One-step DNA melting in the RNA polymerase
cleft opens the initiation bubble to form
an unstable open complex
Theodore J. Griesa, Wayne S. Konturb,2, Michael W. Cappa, Ruth M. Saeckera,1, and M. Thomas Record, Jr.a,b,1
aDepartment of Biochemistry, University of Wisconsin, 433 Babcock Drive, Madison, WI 53706; and
1101 University Avenue, Madison, WI 53706
bDepartment of Chemistry, University of Wisconsin,
Edited* by E. Peter Geiduschek, University of California at San Diego, La Jolla, CA, and approved April 13, 2010 (received for review January 24, 2010)
Though opening of the start site (þ1) region of promoter DNA is re-
is known about how and when this occurs in the mechanism. Early
events at the λPRpromoter load this region of duplex DNA into
theactivesitecleftofEscherichiacoliRNAP, formingthe closed,per-
manganate-unreactive intermediate I1. Conversion to the subse-
quent intermediate I2overcomes a large enthalpic barrier. Is I2
is the same as the RPocontrol, whereas nontemplate (nt) thymines
are significantly less reactive than in RPo. We propose that: (i) the
þ1ðtÞ thymine is in the active site in I2; (ii) conversion of I2to RPore-
positions the nt strand in the cleft; and (iii) movements of the nt
strand are coupled to the assembly and DNA binding of the down-
stream clamp and jaw that occurs after DNA opening and stabilizes
RPo. We hypothesize that unstable open intermediates at the λPR
promoter resemble the unstable, transcriptionally competent open
complexes formed at ribosomal promoters.
bottleneck step ∣ transcription regulation ∣ burst experiment ∣
protein nucleic acid interactions
changes in both biomolecules. Taken together, these steps consti-
tute the mechanism of DNA opening and the start of the
transcription cycle. For Escherichia coli RNA polymerase holoen-
zyme (RNAP, subunit composition: α2ββ0ωσ70), binding free
energy drives opening of the initiation bubble (−11 to þ2, num-
bering relative to the start site base þ1) in promoter DNA,
placement of þ1 template base in the active site of the enzyme,
and subsequent conformational changes to form the stable open
complex RPo. Each of these steps provides a checkpoint for
Over the past decade, structural (X-ray, FRET), single-
molecule, and rapid mixing kinetic studies have greatly advanced
the understanding of this machinery and these steps. Key ad-
vances include: (i) elucidation of the RNAP architecture at
atomic resolution (1–4); (ii) dissection of composite forward
and backward rate constants for RPoformation into individual
rate and/or equilibrium constants for the steps leading to RPo
(5, 6); (iii) single-molecule measurements of DNA topological
changes (7); (iv) real-time determination of hydroxyl radial
(HO•) protection patterns of DNA during stable open complex
(RPo) formation (8–10); and (v) finding that unstable open
complexes are stabilized by binding the initiating nucleoside
triphosphate and greatly destabilized by the stress response fac-
tors ppGpp and DksA (11–13). These advances make it possible
to address the key unresolved questions of initiation. How is the
nteractions between RNA polymerase and specific promoter
DNA sequences trigger a precise progression of conformational
opening of 12–14 base pairs distributed between the steps of
RPoformation? Does RNAP disrupt the DNA duplex in the
active site cleft or does DNA melt outside the channel and enter
as individual strands?
Evidence for at least two kinetically significant intermediates
(generically designated I1and I2) preceding RPoexists for a
variety of promoters recognized by E. coli RNAP (cf. refs. 6,
8, and 14–17). Conversion of I1to I2is the rate-determining
(bottleneck) step in forming the open complex at the λPRpromo-
ter and exhibits a 34-kcal activation enthalpy barrier (17). The
reverse direction of this step is the bottleneck step in dissociation
of RPo(6). Because I1is a closed complex (9), determining when
DNA opens requires trapping I2.
The minimal mechanism of RPoformation is formally analo-
gous to minimal mechanisms of solute transport through mem-
branes and enzyme catalysis. All involve three steps in which the
initial step in each direction is rapidly reversible and a middle step
that is the bottleneck in both directions. In RPoformation, as in
mechanisms of catalysis and transport, ligands and solutes pri-
marily act on the rapidly reversible steps and not on the central
bottleneck step (6, 18). Given these analogies, is the central bot-
tleneck step of open complex formation indeed DNA opening,
just as transport is the central step in transporter mechanisms
and catalysis is in enzyme mechanisms?
We address this fundamental mechanistic question by using a
powerful method from physical enzymology, the burst experi-
ment, which forms a transiently high concentration of an other-
wise unobservable intermediate (preceding the bottleneck step).
In the forward direction, a burst of I1is generated by rapid mixing
with a sufficiently high [RNAP] (cf. Fig. 1A). HO• and perman-
DNA in such a forward burst experiment demonstrate that I1is a
protected to þ20 from HO• attack (9). Structural modeling on
the basis of these data and the X-ray structures of the bacterial
RNAP (1, 2) indicates that duplex DNA in I1is loaded in the
active site cleft of RNAP but not yet open (9).
Because the rate limiting forward step is the conversion of I1
to I2, no subsequent burst of I2occurs in the forward direction
experiment (Fig. 1A). Hence the dissociation direction must be
investigated to obtain a sufficient population of I2to characterize.
The time course of a standard dissociation experiment in which a
4) footprints of the population of the λPRpromoter
4-unreactive complex in which downstream duplex DNA is
Author contributions: T.J.G., W.S.K., R.M.S., and M.T.R. designed research; T.J.G., W.S.K.,
M.W.C., and R.M.S. performed research; T.J.G., R.M.S., and M.T.R. analyzed data; and
T.J.G., R.M.S., and M.T.R. wrote the paper.
The authors declare no conflict of interest.
*This Direct Submission article had a prearranged editor.
1To whom correspondence may be addressed. E-mail: firstname.lastname@example.org or mtrecord@
2Present address: Department of Bacteriology, Great Lakes Bioenergy Research Center,
University of Wisconsin, 1550 Linden Drive, Madison, WI 53706.
This article contains supporting information online at www.pnas.org/lookup/suppl/
10418–10423 ∣ PNAS ∣ June 8, 2010 ∣ vol. 107 ∣ no. 23www.pnas.org/cgi/doi/10.1073/pnas.1000967107
ible is shown in Fig. 1B. Because I2is an unstable intermediate,
rapidly converting back to the stable open complex on the time
scale of its conversion to I1, it never accumulates to a signifi-
cant level in this experiment. Kontur et al. (6) discovered that
rapid destabilization of the stable open complex with moder-
ately high concentrations of urea or salt generates a dramatic
transient buildup (burst) of I2∼0.5 s after mixing (Fig. 1C). (A
RPo-destabilizing temperature downshift cannot be performed
rapidly enough to detect such bursts.) Because the rate of con-
version of I2to I1is found to be independent of urea or salt
concentration, the burst of I2persists for a period of approxi-
mately 1 s, ample time for characterization of the extent of
opening of bases in I2and of the decay of I2to I1by fast
In Fig. 1 (all panels) at 10°C, the stable open complex (labeled
RPo) may be an equilibrium mixture containing some of the
intermediate complex I3identified previously (6). An increase
in the population of I3is expected early in the solute upshifts
in Fig. 1C, decaying to I2in less than 100 ms. Simulations on
the basis of kinetic data (6) show that the time resolution of
the three-syringe burst/fast footprinting experiments reported
in this research is insufficient to investigate I3, and its population
is therefore combined with that of RPoin Fig. 1.
To determine whether the DNA in I2is closed or partially or
fully open in the region of the initiation bubble, we footprinted
thymines with a constant dose of MnO−
probed as a function of time after rapidly destabilizing open
complexes with 1.1 M NaCl or after mixing them with 0.12 M
NaCl (control reaction). The sequencing gels in Fig. 2 compare
the time-dependent behavior of all MnO−
each strand after the upshift with that of the control reaction.
For the template (t) strand (Fig. 2A) the control lanes indicate
that thymines at positions þ1, −8∕ − 9 (doublet band), and −11
monotonic decay of MnO−
in the time range 0.1–10 s. For the nontemplate (nt) strand
(Fig. 2B), the control lanes indicate that thymines at positions
þ2 and −3∕ − 4 (doublet band) are MnO−
the kinetics of the decay in MnO−
4(19). Complexes were
4-reactive positions on
4-reactive in RPo. After the upshift to 1.1 M NaCl, a
4reactivity of these bands is observed
4-reactive in RPo;
4reactivity with time after
P + R
0.12 M Salt
1.1 M Salt
P + R
0.12 M Salt
unstable intermediate I2. Simulations of changes in the populations of RPo
(Red), I2(Green), I1(Blue), and free λPRpromoter DNA (P, Gray) at 10 °C,
0.01 M Mg2þas a function of time are shown for: (A) reversible association
at high [RNAP] (100 nM) and 0.12 M salt (KCl or NaCl); (B) irreversible (in ex-
cess competitor) dissociation at 0.12 M salt; and (C) irreversible dissociation
after a rapid upshift to 1.1 M salt. Rate and equilibrium constants for the
association simulation (A) are from ref. 17; at no point on theassociation time
course is there a significant population of I2. Rate and equilibrium constants
for the dissociation simulations are from ref. 6. The salt upshift (C) produces a
large transient burst of I2, not observed in the low salt dissociation experi-
ment (B). The species designated RPoalso includes a minor population of
the late intermediate I3(6), which is not resolved from RPoin the present
studies. Values of the parameters used to generate these simulations are
given in Table S1.
A burst dissociation experiment is required to characterize the
1.1 M NaCl upshift
during the burst and subsequent decay of intermediate promoter complex
I2after an upshift of RPoto 1.1 M NaCl. Sequencing gels (representative
of three independent experiments) show the decay of permanganate reac-
tivity of individual open thymine bases (left lanes) on the template (A) and
nontemplate (B) strands as a function of time after upshift, probed with a
constant dose of NaMnO4(see Material and Methods). Right lanes show
corresponding low salt RPoreactivity controls for both strands as a function
of time after mixing with 0.12 M NaCl.
Visualizing open thymines on template and nontemplate strands
Gries et al.PNAS
June 8, 2010
the NaCl upshift is very similar to that of the t strand. Thymines at
−7 and −10 on the nt strand are not detected, though the DNA is
open in this region (as judged by reactivity at positions −11 and
−8∕ − 9 on the t strand). We infer that interactions of these up-
stream nt strand thymines with σ70region 2 (cf. ref. 20 and refer-
ences therein) protect them from reacting with MnO−
well as in RPo.
In addition to providing visual demonstrations of the positions
of reactive thymines in I2and of the time course of their decay as
I2converts to products (I1and then free promoter DNA), Fig. 2
allows a visual comparison of the reactivities of these thymines in
I2[judged by the early time points (0.1–0.25 s) after the upshift],
both relative to other thymines in I2and to the RPocontrol. At
0.25 s, the population distribution of promoter DNA (Fig. 3A) is
80% I2, 10% RPoand I3, and 10% closed I1and free promoter
DNA. Strikingly, in the early time lanes where I2is the major
species, thymines at both þ2 and −3∕ − 4 on the nt strand appear
much less reactive than in the RPocontrol, whereas reactive
thymines at þ1 and other positions on the t strand appear nearly
as reactive as in the RPocontrol.
To explore the behavior of each MnO−
ing dissociation, we quantified their individual decay kinetics
and reactivities relative to the RPocontrol (Fig. 3 and Table 1).
In Fig. 3A, the populations of I2and of RPo(including I3) are
plotted as a function of time after the upshift on a logarithmic
time scale (two decades, from 0.1 to 10 s) to compare with the
analysis of the individual thymines shown in the subsequent pa-
nels. For all thymines, the observed change in MnO−
well fit by a single exponential decay (Solid Curve, Fig. 3 B–F).
Although I2is initially the dominant species after the upshift,
the presence of some RPo, I3, and closed promoter DNA requires
that the data be deconvoluted to quantify the MnO−
each thymine in I2. A series of simulations of the observed decay
kinetics were performed in which these reactivities were system-
atically varied from 0% (unreactive in I2) to 100% (as reactive in
I2as in RPo). Simulated time courses calculated by using the
best-fit reactivities of each thymine in the bubble in I2are com-
pared with the 0% and 100% limiting cases and with the experi-
mental data in Fig. 3 B–F. Best-fit reactivities are listed in Table 1
together with the rate constants for decay of each reactive
thymine in the conversion of I2to I1.
Comparison of the results reveals that:
4-reactive thymine dur-
i. Decay rate constants are the same for all thymines
(0.6 ? 0.1 s−1) and agree within the uncertainty with the rate
constant for the conversion of I2to I1(k−2¼ 0.72 ? 0.07 s−1)
determined by filter binding of quenched samples as a func-
tion of time after exposure to a 1.1 M salt upshift (6).
ii. The MnO−
strand in I2is the same as in the RPocontrol. MnO−
ities of upstream thymines on the t strand in I2are about 80%
as large as in the RPocontrol. MnO−
stream thymines on the nt strand in I2are 50–55% as large
as in the RPocontrol. (These conclusions are insensitive to
assumptions regarding the amount and MnO−
the I3 present in the residual population labeled RPo in
Fig. 3A) Thymines at −7 and −10 on the nt strand are fully
protected (zero MnO−
4reactivity of the start site (þ1) thymine on the t
4reactivities of down-
4reactivity) in both I2and RPo.
Opening of the Initiation Bubble Occurs in the Active Site Cleft to
Convert the Closed Complex I1to I2. The architecture of multisubu-
nit RNA polymerases appears to have evolved a series of steric
blocks to prevent access of nonpromoter DNA to the active site
(2–4, 9, 21, 22). For example, bacterial RNAP recognizes promo-
ter DNA by interactions between the σ subunit and hexameric
sequences (−10 and −35 regions) that are upstream of þ1
(cf. ref. 23 and references therein). Placement of σ70with respect
to the channel requires the DNA to bend sharply at −11∕ − 12 to
enter the active site cleft formed by the β and β′ “pincers”
(or jaws) (2, 17). Does double-stranded DNA bind in the cleft
at this point in the mechanism prior to being opened by RNAP?
Or does the DNA open above the cleft, allowing the template
strand to then descend down to the active site “floor” (2, 23)?
The relatively “closed” state of the pincers (less than 25 Å
apart) observed in crystal structures of the bacterial RNAP (2, 22)
and the transcription factor TFIIB bound to the 12-subunit
eukaryotic RNAP (24, 25) has motivated proposals that DNA
must open outside of the cleft. For the bacterial RNAP, which
lacks a helicase cofactor, opening outside the active site cleft
is proposed to be nucleated by a thermal breathing mechanism
(23). In this proposal, transient opening and closing of the A/
T-rich −10 hexamer leads to capture of the nt strand by σ70region
2 at the upstream entrance (23), followed by entry of only the t
strand into the cleft (2, 23). Indeed an RNAP subassembly con-
sisting of σ70region 2 and an N-terminal fragment of β′ was
observed to form an open (MnO−
A/T-rich promoter set in negatively supercoiled DNA; this
4-reactive) complex with a highly
the burst population of I2decays to I1and promoter DNA. (A) Predicted
populations of RPoand I2(see SI Text) are plotted versus time (log scale) after
NaCl upshift (0.1–10 s). (B–F) Relative MnO−
let or doublet band (calculated from three independent experiments on each
strand; see Fig. 2) as a function of time (log scale). Solid curves are single
exponential fits of these data, yielding the rate constants for the decay of
I2in Table 1. Simulations of the decay using the population distribution in
Fig. 1C and varying the MnO−
in I2relative to the RPocontrol are shown as dashed lines. The best fit is
shown in red; the limits of the fit (I2closed (0% reactive) and I2as open
as RPo(100%)) are shown in gray.
Kinetics of the open to closed transition of individual thymines as
4reactivities of each thymine sing-
4reactivity of each thymine singlet or doublet
www.pnas.org/cgi/doi/10.1073/pnas.1000967107Gries et al.
polymerase subassembly did not open this promoter on linear
DNA (26). Whereas these structure- or equilibrium-based me-
chanistic hypotheses are appealing and could apply to promoters
under highly negative supercoiling stress and/or high tempera-
ture, E. coli RNAP readily forms open complexes on linear pro-
moter fragments in vitro.
Kinetic-mechanistic studies of RPoformation and dissociation
at the λPRpromoter combined with footprinting data argue that
interactions of regions of E. coli RNAP with promoter DNA
bound in the cleft actively distort and open the initiation bubble.
HO• and DNase I cleavage of the DNA backbone of the early
intermediate I1demonstrate that both the t and nt DNA strands
are protected without interruption in this region (−15 to þ25),
(9, 27, 28). For the T7A1 promoter, time-resolved HO• foot-
prints of populations of intermediates during formation of stable
open complexes have been interpreted in terms of a mechanism
involving three classes of intermediates before the rate-determin-
ing step (10). Comparison of the kinetics of development of
HO• protection led the authors to propose that DNA opening
occurs outside the cleft and before the rate-determining step (10).
Further research is needed to determine the nature of the rate-
determining step at T7A1 and to understand the origins of the
apparent mechanistic differences between these two promoters.
Experiments presented here demonstrate that MnO−
thymines appear in the conversion of I1to I2at λPR. We conclude
from these results that the pincers of RNAP are sufficiently flexi-
ble in solution to allow DNAto enter as a double helix, where it is
then opened via binding interactions with elements on RNAP.
Additional evidence that DNA opening occurs in the active site
cleft and not in solution is provided by the observation that the
[salt] dependence of the DNA opening step (conversion of I1to
I2) is much smaller in magnitude than that of melting 12–14 base
pairs of DNA in solution (18). We proposed that the N-terminal
polyanionic domain of σ70(σ region 1.1) must move in the cleft,
allowing the duplex DNA to descend (18). These movements
would position −2∕ − 1ðtÞ near the highly conserved region on
the downstream lobe of β known as fork loop 2. By analogy with
base-flipping enzymes, we speculate that fork loop 2 inserts in the
minor groove, creating a 90° bend and unwinding the DNA helix
to form the I1-I2transition state. Interactions between the DNA
phosphate backbone and positive regions in the cleft provide an
additional driving force for opening. A bind-bend-open mecha-
nism has also been proposed for RPoformation by the single
subunit T7 phage RNAP and tested by stopped flow kinetic
and FRET experiments (29, 30).
Single-molecule DNA magnetic tweezer experiments revealed
that unwinding of ∼1 turn of the helix in an E. coli RNAP–
promoter complex occurred in a single process at all promoters
studied on both negatively and positively supercoiled DNA (7).
Because the time resolution of the assay was ∼1 s, these experi-
ments could not resolve whether untwisting occurred in a single
kinetic step, or whether the mechanism involved a sequence of
unwinding steps. Our results show that opening (unpairing, par-
4does not detect any unstacked thymine bases
4reactivity with the kinetics of downstream
tial unstacking, and therefore unwinding) occurs in the bottleneck
step of RPoformation (I1to I2) at the λPRpromoter.
Proposed DNA Conformations in the Steps in Open Complex
Formation. Fig. 4 shows a schematic of the proposed series of
conformational changes in the downstream DNA in the RNAP
active site channel as the reaction proceeds from the closed in-
termediate I1to the unstable open complex I2and finally to the
stable open complex RPo. DNA in I1is shown as double-stranded
with the exception of a distortion at −11ðtÞ, which flips out
−11AðntÞ (20). However, because −11ðtÞ is not MnO−
we infer that it remains stacked with other DNA bases and/or is
interacting with RNAP (9). In our model of I1, the þ1 base pair
lies ∼40 Å above the active site Mg2þ(9, 17). Further descent of
duplex DNA in I1is blocked by σ region 1.1 and the β′ “bridge”
helix that spans the channel.
Because the MnO−
both I2and RPo, we propose that opening of the bubble in I1→ I2
places the þ1ðtÞ base in the active site (near the catalytic Mg2þ).
However, the reduced reactivities of other thymines, especially
those on the nt strand (Table 1), motivate the proposal that the
surrounding protein environment and/or degree of stacking of
these thymines in I2differs from that in RPo. Large conforma-
tional changes in RNAP accompany the conversion of I2to
RPo(5, 15). Very large solute effects on the dissociation rate con-
stant kdand the large activation heat capacity of kdprovide evi-
dence that the downstream clamp/jaw is assembled and tightened
on the downstream DNA duplex (þ5 to þ20) in these late steps
of RPoformation, after the DNA has been opened (6, 18, 31) (see
Fig. 4). The twofold increase in permanganate reactivity of nt
strand thymines at −4∕ − 3 and þ2 in the conversion of I2to RPo
indicates that the nt strand is repositioned and/or unstacked in
these steps. These local changes in the nt strand in the cleft
may be coupled to changes in positioning of σ70region 1.1 and
the switch regions (3, 32) and to the large scale downstream
conformational changes involved in assembly and DNA binding
4reactivity of þ1 appears to be the same in
Table 1. Permanganate reactivities of thymine bases in the open region of I2and rate constants for closing these
positions in I2to I1
Strand Template Nontemplate
Rate constant (s−1)*
Predicted reactivity of I2(relative to RPocontrol)†
*Fit parameters from the decay kinetics (solid lines) in Fig. 3. The rate constant for the conversion of I2to I1(k−2) determined by
nitrocellulose filter-binding experiments at 10 °C is 0.72 ð?0.07Þ s−1(6).
†Uncertainty in these reactivities, estimated from the range of triplicate determinations at a given time point in the range from 0.1 to
0.75 s, is ?10%. See Material and Methods.
opening and stabilizing the initiation bubble at the λPRpromoter. Proposed
location (relative to the active site Mg2þ, shown as a red dot) and extent of
bending and/or opening of DNA strands (template, blue; nontemplate,
green; −20 to þ20) in the bent and wrapped closed complex I1, in the rela-
tively unstable initial open complex I2, and in the stable open complex RPo.
The transcription start site thymine is shown as a red rectangle. −11A (Green
Rectangle) on the nontemplate strand is proposed to be flipped out in the
90° bend that directs the duplex into the active site cleft in I1. Formation of I1
loads downstream duplex in the active site cleft. RNAP opens the DNA in the
cleft and positions the t strand start site base (þ1) in the active site, forming
I2. Final loading of the nt strand and assembly of a clamp/jaw (Gray Rectan-
gles) on the downstream duplex DNA greatly stabilize RPorelative to I2.
Proposal for DNA conformational changes during the steps of
Gries et al.PNAS
June 8, 2010
of the clamp/jaw. Together these events greatly stabilize RPo
relative to I2.
Opening of the entire bubble (−11 to þ2) is rate-determining
at the λPRpromoter and has the properties of a single (elemen-
tary) kinetic step. At other promoters, opening may be separated
into several kinetically distinguishable steps and may or may not
be rate-determining (10, 20). For the λPRpromoter, the activa-
tion enthalpy barrier is very high for the forward direction of the
opening step [34 kcal; (17)]; and the overall enthalpy change for
this step is also large [∼24 kcal; (6)]. For comparison, the enthal-
py of melting 13 bp of DNA in solution is approximately 75 kcal at
25°C (33). One interpretation of the 34-kcal activation barrier is
that approximately half the bubble is open and unstacked in the
transition state and that few enthalpically favorable interactions
with RNAP have formed at this stage. Alternatively, the majority
of bases in the bubble may be unpaired but not unstacked or, if
unstacked, may be engaged in enthalpically favorable interactions
with RNAP. Opening in the cleft is likely initiated at the −11 bend
by interactions between the −10 region of the nt strand and
aromatic residues of σ70region 2 (20). Because these interactions
presumably are enthalpically favorable, we infer that the majority
of the bubble is open in the transition state. Conversion of this
very unstable transition state to I2likely establishes additional
interactions with RNAP, possibly including those between the t
strand and switch 2 of β′ (32).
Proposed Physiological Relevance of the Unstable Open Complex I2.
Under physiological conditions, I2is highly unstable relative to
RPoat the λPRpromoter and converts to RPoso rapidly that it
never accumulates (Fig. 1 A and B). Yet the conversion of I1to I2
opens the entire transcription bubble and may correctly load the
start site base in the active site. Do multiple open complexes
(RPo, I3, and I2) with large differences in stability play distinct
functional roles in the regulation of transcription initiation? We
propose that the answer to this question is strongly affirmative.
Extensive studies of transcription initiation at the ribosomal
rrnB P1 promoter by Gourse, Ross, and coworkers reveal that the
open complex formed at this promoter in the absence of NTPs
and negative supercoiling is highly unstable, with a short lifetime,
existing in an equilibrium shifted toward closed complexes (cf.
ref. 34 and references therein). If open complex formation at λPR
were blocked at I2or I3, unable to progress to the stable RPo,
then a similar situation would be observed. An equilibrium would
exist between these unstable open intermediates and the closed
complex I1shifted toward I1at 25 °C and with a lifetime of I2of
only a few seconds. On the basis of this kinetic/thermodynamic
analogy, and because I2at λPRand the functional open complex
at rrnB P1 both appear to be the first open complex formed from
a closed complex at these promoters, we propose that the
unstable intermediate(s) I2(and/or I3) at λPRbehave functionally
and structurally like the unstable open complex characterized at
A key functional property of the open complex at rrnB P1 is the
ability to initiate synthesis of a full-length transcript rapidly and
efficiently, without any short, abortive product synthesis, upon
addition of all four NTPs (35). A key structural property of
the rrnB P1 open complex is its shorter downstream boundary
of the hydroxyl radical footprint (∼ þ 12 to þ15 of the (t) strand
for the rrnB P1 (36), compared to þ20 to þ25 for RPoat λPR(9).
This difference in downstream boundaries likely reflects a differ-
ence in the extent of assembly and DNA binding of downstream
mobile domains in β′ including the clamp, jaw, and sequence
insertion 3, which we propose stabilize RPoat λPR(6, 18, 31).
We hypothesize that these hallmarks of the open complex at rrnB
P1 will be observed for I2(and/or I3) at λPR.
The instability (relative to the closed complex) and short life-
time of the open complex at ribosomal promoters makes it a tar-
get of regulation by proteins like DksA (12) and by ligands
including the stress factor ppGpp (37, 38) and the initiating
NTP (11). Clearly, ribosomal expression has been tuned to re-
spond rapidly to changes in conditions, including changes in
NTP concentrations. We hypothesize that complete assembly
and tightening of the clamp/jaw on downstream DNA character-
istic of the stable open complex RPoat λPRis disfavored at the
rrnB P1 promoter [possibly because of differences in interactions
with the nt strand in the cleft (39, 40)], explaining its instability
relative to λPR. If so, tight downstream interactions would not
need to be broken for RNAP to escape from the rrnB P1 promo-
ter, allowing highly efficient production of full-length transcripts
under exponential growth conditions. In contrast, a fully as-
sembled and tightened clamp at λPRin RPomay impede escape
and favor abortive initiation, consistent with the observation that
downstream DNA from þ1 to þ20 plays a key role in determining
the efficiency of the transition to elongation (41).
For a series of promoters that form stable binary open com-
plexes, Hsu, Chamberlin and colleagues obtained evidence for
two ternary initial transcription complexes (ITCs): One ITC
makes only abortive products and the other primarily makes full
length transcripts. For these promoters, the overall abortive:pro-
ductive ratio is large and promoter-sequence-dependent [(42);
see also ref. 43]. Abortive transcripts are made in vivo (44); their
cellular function as small RNAs is not yet known.
What is the relationship between multiple binary open com-
plexes (I2, I3, and RPo) and multiple ITCs? We propose that
stable RPo,with its tightly gripping downstream clamp/jaw, is cap-
able of only abortive synthesis, whereas less-stable I2and I3, in
which the clamp/jaw is less assembled and/or less tightly bound
to downstream DNA, are capable of productive initiation. In this
model, the relative populations of stable (RPo) and less stable (I2
and I3) open complexes determine the observed abortive:produc-
tive ratio. The fact that the clamp/jaw assembles only after DNA
opening and untwisting is complete indicates that its grip likely
impedes the movement of duplex DNA within it. Promoter se-
quence, sigma factor, concentrations of regulatory ligands, and
solution conditions all must affect the relative stability and life-
time of each initiation intermediate and hence are determinants
of initiation rate and the abortive:productive ratio, providing
potent and relatively unexplored avenues for the regulation of
Material and Methods
Solutions and Materials.Standard methods of purifying RNAP and
of obtaining32P-labeled DNA fragments were used. See SI Text.
High [NaCl] Fast-Kinetic MnO4DNA Footprinting. Labeled DNA pro-
moter fragments in BB were mixed with RNAP in SB at a final
concentration of 10 nM and incubated at room temperature
(∼20°C) for 90 min to preform open complexes. These complexes
were loaded into the sample A tube of a KinTek Corporation
RQF-3 Rapid Chemical Quench-Flow instrument cooled to 10 °C
by a circulating water bath. Push syringe A was loaded with BB
supplemented to 4% SB. The sample B tube and push syringe B
400 μg∕mL heparin, and 4% SB. Push syringe C was loaded with
a solution containing 200 mM NaMnO4and 900 mM NaCl.
Collection tubes were filled with 300 μL of a quench solution con-
taining 3.75 M ammonium acetate and 7.1 M β-mercaptoethanol.
The quench-flow instrument was operated in a push-pause-push-
pause-push mode. The first push rapidly mixed the preformed
open complexes with the high salt solution resulting in a final
[NaCl] of 1.1 M. This solution was held in reaction loop 7 for
the desired perturbation time. The second push mixed the con-
tents of reaction loop with the solution in push syringe C resulting
in a final [MnO−
exittubefor150msbeforethefinal pushexpelledthesolution into
the collection tube containing quench solution. Quenched reac-
4] of 66.7 mM. This solution was held in the
www.pnas.org/cgi/doi/10.1073/pnas.1000967107Gries et al.
tions were immediately ethanol precipitated. Each load–reaction
cycle took 250 min. Low salt control reactions were performed as
above with the exception of loading solutions that keep [NaCl] at
120 mM. DNA fragments were washed with 70% ethanol,
resuspended, reprecipitated, and washed. Modified fragments
were cleaved by incubation at 90 °C in 1 M piperidine. Reactions
were evaporated and resuspended in TE buffer (10 mM Tris HCl,
1 mM EDTA, pH 8.0) three times. The resulting DNA was resus-
pended in urea loading buffer and resolved on an 8% acrylamide
gel. The gel was dried and exposed to a storage phosphor screen.
The screen was scanned on a Typhoon scanner, and the resulting
data analyzed with ImageQuant software. Phosphoimager
intensities of each MnO−
a function of time after the salt upshift were fit to a first order
(single exponential) rate equation in which the long-time plateau
value was floated. To obtain normalized θ values [fraction of
4reactive thymine band (or doublet) as
that base remaining in an open (I2) condition], this long-time
plateau intensity, arising from background/duplex reactivity of
the thymine, was subtracted from the observed phosphoimager
intensity at each time and divided by the background corrected
intensity of that position in a low salt RPocontrol. Data fitting
was performed by using IgorPro 5 software.
Population Modeling. See SI Text.
ACKNOWLEDGMENTS. We thank Dr. C. Davis for preliminary KMnO4experi-
ments. We are grateful to our colleagues for many fruitful discussions and
to the reviewers and the editor for their comments on the manuscript.
This work was supported by National Institutes of Health Grant GM23467
(to M.T.R.). T.J.G. gratefully acknowledges the support of the William R. and
Dorothy E. Sullivan Distinguished Graduate Fellowship. W.S.K. acknowledges
support from National Institutes of Health Biotechnology Training Grant
(NIH 5 T32 GM08349).
1. Murakami KS, Masuda S, Darst SA (2002) Structural basis of transcription initiation:
RNA polymerase holoenzyme at 4 Å resolution. Science 296:1280–1284.
2. Vassylyev DG, et al. (2002) Crystal structure of a bacterial RNA polymerase holoenzyme
at 2.6 Å resolution. Nature 417:712–719.
3. Cramer P, Bushnell DA, Kornberg RD (2001) Structural basis of transcription: RNA
polymerase II at 2.8 Å resolution. Science 292:1863–1876.
4. Zhang G, et al. (1999) Crystal structure of Thermus aquaticus core RNA polymerase at
3.3 Å resolution. Cell 98:811–824.
5. Saecker RM, Record MT, Jr (2002) Protein surface salt bridges and paths for DNA
wrapping. Curr Opin Struct Biol 12:311–319.
6. Kontur WS, Saecker RM, Capp MW, Record MT, Jr (2008) Late steps in the formation of
E. coli RNApolymerase—λPRpromoteropen complexes: characterizationof conforma-
tional changes by rapid [perturbant] upshift experiments. J Mol Biol 376:1034–1047.
7. Revyakin A, Ebright RH, Strick TR (2004) Promoter unwinding and promoter clearance
by RNA polymerase: detection by single-molecule DNA nanomanipulation. Proc Natl
Acad Sci USA 101:4776–4780.
8. Sclavi B, et al. (2005) Real-time characterization of intermediates in the pathway to
open complex formation by Escherichia coli RNA polymerase at the T7A1 promoter.
Proc Natl Acad Sci USA 102:4706–4711.
9. Davis CA, Bingman CA, Landick R, Record MT, Jr, Saecker RM (2007) Real-time foot-
printing of DNA in the first kinetically significant intermediate in open complex for-
mation by Escherichia coli RNA polymerase. Proc Natl Acad Sci USA 104:7833–7838.
10. Rogozina A, Zaychikov E, Buckle M, Heumann H, Sclavi B (2009) DNA melting by RNA
polymerase at the T7A1 promoter precedes the rate-limiting step at 37°C and results
in the accumulation of an off-pathway intermediate. Nucleic Acids Res 37:5390–5404.
11. Gaal T, Bartlett MS, Ross W, Turnbough CL, Jr, Gourse RL (1997) Transcription
regulation by initiating NTP concentration: rRNA synthesis in bacteria. Science
12. Paul BJ, et al. (2004) DksA: a critical component of the transcription initiation
machinery that potentiates the regulation of rRNA promoters by ppGpp and the
initiating NTP. Cell 118:311–322.
13. Paul BJ, Berkmen MB, Gourse RL (2005) DksA potentiates direct activation of amino
acid promoters by ppGpp. Proc Natl Acad Sci USA 102:7823–7828.
14. Buc H, McClure WR (1985) Kinetics of open complex formation between Escherichia
coli RNA polymerase and the lacUV5 promoter. Evidence for a sequential mechanism
involving three steps. Biochemistry 24:2712–2723.
15. Roe JH, Burgess RR, Record MT, Jr (1985) Temperature dependence of the rate
constants of the Escherichia coli RNA polymerase—λPR promoter interaction.
Assignment of the kinetic steps corresponding to protein conformational change
and DNA opening. J Mol Biol 184:441–453.
16. Li XY, McClure WR (1998) Characterization of the closed complex intermediate
formed during transcription initiation by Escherichia coli RNA polymerase. J Biol Chem
17. Saecker RM, et al. (2002) Kinetic studies and structural models of the association of
E. coli σ70RNA polymerase with the λPRpromoter: Large scale conformational changes
in forming the kinetically significant intermediates. J Mol Biol 319:649–671.
18. Kontur WS, Capp MW, Gries TJ, Saecker RM, Record MT, Jr (2010) Probing DNA
binding, DNA opening and assembly of a downstream clamp/jaw in Escherichia coli
RNA polymerase—λPRpromoter complexes using salt and the physiological anion
glutamate. Biochemistry, in press.
19. Borowiec JA, Zhang L, Sasse-Dwight S, Gralla JD (1987) DNA supercoiling promotes
formation of a bent repression loop in lac DNA. J Mol Biol 196:101–111.
20. Schroeder LA, et al. (2009) Evidence for a tyrosine-adenine stacking interaction
and for a short-lived open intermediate subsequent to initial binding of Escherichia
coli RNA polymerase to promoter DNA. J Mol Biol 385:339–349.
21. Bushnell DA, Kornberg RD (2003) Complete, 12-subunit RNA polymerase II at 4.1 Å
resolution: Implications for the initiation of transcription. Proc Natl Acad Sci USA
22. Murakami KS, Masuda S, Campbell EA, Muzzin O, Darst SA (2002) Structural basis
of transcription initiation: An RNA polymerase holoenzyme-DNA complex. Science
23. Murakami KS, Darst SA (2003) Bacterial RNA polymerases: The wholo story. Curr Opin
Struct Biol 13:31–39.
24. Kostrewa D, et al. (2009) RNA polymerase II-TFIIB structure and mechanism of
transcription initiation. Nature 462:323–330.
25. Liu X, Bushnell DA, Wang D, Calero G, Kornberg RD (2010) Structure of an RNA
polymerase II-TFIIB complex and the transcription initiation mechanism. Science
26. Young BA, Gruber TM, Gross CA (2004) Minimal machinery of RNA polymerase
holoenzyme sufficient for promoter melting. Science 303:1382–1384.
27. Craig ML, et al. (1998) DNA footprints of the two kinetically significant intermediates
in formation of an RNA polymerase-promoter open complex: evidence that
interactions with start site and downstream DNA induce sequential conformational
changes in polymerase and DNA. J Mol Biol 283:741–756.
28. Davis CA, Capp MW, Record MT, Jr, Saecker RM (2005) The effects of upstream DNA
on open complex formation by Escherichia coli RNA polymerase. Proc Natl Acad Sci
29. Tang GQ, Patel SS (2006) Rapid binding of T7 RNA polymerase is followed by simul-
taneous bending and opening of the promoter DNA. Biochemistry 45:4947–4956.
30. Tang GQ, Patel SS (2006) T7 RNA polymerase-induced bending of promoter DNA is
coupled to DNA opening. Biochemistry 45:4936–4946.
31. Kontur WS, Saecker RM, Davis CA, Capp MW, Record MT, Jr (2006) Solute probes of
conformational changes in open complex (RPo) formation by Escherichia coli
RNA polymerase at the λPRpromoter: Evidence for unmasking of the active site in
the isomerization step and for large-scale coupled folding in the subsequent conver-
sion to RPo. Biochemistry 45:2161–2177.
32. Belogurov GA, et al. (2009) Transcription inactivation through local refolding of the
RNA polymerase structure. Nature 457:332–335.
33. Holbrook JA, Capp MW, Saecker RM, Record MT, Jr (1999) Enthalpy and heat capacity
changes for formation of an oligomeric DNA duplex: interpretation in terms of
coupled processes of formation and association of single-stranded helices. Bio-
34. Haugen SP, Ross W, Gourse RL (2008) Advances in bacterial promoter recognition and
its control by factors that do not bind DNA. Nat Rev Microbiol 6:507–519.
35. Gourse RL (1988) Visualization and quantitative analysis of complex formation
between E. coli RNA polymerase and an rRNA promoter in vitro. Nucleic Acids Res
36. Rutherford ST, Villers CL, Lee JH, Ross W, Gourse RL (2009) Allosteric control of
Escherichia coli rRNA promoter complexes by DksA. Genes Dev 23:236–248.
37. Barker MM, Gaal T, Gourse RL (2001) Mechanism of regulation of transcription
initiation by ppGpp. II. Models for positive control based on properties of RNAP
mutants and competition for RNAP. J Mol Biol 305:689–702.
38. Barker MM, Gaal T, Josaitis CA, Gourse RL (2001) Mechanism of regulation of
transcription initiation by ppGpp. I. Effects of ppGpp on transcription initiation in vivo
and in vitro. J Mol Biol 305:673–688.
39. Haugen SP, et al. (2006) rRNA promoter regulation by nonoptimal binding
of sigma region 1.2: An additional recognition element for RNA polymerase. Cell
40. Feklistov A, et al. (2006) A basal promoter element recognized by free RNA polymer-
ase sigma subunit determines promoter recognition by RNA polymerase holoenzyme.
Mol Cell 23:97–107.
41. Hsu LM, Vo NV, Kane CM, Chamberlin MJ (2003) In vitro studies of transcript initiation
by Escherichia coli RNA polymerase. 1. RNA chain initiation, abortive initiation, and
promoter escape at three bacteriophage promoters. Biochemistry 42:3777–3786.
42. Vo NV, Hsu LM, Kane CM, Chamberlin MJ (2003) In vitro studies of transcript initiation
by Escherichia coli RNA polymerase. 2. Formation and characterization of two distinct
classes of initial transcribing complexes. Biochemistry 42:3787–3797.
43. Kubori T, Shimamoto N (1996) A branched pathway in the early stage of transcription
by Escherichia coli RNA polymerase. J Mol Biol 256:449–457.
44. Goldman SR, Ebright RH, Nickels BE (2009) Direct detection of abortive RNA transcripts
in vivo. Science 324:927–928.
Gries et al. PNAS
June 8, 2010