(Received for publication, February 19, 1997)
From the Department of Molecular Biophysics and
Biochemistry and ¶ Genetics, Yale University, New Haven,
Connecticut 06510 and § Groupe d'Etude
Mutagénèse et Cancerogenèse, Unite Mixte de Recherche
216 CNRS, Institut Curie, Bâtiment 110, Université Paris
Sud, F-91405 Orsay, France
RecA is a 38-kDa protein from Escherichia coli that polymerizes on single-stranded DNA, forming a nucleoprotein filament that pairs with homologous duplex DNA and carries out strand exchange in vitro. In this study, we measured RecA-catalyzed pairing and strand exchange in solution by energy transfer between fluorescent dyes on the ends of deoxyribo-oligonucleotides. By varying the position of the dyes in separate assays, we were able to detect the pairing of single-stranded RecA filament with duplex DNA as an increase in energy transfer, and strand displacement as a decrease in energy transfer. With these assays, the kinetics of pairing and strand displacement were studied by stopped-flow spectrofluorometry. The data revealed a rapid, second order, reversible pairing step that was followed by a slower, reversible, first order strand exchange step. These data indicate that an initial unstable intermediate exists which can readily return to reactants, and that a further, rate-limiting step (or steps) is required to effect or complete strand exchange.
RecA is a 38-kilodalton protein from Escherichia coli,
which has been shown to be necessary for conjugal homologous
recombination in vivo (1). In vitro, RecA protein
polymerizes on single-stranded DNA in the 5 to 3
direction to form a
right-handed helical structure in which the DNA is extended to 1.5 times its original length (2, 3). Pairing with homologous duplex DNA
results in a rapid uptake of the double-stranded DNA into a
three-stranded complex, which can be kilobases in length (4-6). Strand
exchange results in displacement of the strand of duplex DNA that has
the same sequence as the filament strand; the strand is displaced in
the 5
to 3
direction (7-9). Homologs of RecA exist in eukaryotes from yeast to man and have been found to hydrolyze ATP, to form nucleoprotein filaments, to pair homologous DNA, and to carry out
strand exchange in ways that are qualitatively similar to RecA protein
(10-15). Studies of the mechanisms of E. coli RecA may help
to shed light on eukaryotic as well as prokaryotic recombination.
Most studies on the kinetics of RecA-catalyzed strand exchange have
used phage DNA that is several kilobases in length (16-20). RecA
filaments formed on long single-stranded DNA generate coaggregates with
duplex DNA that concentrate the DNA but also limit diffusion (21).
Despite this complication, joint molecule formation displayed the
saturation of rates with increasing substrate concentration that is
typical of Michaelis-Menten kinetics (17). This observation suggested
the existence of a reversible pairing step and a second, rate-limiting
step in the reaction. Yancey-Wrona and Camerini-Otero (22) developed a
solution assay for pairing and stable synapsis of a single-stranded
oligonucleotide with duplex DNA in which a RecA filament formed in the
presence of ATPS1 protects a restriction
site in the duplex target molecule. With this assay, they found that
pairing was second order, reversible, and independent of the complexity
of the target. They were also able to determine the equilibrium
constant for pairing (22).
Several studies have used fluorescence resonance energy transfer to
measure the kinetics of the formation or disruption of particular
nucleic acid structures (23-26). Energy transfer between two
fluorescent dyes indicates their proximity, and in the case of DNA, the
proximity of any two strands labeled with the dyes (27-29). We
developed three assays based on energy transfer. These assays can be
monitored in real time and presumably do not perturb the reaction. They
allow the observation, by means of stopped-flow spectrofluorometry, of
pairing, strand exchange, or both, depending on the placement of the
two dyes (Fig. 1). An oligonucleotide system was chosen because of the
ease of labeling the DNA with fluorescent dyes and the high yield of
labeled DNA. Oligonucleotides also have the advantage of avoiding some
of the more complicated aspects of strand exchange with long DNA
substrates, such as the effects of coaggregate formation and the
topological difficulties of strand displacement (18, 21, 30, 31).
Fitting of the stopped-flow data to a model with a reversible pairing
step, followed by a reversible strand exchange step, allowed the
determination of rate constants for the whole strand exchange
reaction.
RecA was purified as described (32). T4 polynucleotide kinase was supplied by New England Biolabs. ATP, phosphocreatine, and phosphocreatine kinase were from Sigma. Bovine serum albumin was purchased from Boehringer Mannheim. Dithiothreitol was provided by Promega. Proteinase K was from American Bioanalytical.
Preparation of Oligonucleotide SubstratesAll DNA concentrations are given in terms of moles of nucleotides. Oligonucleotides were synthesized on an Applied Biosystems DNA synthesizer (model 380B) at the Keck Biotechnology Resource Laboratory at Yale. Primary amines on C6 linkers (Glen Research) were added to 83-mer oligonucleotides during synthesis for labeling with fluorescent dyes.
Fluorescent oligonucleotides were produced by reacting 360 µl of 5 mM DNA with 4 mg of 5-carboxyfluorescein or 5- and 6- carboxytetramethylrhodamine succinimidyl ester (Molecular Probes, Inc.) dissolved in 40 µl of anhydrous dimethyl sulfoxide. The reaction was carried out in 250 mM carbonate buffer, pH 9, overnight at room temperature in the dark. Oligonucleotides were then precipitated in ethanol, dissolved in formamide, and purified by electrophoresis on a 12% denaturing polyacrylamide gel (33). The absorbance at 260, 496, and 558 nm of the oligonucleotides was measured to determine, respectively, the concentrations of DNA, fluorescein, and rhodamine (34). Duplex oligonucleotides were prepared as described (33).
Duplexes were 5-end-labeled with 32P using T4
polynucleotide kinase (35). Duplexes were checked for complete
annealing by electrophoresis on an 8% nondenaturing polyacrylamide gel
and quantitated by use of a Molecular Dynamics PhosphorImager. All duplexes used contained less than 5% single strands.
Reactions were
conducted at 37 °C in buffer containing 33 mM PIPES
acetate, pH 7.0, 1 mM magnesium acetate, 1.2 mM
ATP, 2 mM dithiothreitol, and 100 µg/ml bovine serum
albumin. Stopped-flow fluorescence assays had no bovine serum albumin
(to prevent bubbles) and had a final concentration of 16 mM
phosphocreatine and 10 units/ml creatine phosphokinase for ATP
regeneration. Filaments were formed on 10 µM M() or R83
oligonucleotides (Sequences 1 and 5, Table I) by incubation with 3.33 µM RecA for a minimum of 2 min. Magnesium was increased
to 16 mM, and duplex 83-mer was added to a final
concentration of 20 µM. Unlabeled oligonucleotides still
bore the primary amine linkage.
|
For the pairing assays (Fig. 1), 10 µM
M()·3
F2 (Sequence 1, Table I) was
incubated with 3.33 µM RecA for 2 min under standard reaction conditions for filament formation. The magnesium was increased
to 16 mM and a final concentration of 20 µM
duplex (M(
)·3
NH2/M(+)·5
R or
M(
)·3
R/M(+)·5
NH2; Sequences 1 and 2, Table I) was
added and reacted for 2 min. In the case of the strand displacement assay, 25 µM RecA was incubated with 75 µM
M(
)·3
NH2 for 5 min in standard reaction buffer. The
filament was added to M(
)·3
F/M(+)·5
R duplex in reaction buffer
with 16 mM magnesium acetate to produce 7.5 µM M(
)·3
NH2 filament with 2.5 µM RecA and 10 µM M(
)·3
F/M(+)·5
R duplex DNA, which was reacted for 2 min. Two additional reactions were
carried out for each assay with either the fluorescein- or the
rhodamine-labeled strand replaced by an equivalent unlabeled, primary
amine-tagged oligonucleotide.
Fluorescence emission spectra from 502 nm to 620 nm were taken with excitation at 493 nm on an SLM8000C spectrofluorometer (Spectronic Instruments, Inc.). Excitation and emission polarizers were aligned at 54.7° with respect to each other to help eliminate polarization artifacts (36). Reactions containing rhodamine were also observed from 565 nm to 620 nm with excitation at 558 nm. Buffer spectra under both conditions were subtracted from the data. The increase in emission by rhodamine due to FRET was calculated by subtracting both the background emission of the sample containing only fluorescein-labeled DNA and the background emission of the sample with only rhodamine-labeled DNA. The spectrum of the fluorescein-only reaction was normalized at 525 nm to the height of the energy transfer reaction. The spectrum of the rhodamine-only reaction was normalized to the energy transfer reaction using the emission at 585 nm with excitation at 558 nM. The sum of the normalized spectra was subtracted from the spectrum of the energy transfer reaction. The normalizations and subtraction of background were performed according to the following formula.
![]() |
(Eq. 1) |
Assays were performed on
an Applied Photophysics DX.17MV sequential stopped-flow
spectrofluorometer. One syringe contained an 83-mer
oligonucleotide-RecA filament with an ATP-regeneration system, and the
other contained duplex in buffer plus magnesium for the magnesium
shift. The reaction conditions were standard, as described above. The
contents of the syringes were mixed in equal proportions with a dead
time of 50 ms or less. Excitation was at 493 nm, and an interference
filter with maximum transmission at 520 nm and a 10-nm bandwidth
(Corion) was used to select the emission wavelength observed. The time
base for collection of data was split, so that half the data points
were collected in the first 20 s. The control for photobleaching
was performed with 10 µM M()·3
F-RecA filament. The
control for nonspecific quenching of fluorescein was conducted with 10 µM M(
)·3
F-RecA filament and 20 µM
unlabeled duplex DNA. Heterologous controls were done with 10 µM R83·3
F or R83·3
NH2 as the filament
strand and 20 µM of the appropriate duplex. The data in
Table II were generated using curve-fitting software supplied with the
DX.17MV. The data in Figs. 5 and 7 were plotted and fit, and residuals
were calculated using KaleidaGraph software (Abelbeck Software).
|
Mathematical Modeling of Stopped-flow Data
The kinetics of
the reactions were further analyzed by comparison of the stopped-flow
data with theoretical curves generated from the reaction scheme
outlined in Fig. 1. Theoretical curves were computed by numerically
resolving the system of derivative equations by the fourth order
Runge-Kutta method. Starting at time t1, this
method calculates the concentration change of each species during a
short interval, t, based on the concentrations of all
species at t1. In this way, one can calculate
the concentration of all species at time t2 from the
concentrations at time t1 by adding the
concentration changes during the interval
t. By repeating this operation from time t = 0, one can compute the
concentration of all species as a function of time. The method was
numerically stable within a large range of
t values. In
general, we did not observe numerical instability using
t = 0.01 s. Furthermore, the concentrations
approached the theoretical equilibrium values after sufficient reaction
time, which supported the validity of the resolution method.
The concentrations of the species calculated by this method were then
converted to fluorescence intensity for comparison with the
experimental data. The maximum fluorescence change possible for either
pairing assay 1 or the strand displacement assay was modeled by
annealing of M()·3
F-RecA filament and M(+)·5
R (see "Results"). The relationship between fluorescence changes and concentration changes assayed by gel electrophoresis was linear for
both assays, so the change in concentration at any point on the
theoretical curves could be converted to a change in fluorescence by
multiplying it by the ratio of the maximum fluorescence change to the
maximum concentration change. For the pairing assays, we assumed that
the energy transfer for the intermediate was the same as for the
annealed DNA. A better fit of the data for the strand displacement
assay was obtained when a slightly smaller energy transfer for the
intermediate was assumed.
The comparison of theoretical and experimental data was made by least
square analysis. To find the best fit, the rate constants were
systematically varied and the corresponding theoretical data were
computed as described above. To reduce the number of parameters to be
fitted and to estimate the value of the rate constants for the back
reaction (k1 and k
2),
the equilibrium constants Keq1 for pairing and
Keq2 for strand exchange were determined from
the amount of strand exchange products determined by gel assay.
Keq1 = k1/k
1 = C/(A × B) and
Keq2 = k2/k
2 = (D × E)/C, where A
and B represent the concentrations of the reactants,
C is the concentration of the intermediate, and D
and E are the products. This system of equations was also
analyzed by the least square method to find the values of the
equilibrium constants. The theoretical values were computed by
numerical resolution of the equation system with the half-interval
method (38).
The same DNA
and RecA concentrations as in the FRET assay described above were used
for the gel electrophoresis assay shown in Fig. 6. The M()·3
F
strand (Sequence 1, Table I) of the duplex substrate used was
5
-end-labeled with 32P as described above. Aliquots (10 µl) of the strand exchange reaction were taken at different times and
deproteinized as described (35). Loading buffer was added to produce a
final concentration of 5% glycerol, 0.05% bromphenol blue, 0.05%
xylene cyanol. Samples were loaded on an 8% native polyacrylamide gel
and run 3 h at 275 V and room temperature. The gel was dried and
quantitated by use of the Molecular Dynamics PhosphorImager to
determine the extent of strand displacement.
The other gel electrophoresis assays that were used to calculate the concentrations of the products of strand exchange with the substrates for the three fluorescence assays were performed with 10 µM single-stranded DNA, 3.33 µM RecA protein, and 20 µM duplex DNA under standard conditions. Two-fold, 3-fold, and 5-fold increases in the preceding concentrations were made for the substrates of the fluorescent strand displacement assay. In another set of titrations, the filament concentration was held constant at 10 µM and the duplex concentration was 5, 7, 10, 20, or 30 µM. In all cases, the final concentration of products was determined from an aliquot of the reactions taken at 3 min after the addition of duplex DNA. The aliquots were deproteinized, electrophoresed, and quantitated as described above.
When fluorescein comes into proximity with tetramethylrhodamine, the overlap between its emission spectrum and the excitation spectrum of rhodamine allows the non-radiative transfer of energy to rhodamine by a process called fluorescence resonance energy transfer, or FRET (27-29). The maximum emission of fluorescein is around 525 nm in the presence of RecA. The emission of fluorescein at 558 nm, which coincides with the peak of the absorption spectrum of tetramethylrhodamine, is still half the maximum intensity observed at 525 nm; thus energy transfer can occur. Tetramethylrhodamine has a maximum of emission at 582 nm.
By varying the placement of the fluorescent probes attached to 83-mer oligonucleotides, we devised assays that separately measure homologous pairing and strand displacement promoted by RecA protein. We use the term strand displacement to indicate the apparent separation of the single-stranded product from the heteroduplex product of strand exchange without any deliberate deproteinizing treatment. We are unable to specify, however, if the displaced strand has completely dissociated from the heteroduplex, or if it is still in a common nucleoprotein complex, but is sufficiently removed or shielded from the duplex so that the dyes are unable to interact.
Fig. 1 depicts the design of the three energy transfer
assays. Two schemes were used to observe pairing. In the first, dubbed pairing assay 1, the RecA nucleoprotein filament was formed
on the M() oligonucleotide to which fluorescein was attached at the
3
end via a 6-carbon linker arm (M(
)·3
F: Sequence 1, Table I). This filament was reacted with a duplex molecule
consisting of the M(
) oligonucleotide that had no dye attached, but
still bore a primary amine on a linker arm, and the M(+)
oligonucleotide with rhodamine on a linker at the 5
end
(M(
)·3
NH2/M(+)·5
R; Sequences 1 and 2, Table I).
When attached in these positions, fluorescein and rhodamine should come
together during pairing and remain together after strand exchange takes
place, resulting in the quenching of fluorescein emission and the
enhancement of rhodamine emission (Fig. 1). In pairing assay 2, the
rhodamine was located on the 3
end of the M(
) strand of the duplex
oligonucleotide, so that the emission of fluorescein should be quenched
and the emission of rhodamine enhanced only during the phase of the
reaction in which all three strands are in proximity within the
filament (M(
)·3
F + M(
)·3
R/M(+)·5
NH2: Sequences
1 and 2, Table I; Fig. 1). Subsequently, as the rhodamine-labeled
strand is displaced during strand exchange, energy transfer should
decrease, resulting in the recovery of fluorescein emission and a
decrease in rhodamine emission. Thus, pairing assay 2 should detect
both the formation of the pairing intermediate and the displacement of
the M(
) strand as a result of strand exchange.
The third assay was designed to measure only strand displacement.
Unlabeled M() filament was reacted with a duplex oligonucleotide consisting of the M(
) oligonucleotide that was labeled at the 3
end
with fluorescein and the M(+) oligonucleotide that was labeled with
rhodamine at the 5
end (M(
)·3
NH2 + M(
)·3
F/M(+)·5
R: Sequences 1 and 2, Table I). Before the
reaction with single-stranded RecA filament, the dyes on the duplex DNA
should undergo energy transfer because they are together. Energy
transfer should decrease as the fluorescein-labeled strand leaves the
three-stranded intermediate upon strand exchange, resulting in an
enhancement of fluorescein emission and a decrease in rhodamine
emission (Fig. 1). With this set of three assays, pairing, pairing and
strand displacement, or just strand displacement can be monitored by
changes in energy transfer.
Energy transfer can be observed either as quenching of the donor
(fluorescein) or enhancement of the acceptor (rhodamine). The quenching
of fluorescein emission is much easier to measure than the enhancement
of rhodamine emission, as can be seen from the steady-state spectra for
pairing assay 1 in Fig. 2, which will be described
further below. However, fluorescein is sensitive to its environment,
and its emission may be quenched by other means than energy transfer,
such as the formation of a filament on DNA by RecA protein (data not
shown). In contrast, the increase in rhodamine emission due to energy
transfer, called sensitized emission, cannot arise from any
other source. These properties of the spectra led us to the following
compromise. We studied the rapid kinetics of the reactions by
monitoring the emission from fluorescein, but we measured the
steady-state emission spectra to confirm that changes in fluorescein
emission corresponded to changes in the sensitized emission of
rhodamine.
Steady-state Emission Spectra
Fig. 2 contains the
steady-state spectra for pairing assay 1, depicted schematically in
Fig. 1. Pairing assay 1 is used as an example of the energy transfer
calculations carried out for all three assays. We reacted M()·3
F
with one RecA monomer per three nucleotide residues for 2 min at
37 °C for filament formation (Sequence 1, Table I). The filament was
then reacted with an equimolecular quantity of
M(
)·3
NH2/M(+)·5
R duplex for 2 min (Sequences 1 and
2, Table I). Two emission spectra were taken, one with an excitation
wavelength of 493 nm, which primarily excited fluorescein (Fig.
2A), and another with an excitation wavelength of 558 nm,
which exclusively excited rhodamine (data not shown). Since both
fluorescein and rhodamine emit in the range of 580 to 585 nm when
excited at 493 nm, the contribution by fluorescein at those wavelengths
had to be subtracted to observe the sensitized emission of
rhodamine.
The sensitized emission of rhodamine was calculated by the use of
singly labeled controls as described (37) (see "Experimental Procedures" for details). A fluorescein-only control (M()·3
F filament + M(
)·3
NH2/M(+)·5
NH2) and a
rhodamine-only control (M(
)·3
NH2 filament + M(
)·3
NH2/M(+)·5
R) were carried out under the same
conditions as the energy transfer reaction that contained both dyes
(Fig. 2A). The sum of the normalized spectra of the fluorescein- and rhodamine-only controls is plotted in Fig.
2B along with the spectrum of the energy transfer reaction,
which contains both fluors. The difference between the two spectra in Fig. 2B is the sensitized emission of rhodamine, which is
plotted in Fig. 2D. The difference between the
fluorescein-only control and the energy transfer reaction at 525 nm is
the quenching of fluorescein as a result of energy transfer (Fig.
2A), which is larger and much easier to observe than the
sensitized emission. However, as indicated above, calculation of the
sensitized emission was necessary to confirm that the quenching of
fluorescein was not solely due to interactions with RecA protein or
other means not related to energy transfer.
A heterologous control for pairing assay 1 was performed to ensure that
energy transfer was homology-dependent, and not the result
of nonspecific binding. A mixture of 83-mers of random sequence, R83,
was labeled with fluorescein and used in place of M() as the filament
strand (R83·3
F + M(
)·3
NH2/M(+)·5
R: Sequences 1, 2, and 5, Table I). The emission spectra of this reaction and the sum
of the normalized singly labeled control reactions are shown in Fig.
2C. In the case of the heterologous control, the two spectra
almost completely overlap, and the difference between them, which is
plotted in Fig. 2D, is less than one-sixth of the sensitized
emission of the homologous reaction. Thus, pairing assay 1 resulted in
energy transfer that depended on homology, as expected from the design
of this assay (Fig. 1).
Pairing assay 2, depicted in Fig. 1, was performed under the same conditions as pairing assay 1. Steady-state spectra confirmed that the dyes behaved as expected and that a heterologous control led to no energy transfer (spectrum not shown). Because of the biphasic nature of the changes in energy transfer in this assay, we defer further consideration to the section below on stopped-flow analysis.
The strand displacement assay involved two sets of energy transfer
calculations. First, the sensitized emission of the
M()·3
F/M(+)·5
R duplex was calculated with duplexes in which
either the fluorescein- or the rhodamine-labeled strand had been
replaced with an unlabeled strand of the same sequence. The spectrum of
the doubly labeled duplex is shown in Fig.
3A, and the sensitized emission of that duplex is plotted in Fig. 3B. Then,
M(
)·3
NH2 that had been incubated with RecA protein in
a separate Eppendorf tube was added in 1.5-fold excess to each of the
three duplexes. After strand exchange had occurred, spectra were
collected and the sensitized emission of rhodamine was calculated once
more. The spectrum of the energy transfer reaction is shown in Fig.
3A for comparison with the spectrum with the original duplex
DNA. The decrease in sensitized emission as a result of the separation
of the strands of the duplex is evident from the plot of the sensitized
emission before (duplex only) and after strand exchange (Fig.
3B). A heterologous control was performed with the Het(+)
oligonucleotide as the filament strand, and no change took place in the
spectrum (Sequence 3, Table I, no spectrum shown). Thus, in the strand
displacement assay, the energy transfer decreased as expected in a
homology-dependent manner (Fig. 1).
Correlation of Sensitized Emission and Changes in Fluorescein Emission
Although the changes in sensitized emission occurred in
all three assays, we needed to be able to compare the sensitized
emission between assays and to determine how it was related to the
changes in fluorescein emission. To compare energy transfer
quantitatively in different assays, the ratio of the sensitized
emission of rhodamine to the normalized emission of the rhodamine-only
control was taken. This ratio, expressed as a percentage, should be
independent of variations in concentration. The normalized sensitized
emission is the percent increase in rhodamine emission for the two
pairing assays, shown in Fig. 4 (A and
B). For the strand displacement assay, shown in Fig.
4C, the percent decrease in rhodamine emission was
calculated from the difference in the normalized sensitized emission
before and after the M()·3
F/M(+)·5
R duplex was reacted with
M(
)·3
NH2.
To assess the correlation between changes in the sensitized emission of rhodamine and the emission of fluorescein, we had to know what portion of the changes in fluorescein emission was specific to energy transfer. In the case of the pairing assays, the quenching of fluorescein that was specific to energy transfer was determined from the difference in intensity at 525 nm between the fluorescein-only control and the reaction containing both dyes. Those two reactions were identical except for the presence of rhodamine in the doubly labeled sample, so the differences in emission should have been solely due to energy transfer. The nonspecific quenching was reflected in the difference between the total quenching observed during the course of the reaction containing both dyes and the specific quenching (Fig. 4, A and B). For the strand displacement assay, the nonspecific quenching was calculated from the change in emission of the fluorescein-only control before and after strand exchange. By comparing the sensitized emission, the specific quenching, and the nonspecific quenching of fluorescein, the reliability of the quenching of fluorescein as an indicator of energy transfer was determined.
Pairing assay 1 gave 37% specific quenching of fluorescein and 94% enhancement of rhodamine emission (Fig. 4A). The total quenching of fluorescein was 30%, indicating that all the quenching was the result of energy transfer.3 The heterologous control for pairing assay 1 produced 5% nonspecific and 1% specific quenching of fluorescein, with 6% enhancement of rhodamine emission. These data demonstrated that there was a good correlation between the quenching of fluorescein and energy transfer as assessed by the sensitized emission of rhodamine for pairing assay 1. There was no energy transfer to rhodamine in the absence of the cofactor ATP, but there was 8% quenching of fluorescein, 5% of which appeared to be specific (Fig. 4A). This specific component might be an artifact produced by slight differences in concentration between the fluorescein-only sample and the doubly labeled sample. The overall quenching was probably the result of RecA binding DNA even in the absence of a cofactor (39, 40). In any event, the controls for the requirement of homology and ATP were satisfactory.
A further control was performed to demonstrate that the energy transfer
observed for homologous substrates was not due to an artifact generated
by end-to-end aggregation rather than homologous alignment. In this
control, rhodamine was placed at the 3 end of the M(+) in the duplex
DNA, so that rhodamine and fluorescein would be on opposite ends of the
heteroduplex product of strand exchange (M(
)·3
F + M(
)·3
NH2/M(+)·3
R). This would make energy transfer
impossible unless there was nonspecific aggregation of ends. Only 5%
quenching of fluorescein and 8% enhancement of rhodamine was observed,
which was comparable to the heterologous background, suggesting that
end-to-end aggregation may have been responsible for most of the
heterologous signal at those concentrations of DNA and RecA (Fig.
4A).
When a 10-fold excess of unlabeled heterologous duplex oligonucleotide
(Sequences 3 and 4, Table I) was added to filament formed on
M()·3
F before the
M(
)·3
NH2/M(+)·5
R duplex DNA was added, 3% nonspecific and 29% specific quenching of fluorescein, as
well as 41% enhancement of the rhodamine emission still occurred. This
demonstrated that the RecA filament took up heterologous sequences, but
that homologous ones were able to compete with the heterologous DNA
(Fig. 4A). This competition might have been more effective
if the reaction time were longer and ATP regeneration had been
used.
Pairing assay 2 produced 7% specific quenching, 14% nonspecific quenching of fluorescein, and 20% enhancement of rhodamine emission (Fig. 4B). The nonspecific component of the quenching of fluorescein was larger relative to the specific quenching than in pairing assay 1, apparently because the energy transfer was far less (compare Figs. 4A and 4B). There were no energy transfer-related changes in fluorescence for the heterologous control for pairing assay 2 (Fig. 4B). Since the nonspecific quenching in pairing assay 2 was similar in magnitude for homologous and heterologous DNA, most of the nonspecific quenching was probably caused by interaction of the duplex DNA with the RecA filament that was independent of homology, such as end-to-end aggregation. The correlation between the quenching of fluorescein and energy transfer seems less good for pairing assay 2 than for pairing assay 1, but the spectra could not be taken when energy transfer was the greatest in pairing assay 2 because of the biphasic changes that occur in that assay (see below).
In the strand displacement assay, the fluorescein emission increased 370%, 350% of which was specific to energy transfer, and the sensitized emission of rhodamine decreased 62% (Fig. 4C). The heterologous control for strand displacement produced no changes at all. Since no strand exchange could take place with these substrates, this result was expected. In general, pairing assay 1 and the strand displacement assay were better than pairing assay 2 in terms of having few or no changes in fluorescein emission that were not related to energy transfer.
Stopped-flow Analysis of Pairing and Strand Displacement RatesHaving established that changes in fluorescein emission
were correlated with homology-dependent changes in energy
transfer, we used changes in fluorescein emission as observed by
stopped-flow spectrofluorometry to study the rapid kinetics of
homologous pairing and strand exchange. Samples were excited at 493 nm,
and the emission of fluorescein at 520 nm was observed. A control for
photobleaching of fluorescein showed less than 2% of the change in
fluorescence observed for any of the assays. Heterologous controls for
the assays performed with R83·3F or R83·3
NH2 produced
no fluorescence change beyond background noise. A control for
nonspecific quenching of fluorescein yielded a change in fluorescence
that was 6% of the amplitude change in pairing assay 1 and 16% of the
change in pairing assay 2 (see "Experimental Procedures" for
further details of the controls).
Pairing assay 1 yielded a single, rapid decrease in fluorescein
emission, as predicted by the scheme in Fig. 1 (Fig.
5A). Pairing assay 1 was performed with a
range of concentrations of single-stranded oligonucleotide ranging from
5 µM to 50 µM and duplex DNA from 10 µM to 100 µM in 1-to-1 ratios of filament
to duplex. Also, the filament concentration was held constant at 10 µM and the concentration of duplex DNA was varied 4-fold.
For initial analysis of rates, the time courses for pairing assay 1 at
different concentrations were fit with single exponential functions or
the sum of two exponential functions. The general equations used for
fitting the data were I = A1·exp(k1·t) + C or
A1·exp(
k1·t) + A2·exp(
k2·t) + C, where I = fluorescence intensity, t = time, k = first order rate
constant, A = amplitude, and C = end
point of the curve. The results are listed in Table II. Residuals for the single or double exponential fits are plotted beneath
the data to indicate how well the data are described by exponential
functions (Fig. 5A). The residual for the single exponential fit for pairing assay 1 deviated considerably from zero in the first
30 s, whereas the residual for the double exponential fit deviated
less and at later times (Fig. 5A). The failure of a single exponential to fit the data indicated that pairing assay 1 was not
detecting a first order process. The apparent first order rate
constants, kobs(a) and
kobs(b), varied about 3-fold
with concentration (Table II), indicating that the sum of two
exponential functions also did not adequately describe the
reaction.
Controls were performed with constant concentrations of DNA (10 µM single strand and 20 µM duplex) and with
one RecA monomer to six nucleotides or one nucleotide (data not shown).
The rates were the same as for the reaction with one RecA monomer to
three nucleotides, within the error of the measurements. These controls demonstrated that the concentration dependence of the rate constants was not just the result of changes in protein concentration.
Additionally, when the control for nonspecific quenching was subtracted
from the data for 10 µM filament and 20 µM,
kobs(a) remained the same
and kobs(b) increased by
only 0.01 s1. The lack of fit to a single exponential
function and the variation in constants with concentration confirmed
that the pairing reaction was not first order, which is consistent with
the pairing reaction being a bimolecular process.
In contrast to pairing assay 1, pairing assay 2 demonstrated two phases: a decrease in fluorescein emission upon pairing and increase with displacement of the rhodamine-labeled strand, as predicted in Fig. 1 (Fig. 5B). Also in contrast to pairing assay 1, pairing assay 2 was well fit by the sum of two exponentials. The residual shows the quality of the fit, with no significant deviations from zero (Fig. 5B). The apparent first order rate constants kobs(c) and kobs(d) varied little with concentration (Table II). These observations suggest that pairing assay 2 may be reflecting two first order processes. This is difficult to reconcile with pairing assay 1, unless two or more intermediates exist, and pairing assay 2 is scoring a different, later intermediate from assay 1. See "Discussion" for further details.
The strand displacement assay resulted in a single increase in
fluorescein emission, as expected (Figs. 1 and 5C). The time course of the strand displacement assay was described as well by a
single exponential as by the sum of two exponentials (Fig. 5C). The residuals for both fits show trends away from zero
at early times (Fig. 5C). This may be the result of changes
in fluorescence that take place as the duplex is taken up into the RecA
single-stranded DNA filament. The rates varied only from 0.05 ± 0.02 to 0.07 ± 0.01 s1 over a 10-fold concentration
change, which suggested a first order reaction mechanism, despite the
problems with fitting the data to a single exponential (Table II). A
first order mechanism is consistent with the proposed model, in which
the pairing intermediate goes on to complete strand exchange and
displacement. The change in fluorescence for the strand displacement
assay was slower than for pairing assay 1 or the first phase of pairing
assay 2, but was almost the same as the second phase of pairing assay
2. This indicates that the strand displacement assay and the latter
half of pairing assay 2 probably detect the same step of strand
exchange. Controls with different RecA to nucleotide ratios, like those performed for pairing assay 1, indicated that the rates changed only
when there was less than one monomer of RecA to three nucleotides (data
not shown). At one RecA to six nucleotides, the rate dropped to 0.02 s
1, which suggested that incomplete filament formation
had more of an effect on strand exchange than on pairing.
The preliminary analysis above indicated that at least one step in the strand exchange process is concentration-dependent. A link between the fluorescence assays and an independent assay for the concentrations of the species had to be established for further kinetic analysis, described below. The fluorescence assay for strand displacement was compared with the more traditional gel electrophoresis assay to establish whether they both detected the same step of the strand exchange reaction. We used the same substrates in both assays to observe the time course of strand exchange. In the fluorescence assay, the emission of fluorescein at 525 nm was monitored; in the gel electrophoresis assay, aliquots of the reaction were taken at various times and deproteinized with SDS and proteinase K. The deproteinized DNA was then run on a nondenaturing polyacrylamide gel, and the percent strand exchange determined. The two assays were independently normalized to a maximum of 100% (Fig. 6). The time courses of strand exchange measured by fluorescence assay and by gel assay coincided. According to the simplest interpretation, the two assays measure the same step of strand exchange.
The gel assay was also used to measure the absolute yield of strand
exchange with the same substrates that were used for the fluorescence
assays. Knowledge of the equilibrium constants for the strand exchange
reaction reduced the number of rate constants that had to be
independently varied in the mathematical modeling of the stopped-flow
data below. Two sets of experiments were conducted to discover the
effect on yield of varying substrate concentration. When the filament
concentration was held constant at 10 µM and the duplex
DNA concentration increased from 5 µM to 30 µM, the yield of heteroduplex increased from 16 to 68%
(data not shown). The yield at one filament to one duplex was 56%.
These results indicate that pairing is reversible
(k1
0); otherwise, the yield should have
reached maximum at one filament to one duplex, rather than continuing
to increase even though the duplex was no longer the limiting
substrate. The data from this experiment were used to calculate the
relationship between the equilibrium constants for pairing and strand
exchange as described under "Experimental Procedures" under
"Mathematical Modeling of Stopped-flow Data."
In other experiments, the concentration of DNA and RecA protein was
increased up to 5-fold while the 1-to-1 ratio of single-stranded to
double-stranded DNA was maintained, and the percent of strand exchange
remained constant at an average of 54 ± 3% (data not shown).
This result also suggested that an equilibrium existed in the overall
reaction, including the strand exchange step, which supported the
modeling of strand exchange as reversible, with k2
0 (see below).
We measured the yield of strand exchange with the substrates for the different assays to see if the dyes themselves affected the reactions. Under the conditions of the steady-state fluorometric assays, in the absence of an ATP regeneration system, the gel assay revealed that the respective yields were 40 ± 10%, 30 ± 10%, and 40 ± 10% for pairing assays 1 and 2 and the strand displacement assay. Substrates that lacked any modification, even the primary amine, gave 30 ± 10% products. The dyes did not seem to significantly affect the yield of strand exchange.
We also wanted to confirm that fluorescence changes in the reactions
were proportional to concentration changes, so that we could convert
concentration to fluorescence to compare theoretical curves generated
by mathematical modeling of the reactions with the stopped-flow data.
To calibrate changes in fluorescein emission with changes in
concentration, we used RecA-catalyzed annealing of 10 µM
M()·3
F filament and 10 µM M(+)·5
R to model the
maximum amplitude for pairing assay 1 and the strand displacement
assay. The annealing reaction was performed under the same conditions as the other stopped-flow fluorescence measurements. The amplitude of
the change in fluorescein emission observed with the strand displacement assay was 62% of the amplitude observed with the annealing reaction, which is close the yield of strand exchange assayed
by gel electrophoresis (54%). Pairing assay 1 produced an amplitude
that was 75% of the annealing amplitude, which was higher than
expected from the results of the gel assay. The pairing assay may have
detected intermediates that had not gone on to complete strand
exchange, and thus were not detected by the gel assay. The amplitudes
of the assays relative to the annealing reaction were used to convert
concentrations to fluorescence emission, as described under
"Experimental Procedures."
From the preliminary analysis of the data by fitting
with exponential expressions, it was clear that there were at least two steps in the strand exchange reaction and the first step was probably a
second order process. We have further analyzed the kinetic data based
on the reaction scheme shown in Fig. 1 by comparison of experimental
data with theoretical curves generated as described under
"Experimental Procedures." The analysis was done with
concentrations in terms of molecules of DNA, so all constants derived
are in those terms. The strand exchange reaction was modeled as a
completely reversible reaction. This assumption was made on the basis
of the observations noted above in the results of the gel assays. The
equilibrium constants for pairing and strand exchange,
Keq1 and Keq2, were
determined from the analysis of the variation in the amount of products
with substrate concentrations as described under "Experimental
Procedures" and elsewhere (38). The fit was reasonable when the
Keq1 value was between 1 × 106
and 1 × 107 M1, and the
best fit was obtained with Keq1 = 5 × 106 M
1 (data not shown). This
analysis provided the relationship between Keq1
and Keq2 values that was used to estimate the
rates of the back reaction (k
1 and
k
2), and thus decrease the number of kinetic
parameters to be fitted from 4 to 3.
Since the preliminary exponential analysis indicated that the strand
displacement step followed first order kinetics and the k2 value was about 0.06 s1,
regardless of the concentration of substrate, we have analyzed the data
from pairing assay 1 using this k2 value. To
find the best fit, we have systematically varied the
Keq1 and k1 values. The
Keq2 value, and thus the
k
2 value, was estimated from the relationship
between the Keq1 and Keq2
values. This procedure was performed on the data for all the
concentrations listed in Table II except for the lowest concentration,
which was at the limit of detection and had a low signal-to-noise
ratio. We obtained a good fit for all cases, and Fig.
7A is one example. The
Keq1 and k1 values
calculated from each data set were independent of substrate
concentrations (Table III). The ability to fit the data from pairing assay 1 to a model with a second order pairing step and to
obtain rate and equilibrium constants that are independent of
concentration supports a model in which the pairing step is bimolecular.
|
The best fit for k1 was about 1 × 106 M1 s
1 and for
Keq1 was 1 × 107
M
1. The Keq1 value
thus determined was close to the value determined from the analysis of
amount of strand exchange products at the end of the reaction (5 × 106 M
1). This consistency in
the value of Keq1 obtained by different methods
further supports the validity of our analysis. Uncertainty in the value
of the yield of strand exchange (±10%), and thus the change in
fluorescence intensity, affected the Keq1 value, but did not significantly alter the k1 value
obtained. In some cases we also varied the k2
value to examine the effect of its uncertainty on the determination of
k1 value. The variation of k2 value from 0.04 to 0.12 s
1 did
not significantly affect the analysis, showing that pairing assay 1 provides mainly information about the first step of the reaction, as we
expected from the design of the assay (Fig. 1).
We then analyzed the data from the strand displacement assay, taking
into account the first step of the reaction to more accurately estimate
the k2 value. We searched for the best fit by
systematically varying k2 value and using the
values for Keq1 and k1
determined above. The average of the best fits for all concentrations
(except the highest) was k2 = 0.06 ± 0.02 s1 (Table III). The fit became better when one assumed
that the energy transfer between two strands of the duplex DNA
decreased by about 20% upon binding to the RecA-single-stranded DNA
filament (Fig. 7C). Several observations have demonstrated
that the structure of duplex DNA is modified upon the binding to
RecA-single-stranded DNA complex (41-45). Our results indicated that
the modification occurred immediately after the binding, or perhaps
only the duplexes that had randomly assumed a different conformation
interacted with RecA (see Fig. 1). Uncertainty in the yield of strand
exchange and Keq2 values (2 × 10
7 to 3 × 10
7 M)
affected the determination of the value of
k2.
Finally, we analyzed the data from pairing assay 2 using the values of the constants determined above from the other two assays. We obtained a relatively good fit with these values, indicating that the reaction scheme is roughly correct. The best fit for the data from pairing assay 2 was obtained with smaller k1 and k2 values than those determined from the data for the other assays (Fig. 7B, Table III). However, the analysis of the data from pairing assay 2 is complicated by the smaller amplitude of the fluorescence changes and the larger nonspecific changes in fluorescence relative to pairing assay 1 and the strand displacement assay. Consequently, the rate constants cannot be estimated more precisely from the data for pairing assay 2.
From the beginning of studies in vitro on the recombination activities of RecA protein, it has been clear that there is an initial phase of the reaction in which homologous recognition occurs, followed by a slower phase during which extensive strand exchange occurs (9). The observation that homologous recognition occurred without any free ends in the DNA substrates demonstrated that recognition does not require that the strands of nascent heteroduplex DNA be truly be intertwined, as in Watson-Crick duplex DNA (41-44, 46-48). However, little or no success has attended efforts to isolate the initial intermediate or even to demonstrate that strand exchange at or near the site of recognition is an event that is distinct and separable from recognition. Indeed, studies from several laboratories have supported the hypothesis that localized switching of bases from parental to recombinant pairs is the mechanism of homologous recognition (49-51). The present kinetic studies, which make use of fluorescence energy transfer and topologically simple oligonucleotides, provide insights on the staging of events in the overall reaction, demonstrating two or possibly three distinct steps.
We developed a set of three assays that detected pairing, strand displacement, or both, depending on the location of the fluorescent dyes fluorescein and tetramethylrhodamine (Fig. 1). These assays are performed in solution without the disruption of intermediates, and stopped-flow analysis can be used to measure the rates of the separate phases of the reaction. Although we cannot exclude steric effects of the probes themselves, the yield of strand exchange did not differ significantly between dye-labeled and unlabeled substrates. Since the fluorescence assays use oligonucleotides, they suffer the same drawbacks as any assay that uses RecA and oligonucleotides, namely that RecA does not bind as well to oligonucleotides as to long DNA (52, 53). However, the length of the oligonucleotides is more than sufficient for homologous pairing and strand exchange (33, 54-56). Since reactions with oligonucleotides are not complicated by the formation of coaggregates (18, 21, 30) or by topological barriers to strand displacement (31), they provide a simplified model system that is amenable to kinetic analysis.
Model of Pairing and Strand ExchangeWhen the stopped-flow data were fit to the model in Fig. 1, the assumption was made that the strand exchange reaction with oligonucleotides is reversible, which is supported by several lines of evidence. Rosselli and Stasiak (55) demonstrated that, when the products of strand exchange with 52-mer oligonucleotides were fixed with glutaraldehyde, protein-free heteroduplex was released as soon as heteroduplex appeared upon deproteinization, while the displaced single-strand remained bound by RecA. These observations were confirmed by us with a filter assay in which protein-bound DNA was retained by a filter on the basis of size.4 As a result, the displaced strand was available for another round of strand exchange (55). In addition, we were able to show that in a strand exchange reaction at completion, the addition of excess single-stranded product to drive the reaction back established a new equilibrium governed by the same Keq as the initial reaction.4
Further evidence of reversibility came from the experiments reported
here. As noted under "Results," the yield of strand exchange remained near 50% over a 5-fold change in the concentration of the
substrates while the molecular ratio of filament to duplex was kept
constant, which indicated that an equilibrium existed for the reaction
and that k2 was significant (Fig. 1). Also,
the yield of heteroduplex continued to increase as the
duplex/single-strand ratio exceeded 1, which demonstrated that
k
1
0 (Fig. 1). The assumption of
reversibility was supported by previous studies, gel electrophoresis
data in the current study, and most of all by the fact that the
stopped-flow data could be fit to such a model.
The kinetic order of the steps in strand exchange was also supported both by the fit of the data to the model and by the more qualitative preliminary analysis of the stopped-flow data with exponential functions. When the stopped-flow data from pairing assay 1 were fit to a model with a second order, reversible pairing step, the rate constants and the equilibrium constant Keq1 varied little with substrate concentration, which supported the model (Table III). Likewise, the first order rate constants determined by fitting the data from the strand displacement assay to a reversible first order step were constant (Table III). The model in Fig. 1 is supported not only by general observations but also by the quality of the fit of the data from pairing assay 1 and the strand displacement assay to the model.
The equilibrium and rate constants derived from the data for pairing
assay 1 and the strand displacement assay describe a two-step process
for the detection of homology. The value of k1, 1 × 106 M1
s
1, determined from the stopped-flow data for pairing
assay 1, is about the same as the rate constant for the annealing of
complementary 16-mer oligonucleotides in the absence of protein (26).
The value of k
1 (0.10 s
1) is
close to the value of k2 for strand displacement
(0.06 s
1). Thus, the pairing intermediate has an equal
probability of dissociating or going on to strand displacement. The
back reaction for strand displacement (k
2 = 2-3 × 105 M
1
s
1) is significant compared with the rate constant,
k1, for the initial pairing step. In light of
the reversibility of both steps in strand exchange, discrimination
between homologous and heterologous substrates could occur at either or
both steps.
Pairing assay 2, however, did not completely fit the kinetic scheme outlined in Fig. 1. Pairing assay 2 was biphasic, as expected (Fig. 5B). The second phase of pairing assay 2 was first order and had almost the same rate as strand displacement (Tables II and III); thus, it probably also reflected strand displacement, as predicted. Surprisingly, however, the first phase of pairing assay 2 also appeared to be first order. The data were perfectly fit by a single exponential function (Fig. 5B), and the rate constants were independent of concentration (Table II). In clear contrast, the data for pairing assay 1 were well described by a second order model. Since the initial homologous interactions should be second order, we infer that the first exponential phase of pairing assay 2 represents events that occur after those detected by assay 1. The simplest explanation is that pairing assay 2 detects a third step in the strand exchange reaction between initial pairing and strand displacement.
The two pairing assays might have detected different intermediates because of differences in the placement of the fluorophores. A priori, there is no reason why pairing assay 1 and pairing assay 2 should detect the same intermediate. In pairing assay 1, rhodamine was on the complementary strand, and the expected second order kinetics were observed (Fig. 1). On the other hand, in pairing assay 2, rhodamine was on the identical strand of the duplex, and first order kinetics were seen. The identical strand of the duplex may have been just outside the radius of energy transfer during the initial pairing step. Perhaps the filament strand is close to its complement during the initial second order pairing step, which is followed by a first order conformational change that brings identical strands into proximity. Finally, the second pairing intermediate goes on to complete strand exchange in a first order reaction. Further studies of pairing assay 2 are needed to evaluate this three-step model.
Comparison with Rates for Human Rad51Preliminary studies have been done on the strand exchange reaction catalyzed by Rad51 protein, a human homolog of RecA, using the assay system described in this paper (15). Surprisingly, the apparent first order rate constants for pairing assay 1 and the strand displacement assay were the same.5 From these initial studies of Rad51, it seems that either pairing assay 1 does not detect the initial pairing step catalyzed by human Rad51, or pairing and strand exchange are the same step, and all discrimination of homology occurs at the strand exchange step. Further studies are needed to distinguish between these possibilities. If pairing and strand exchange are simultaneous, Rad51 might be less selective, or the process of finding homology might be less efficient than for RecA.
Comparison with Previous Studies of KineticsIn previous
studies with long DNA substrates, joint molecule formation was assayed
by a nitrocellulose filter assay that favored the survival of joints
that had already undergone limited strand exchange (16-20, 57). Thus,
the formation of such joints was probably more closely related to
strand exchange as measured in the present study. Published data
support this interpretation; the first order rate constant for joint
molecule formation determined by Julin et al. (20) was 0.014 s1, which is only slightly less than the
k2 value observed here (0.06 s
1).
Yancey-Wrona and Camerini-Otero (22) developed an assay for the stable
synapsis of an oligonucleotide with a longer duplex target. In their
assay, the protection of a restriction site in a 57-mer oligonucleotide
duplex by a shorter RecA-oligonucleotide complex formed in the presence
of ATPS was measured following an 8-min digestion with restriction
endonuclease. They used a standard Michaelis-Menten model to analyze
the data and found that the equilibrium constant for the homologous
pairing ranged from 7.7 × 106 to 1.3 × 107 M
1, which is in agreement
with the value of Keq1 determined in this study
(1 × 107 M
1). However, the
rate constant they measured for stable synapsis was 2 orders of
magnitude lower than the strand exchange rate observed in this study,
which may have been related to the use of ATP
S as a cofactor.
Reversibility
is presumably advantageous in vivo. In the cell, RecA or
some other protein has to be able to dissociate whatever joint molecule
has been formed in the face of topological constraints on strand
displacement or in the event that the substrates are not completely
homologous. This has been observed in vitro with long DNA
substrates that do not have a free end to displace, such as distal
joints or medial joints formed with 3 single strand ends, which
undergo cycles of formation and dissociation (6, 58, 59). These joints
may involve localized base pair switching because they can be isolated
following deproteinization. Uptake of duplex DNA into the RecA filament
unwinds the DNA and eliminates favorable stacking interactions between
bases (41-48). Perhaps the role of the unique structure of the RecA
filament is to ensure the instability of the heteroduplex, without
losing the specificity of Watson-Crick base pairing, so that the
energetic barrier in either direction of pairing and strand exchange is
lower. The reduction of the stability of heteroduplex may raise the
threshold for stabilization of joints, ensuring a more perfect fit
before strand exchange is allowed to proceed. The structure of the
filament itself is thus the instrument of reversibility and
fidelity.
We thank S. O'Malley, K. Anderson, and M. Ibba for assistance in learning fluorescence techniques. We also thank E. Folta-Stogniew and R. Gupta for their critical reading of the manuscript, and D. Crothers and T. Jovin for their helpful suggestions. We are grateful for technical assistance from Z. Li and J. Zulkeski.