ABSTRACT
We have developed a new helicase assay that overcomes many limitations of other assays used to measure this activity. This
continuous, kinetic assay is based on the displacement of fluorescent dyes from
dsDNA upon DNA unwinding. These ligands exhibit significant fluorescence
enhancement when bound to duplex nucleic acids and serve as the reporter molecules of DNA unwinding. We evaluated the potential of several dyes [acridine orange, ethidium bromide, ethidium homodimer, bis-benzimide (DAPI), Hoechst 33258 and thiazole orange] to function as
suitable reporter molecules and demonstrate that the latter three dyes can be
used to monitor the helicase activity of
Escherichia coli
RecBCD enzyme. Both the binding stoichiometry of RecBCD enzyme for the ends of duplex DNA and the apparent rate
of unwinding are not significantly perturbed by two of these dyes. The effects
of temperature and salt concentration on the rate of unwinding were also
examined. We propose that this dye displacement assay can be readily adapted
for use with other DNA helicases, with RNA helicases, and with other enzymes
that act on nucleic acids.
Helicases serve a variety of functions in DNA metabolism (for reviews, see
1
,
2
). Cellular (
Escherichia coli
DnaB, PriA and Rep proteins), phage (T4 gene
41
and dda proteins; T7 gene
4
protein), and viral (SV40 large T antigen; HSV-1 UL5/UL52 complex and UL9 protein) helicases are involved in the initiation of replication, by
unwinding DNA so that other proteins of the replication apparatus can assemble
on the ssDNA. These proteins also participate in the elongation phase of
replication, by unwinding DNA ahead of this complex to provide the required template. Other helicases
(e.g. the
E.coli
RecBCD and RecQ proteins) are implicated in recombination by genetic criteria.
In vitro
, RecBCD enzyme helicase activity generates the ssDNA substrate to which the
RecA protein of
E.coli
can bind to initiate heteroduplex DNA formation (
3
,
4
). Helicases such as the
E.coli
RuvAB complex function in recombination by promoting the branch migration of
crossover intermediates (
5
). Another class of helicases includes the
E.coli
UvrAB and UvrD,
Saccharomyces cerevisiae
Rad3 and Rad25, and human ERCC-2, -3 and -6 proteins (reviewed in
6
). These helicases act in nucleotide excision repair (
7
,
8
) or methyl-directed mismatch repair (
9
) during both pre-incision (recognition of DNA damage or alteration) and post-incision (displacement of damaged fragment) steps. Various aspects
of RNA metabolism are also dependent upon the action of helicases (for review,
see
10
). The unwinding of RNA templates is required in processes as diverse as
transcription termination (
11
), translation initiation (
12
,
13
) and RNA processing (
14
-
16
).
Many assays have been developed to measure the unwinding of duplex nucleic acids
by helicases. The first assays measured the sensitization of labeled duplex DNA to single-strand specific nucleases such as S
1
or exonuclease I, a result of the production of ssDNA during unwinding (
17
-
20
). Electron microscopy was employed to visualize directly the regions of DNA
unwound by proteins such as RecBCD enzyme (
21
), Rep protein (
22
),
E.coli
helicases I and II (
23
,
24
) and SV40 large T antigen (
25
). Currently, the most common assay measures the ability of a helicase to
displace a labeled fragment which is annealed to a single-stranded DNA or RNA molecule; this displacement is detected by gel
electrophoresis, as a band which has altered mobility (
26
,
27
). Several continuous fluorometric assays have also been developed. The first
described exploits ssDNA binding proteins [e.g.
E.coli
single-strand DNA binding protein (SSB) protein or phage T4 gene
32
protein] as the reporter molecule (
28
). As the dsDNA is unwound, the SSB protein binds to the ssDNA formed, and its
intrinsic fluorescence is quenched. A second method employs duplex DNA strands
whose complementary ends are labeled with different fluorescent moieties; when
these ends are separated by the helicase, the fluorescence quenching which
occurs when the labels are in close proximity is abolished (
29
,
30
). The third fluorescent assay utilizes an oligonucleotide which contains 2-aminopurine substituted for adenine (
31
). This analog exhibits fluorescence quenching when hydrogen bonded to thymine
residues, and thus unwinding can be measured as an alleviation of this
quenching.
pBR322 DNA was prepared by alkaline lysis and double-banding in CsCl-ethidium bromide density gradients (
37
). Phage M13 ssDNA was prepared according to standard procedures (
38
). Nucleotide concentrations were determined at 260 nm using extinction
coefficients of 6500 and 8784 M
-1
[middot]cm
-1
for dsDNA and ssDNA, respectively.
RecBCD enzyme was purified as previously described (
28
,
39
) and was quantitated at 280 nm using an extinction coefficient of 4 * 10
5
M
-1
[middot]cm
-1
(
28
). The specific activity of the enzyme preparation was 3.1 * 10
5
nuclease units/mg protein (
40
) and 4.1 * 10
4
helicase units/mg protein (
28
); its apparent binding stoichiometry was ~3 molecules/end (
28
). SSB protein was purified as described (
41
) and was quantitated using an extinction coefficient of 3 * 10
4
M
-1
[middot]cm
-1
at 280 nm (
42
).
EB was purchased from Sigma. AO, DAPI, EHD, H33258 and TO were obtained from
Molecular Probes (Eugene, OR). Solid TO was also generously provided by Becton
Dickinson. Each dye, except TO, was dissolved as a concentrated stock solution
(~5 mg/ml) in H
2
O, and serial dilutions were made from this stock. TO was dissolved as a
concentrated stock solution (~2 mg/ml) in 100%
N
,
N
, dimethyl formamide (DMF); serial dilutions were made from this stock into TO
dilution buffer (10% DMF, 0.1 mM 2-mercaptoethanol) to keep the dye soluble. Concentrations of the following
dyes were determined in H
2
O using the following extinction coefficients supplied by Molecular Probes
(given in M
-1
[middot]cm
-1
): AO, 65 * 10
3
at 488 nm; DAPI, 33 * 10
3
at 345 nm; EB, 5.5 * 10
3
at 546 nm; EHD, 8.9 * 10
3
at 528 nm. The concentrations of H33258 and TO were determined in methanol
using extinction coefficients of 46 * 10
3
and 77 * 10
3
M
-1
[middot]cm
-1
at 344 and 502 nm, respectively.
The standard reaction buffer consisted of 25 mM Tris-acetate (pH 7.5), 1 mM Mg(OAc)
2
and 1 mM DTT. When SSB protein was included, it was present at a 3-fold molar excess, assuming a site size of 15 nucleotides/monomer under
stoichiometric conditions; this corresponds to a protein concentration which is
20% of the DNA (nucleotide) concentration. The DNA substrate was 10 [mu]M nucleotide (2.1 nM ends)
Bam
HI-digested pBR322.
The reaction (350 [mu]l total volume) contained standard buffer, linearized pBR322 DNA, and the
indicated concentration of RecBCD enzyme. [For experiments in which the
concentration of TO was varied, an appropriate amount of TO dilution buffer was
added to the reaction so that the final concentration of DMF was constant (<0.3%). This was necessary because control experiments showed that DMF quenched
fluorescence significantly (unpublished observation).] SSB protein was added at
this point for dye fluorescence measurements. The fluorescence signal due to
these components was zeroed out. The fluorophore (either SSB protein or dye)
was then added, and the reaction was allowed to equilibrate to the indicated
temperature. Unwinding was initiated by the addition of ATP to 3 mM final
concentration. Under these conditions (i.e. ATP concentration in excess of Mg
2+
ion concentration), the dsDNA exonuclease activity of RecBCD enzyme is largely
suppressed (
39
,
43
).
Fluorescence measurements were carried out on a Shimadzu RF5000U
spectrofluorophotometer. When the intrinsic fluorescence of SSB protein was
measured, the excitation and emission wavelengths were set at 290 and 340 nm, respectively. The excitation and emission wavelengths for each dye were determined experimentally in the
presence of dsDNA. For AO, the values were 487 and 510 nm; for DAPI, 345 and
467 nm; for EB, 546 and 590 nm; for EHD, 552 and 596 nm; for H33258, 344 and 487 nm; and for TO, 504
and 528 nm. When SSB protein fluorescence was monitored, the band widths were 1.5 and 10 nm for the
excitation and emission slits, respectively; for all dye fluorescence
experiments, these values were 5 and 10 nm, respectively. There was no indication of energy transfer between SSB protein
and the dyes.
For SSB protein fluorescence measurements, the value for 100% unwinding was
calculated as described previously (
28
,
44
). For dye fluorescence experiments, the value for complete unwinding was obtained
by subtracting the previously determined fluorescence of an equimolar amount of
ssDNA (i.e. heat-denatured pBR322) (F
ssDNA
) from the initial fluorescence (F
dsDNA
); these quantities were determined for every set of reaction conditions that
were examined. The difference in these values (F
dsDNA
- F
ssDNA
) was taken to be the maximum fluorescence change possible if all of the DNA
molecules were fully unwound (F
exp
). The observed fluorescence change (F
obs
) was divided by this difference to obtain the extent of unwinding (i.e. the
percentage of total DNA unwound). Initial rates were then calculated as
described previously (
28
). Briefly, this involves drawing a line corresponding to the initial slope and
determining the point at which it intersects a line drawn through the plateau
at the end of the reaction. This intercept defines the time required for
complete unwinding. The extent of unwinding was multiplied by the concentration
of base pairs in the reaction to calculate the total concentration of unwound
base pairs, and this value was divided by the time required for complete
unwinding to yield the apparent rate.
For helicase activity to be detected, dyes must exhibit relative enhancement of
fluorescence upon binding dsDNA as compared to that obtained when the fluorophore is either in solution or bound to ssDNA.
Although each dye tested displays greater fluorescence in the presence of dsDNA
than in the presence of ssDNA, the fluorescent quantum yield and the dsDNA-specific enhancement are dependent upon the dye used (Fig.
2
A). A dye titration was performed to determine both the magnitude of the
unwinding signal (i.e. how much fluorescence is emitted in the presence of
dsDNA as compared with that either in the presence of ssDNA or when free in
solution) and the useful range of dye concentration. In Figure
2
A, a wider separation in the values for dsDNA (-) and those for ssDNA ([circle]) or free ligand (
+
) indicates that there will be greater discrimination between the duplex
substrate and unwound products. With AO, for example, the signal difference is
small (i.e. only ~15% of the initial fluorescence signal can be lost as a result of
unwinding). For TO, the signal from ssDNA-bound ligands is greater than for AO, but this fact is offset by a much
greater increase in the fluorescence in the presence of dsDNA.
Initial experiments indicated that DNA unwinding could be monitored using the
signal produced by the binding of fluorescent dyes to duplex DNA (examples of
typical unwinding traces are shown in Figs
3
A and
4
A). To determine whether the presence of dyes affected the observed rate of
unwinding, the reaction [including SSB protein to allow comparison with
previous studies (
28
)] was performed at several concentrations of DAPI, H33258 and TO. We noted that
unwinding reactions in the presence of either DAPI or H33258 (but not TO)
display lag behavior at concentrations of dye <100 nM (data from a H33258 titration are shown in Fig.
3
A); at or above 100-150 nM dye, lag behavior is not observed (data not shown). We presume
that the lag behavior at low dye concentration is due to redistribution of the
relatively few displaced dye molecules to vacant binding sites in regions of
duplex DNA which have not yet been unwound. Clearly, this non-linear behavior of the fluorescence signal at these low dye concentrations
precludes use of these dye concentrations as direct indicators of DNA unwinding. Furthermore, the observed reduction in the
fluorescence, as a fraction of the initial value (which is related to the
apparent extent of DNA unwinding), is apparently less at very low dye
concentrations. This artifact is not attributable to binding of dye molecules
to the walls of the cuvette, for example, because the addition of dye at either
the beginning or the end of the reaction results in the same fluorescence
signal (data not shown). This phenomenon, instead, can be attributed to the
rebinding of dye molecules to regions of secondary structure in the unwound
ssDNA. Such rebinding would contribute a greater percentage of the total
fluorescence difference at low dye concentrations and would therefore account
for a lower apparent extent of unwinding. In contrast to the limitations of
data obtained at low dye concentrations, at higher dye concentrations (>50 nM),
both the observed rate and the apparent extent of DNA unwinding are, as
expected, independent of dye concentration (Fig.
3
A).
RecBCD enzyme helicase activity demonstrates saturation behavior with regard to
DNA concentration. The apparent rate of unwinding increases with enzyme
concentration until all of the dsDNA ends present are bound by active enzyme (
28
). The protein concentration at which saturation is achieved can be used to
determine the apparent binding stoichiometry (i.e. the number of enzyme
molecules/end), defined as the concentration of protein just at saturation
divided by the concentration of DNA ends in the reaction. If the presence of
fluorophores does not affect the interaction of RecBCD enzyme with the ends of
the DNA substrate, this value should be invariant.
In Figure
4
A, raw data from an experiment in which RecBCD enzyme concentration was varied
at a given concentration of TO are shown; SSB protein was included to allow
comparison with previous results. From results such as these, using either
DAPI, H33258 or TO, the data in Figure
4
B were generated. As is readily apparent, the enzyme concentration at which the
reaction saturates does not vary significantly among the dyes. For DAPI,
saturation is achieved at a protein concentration of 7.8 nM. Given that the
concentration of DNA ends in the reaction is 2.1 nM, this corresponds to an
apparent binding stoichiometry of 3.7 +- 0.5 enzyme molecules/end. Similarly, protein titrations using H33258 and TO yield apparent binding stoichiometries of 3.5 +- 0.5 and 3.5 +- 0.5 molecules/end, respectively. For comparison, the
SSB protein-fluorescence assay (in the absence of dyes) yields kinetic traces that are
indistinguishable from those obtained from the dye-displacement assay and which yield an apparent binding stoichiometry of
3.1 +- 0.3 molecules/end (data not shown). Thus, dye molecules bound to the
substrate do not affect the apparent binding stoichiometry of RecBCD enzyme to
dsDNA ends.
Enzyme titrations also yield
V
max
for unwinding in the presence of each dye. The value obtained using the SSB protein fluorescence-quenching-assay is 250 +- 25 nM bp/s (data not shown). For DAPI, H33258 and TO, the
observed rates of unwinding under these conditions are 180 +- 20, 221 +- 25 and 211 +- 15 nM bp/s, respectively. Thus, at 20oC, the
k
cat
for DNA unwinding is 119 bp unwound/functional enzyme molecule/s as measured by
the SSB protein intrinsic fluorescence assay, whereas for the DAPI, H33258 and TO assays, it is 86, 105 and 100 bp unwound/functional enzyme molecule(s),
respectively. It should be noted that because the DAPI and H33258 reactions
were performed at dye concentrations which gave less than maximal rates of unwinding (see Fig.
3
B), the values for these dyes are underestimates. When higher concentrations of
these dyes were used (400 nM DAPI or 300 nM H33258), the rates are, within
experimental error, equal to those obtained using the SSB protein fluorescence-quenching-assay (data not shown); the rate in the presence of TO is ~16% lower.
Accurate measurement of DNA unwinding rates of saturating enzyme concentrations
is difficult, even at 20oC, due to the rapid rates of unwinding. To confirm that the inclusion of
these dyes did not substantially affect unwinding, reactions were performed at low enzyme concentration, in the presence or absence of dye, and the apparent
rate of unwinding was monitored by the SSB protein-fluorescence assay (data not shown). As expected, the time course of
unwinding as measured by SSB protein-fluorescence in the presence of dye (e.g. 250 nM H33285) was identical,
within experimental error, to that obtained when dye fluorescence was monitored
in the presence of SSB protein. The addition of H33258 to a final concentration
of 100 or 250 nM did not affect the observed rate of unwinding, whereas the
addition of TO had a slight effect (~20% decrease at 250 nM dye; Table
1
and see above).
Table 1
By performing the unwinding assay at different temperatures, the apparent energy of activation (
E
a
) can be determined. This parameter was determined for reactions containing
DAPI, H33258 or TO in the presence of SSB protein. Using data from reactions performed at 20, 25, 32 and 37oC, Arrhenius plots were generated (data not shown). The slopes of these
plots yield
E
a
values of 12.7 +- 1.5, 9.9 +- 2.4 and 9.7 +- 2.5 kcal/mol for DAPI, H33258 and TO, respectively. The
values for H33258 and TO are equivalent to that determined using the SSB
fluorescence assay (9.7 kcal/mol;
28
), whereas that for DAPI is somewhat greater but still within the experimental
error of these data.
We have devised a helicase assay that is amenable to kinetic studies and that
does not rely upon the presence of additional protein components. With this
assay, helicase activity can be measured by following the change in fluorescence upon displacement of nucleic acid-binding dyes. When the results of this assay are compared with those from
the SSB protein fluorescence-quenching assay, it is found that the basic enzymatic parameters, such as
the observed rate of unwinding and the apparent binding stoichiometry of RecBCD
enzyme to duplex DNA ends, as well as the energy of activation, are largely
unaffected for a subset of these dyes.
This assay could be broadly applicable for measurement of unwinding activity of a
variety of helicases, regardless of substrate requirements, provided that a
suitable, non-inhibitory dye ligand is selected. Since the choice of a non-inhibitory dye appropriate for use with a new helicase will be
empirical, we have surveyed the characteristics of several candidate dyes.
(Even if the presence of these ligands results in some inhibition of unwinding,
it should be possible to calibrate the dye displacement assay against another,
more direct type of helicase assay.) Our assay should be adaptable to helicases
which require ssDNA tails for initiation, even when the concentration or length
of the duplex region is low, because the fluorescence enhancement upon binding
duplex nucleic acids of several of these dyes is so great. Another advantage is
that the effect of ssDNA binding proteins on unwinding reactions can be easily
examined. Using RecBCD enzyme, we found that in the absence of SSB protein, a
decrease in dye fluorescence is still observed (data not shown), despite the
potential for reannealing of the ssDNA. The unwinding rates obtained in the
absence of SSB protein are somewhat higher (25-50%) than those obtained in its presence, as measured by the fluorescent
dye-displacement assay. This effect was unanticipated, since the rate of DNA
unwinding as measured by substrate disappearance by agarose gel electrophoresis reveals no significant difference regardless of whether SSB protein or dye is present (data not
shown). Unwinding reactions monitored by fluorescent dye-displacement and performed in the absence of SSB protein, however, do
display a linear dependence of apparent unwinding rate over a broad range of
subsaturating helicase concentration, yield a constant
V
max
at saturating RecBCD enzyme concentration, and exhibit the same stoichiometry
of binding of the enzyme to DNA ends (data not shown). These features allow
qualitative data to be obtained in the absence of SSB protein, although the
apparent rate values may vary somewhat in magnitude from those obtained in its presence. At present, we cannot provide a
ready explanation for the quantitative difference observed for the two assays
(fluorescent and electrophoresis) when performed in the absence of SSB protein, although each measures different
facets of the unwinding process. Preliminary experiments suggest that the addition of a ssDNA-specific nuclease (e.g. P1) may help to eliminate this apparent
discrepancy in the measured rates. That is, the rates in the presence of P1
nuclease and the absence of SSB approach the lower rates observed only in the
presence of SSB (unpublished observation). Finally, although not demonstrated here, this assay
can be adapted to measure the processivity of helicase action. Specifically,
such assays can be performed if a suitable trapping agent such as heparin is
added to confine the fluorescent signal to the results of a single round of
unwinding (
46
).
To be generally useful, an important premise of the dye-displacement assay is that the ligands do not inhibit the activity of the helicase
under study. The potential generality of this requirement is substantiated by
the following observations. First, although only results with RecBCD enzyme are
reported herein, we have also determined that the helicase activity of the RecQ
protein can be measured by this assay (unpublished observations). Secondly, the
fact that we observed no significant inhibition of RecBCD enzyme helicase
activity in the presence of some of these dyes is consistent with studies by
George
et al
. which used other DNA helicases (
36
). Using the fragment-displacement assay, the effect of various DNA-interacting ligands on the ATPase and unwinding activities of
several helicases, including
E.coli
helicase I, helicase II (UvrD) and Rep protein, was examined. They found that,
in general, DNA-binding ligands which occupy the minor groove of duplex DNA, such as AO,
DAPI and H33258, do not greatly affect the activities of those enzymes (
K
i
~1-10 [mu]M). It should be noted that the
K
i
values determined by George
et al
. are significantly greater than the concentrations of dyes we have utilized (<400 nM). In contrast, those authors found that ligands such as mitoxantrone,
which not only intercalate but also position functional groups within the major
groove of DNA, display potent but differential inhibition which is dependent
upon both the ligand and the enzyme. Other DNA ligands which inhibit helicase
activity include the antitumor drug CC-1065 (
29
,
47
,
48
) and anthracycline antibiotics (
49
). Thus, since the dyes we have focused upon - DAPI, H33258 and TO - belong to the former class of non-intercalating ligands which typically do not inhibit a representative set
of helicases, it is expected that they will be generally useful for this type
of assay. As mentioned above, given both the variety of fluorescent dyes that
could conceivably be used and the ease of the assay, it would be a simple task
to initially survey a few of these dyes with a new helicase to determine which
display little or no inhibition of activity.
Though our studies have focused on DNA helicases, the dye displacement assay may
provide a new means by which the unwinding activity of RNA helicases can be
examined. The measurement of RNA helicase activity has until now relied upon variations on
the fragment displacement assay, but our results suggest that fluorescent
detection of unwinding is also possible. Even though it is an intercalator, EB
may be expected not to have an inhibitory effect on helicase function, based
upon the results of George
et al
. (
36
); indeed, we have determined that EB, which has a relatively high fluorescence
enhancement upon dsDNA (Fig.
2
B), can also be used to measure DNA unwinding (unpublished observation). Since
this dye binds to RNA in addition to DNA, it is readily conceivable that RNA
helicases may be amenable to this assay if an appropriate ligand, such as EB
or, perhaps, propidium iodide, is utilized.
Another application for which this approach may be useful is in the measurement
of DNA synthesis. A variation of the SSB protein-fluorescence assay (
28
) has been used to monitor the synthetic activity of
E.coli
DNA polymerase III (
50
). In this adaptation, an increase in fluorescence signal due to displacement of
SSB protein from ssDNA templates indicates that duplex DNA has been
synthesized. It is not unreasonable to propose that a fluorescent dye assay
could be developed which would accomplish a similar purpose; in this instance,
the binding of dye to dsDNA would provide a real-time measurement of DNA synthesis. It is also possible that the same
reasoning could be used to develop a spectrophotometric assay to measure the
degradation of duplex DNA by nucleases.
In conclusion, the dye-displacement assay can be adapted for use with any helicase, whether it
utilizes a DNA or RNA substrate, provided that a suitable, non-inhibitory nucleic acid binding dye is selected; though selection is
empirical, many choices exist. In our experience, the most important parameter
to optimize is the concentration of dye: concentrations which are too low are subject to artifacts due to rebinding of displaced dye molecules to regions which have not been unwound, whereas concentrations which are too high will be
inhibitory to enzyme function. Once this parameter is optimized for a
particular substrate and enzyme, the dye-displacement assay gives results which are consistent with those obtained
from other types of assays.
We wish to thank members of our laboratory, particularly Jim New, Bill Rehrauer
and Cliff Ng, for their input and criticisms, and Mr Burt Houtz of Becton-Dickinson for providing a sample of thiazole orange. A preliminary account
of this work was presented at the FASEB conference `Genetic Recombination and
Genome Rearrangements' held at Copper Mountain, Colorado on July 25-30, 1993. These studies were submitted in partial fulfilment of the
doctoral requirements of Northwestern University (
51
). This work was supported by funds from the National Institutes of Health grant
GM-41347.
+
Present address: Imperial Cancer Research Fund, Clare Hall Laboratories, South
Mimms, Herts, EN6 3LD, UK
SSB protein
Fluorescent dye
Rate
(nM bp/s)
+
none
40.7 +- 4.1
+
H33258 (100 nM)
36.1 +- 4.5
+
H33258 (250 nM)
35.5 +- 1.9
+
TO (100 nM)
35.6 +- 1.3
+
TO (250 nM)
31.9 +- 0.7
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