y782F compared with wild type topoisomerase II.
In order to determine which rate constants are decreased for the mutant
enzyme, further analysis of the data is required. For the wild type
enzyme, the clear distinction between the burst and steady-state phases
of the data allowed the use of singular perturbation theory to
determine rate constants for each step of the pathway shown in Scheme 1 (15). This distinction no longer exists for the data obtained with
Y782F. Therefore the data were analyzed using a simplified reaction
pathway shown in Scheme 2, where ka' = ka[S].
Using this simplified reaction pathway, the values determined by
fitting the rapid quench data to the burst equation can be directly
related to rate constants (32). Under conditions where the enzyme is
saturated with ATP, ka' values
determined from chemical quench data will reflect the rate of
hydrolysis of the first ATP by topoisomerase II (15). For this
simplified mechanism, the following equation is used (32).
|
(Eq. 5)
|
The calculated values for ka' are 25 ± 7 and 7 ± 2 s
1 for the wild type and Y782F
enzymes, respectively. Therefore, this mutant hydrolyzes the first ATP
3-4-fold more slowly than the wild type enzyme. Although the rate of
hydrolysis of the first ATP is slower for the mutant, it is not
rate-determining, as evidenced by the burst seen in the chemical quench
data. The decrease in the steady-state rate seen for the mutant
indicates that at least one step beyond the first hydrolysis must also
be occurring severalfold more slowly for the mutant enzyme. This slower
step could be release of ADP and/or Pi or the rate of
hydrolysis of the second ATP. It is presently not possible to
distinguish these possibilities.
In order to demonstrate that the inability to cleave DNA is the
property of the Y782F mutant that causes altered ATPase activity, and
not an unrelated effect of the mutation, a second system was sought for
comparison. Unfortunately, there are no known DNA analogs that bind
topoisomerase II identically to DNA but cannot be cleaved. However,
there is another amino acid known to be required for covalent cleavage
by topoisomerase II, Arg690 (19). The mutant R690A was
found to have steady-state ATPase parameters identical to the wild type
enzyme in the absence of DNA, but altered in a way similar to Y782F in
the presence of DNA (see Table I). Results of pre-steady-state chemical
quench and pulse-chase time courses of R690A bound to DNA are shown in Fig. 4. These results are very similar to
those found with Y782F. The value of ka'
calculated from the R690A chemical quench data is 8 ± 3 s
1, again reduced 3-fold from the wild type value. As
with Y782F, R690A binds two ATP in a similar fashion to wild type, but
hydrolysis of the first ATP occurs at a slower rate for the mutant
enzyme. The fact that both DNA cleavage mutants have similar ATPase
kinetics indicates that it is the inability to cleave DNA that causes a perturbation in ATP hydrolysis by topoisomerase II.

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Fig. 4.
Pre-steady-state ATPase analysis of R690A
topoisomerase II. Chemical quench ( ) and pulse-chase ( ) time
courses were performed with R690A mutant topoisomerase II at final
concentrations of 5.5 µM enzyme dimer, 1.5 mM
DNA bp, and 350 µM ATP. The data were fit as described
under "Experimental Procedures" with values for the chemical quench
reaction of A = 4.0 ± 0.6 µM,
B = 9 ± 3 s 1, and C = 10 ± 1 µM s 1, and for the
pulse-chase reaction of A = 10 ± 1 µM, B = 60 ± 15 s 1,
and C = 9 ± 1 µM
s 1.
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DISCUSSION |
Topoisomerase II mutants that cannot covalently cleave DNA retain
a DNA-stimulated ATPase activity but hydrolyze ATP with a slower
maximum turnover rate than the wild type enzyme. Wild type and Y782F
mutant proteins bind a single segment of DNA with equal affinities.
Y782F can topologically trap a single circle of DNA in the presence of
AMPPNP under conditions where the wild type protein can catenate two
circles. Pre-steady-state ATPase kinetics show that the Y782F and R690A
mutants bind two ATPs with a high commitment for catalysis and without
a significant decrease in ATP affinity or binding rates. However, the
noncleaving mutants hydrolyze the first ATP 3-4-fold slower than the
wild type enzyme. A step or steps later in the reaction pathway,
potentially steps associated with ADP or Pi release or
hydrolysis of the second ATP, also occur more slowly for these mutant enzymes.
There are several potential mechanistic implications of these results.
The two mutations studied, Y782F and R690A, are not in the ATPase
domain, and it is unlikely that these residues come within ~20 Å of
the ATP. Therefore, it is doubtful that these mutations directly affect
the rate of the chemical step of ATP hydrolysis. One possibility is
that the ATP hydrolysis rates of these mutant proteins are perturbed
because they cannot undergo the normal pathway of conformational
changes. In order to allow transport of the T segment of DNA through
the G segment, the active site tyrosines and attached 5' phosphoryl
ends of the G segment must move 35-40 Å relative to each other (33).
However, the DNA must be cleaved first for the enzyme-DNA complex to
undergo this conformational change. Neither Y782F nor R690A is capable of cleaving DNA and cannot undergo this conformational change unless
their noncovalent interactions with DNA are broken. Because the rates
and affinities of ATP binding are not appreciably lower for the mutant
proteins, this conformational change is probably not coupled to the
steps of ATP binding. However, the rate of hydrolysis of the first ATP
is measurably reduced in the mutants. This suggests that either a DNA
cleavage-dependent conformational change occurs after ATP
binding but before hydrolysis or that hydrolysis is directly coupled to
the conformational change. The rates of product release, or
conformational changes associated with product release, are also
decreased severalfold for these mutant proteins.
An alternative hypothesis involves altered binding of the T segment of
DNA to the mutant proteins. When wild type topoisomerase II binds
circular DNA, followed by AMPPNP, it can catenate two circles and
remain directly topologically linked to one of them (28). The circle
that remains directly linked to the enzyme presumably contains the G
segment cleaved during the reaction. The other circle in the catenane
is not directly linked to the topoisomerase and contains the T segment
that was transported through the cleaved DNA and the enzyme. It is
apparently the G segment that binds to the topoisomerase with
sufficient affinity to be measured by the double-filter binding assay
described above. Interactions between the T segment and the enzyme are
not well understood. It has been postulated that the T segment binds
within the cavity formed by the ATP-induced dimerization of the ATPase domains (34). The lack of topological trapping of the T segment to the
wild type protein is probably due to its efficient transport through
the G segment and out of the protein. A DNA cleavage mutant cannot
transport a T segment because it cannot open the gate in the G segment.
We find that under conditions where the wild type enzyme can clearly
interact with two circles and catenate them, Y782F can topologically
trap only one circle. Similar conclusions have been reported elsewhere
as unpublished results (3). These results are interpreted as showing
that Y782F binds the G segment of DNA normally but fails to
productively bind the T segment. Results of previous kinetic studies
suggested that DNA stimulates the ATPase activity of topoisomerase II
by increasing the rates or affinities of ATP binding and potentially
the rate of hydrolyzing the first ATP (15). Y782F binds ATP normally
but hydrolyzes at least the first ATP slowly. Therefore, a second
simplified hypothesis to explain these results is that binding of the G
segment of DNA primarily stimulates ATP binding, whereas binding of the T segment stimulates hydrolysis of at least the first ATP. These two
proposed hypotheses are not mutually exclusive; the topoisomerase may
need to undergo the conformational change to open the G segment in
order to accommodate productive binding of the T segment. Either the
conformational change or the T segment binding may be the direct cause
of increased ATP hydrolysis.
A double-filter binding method has been used to quantitatively
determine the binding parameters of wild type and Y782F mutant topoisomerase II to DNA. In the absence of Mg2+, when
neither enzyme can covalently attach to the DNA, they have identical
Kd values of 29 nM. In the presence of 5 mM Mg2+, the Kd value for
each protein decreases to 11 nM. The ionic strengths were
kept constant between the plus and minus Mg2+ reactions, so
that the 2-3-fold decrease in Kd values should be
specifically due to the presence of the divalent cation. Previous
binding studies with type II topoisomerases and DNA have provided
varied results on the effects of Mg2+. Tight binding of
E. coli DNA gyrase (35) and yeast topoisomerase II (36) were
only detected in the presence of Mg2+, whereas binding of
the Drosophila enzyme was minimally affected by
Mg2+ (37, 38). Because noncovalent and potential covalent
interactions can be clearly distinguished in the present study with the
Y782F mutant, these data can unambiguously be interpreted to show that Mg2+ increases the noncovalent binding affinity of yeast
topoisomerase II 2-3-fold for DNA. Additionally, because under both
conditions the wild type and mutant have identical dissociation
constants, only a negligible fraction of wild type topoisomerase II is
covalently attached to DNA at equilibrium in the presence of
Mg2+. These results are in agreement with quenched DNA
cleavage results reported from many laboratories (see Ref. 16 for a
review). One potential concern with the cleavage reactions is that the quench solutions used could drive the enzyme-DNA cleavage-religation equilibrium toward religation. The present equilibrium binding studies
do not use a quench and support the idea that the true DNA
cleavage-religation equilibrium strongly favors religation.
Previous studies to determine the effects of perturbing DNA cleavage
and religation on the ATPase activity of topoisomerase II have produced
a variety of results. Preliminary data for the yeast Y782F mutant
enzyme (referred to in a previous study as Y783F in accordance with the
older sequence information) have been described as showing that this
mutant has no DNA-stimulated ATPase activity (39). In the present
results, the kcat for ATP hydrolysis of this mutant
is shown to increase 2-3-fold in the presence of DNA, whereas the
Km decreases almost 10-fold. Depending on how the
previous experiments were done, the rather subtle effect on
kcat might have been missed. One result of the DNA
stimulation on the ATPase activity is an easily detectable burst in
hydrolysis equal to half the ATP active site concentration; no burst is
seen in the absence of DNA (14). Both the Y782F and R690A cleavage
mutants show a similar burst in ATPase activity when bound to DNA,
indicative of DNA stimulating the rates of ATP binding and/or
hydrolysis of the first ATP. A variety of drugs perturb the DNA
cleavage/religation equilibrium catalyzed by topoisomerase II. Fortune
and Osheroff (40) found that merbarone blocks DNA cleavage yet has no
effect on the ATPase activity of human topoisomerase II
. It is
presently unclear why a drug that prevents DNA cleavage would have such
a different effect on the ATPase activity than the DNA cleavage
mutants. Etoposide, a drug that is thought to slow the rate of DNA
religation by topoisomerase II, was shown to have essentially no effect
on the ATPase activity of Drosophila topoisomerase II (41),
whereas it dramatically inhibited the ATPase activity of human
topoisomerase II
(39). The mechanistic implications of the
differences in these results are presently unclear. It should be
interesting to determine the pre-steady-state effects of these drugs to
more clearly define where they perturb the mechanistic pathway of DNA
topoisomerase II.