Peculiar 2-Aminopurine Fluorescence Monitors the Dynamics of Open
Complex Formation by Bacteriophage T7 RNA Polymerase*
Rajiv P.
Bandwar and
Smita S.
Patel
From the Department of Biochemistry, Robert Wood Johnson Medical
School, Piscataway, NJ 08854
Received for publication, December 14, 2000
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ABSTRACT |
The kinetics of promoter binding and open complex
formation in bacteriophage T7 RNA polymerase was investigated using
2-aminopurine (2-AP) modified promoters. 2-AP serves as an ideal probe
to measure the kinetics of open complex formation because its
fluorescence is sensitive to both base-unpairing and base-unstacking
and to the nature of the neighboring bases. All four 2-AP bases in the TATA box showed an increase in fluorescence with similar kinetics upon
binding to the T7 RNA polymerase, indicating that the TATA sequence
becomes unpaired in a concerted manner. The 2-AP at
4 showed a
peculiarly large increase in fluorescence upon binding to the T7 RNA
polymerase. Based on the recent crystal structure of the T7 RNA
polymerase-DNA complex, we propose that the large fluorescence increase
is due to unstacking of the 2-AP base at
4 from the guanine at
5,
during open complex formation. The unstacking may be a critical event
in directing and placing the template strand correctly in the T7 RNA
polymerase active site upon promoter melting for template directed RNA
synthesis. Based on equilibrium fluorescence and stopped-flow kinetic
studies, we propose that a fast form of T7 RNA polymerase binds
promoter double-stranded DNA by a three-step mechanism. The initial
collision complex or a closed complex, EDc is formed
with a Kd of 1.8 µM. This complex
isomerizes to an open complex, EDo1, in an energetically
unfavorable reaction with an equilibrium constant of 0.12. The
EDo1 further isomerizes to a more stable open complex, EDo2, with a rate constant around 300 s
1. Thus, in the absence of the initiating
nucleotide, GTP, the overall equilibrium constant for closed to open
complex conversion is 0.5 and the net rate of open complex formation is
nearly 150 s
1.
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INTRODUCTION |
The bacteriophage RNA polymerases, such as the T7 RNA polymerase,
are single subunit enzymes capable of catalyzing all the processes such
as initiation, elongation, and termination of transcription. The
structural simplicity and the template specificity of the phage RNA
polymerases make them attractive for exploring the mechanism of
transcription and for understanding transcription regulation at the
level of protein-DNA interactions. Various crystal structures of T7 RNA
polymerase as a complex with the promoter DNA and initiating nucleotides are available, and these inspire detailed studies to
understand the dynamics of the transcription initiation and elongation
mechanisms (1, 2).
The initiation of transcription is a multistep process that directs the
polymerase to the promoter region, where RNA synthesis is initiated.
After the RNA polymerase recognizes a consensus DNA sequence, the
specific binding energy is used to melt a region of the promoter, part
of which serves as a template for the initiation of RNA synthesis (3).
Several studies including the crystal structures show that T7 RNA
polymerase recognizes a consensus sequence that extends from
17 to +4
relative to the transcription start site. In the absence of the
initiating nucleotide, GTP, T7 RNA polymerase melts
4 to +2 region,
which includes the TATA sequence, and in the presence of GTP, the
unpaired region extends from
4 to +4 (4). Our goal in these studies
was to determine the kinetic and thermodynamic parameters that govern
the formation of the closed and open complexes. These studies provide
the framework to understand how the efficiency of initiation can be
regulated by protein-DNA interactions during initiation.
Previously, we have used the fluorescent adenine analog
2-AP,1 which was incorporated
in the promoter DNAs, to monitor the dynamics of T7 RNA polymerase
interactions with the T7 promoters (5). These studies were carried out
with the promoter DNAs in which all five adenines in the
4 to +4
region were substituted with 2-AP. These previous studies had indicated
that T7 RNA polymerase exists in two forms (5). The fast form binds
dsDNA at close to a diffusion-limited rate and melts the promoter at a
fast rate (5). The slow form is observed only under excess DNA
concentrations, and the kinetics of open complex formation are limited
by the conversion of the slow form to the fast form. At that time, two other studies reported the kinetics of open complex formation in T7 RNA
polymerase using 2-AP fluorescence (6, 7). Our conclusions were similar
to one study in the literature (7), and it was proposed that open
complex formation occurred very rapidly, but a second study indicated
much slower rates of open complex formation (6). In the present paper,
we have systematically substituted each of the adenines in the
4 to
+4 region with 2-AP, and measured the kinetics of the fast form of the
T7 RNA polymerase with these DNAs. The fluorescence stopped-flow
studies have provided important structural insights and a more detailed
multistep mechanism of the initiation process, including the rate
constant and equilibrium constant of promoter opening and the
associated conformational changes.
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MATERIALS AND METHODS |
Protein--
T7 RNA polymerase was isolated and purified
according to established procedures as described in our earlier papers
(5, 8, 9). The purified T7 RNA polymerase was found to be >95% pure
by Coomassie staining of protein resolved by SDS-polyacrylamide gel
electrophoresis. The purified T7 RNA polymerase was stored at
80 °C in a buffer consisting of Na3PO4 (20 mM) pH 7.7, Na3-EDTA (1 mM),
dithiothreitol (1 mM), NaCl (100 mM), and
glycerol (50% v/v). The protein concentration was determined by
measuring the absorbance at 280 nm in 8 M urea using the
molar extinction coefficient of 1.4 × 105
M
1
cm
1.
Synthesis and Purification of 2-AP Incorporated
Oligonucleotides--
Oligodeoxynucleotides of T7
10 promoter
sequence containing 2-AP were custom synthesized by Integrated DNA
Technologies (Coralville, IA) and supplied as desalted samples. The
desalted oligonucleotides were further purified by denaturing 16%
polyacrylamide gel electrophoresis. Upon electrophoresis, the major
band corresponding to the DNA of interest was visualized by UV
shadowing and excised. The DNA was extracted from the gel by
electroelution using an ELUTRAP® electro-separation system
from Schleicher & Schuell. The electroeluted DNA was precipitated using
ethanol, dried, and resuspended in deionized water. The concentration
of the stock solution was determined by measuring the absorbance at 260 nm and using the integrated extinction coefficients of the bases at 260 nm: dA = 15,200, dT = 8,400, dG = 12,010, dC = 7,050 and 2-AP = 1,000 M
1
cm
1. The template and non-template ssDNAs
were annealed as reported previously (8), to yield the synthetic T7
promoter dsDNA containing only one 2-AP either on a template or
non-template strand.
Absorption and Emission Spectrophotometric Studies--
The
absorption spectra were measured using an HP8452A diode-array
spectrophotometer. The solutions (1 µM) of the free base 2-AP riboside and DNA (both normal and single 2-AP incorporated, single-stranded and double-stranded) were prepared in the buffer (50 mM Tris acetate, pH 7.5, 50 mM sodium acetate,
10 mM magnesium acetate, 5 mM dithiothreitol).
The fluorescence excitation and emission spectra were measured at
25 °C on a FluoroMax-2 spectrofluorometer (Jobin Yvon-Spex Instruments S.A., Inc.) using the DataMax program for spectra acquisition and analysis. The samples were excited at 315 nm and the
fluorescence emission spectra were collected in the wavelength range
325-425 nm. The fluorescence intensity of each of the singly 2-AP
incorporated ssDNA (1 µM), dsDNA (1 µM),
and dsDNA (1 µM) with T7 RNA polymerase (4 µM) equilibrated for 90 s, was measured at 370 nm
upon excitation at 315 nm. The fluorescence of the p-dsDNA with 2-AP at
t(
4) and t(
2) was also measured in a similar manner. The
fluorescence intensity of T7 RNA polymerase (4.0 µM) was
also measured in the same assay. After subtracting the fluorescence contribution of buffer from the DNAs (ssDNA, dsDNA, and p-dsDNA) and T7
RNA polymerase, the resulting fluorescence
(Fobs) was divided by respective concentrations
(x) to obtain a fluorescence coefficient (µM
1)
F(x) for T7 RNA polymerase and each
2-AP DNA.
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(Eq. 1)
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Similarly, the fluorescence coefficient,
F(ED), for each of the T7 RNA polymerase-DNA
complex was calculated after correcting for the fluorescence
contribution from all unbound species at equilibrium.
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(Eq. 2)
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F(E) and F(D) are
the fluorescence coefficients of the T7 RNA polymerase and DNA,
respectively, as mentioned above, and [E]free and
[D]free are the corresponding concentrations of the
unbound T7 RNA polymerase and DNA at equilibrium. The concentrations of ED for T7 RNA polymerase complexes of dsDNA and p-dsDNA were determined to be 0.90 and 1.0 µM, respectively, at the T7 RNA
polymerase and DNA concentrations mentioned above, and considering the
respective Kd values of 0.83 µM and 2 nM measured from nitrocellulose-DEAE filter binding
experiments.2
Stopped-flow Kinetics--
The stopped-flow experiments were
carried out using SF-2001 stopped-flow spectrophotometer equipped with
a photomultiplier detection system from KinTek Corp. (Austin, TX). The
T7 RNA polymerase and the singly-labeled 2-AP promoter DNA (40 µl
each) in the buffer described above were rapidly mixed from separate
syringes at a flow rate of 6.0 ml/s. The sample was irradiated
continuously with light of wavelength 315 nm from a high pressure xenon
compact arc lamp sampled through the excitation monochromator with a
slit width of 5 mm. The progress of the reaction was monitored from the
time dependent changes in the fluorescence emission intensity of 2-AP,
detected by photomultiplier tube at
360 nm using a cut-off
filter (WG 360, serial no. 273129 from Hi-Tech Scientific). The
photomultiplier tube high voltage was kept constant throughout the
experiment, and the kinetic traces from 10-15 experiments were
collected and averaged to optimize the signal.
Kinetic Data Analysis--
The kinetic traces obtained under
pseudo-first-order conditions from the average of 10-15 traces in each
experiment were fit to a single exponential.
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(Eq. 3)
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F is the fluorescence signal (observed at
360 nm) at time t (seconds); A and
kobs correspond to amplitude (i.e.
the change in fluorescence intensity from the beginning to the end of
the phase), and observed rate constant of the phase respectively, and
C indicates the background fluorescence intensity at
t = 0. The pseudo-first-order rate constants thus
obtained were plotted as a function of the T7 RNA polymerase
concentration, and the corresponding plots were fit by non-linear
regression analysis, using SigmaPlot from Jandel Scientific.
In order to interpret the stopped-flow fluorescence kinetic data in
terms of a DNA binding mechanism, a global nonlinear least-squares fitting was performed using the software Scientist from MicroMath Scientific Software (Salt Lake City, UT). A DNA binding model was
selected, and a set of differential equations was written for each of
the kinetic species in the mechanism. One such set of equations was
written for every concentration of the T7 RNA polymerase and DNA.
During the global fitting, individual sets were distinguished by a
letter suffix for each T7 RNA polymerase concentration. The
stopped-flow fluorescence kinetic traces were fit directly by assigning
the observed fluorescence, F(t) (the dependent variable) at
any given time (the independent variable), as the sum of the background
fluorescence, Fb, which is free DNA and protein
fluorescence, and the fluorescence of each ED species in the mechanism
(EDi), as shown in Equation 4.
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(Eq. 4)
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Fi is the specific fluorescence of each
ED species, Fb is the background fluorescence
due to buffer and free protein. EDi is the amount of each T7 RNA polymerase-DNA species at any given time,
which changes during the time course of DNA binding. The initial
estimates for the rate constants (as parameters) during global fitting
of the kinetic data were obtained from the quantitative analysis of the
observed rate constants versus T7 RNA polymerase concentrations. The process of global fitting involved first, fixing
only one set of intrinsic rate constants and determining the specific
fluorescence values (Fi) (as parameters) for
various sets of T7 RNA polymerase concentrations. Subsequently, one of the parameter sets (either rate or fluorescence value) was fixed, and
global fitting was used to optimize the other floating parameter set.
Eventually, all the parameters were floated to fit the data to a single
mechanism. The fitting process was governed by a modified Marquard-Levenberg algorithm making use of the analytical Jacobian matrix. The quality of the fit was judged by visual examination of
overlays of the fitted curves and the data as well as inspection of the residuals.
 |
RESULTS |
Fluorescent Properties of the 2-AP-Containing
10 Promoter
DNAs--
2-Aminopurine was incorporated at specific positions in the
template (t) or the non-template (nt) strands of the 40-bp DNA, with
the consensus sequence of the
10 promoter from
21 to +19 (Fig.
1). Five DNAs were prepared in which the
individual adenines at positions nt(
1), nt(
3), nt(+4), t(
2),
t(
4) were substituted with 2-AP. The absorption and emission spectra
of these DNAs were measured both in the ssDNA and dsDNA, and compared
with one another and to the free 2-AP riboside. The absorption spectra
of the DNAs were identical, exhibiting a strong peak at 260 nm,
corresponding to the absorption of normal bases, and a shoulder in the
region 305-315 nm, corresponding to 2-AP absorption. The absorbance
ratio (A315/A260) was
consistent with the presence of a single 2-AP in the 40-mer ssDNA. Upon
excitation at 315 nm, the 2-AP containing ssDNAs showed a fluorescence
spectra, with a peak at 370 ± 1 nm. Each of the ssDNAs showed
varying emission intensities (Fig. 1, lower left
panel), and the fluorescence was quenched when the ssDNAs
were annealed to the complementary strand. On an average, the
fluorescence of free 2-AP base was quenched ~95% upon incorporation into ssDNA, and an additional 30-90% quenching was observed when the
ssDNA was converted to dsDNA (Fig. 1, lower left
panel). Therefore, the 2-AP fluorescence is sensitive to the
structure of the DNA, with the fluorescence of 2-AP being higher in the
ssDNA versus the dsDNA.

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Fig. 1.
Positional effect of the 2-AP fluorescence in
the T7 10 promoter and the T7 RNA
polymerase-DNA complex. The 2-AP base was incorporated at the
indicated positions in the 40-bp dsDNA T7 10 promoter (consensus
sequence from 21 to +19). The number in
parentheses refers to the base position relative to the
transcription start site at +1, shown in boldface
letters. The fluorescence coefficients of the singly 2-AP
modified DNAs in the ssDNA, dsDNA, and in the T7 RNA polymerase-DNA
complex (ED) are listed in lower left
panel, and the corresponding values are plotted in a bar
chart (lower right panel).
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Our studies also showed that the 2-AP fluorescence in ssDNA was
dependent on the neighboring bases. The fluorescence intensity of 2-AP
was the least at position nt(+4) greatest at nt(
3) and t(
2)
positions, and intermediate at nt(
1) and t(
4) positions (Fig. 1,
lower left panel). Upon examination of
the DNA sequence (Fig. 1), we found that when the 2-AP base was flanked
by two guanines, the fluorescence was the least (at position nt(+4)). When 2-AP was adjacent to one guanine, the fluorescence was
intermediate (at position nt(
1) and t(
4)), and when the flanking
bases were not guanine, then the fluorescence was the highest (at
position t(
2) and nt(
3)). Thus, guanine residues quench the
fluorescence of 2-AP, especially when the 2-AP is stacked next to the guanine.
Fluorescent Properties of the T7 RNA Polymerase-DNA Complex at
Equilibrium--
Each of the five dsDNA promoters (1 µM)
containing 2-AP at the various positions was incubated with T7 RNA
polymerase (4 µM), under these conditions, most of the
dsDNA should be bound to the T7 RNA polymerase. Upon complex formation,
an increase in the fluorescence intensity of 2-AP was observed with all
dsDNAs, except the dsDNA with the 2-AP at nt(+4) (Fig. 1). Because the
increase in the 2-AP fluorescence is most likely due to base-unpairing or base-unstacking or both, these results indicate that the +4 position
is not melted in the open complex formed in the absence of the
initiating GTP. The fluorescence of 2-AP at nt(
1), nt(
3), and
t(
2) positions was 1.2-1.7 times lower than the respective ssDNA
fluorescence. Thus, either the bases at these positions were not
completely unpaired in the open complex or, if they were completely
unpaired, their fluorescence was quenched in the ED complex. A
peculiarly large increase in the fluorescence of 2-AP at t(
4) was
observed in the ED complex. The fluorescence of 2-AP at t(
4) was
nearly 4 times higher than the free ssDNA fluorescence. This indicated
a unique conformation of the 2-AP base at t(
4) in the ED complex. The
fluorescence of 2-AP at t(
4) increased about 41-fold from the dsDNA
fluorescence upon ED formation, compared with the 3-6-fold increase
that was observed in the other three positions.
The peculiar increase in the fluorescence of 2-AP at t(
4) was also
observed in the p-dsDNA (Fig. 4). The p-dsDNA was prepared by annealing
a 17-mer non-template strand (
21 to
5) to the 40-mer template
strand (
21 to +19), with 2-AP at either t(
4) or t(
2) position.
Although the 2-AP at t(
4) is already unpaired in the p-dsDNA, an
8-fold increase in fluorescence was observed when the p-dsDNA was bound
to T7 RNA polymerase. We have rationalized these results based on the
crystal structure of the ED complex and the fluorescent properties of
the 2-AP base. The crystal structure showed that the adenine at t(
4)
was flipped out and unstacked from the adjacent guanine residue at
t(
5) in the ED complex (1). Because the guanine quenches the
fluorescence of the stacked 2-AP base, the unstacking of 2-AP from the
guanine results in a large increase in fluorescence relative to its
fluorescence in the uncomplexed dsDNA. Thus, the increase in
fluorescence of 2-AP at t(
4) in the complexed dsDNA is both due to
base-unpairing and base-unstacking. Whereas in the p-dsDNA promoter
that is premelted, the increase in fluorescence must be due to
base-unstacking alone.
Stopped-flow Kinetics Reveals the Pathway of Open Complex
Formation--
Previous stopped-flow kinetic studies have shown that
there are two forms of T7 RNA polymerase (5). The fast form binds and
melts the dsDNA promoter with a relatively tight Kd and with fast kinetics. At high DNA concentrations and under limiting polymerase conditions, the slow form converts to the fast form at a
relatively slow rate. However, under conditions of excess polymerase
over DNA, the DNA binding kinetics from the slow form is not observed.
We are still investigating the significance of the two forms of the
polymerase; hence, the experiments in this paper were designed to
measure the kinetics and equilibrium of only the fast form of T7 RNA polymerase.
To measure the kinetics of DNA binding and strand separation at each of
the four positions in the TATA box, the stopped-flow kinetic studies
were carried out with the four 2-AP-modified dsDNAs. T7 RNA polymerase
(0.4 µM) was mixed with the 2-AP dsDNA (0.1 µM), and the fluorescence at
360 nm was monitored
as a function of time with continuous excitation at 315 nm. All four
2-AP-modified dsDNA promoters showed a time-dependent
increase in fluorescence that fit to a single exponential (Fig.
2a). The observed rate of the
fluorescence increase was the same regardless of the position of the
2-AP within the TATA sequence. The amplitudes were different and
followed the same trend as observed in the equilibrium fluorescence measurements (Fig. 2b). A large fluorescence increase at
t(
4) was observed, and successively smaller changes were observed at nt(
3), t(
2) and nt(
1) positions.

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Fig. 2.
Kinetics of T7 RNA polymerase binding to 2-AP
modified promoter DNAs. a, stopped-flow kinetic traces
(dotted) show the time-dependent increase in the
fluorescence of 2-AP at t( 4), nt( 3), t( 2) and nt( 1) positions
in the 40-bp dsDNA promoter upon binding to T7 RNA polymerase. The
dsDNA promoter (0.1 µM) was mixed with T7 RNA polymerase
(0.4 µM, final concentrations) at 25 °C in a
stopped-flow instrument. The sample was excited at 315 nm, and the
time-dependent fluorescence was measured at 360 nm. Each trace is an average of 10-15 measurements, and the
solid smooth line is the fit to a
single exponential with a rate constant of 45 ± 12 s 1 (Equation 3). b, the amplitude
or the fluorescence change during the time course of the kinetic
measurement is shown for each of the four positions. The
black bars (left) represent the
amplitudes from the single exponential fits of the stopped-flow
kinetics (shown in a) relative to the t( 4) position. The
white bars (right) represent the
fluorescence change measured from equilibrium studies, shown in Fig.
1.
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To dissect each step in the kinetic pathway of DNA binding, the
stopped-flow experiments were conducted at varying T7 RNA polymerase
concentrations, with each of the dsDNA promoters. The resulting time
courses fit to a single exponential, and the observed rates increased
with increasing T7 RNA polymerase concentration in a hyperbolic manner.
In the previous studies, we had observed only a linear increase because
higher T7 RNA polymerase concentrations were not employed (5). The rate
versus T7 RNA polymerase concentration dependences were
similar for each of the four positions, and a representative kinetic
trace and the polymerase concentration dependence is shown for the
dsDNA with 2-AP at t(
4) (Fig. 3, a and b).

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Fig. 3.
Stopped-flow kinetics of T7 RNA polymerase
binding to dsDNA and p-dsDNA promoters. a, the dsDNA
promoter modified with 2-AP at t( 4) (0.05 µM) was mixed
with T7 RNA polymerase (0.4 µM) at 25 °C, and 10-15
traces were averaged to obtain the dotted trace
shown. The kinetic trace was fit to a single exponential
(solid line), and the corresponding residuals are
shown on top. b, the experiment in a
was repeated at varying [polymerase] at a constant [dsDNA] = 0.05 µM. At each concentration of T7 RNA polymerase, 10-15
traces were averaged and the observed rates were plotted against T7 RNA
polymerase concentration. The rate dependence was fit to the hyperbolic
equation (kobs = k 2 + k2*[polymerase]/(Kd + [polymerase]) with k2 = 152 ± 11 s 1, Kd = 1.9 ± 0.4 µM, k-2 = 6 ± 3 s 1. c, the T7 p-dsDNA promoter
modified with 2-AP at t( 4) (0.05 µM) was mixed with the
T7 RNA polymerase (0.4 µM) and the resulting kinetic
trace (dotted) is shown along with the fit to a single
exponential (solid line). The residuals from the
fit are shown on top. d, the experiment in
c was repeated at varying [polymerase] at a constant
[p-dsDNA] = 0.05 µM. The averaged kinetic trace at each
polymerase concentration was fit to a single exponential (Equation 3,
solid smooth line), and the observed
rates were plotted against T7 RNA polymerase concentration. The rate
dependence was fit to the hyperbolic equation with
k2 = 303 ± 31 s 1, Kd = 1.3 ± 0.6 µM, k 2 = 17 ± 23 s 1.
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The fluorescence increase during the time course of DNA binding
indicates that the ED species has a higher fluorescence relative to the
uncomplexed dsDNA. The single exponential increase indicates that only
one ED species is populated. However, the hyperbolic dependence of the
observed rate versus T7 RNA polymerase concentration indicates that there are at least two steps in the DNA binding pathway
and two ED species that are formed sequentially, as shown.
E is the T7 RNA polymerase, D is the dsDNA promoter, and
EDc and EDo1 are the two ED complexes. The
values of the intrinsic rate constants for the above mechanism were
obtained by fitting the dependence to the hyperbolic equation:
kobs = k
2 + k2*[polymerase]/(Kd + [polymerase]), which provided a Kd = 1.9 ± 0.4 µM, k2 = 152 ± 11 s
1, and k
2 = 6 ± 3 s
1. Similar values for the
Kd and the maximum rate constant (k2) were observed for all the dsDNA promoters
(data not shown). The kinetics indicate that the EDc
complex is in rapid equilibrium with free E and D. The EDo1
that results from the isomerization of EDc has a higher
fluorescence than the free dsDNA and makes a major contribution to the
observed fluorescence change. The observed rate, at low concentrations
of E, is limited both by the amount of EDc and by the rate
of EDc to EDo1 conversion (i.e. k2). At high concentrations of E, the observed
rate is limited by the rate of EDc to EDo1
conversion. Hence, the maximum rate (152 ± 11 s
1) provides an estimate of the
intrinsic rate constant, k2 for EDc to EDo1 conversion, the intercept of 6 ± 3 s
1 provides an estimate of
k
2, and the K1/2 an
estimation of the Kd for the EDc complex.
To understand the nature of the EDc and EDo1
complexes, we conducted stopped-flow studies with the 17/40 p-dsDNA
promoter. The p-dsDNA is premelted in the
4 to
1 region, and we
believe that the fluorescence increase in the p-dsDNA with 2-AP at
t(
4) originates mainly from base-unstacking. Thus, the kinetics of fluorescence change with the p-dsDNA promoter should provide the rate
of base-unstacking and enable us to isolate this step from the strand
separation or the base-unpairing step. A constant amount of 17/40
p-dsDNA (0.05 µM) was mixed with varying concentrations of T7 RNA polymerase (0.1-4.0 µM). The resulting
increase in fluorescence with time was fit to a single exponential
(Fig. 3c). The observed rates increased in a hyperbolic
manner as the T7 RNA polymerase concentration was increased (Fig.
3d). The hyperbolic fit provided a maximum rate constant,
k2, of 303 ± 31 s
1, Kd of 1.3 ± 0.6 µM, and k
2 of 17 ± 23 s
1. The observed kinetics are consistent with
the following two-step mechanism for DNA binding.
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pD represents p-dsDNA promoter. The above mechanism
indicates that the EpDc is in rapid equilibrium with E and
pD with a Kd similar to the dsDNA, indicating the
similar nature of the collision complexes with the two types of
promoters. The EpDc isomerizes to EpDo2 in the
p-dsDNA promoter at a rate close to 300 s
1, which is 2 times faster than the
isomerization rate of the dsDNA promoter. Although the TATA
sequence in the p-dsDNA promoter is premelted, the template strand
of the p-dsDNA still needs to bind within the active site pocket of
the T7 RNA polymerase in the manner shown by the crystal structure and
in Fig. 6 (1). The isomerization therefore represents a conformational
change in the collision complex that results in the open complex,
EpDo2. This conformational change is likely to be the
base-unstacking step in the p-dsDNA promoter, whereas, in the dsDNA
promoter, the conformational change likely involves both base-unpairing and base-unstacking reactions.
Extent of Open Complex Formation--
The overall equilibrium
constant of the two-step DNA binding mechanism (shown in Reaction 1)
derived from the stopped-flow kinetic studies indicated that most of
the closed complex, EDc, should be converted to the open
complex, EDo1, at equilibrium. We therefore expected the
dsDNA fluorescence to increase to the same level or higher than the
fluorescence of p-dsDNA in the ED complex. When we compared the
fluorescence values of the complexed dsDNA and p-dsDNA promoters, we
observed that the fluorescence of the complexed dsDNA promoter was
3-3.5 times lower than the fluorescence of complexed p-dsDNA (Fig.
4). The lower fluorescence of complexed
dsDNA can be explained in several ways. First, the template strand in
the premelted p-dsDNA may be in a conformation that is different from
the "polymerase-induced" conformation of the melted dsDNA promoter.
To see a larger fluorescence of the p-dsDNA in the ED complex, the
bases in the p-dsDNA would have to be unstacked to a larger extent
relative to the melted bases in the dsDNA. Note that our previous
studies have shown that the dsDNA and p-dsDNA promoters have similar
pre-steady state rates of RNA synthesis, suggesting that they form
kinetically similar open complexes (9). The second reason for the lower
fluorescence of the complexed dsDNA may be that the EDc
complex is not converted stoichiometrically to the open complex,
EDo1. In other words, the complexed dsDNA may exist as a
mixture of closed and open structures. Such an idea has been suggested
for T7 RNA polymerase based on steady state analysis of RNA synthesis
using dsDNA and p-dsDNA; however, the equilibrium constant was not
measured by direct means (10).

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Fig. 4.
The fluorescence of 2-AP in the p-dsDNA
and the T7 RNA polymerase-p-dsDNA complexes. The 2-AP base was
incorporated at the indicated positions in the 17/40 p-dsDNA promoter.
Lower left panel shows the
fluorescence coefficients of the DNAs modified with 2-AP at t( 4) and
t( 2) in the ssDNA, p-dsDNA, and in the T7 RNA polymerase-DNA
complexes (EpD). The lower right panel
compares the fluorescence coefficients of the p-dsDNAs with the dsDNAs
modified with 2-AP either at t( 4) or t( 2) positions.
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We propose that the equilibrium constant governing the closed to open
complex transition can be determined by comparing the fluorescence
values of the complexed p-dsDNA and dsDNA promoters. We assume that the
complexed p-dsDNA promoter is converted fully to the open complex, and
hence the fluorescence value of complexed p-dsDNA is a measure of the
maximum amount of open complex in the reaction. Equilibrium
fluorescence measurements were carried out with DNAs containing the
2-AP at t(
4) or t(
2) position in the p-dsDNA and dsDNA promoters,
and the fluorescence values are listed in Fig. 4 (lower
left panel). Each of these positions provided a
ratio between 29% and 36%, indicating that about 33% of the complexed dsDNA promoter was in the open form. The fact that both t(
2) and t(
4) provided a similar ratio of open to closed form indicates that this phenomenon is not peculiar to position t(
4). These data provide a ratio of open to closed form equal to
0.33/0.67 = 0.5, which is a measure of the overall equilibrium
constant for the EDc to EDo conversion in the
dsDNA promoter.
Three-step Mechanism of DNA Binding--
The minimal mechanism
that best describes the stopped-flow kinetics is the two-step DNA
binding mechanism (Reaction 1), but it predicts almost stoichiometric
conversion of the closed complex EDc to the open complex,
EDo1. The equilibrium fluorescence measurements, discussed
above, indicated that the EDc was not converted
stoichiometrically to EDo1, and the two species exist in
equilibrium. The equilibrium constant indicated that the formation of
EDo1 was governed by an unfavorable equilibrium, but a
two-step mechanism with an unfavorable equilibrium between
EDc and EDo1 did not fit the stopped-flow kinetic data. To account for the equilibrium between EDc
and EDo1 and to explain the stopped flow data, a third step
was introduced after EDc formation and before
EDo2 was formed, as shown below (Reaction 3).
EDc is a closed complex in equilibrium with the first
open complex EDo1 that isomerizes to EDo2.
Using numerical integration methods, we were able to globally fit the
stopped-flow kinetic data at various T7 RNA polymerase concentrations
to the three-step mechanism, as shown by the solid
lines in Fig. 5a.
The method of global fitting involved writing the differential
equations of the three-step model, which were solved by numerical
methods using the program Scientist, as described in more detail under "Materials and Methods." A single set of rate constants was
obtained from global fitting and shown in Table
I. The derived rate constants indicated
that the dsDNA promoter forms a closed complex EDc with a
Kd of 1.8 µM. The closed complex
isomerizes to an open complex EDo1 with a forward rate
constant (k2) of 208 ± 15 s
1 and a reverse rate constant of
(k
2) 1719 ± 132 s
1. Hence, there is an unfavorable
equilibrium of 0.12 for EDc to EDo1 conversion.
The EDo1 then isomerizes to a more kinetically stable
EDo2 species with a forward rate constant
(k3) of 322 ± 8 s
1 and a reverse rate constant
(k
3) of 54 ± 1 s
1. The same treatment of the p-dsDNA binding
kinetics (Fig. 5b) showed that the data fit only to a
two-step mechanism (Reaction 2), and the kinetic constants derived from
the global fit are shown in Table I. The p-dsDNA (pD) forms an
EpDc collision complex with a Kd of 0.2 µM, the EpDc isomerizes to EpDo2
with a forward rate constant (k2) of 222 ± 47 s
1 and a reverse rate constant
(k
2) of 0.2 ± 51 s
1.

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Fig. 5.
Global least squares fitting of the
stopped-flow kinetic data for the T7 RNA polymerase binding to dsDNA
and p-dsDNA. The averaged kinetic traces at various concentrations
of T7 RNA polymerase (constant [DNA] = 0.05 µM) are
represented as dotted traces, and the
solid lines show the global least squares fit to
the kinetic mechanisms, shown in Table I. The time axis is logarithmic.
a, each kinetic trace (total of 500 points) was collected on
a linear time scale in two time windows, with 350 points in the first
time window (0-0.1 s) and 150 points in the second time window (0.1-5
s). The least squares fit for the binding of dsDNA to the three-step
mechanism provided the following rate constants: Kd = 1.8 µM, k2 = 208 ± 15 s 1, k 2 = 1719 ± 132 s 1, k3 = 322 ± 8 s 1, k 3 = 54 ± 1 s 1. b, the kinetic
traces were collected on a logarithmic time scale. The global least
squares fit for the binding of p-dsDNA to the two-step mechanism
provided the following rate constants: Kd = 0.2 µM, k2 = 222 ± 47 s 1, k 2 = 0.2 ± 51 s 1.
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|
The overall equilibrium constant for EDc to
EDo2 conversion was calculated from the derived rate
constants of the three-step mechanism. This value of 0.7 is in close
agreement with 0.5, which was calculated by comparing the fluorescence
values of complexed dsDNA and p-dsDNAs. The percentage of
EDo complexes (EDo1 + EDo2) in the
dsDNA predicted from the three-step mechanism is 35%, in close
agreement to the amount (29-36%) calculated from the relative fluorescence values of the complexed dsDNA and p-dsDNA, under similar
experimental conditions. Note that the two methods for obtaining the
equilibrium constant are independent. The three-step mechanism also
predicts that the open complex EDo1 does not accumulate to
a significant level. Therefore, the formation of EDo1 was
not observed as a separate phase in the stopped-flow experiments. Based
on the available data, we speculate that the EDo1
represents a complex in which only the TATA sequence is melted and the
EDo2 may represent a complex in which the t(
4) base is
unstacked, the melted region extends to +2, and the template strand is
directed into the T7 RNA polymerase active site making interactions
with the active site residues.
 |
DISCUSSION |
The transient-state fluorescence measurements have provided the
detailed kinetics of the T7 RNA polymerase-promoter interactions. The
interactions were measured by following the fluorescence of the 2-AP
base, an analog of adenine that can be incorporated at specific
positions in the promoter DNA. The normal bases in the DNA have very
short decay times, typically few picoseconds, and as a result a very
weak intrinsic fluorescence of DNA (11, 12). This hampers the use of
normal DNA for studying DNA-protein interactions to monitor real-time
changes in the DNA within the confines of the protein-DNA complex.
However, substitution of a normal base by its modified structural
analog with fluorescent properties, without significantly altering the
native structure of DNA, provides an important handle to study real
time changes in the DNA. The 2-AP is a structural analog of adenine
that forms a stable Watson-Crick type base pair with thymine. The
2-AP:T base pair is only marginally weaker (0.5 kcal/mol) than a normal
A:T pair as reported for a decamer (13). The 2-AP substitution does not
destroy the B-helical structure of the dsDNA, and does not affect the
specific recognition by proteins, in most cases (14-16). The 2-AP can
be selectively excited at longer wavelengths (310-320 nm), in the
presence of tryptophan and tyrosine protein residues, and the 2-AP
fluorescence is highly sensitive to the structure of the DNA.
We have substituted four adenines in the TATA box preceding the
initiation site of the T7
10 promoter and one adenine in the coding
region individually to 2-AP. The absorption and fluorescence emission
studies indicated that, although the 2-AP at all positions absorbed
equally, their fluorescence intensity was dependent on the neighboring
bases in the DNA sequence. The fluorescence intensity was the minimum
when the 2-AP was flanked by guanines. The 2-AP fluorescence quenching
by guanine has been reported (17). The results in the literature showed
that the guanine caused static quenching of 2-AP fluorescence, when it
was immediately adjacent to 2-AP, and the quenching was reduced when
the guanine was placed at longer distances from 2-AP. Most effective
quenching occurred through strong
-stacking when G was located on
the same strand and adjacent to 2-AP. These results are in agreement
with our observations that the 2-AP fluorescence in the DNA is
dependent on the neighboring G residue. The fluorescence of 2-AP was
also sensitive to the structure of the DNA. The fluorescence of
H-bonded 2-AP at t(
4) position, for example, in the dsDNA promoter
was less than one-tenth the fluorescence in the ssDNA (Fig. 1).
Therefore, 2-AP can serve as a sensitive and site-specific probe to
monitor local melting of the dsDNA. In fact, 2-AP has been employed
successfully to monitor local melting of DNA bound to the Klenow
fragment of DNA polymerase I (18) and T7 RNA polymerase (6, 7) and base-flipping/unstacking in EcoRI DNA methyltransferases
(19), and bacteriophage T4 DNA polymerase (20).
Peculiar Increase of 2-AP Fluorescence at t(
4)--
The
fluorescence of all the 2-AP bases in the TATA box increased when the
dsDNA promoter formed a complex with T7 RNA polymerase. The 2-AP at
t(
4), however, showed a large increase in fluorescence that was
4-fold higher than the free ssDNA. This unusually large fluorescence
increase of 2-AP at t(
4) was noted previously (7), but the recent
crystal structure of the T7 RNA polymerase-DNA complex provides a
rationale for the large fluorescence increase specifically at position
t(
4) (1). The crystal structure of the T7 RNA polymerase-DNA open
complex shows that adenine at t(
4) is in a unique conformation
relative to other adenines. As shown in Fig.
6, the adenine at t(
4) is completely
unstacked from the neighboring guanine at t(
5), and such unstacking
is expected to cause a drastic increase in 2-AP fluorescence when
placed at this position. This proposal is also supported by the fact
that the t(
4) 2-AP shows an increase in fluorescence even in the
p-dsDNA promoter. Since the 2-AP in the p-dsDNA promoter is premelted, the fluorescence increase most likely arises from base-unstacking. We
rule out the possibility that specific interactions with the T7 RNA
polymerase such as hydrogen bonding or hydrophobic interactions increase the fluorescence of 2-AP at t(
4), because such interactions are predicted to decrease the 2-AP fluorescence (21).

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Fig. 6.
The structure of the
10 promoter DNA in the T7 RNA polymerase-promoter
complex. The coordinates for the DNA were obtained from the 2.4-Å
crystal structure of the complex (PDB identification code 1CEZ),
and shows the orientation of the bases in the template, t ( 17 to 1)
and non-template, nt ( 17 to 3) regions. The sequence is
schematically represented on the top. In the crystal structure, the
template and the non-template strands are paired from 17 to 5 and
melted from 4 to 1. The melted template strand is twisted between
5 and 4, and the four downstream bases ( 4 to 1) are directed
toward the polymerase active site in the palm. The bases 4 to 2 are
stacked but not paired and also flipped compared with the dsDNA region
upstream. The unpaired 4A (green) is unstacked from 5G
(purple), implying that the 2-AP at 4 might be similarly
unstacked from the neighboring G at 5, causing a large fluorescence
change.
|
|
Three-step Mechanism of T7 RNA Polymerase Binding to Promoter
DNA--
Each of the 2-AP bases in the TATA box showed an increase in
fluorescence upon binding to the T7 RNA polymerase with similar kinetics indicating that the bases in the TATA region melt in a
concerted manner during open complex formation. The simplest model that
explained the stopped-flow kinetic data was the two-step mechanism. In
this mechanism, the initial encounter of the T7 RNA polymerase with the
DNA occurs at close to a diffusion-limited rate to form a closed
complex EDc, which is in rapid equilibrium with free E and
D. The EDc isomerizes at an observed rate close to 150 s
1 to form an open complex. It was
interesting that the kinetics of the p-dsDNA promoter, which is
premelted from
4 onward, showed a two-step DNA binding mechanism. The
initial T7 RNA polymerase-p-dsDNA complex was formed similar to the
dsDNA, but the subsequent isomerization step (~ 300 s
1) was 2-fold faster in the p-dsDNA.
Comparison of the fluorescence values of the p-dsDNA and dsDNA in
the polymerase-DNA complex under conditions where the polymerase was in
excess of the DNA suggested that only 29-36% of the complex dsDNA
promoter was open. We therefore incorporated a third step after
EDc formation in the dsDNA binding mechanism. The kinetic data at various polymerase concentrations were globally fit to the
three-step mechanism, and the intrinsic rate constants are listed in
Table I. The kinetic constants indicate that the EDc complex resulting from the initial encounter of the T7 RNA polymerase with the dsDNA promoter has a Kd of 1.8 µM. This complex isomerizes to EDo1 with an
unfavorable equilibrium constant of 0.12, and subsequently to
EDo2 with a rate constant close to 322 s
1. A schematic representation of the
promoter binding and open-complex formation steps is depicted in Fig.
7. We postulate that, during the
formation of EDc, the T7 RNA polymerase makes specific
interactions with the conserved promoter recognition region from
17
to
5 sequence. The EDc complex then isomerizes to
EDo1, in which the promoter is melted from
4 to
1, but
the interactions of the template strand with the T7 RNA polymerase are
not extensive to stabilize the complex. The EDo1 is
therefore in equilibrium with the closed complex EDc. The
EDo1 then isomerizes to EDo2, in which the
template strand is directed within the active site of T7 RNA polymerase, making more extensive interactions, as seen in the recent
crystal structure of the ED complex (1). The EDo2 is therefore a preinitiation complex, in which the template bases up to +2
are melted to pair with the incoming initiating GTPs for RNA synthesis.
The three-step mechanism predicts an overall Kd for
dsDNA equal to Kd1/(1 + K2 + K2*K3) = 0.98 µM (where numbers in subscript refers to step), which is in agreement with the Kd determined from
fluorimetric titrations and nitrocellulose DNA binding experiments
(data not shown). Similarly, the overall Kd for
p-dsDNA is Kd1/(1 + K2) = 0.2 nM, which is similar
to the Kd of 0.3 nM measured by
equilibrium methods reported in our earlier paper (5).

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Fig. 7.
The proposed model of T7 RNA
polymerase-promoter binding and open complex formation during
transcription initiation. The three-step mechanism was determined
by global least squares fitting of the stopped flow fluorescence data.
The non-template strand of the dsDNA promoter (D) is shown in
blue and template strand in red, and the T7 RNA polymerase
(E) binding pocket in brown. The EDc is the
closed complex that exists in equilibrium with the open complex,
EDo1. The EDo2 represents a more stable, second
open complex. The part of the DNA in the EDo2 complex shown
by the solid line was constructed using the DNA
coordinates from the crystal structure of the initiation complex (PDB
identification code 1QLN). The dotted line is the
hypothetical extension of the two strands that illustrate the
transcription bubble during initiation, with +1 and +2 template bases
(black ellipses) aligned for specific base
pairing, respectively with the initiating and elongating
nucleotides.
|
|
In summary, the present studies have demonstrated a unique fluorescence
change arising from unstacking of the 2-AP base at the junction of
binding and melting regions of the promoter template strand, between
bases
4A and
5G. This unstacking may be a critical event in
directing and placing the template strand correctly in the T7 RNA
polymerase active site upon promoter melting for template directed RNA
synthesis. However, to keep the complexed promoter open in the absence
of the transcription reaction appears to be energetically unfavorable,
as seen from the incomplete conversion of the closed to the open
complex. Nevertheless, the observed rate of open complex formation
(~150 s
1) is not rate-limiting during
transcription initiation.
 |
ACKNOWLEDGEMENTS |
We thank members of the Patel laboratory for
helpful discussions and Natalie Stano and Mikhail Levin for help in
preparing the figures.
 |
FOOTNOTES |
*
This work was supported by National Institutes of Health
Grant GM51966 (to S. S. P.).The costs of publication of this
article were defrayed in part by the
payment of page charges. The article must therefore be hereby marked
"advertisement" in
accordance with 18 U.S.C. Section
1734 solely to indicate this fact.
To whom correspondence should be addressed: Dept. of Biochemistry,
Robert Wood Johnson Medical School, 675 Hoes Ln., Piscataway, NJ 08854. Tel.: 732-235-3372; Fax: 732-235-4783; E-mail: patelss@umdnj.edu.
Published, JBC Papers in Press, January 25, 2001, DOI 10.1074/jbc.M011289200
2
N. Stano and S. S. Patel, unpublished results.
 |
ABBREVIATIONS |
The abbreviations used are:
2-AP, 2-aminopurine;
17/40, 17-mer non-template/40-mer template;
bp, base pair(s);
ds, double-stranded;
ss, single-stranded;
p-ds, partially double-stranded;
nt, non-template;
t, template.
 |
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Copyright © 2001 by The American Society for Biochemistry and Molecular Biology, Inc.