From the Department of Biochemistry and Molecular
Biology,
Department of Chemistry, Michigan State University,
East Lansing, Michigan 48824-1319 and ** Graduate School
of Bioscience and Biotechnology, and
Frontier Collaborative Research Center,
Tokyo Institute of Technology, Yokohama 226-8503, Japan
Received for publication, January 31, 2003, and in revised form, March 10, 2003
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
We report a "running start, two-bond"
protocol to analyze elongation by human RNA polymerase II (RNAP II). In
this procedure, the running start allowed us to measure rapid rates of
elongation and provided detailed insight into the RNAP II mechanism.
Formation of two bonds was tracked to ensure that at least one
translocation event was analyzed. By using this method, RNAP II is
stalled briefly at a defined template position before restoring the
next NTP. Significantly, slow reaction steps are identified both before and after phosphodiester bond synthesis, and both of these steps can be
highly dependent on the next templated NTP. The initial and final
NTP-driven events, however, are not identical, because the slow step
after chemistry, which includes translocation and pyrophosphate
release, is regulated differently by elongation factors hepatitis Pre-steady state kinetic analysis allows the progress of an
enzymatic reaction to be tracked in real time (1, 2), and coupling
enzyme functional dynamics to the structure provides the clearest
insight into the mechanism. In this paper, we compare the first
transient state kinetic studies of human (Homo
sapiens) RNAP II1 to the
x-ray structure of the yeast (Saccharomyces cerevisiae) RNAP
II elongation complex (EC) (3). These studies give new insight into the
RNAP II mechanism and demonstrate the feasibility of a detailed kinetic
study of a highly regulated enzyme that is at the hub of gene control
in human cells.
There is increasing recognition that transcriptional elongation is
highly regulated in eukaryotes (4-8). As an example, hepatitis In this work, we use rapid quench kinetics to demonstrate critical
NTP-dependent steps during RNA synthesis. First, we
analyzed recovery from a stall at a defined template position, in the
presence of TFIIF or HDAg. During stall recovery, two fractions of EC
were clearly observed on the active pathway, and most significantly, these ECs had different requirements for binding and utilizing the
incoming substrate NTP. This observation strongly indicates a substrate
NTP-induced fit mechanism, in which the NTP first binds and then helps
to convert the EC to a fully active form. Second, in the
TFIIF-stimulated mechanism but not in the HDAg-stimulated mechanism, a
slow step after phosphodiester bond formation is also highly dependent
on the incoming NTP. Thus, in the presence of TFIIF, elongation is
NTP-driven at both the beginning and the end of a single bond addition
cycle, but only one of these NTPs can be the substrate for
phosphodiester bond formation at a single position. The other NTP
appears to be the substrate for addition of the subsequent bond. These
observations lead to the following conclusions: 1) RNAP II elongates
according to a substrate NTP-induced fit mechanism; 2) translocation
can be induced by prior NTP binding. Translocation must occur at either
the beginning or the end of each bond addition cycle, and in the
presence of TFIIF, both are highly dependent on the next templated NTP.
Significantly, HDAg eliminates the high NTP dependence of the slow step
after phosphodiester bond formation, demonstrating the unusual nature
of the RNAP II mechanism in the presence of TFIIF. As with TFIIF,
the HDAg-stimulated mechanism shows evidence of substrate
NTP- induced fit during escape from a stall, but, unlike TFIIF, NTP
dependence is not detected with HDAg in the normal processive
transition between bonds.
Comparing the kinetics of RNAP II elongation with the yeast EC
structure reverses the view of how NTPs are loaded, alters our
understanding of the translocation mechanism, and provides new insight
into transcriptional efficiency and fidelity.
Cell Culture, Extracts, and Proteins--
HeLa cells were
purchased from the National Cell Culture Center (Minneapolis, MN).
Extracts of HeLa cell nuclei were prepared as described (19).
Recombinant TFIIF (20, 21) and HDAg (9) were prepared as described.
The Running Start, Two-bond Elongation Assay--
The running
start, two-bond elongation assay is shown in Fig. 1 (16, 23).
Initiation was from the adenovirus major late promoter with a modified
downstream sequence so that a 40-nucleotide transcript can be
synthesized in the absence of ATP and GTP. An extract of human HeLa
cells was the source of transcription factors. C40 (a 40-nucleotide RNA
ending in a 3'-CMP) ECs were synthesized by addition of 10 µM dATP, 300 µM ApC dinucleotide, 5 µCi
per reaction [
After quenching reactions with 0.5 M EDTA, beads were
collected; supernatant was removed, and samples were dissolved in 90% formamide loading dye containing 1% SDS. Samples were boiled for 2 min
and RNAs separated in 14-16% polyacrylamide (20:1
acrylamide/bisacrylamide) gels containing 50% w/v urea and 1× Tris
borate-EDTA. Gels were analyzed using a Amersham Biosciences
PhosphorImager. Each gel lane was analyzed independently for percent of
signal present in G44 plus all longer transcripts or G45 plus all
longer transcripts compared with A43 plus all longer transcripts. The
data were handled in this way to compensate for occasional
inconsistency in recovery or loading of samples.
Quality of RNAP II ECs--
The complex kinetics we report
demonstrate multiple conformers of A43 EC at the time of GTP addition
in the running start, two-bond protocol. Because ECs were isolated on
bead templates from HeLa nuclear extracts, it is reasonable to consider
whether A43 ECs differ in their kinetic properties because of
experimental treatments or because of damage to a subset of ECs during
preparation. However, A43 conformational states are determined by
treatments that occur after EC purification. The initial conformational
states detected at A43 are different in the presence of TFIIF, HDAg, or
in the absence of an elongation factor, showing that protein factors
drive RNAP II between functional modes. Furthermore, increasing GTP
concentration blurs the distinction between different kinetic states,
indicating that RNAP II changes conformation through interactions with
substrate, as expected for an RNAP. Also, the distribution of A43
states is dependent on the time of stalling at A43, demonstrating reversibility between states (23). A43 conformational states, therefore, are selected based on treatments (protein factors, substrate, and time of incubation) after purification and are not an
artifact of preparation. In the purification scheme, RNAP II molecules
are selected for the ability to initiate transcription accurately in
concert with the general initiation factors, and all C40 and A43
complexes are active in elongation. Sarkosyl and salt treatment appears
to strip all contaminating transcription factors and complicating
activities from the EC (12).
Kinetic Models--
Kinetic models were designed based on a
qualitative assessment of rate data, as described under "Results,"
and model independent analysis of rate data (24, 25) (not shown). The
program DYNAFIT (26), which utilizes non-linear least squares curve
fitting to obtain the optimal fit to a kinetic model, was used to
estimate rate constants. Most of the rate constants listed in Fig. 4
are currently under determined experimentally, so the values reported are meant to represent a simulation of the mechanism with the caveat
that future refinement will be necessary to determine fully the
accurate rate constant values. The rate constants used for simulations,
however, give a reasonable qualitative and quantitative description of
the rate equation. The mechanisms we apply fit the primary
characteristics of the rate data sets, whereas alternate schemes
prove inadequate to model the data. Furthermore, the models we espouse
seem consistent with the S. cerevisiae RNAP II EC structure (3).
The kinetic pathway shown in Fig. 4A is the simplest induced
fit mechanism with a pausing pathway that allows access of the active
site after an NTP-induced fit conformational change (see under
"Results" and "Discussion"). This is an adequate kinetic model
for RNAP II elongation stimulated by HDAg. To fit the HDAg-stimulated mechanism requires a minimum of three initial states at A43 as follows:
A43a (23% of total ECs); A43b (27% of total ECs); and A43c (50% of
total ECs). From model-independent analysis (not shown), the fastest
rate is estimated as 1250 s
The pathway shown in Fig. 4B is the simplest induced fit
model (including a pausing pathway), with an open active site, that also confers substrate NTP dependence at both the start and end of the
G44 bond addition cycle (see under "Results" and "Discussion"). This mechanism is adequate to model rates of elongation stimulated by
TFIIF through formation of two phosphodiester bonds. The simple induced
fit model (Fig. 4A) is adequate to fit the TFIIF data set
through formation of the G44 phosphodiester bond (not shown) but not
through formation of the G45 bond. In the presence of TFIIF, a minimum
of three initial conformers at A43 are required: A43a (55% of total);
A43b (10% of total); and A43c (35% of total). Model independent
kinetic analysis indicates that the fastest pathway for G44 synthesis
is >500 s
Residuals (a statistical test; Fig. 3) indicate that these models
converge to experimental data generally within 5 or 10%, indicating
the reliability of the simulations. Particularly with HDAg, the curve
fits are very close to experimental values (residuals ±5%). The rate
constants shown in Fig. 4, A and B, are the best estimates we can offer at this time, although more information is
required about individual steps in the mechanism to accurately assign
rate constant values, and additional experiments will be required to
refine the current models. The rate constants shown in Fig. 4 are
constrained to be in approximate thermodynamic balance. The curve fits
and residuals shown in Fig. 3 were optimized from the set of rate
constants shown in Fig. 4, by relaxing this constraint, so the reported
curve fits and residuals represent a slightly better fit to the data
set than those obtained from the rate constants shown in Fig. 4.
Running Start, Two-bond Assay--
To analyze the mechanism and
regulation of human RNA synthesis, we sought a method to obtain
transient state kinetic measurements of RNAP II elongation rates
through formation of multiple bonds. Precisely stalled RNAP II ECs,
immobilized on magnetic beads, were initiated from the adenovirus major
late promoter and isolated with Sarkosyl and salt washing (Fig.
1). C40 ECs contain a 40-nucleotide, 32P-labeled RNA, ending in a 3'-CMP base. The sequence
downstream of C40 is
40CAAAGG45. Because C40
ECs proved unsuitable for measuring the most rapid elongation rates, a
running start, two-bond protocol was adopted. In the presence of 20 µM CTP and UTP, 100 µM ATP was added to advance C40 ECs to the A43 position. After a brief stall at A43, a
steady state distribution was established between paused and active A43
ECs (23), such that, when GTP was added, rapid rates for G44 and G45
synthesis could be determined reproducibly. In this experimental
design, G44 synthesis rates reflect recovery from a stall at A43, and
initial G45 synthesis rates reflect processive elongation from G44
Because the running start assay allowed us to commit a significant
fraction of A43 ECs to rapid elongation, we adapted this method to
analyze GTP-dependent steps along the forward synthesis path. In Fig. 2, we compare synthesis
through the G44 and G45 positions in the presence of the RNAP II
elongation factors HDAg and TFIIF. After a brief ATP pulse, GTP was
added at the indicated concentrations, and reactions were quenched at
various times. The protocol is indicated in Figs. 1 and 2A.
The ATP pulse time, optimized for each reaction, is 120 s in the
absence of a stimulatory factor (Fig. 2B), 60 s in the
presence of HDAg (Fig. 2C), and 30 s in the presence of
TFIIF (Fig. 2D). In Fig. 2B, elongation is shown
in the absence of a stimulatory factor to confirm that TFIIF and HDAg
enhance elongation in the running start assay (compare Fig. 2,
B-D). Rates of G44 and G45 synthesis were evaluated in terms of the percent of transcript at G44 or G45 plus all longer transcripts. In this way, rates of disappearance of G44 and G45 could
be neglected.
The running start, two-bond protocol reveals significant details of the
RNAP II elongation mechanism. In the presence of HDAg or TFIIF,
synthesis rates for G44 differ from synthesis rates for G45,
demonstrating the importance of monitoring two bonds. With the running
start method, analysis of the G44 bond is expected to provide detail
about RNAP II conformational states and kinetic intermediates and any
effects of recovering from the 30- to 120-s stall at A43. Rates of G45
synthesis, on the other hand, are expected to reveal characteristics of
the approach to processive elongation. The rates of first G45
appearance should provide insight into translocation, because
translocation and pyrophosphate release must occur between synthesis of
the G44 and G45 bonds. If only a single bond is tracked, information
about translocation could be lost, because the state(s) of
translocation prior to addition of substrate cannot be known.
Two Intermediates on the Forward Elongation Pathway with Distinct
GTP Dependence--
In Fig. 3, complete
data sets are shown for elongation through the G44 and G45 positions,
in the presence of HDAg (Fig. 3, A and B) or
TFIIF (Fig. 3, C and D). The
comparison between the HDAg- and TFIIF-stimulated data sets provides
clear insight into many details of the RNAP II mechanism and its
regulation by elongation factors. Curve fits are derived from the
kinetic mechanisms shown in Fig. 4,
A and B. Residuals shown below each plot in Fig.
3 demonstrate how closely experimental data can be fit with these simulations.
Analysis of the rates of G44 synthesis (Fig. 3, A and
C) indicates that RNAP II must utilize an induced fit
mechanism, in which binding substrate GTP modifies the A43 EC
conformationally to become catalytically competent. Notably, during the
60- or 30-s stall at A43, delayed ECs fractionate between at least
three different A43 states, two of which are evident on the forward synthesis pathway and one of which is strongly paused. Furthermore, two
A43 ECs are observed on the active synthesis pathway with differing
dependence on the concentration of the incoming GTP substrate.
In the presence of HDAg, ECs partition between three states (Fig.
3A). Two A43 ECs are found on the active synthesis pathway, but these two EC conformers have different responsiveness to GTP. About
27% of A43 ECs are in the most highly poised state (Fig. 3A, fraction b; 0-27% of total ECs;
kobs,fast ~1250 s
In the presence of TFIIF, three classes of A43 EC are detected but with
different occupancy than those observed with HDAg (compare Fig. 3,
A and C). Two of these classes of A43 EC are on
the active synthesis pathway, and one is paused. About 10% of A43 ECs
elongate rapidly to the G44 position even at GTP concentrations that
are too low to support subsequent rapid elongation from G44 to G45
(Fig. 3C, fraction b; 0-10% of total ECs;
kobs,fast >500 s Regulation of First Appearance of G45 by Substrate
GTP--
Judging from the initial times of G44 and G45 appearance on
gels (Fig. 2, B-D), the most rapid rates of G44 synthesis
must be 5-10-fold faster than the rate of initial G45 appearance,
which is surprisingly slow. If this were not the case, G45 would be detected by the 0.002-0.005-s time points, but G45 is not detected until 0.02-0.05 s, even at high GTP concentration. This conclusion is
further demonstrated by quantitation of gel data (Fig. 3, B and D). Analysis of G45 synthesis rate curves shows that the
initially slow appearance of G45 can be attributed to a slow
conformational step after G44 phosphodiester bond formation. This
conclusion is demonstrated by the sigmoidal shapes of the rate curves
shown in Fig. 3, B and D. The distinctive shape
of these curves can only be described by a slow, normally irreversible
step in the RNAP II elongation mechanism after G44 phosphodiester bond
synthesis but before rapid G45 synthesis can commence (see Fig. 4,
A and B). Note that the interval in which this
slow conformational step occurs must include the translocation event
between G44 and G45 synthesis.
In the presence of HDAg (Fig. 3B), there is a slow step
after G44 phosphodiester bond formation that accounts for the slow first appearance of G45. This slow step is indicated by the sigmoidal shapes of G45 synthesis rate curves. Notice that the sigmoidal rate
curves all approach the time axis at a similar intersection point (Fig.
3B, open arrow). This result shows that, in the presence of
HDAg, the lag in G45 first appearance is not highly dependent on GTP
concentration, although in the presence of TFIIF (Fig. 3D),
the lag duration is highly GTP-dependent. For the
HDAg-stimulated mechanism, the slow step after chemistry can be modeled
by a first order rate constant, lacking GTP dependence, of 15.5 ± 0.5 s
Surprisingly, in the presence of TFIIF, the situation is very
different. At 250 and 100 µM GTP, G45 appears in an
apparent burst (Fig. 3D), but these rate curves are
sigmoidal when plotted with an expanded time axis, indicating the slow
step after chemistry (lag of 0.02-0.05 s (Fig. 2D)). From
10 to 25 µM GTP, G45 synthesis rate curves are notably
sigmoidal in shape, further demonstrating the slow step after G44 bond
formation. Surprisingly, at 1 and 2 µM GTP, almost no G45
synthesis is observed within 5 s, although eventually these ECs
will advance (data not shown). This result demonstrates the extreme GTP
dependence of this slow step in the TFIIF-stimulated mechanism. So,
after a stall at A43 in the presence of TFIIF, both the beginning phase
and the ending phase of the G44 bond addition cycle are highly
dependent on the next incoming GTP substrates (Fig. 3, C and
D). In the running start protocol, this unusual condition
arises because elongation was stalled at the A43 position. Because the
GTP-driven event at the end of the G44 bond addition cycle occurs after
chemistry, this event cannot be attributed to utilization of GTP as a
substrate for G44 bond formation and primarily reflects entry of the
GTP substrate for G45 synthesis.
As with TFIIF, HDAg-mediated recovery from a stall at A43 is
GTP-dependent (Fig. 3A), demonstrating the
GTP-induced fit mechanism. With HDAg, however, the transition between
synthesis of the G44 and G45 bonds is not noticeably dependent on the
incoming substrate GTP (Fig. 3B), as it is in the presence
of TFIIF (Fig. 3D). Therefore, HDAg and TFIIF regulate the
normal processive transition between bonds (translocation) in distinct
ways. HDAg simplifies this step and makes it less dependent on GTP.
Also, recovery from a stall is found to be a distinct process from the
normal processive transition between bonds, because TFIIF and HDAg
regulate these steps differently. One result of HDAg-mediated
stimulation is that elongation is facilitated at GTP concentrations
that are very restrictive in the presence of TFIIF, as if HDAg
facilitates a rate-limiting step (translocation) in the RNAP II
mechanism (compare Fig. 3, A-D).
Kinetic Modeling--
RNAP II appears to extend an RNA chain
according to an induced fit mechanism, in which the active site remains
available for substrate NTP binding, even when the conformational
change precedes substrate loading. Furthermore, RNAP II establishes a
steady state condition between the pausing and active synthesis
pathways that is maintained for minutes at the A43 position (23). The
simplest kinetic model that will satisfy these conditions is shown in
Fig. 4A. Induced fit requires that substrate NTP normally
binds prior to a conformational step in the mechanism (27-29). In this
paper, we argue strongly for an induced fit mechanism of RNAP II
elongation. Specifically, we argue for substrate NTP-driven
translocation of the RNA-DNA hybrid past the RNAP II active site, which
is a version of an induced fit mechanism, in which translocation is the
NTP-directed conformational change. We present our argument assuming
that A43a and A43b states are natural intermediates on the RNAP II
elongation pathway. Formally, however, A43a and A43b could be on
separate pathways. For instance, A43a and A43b could represent distinct
RNAP II ECs with different activities (i.e. because of
differential protein composition or covalent modification). We find
this alternate view untenable for the following reasons: 1) relative
occupancy of A43a and A43b depends on whether TFIIF or HDAg is added to
the reaction, showing that these states are inter-convertible (Figs. 3
and 4); and 2) fast and slow complexes are detected for G44 synthesis,
after the stall at A43, but not for G45 synthesis (Fig. 3), so
intrinsically fast and slow ECs are not indicated. Further discussion
of this issue is found under "Experimental Procedures."
Experimental data, therefore, require two A43 states, designated A43a
and A43b, that are occupied to different extents in the presence of
HDAg or TFIIF. The conversion of A43a
The TFIIF-stimulated mechanism, therefore, was fit using the more
complex kinetic model shown in Fig. 4B. This is the simplest induced fit mechanism that can account for GTP dependence at both the
beginning and end of the G44 bond addition cycle. For the TFIIF-stimulated mechanism, a kinetic scheme is shown for the formation
of two bonds. The scheme used to fit synthesis of the G45 bond is a
subset of the mechanism used to fit G44, eliminating steps that are
peripheral and therefore unnecessary for modeling. We assume that the
mechanisms for G44 and G45 synthesis include all of the same steps and
utilize similar rate constants, but in the running start protocol, many
steps observed for G44 synthesis are largely undetected in G45
synthesis. For instance, the pausing pathway is observed in G44
synthesis, because of the stall at A43, but pausing is not observed in
G45 synthesis, unless the EC is delayed at G44 (i.e. by GTP
limitation). Effectively, RNAP II tends to bypass both the pausing
pathway and perhaps the G44a conformational state during rapid
elongation. Some of the rate constants for G45 synthesis appear
different from those for G44 synthesis. This is attributable to the
simplified mechanism used to model formation of the G45 bond.
The only difference between the mechanisms in Fig. 4, A and
B, is the involvement of the n + 2 GTP substrate
(n = RNA length) in the slow transition from G44
In Fig. 4C a popular model for elongation, which we refer to
as an "equilibrium sliding" model, is shown. In such a model, there
is a rapid equilibrium between the pre- and post-translocated states,
and NTP substrate binding stabilizes the post-translocated EC (30-39).
Our kinetic data argue strongly against the equilibrium sliding model
for H. sapiens RNAP II. A43a and A43b cannot be connected by
a rapid equilibrium, because escape from A43a is more highly
GTP-dependent than escape from A43b. Therefore, there must
be distinct GTP-dependent pathways forward from A43a and A43b and inter-conversion between A43a and A43b must be relatively slow, as indicated in our simulations (Fig. 4, A and
B). Our data clearly indicate that, in the presence of
TFIIF, translocation is GTP-driven in the A43a Translocation Can Be Driven by the Incoming Substrate NTP--
We
have tracked the progress of kinetically homogeneous populations of
human RNAP II molecules through the sequence
40CAAAGG45, concentrating on synthesis of the
G44 and G45 bonds. After a brief stall at A43, the beginning of the G44
bond addition cycle is dependent on the incoming substrate GTP for a
significant fraction of ECs (A43a fraction). After G44 chemistry, the
final phase of the G44 bond addition cycle can also depend on the next
incoming substrate GTP. Because the transition from G44
Dwelling at A43 in the absence of GTP appears to reverse the
translocation step for a significant fraction of A43 ECs (designated A43a). In the presence of HDAg, 23% of A43 ECs are detected that are
on the pathway for synthesis but require a higher concentration of GTP
to progress rapidly (Fig. 3A, fraction a).
Another 27% of A43 ECs are more highly poised on the active synthesis
pathway (Fig. 3A, fraction b). In the presence of
TFIIF, 55% of ECs are on the pathway for rapid synthesis but require a
higher GTP concentration for activation, whereas another 10% of A43
ECs are more highly poised for synthesis (Fig. 3C,
fractions a and b). By assuming that both A43a and A43b
conformers are natural intermediates on the elongation path, the A43b
EC is more highly poised for synthesis than the A43a EC, and the A43b
EC appears to be less dependent on the concentration of GTP substrate
for elongation. Furthermore, because both classes of A43 EC bind GTP
and engage in G44 synthesis, the RNAP II active site must remain open
to substrate binding after the GTP-driven conformational change. This
distinction is important because the dNTP-induced fit conformational
change postulated for elongation by single-subunit DNAPs appears to
involve rotation of a critical Regulation by HDAg and TFIIF--
To account for differences in
HDAg- and TFIIF-regulated elongation, we offer the following
explanation. We suggest that TFIIF supports a tight RNAP II clamp
holding the RNA-DNA hybrid and an optimum geometry of the RNAP II
active site for catalysis. Because the clamp is tight, the RNAP II
bridge Substrate NTP-driven Translocation and Allostery--
Recently,
Erie and colleagues (24) reported transient state kinetic studies of
E. coli RNAP elongation. They proposed an allosteric model
to describe slow rates of RNA synthesis at low substrate NTP
concentrations. We concur with their observation that elongation rates
are unexpectedly slow when substrate NTP concentrations fall below a
threshold value. Because the E. coli RNAP allosteric site
appears to be template-specified in their study, it is difficult to
distinguish the allosteric site from the substrate site, unless the
allosteric conformational change is translocation, as we suggest here.
In such a case, the "allosteric" site becomes the substrate site.
Our data strongly indicate that, at limiting substrate NTP
concentrations, active site translocation is the critical,
rate-limiting step along the RNA synthesis pathway, and translocation
is induced by prior NTP binding. In our model, only template-specified
NTP-loading sites at positions n + 1 and n + 2 (n = RNA length) are required.
To analyze human RNAP II, we were forced to adopt the running start,
two-bond protocol to observe rapid elongation rates and to ensure that
we would observe at least one translocation event. In studies of
E. coli RNAP, however, elongation was from ECs that were
stalled for a longer time, and in those studies, only a single bond was
monitored. In Foster et al. (24), E. coli RNAP
rate curves have approximately hyperbolic shapes, unlike the biphasic curves observed for H. sapiens RNAP II in our study. In more
recent work, however, Erie and colleagues have produced rate data at new template positions that produce notably biphasic rate
curves.2 Although E. coli RNAP and H. sapiens RNAP II both can yield rate curves with biphasic shapes, there are significant qualitative differences between the data sets produced in the bacterial and human
systems. Notably, in the bacterial system, the amplitude of the fast
phase is more highly dependent on the NTP substrate concentration,
consistent with a single primary starting conformation of RNAP at the
time of NTP addition and perhaps consistent with the allosteric
mechanism from Erie and co-worker (24). In our data set, biphasic rate
curves appear to reflect two distinct EC conformations on the active
synthesis pathway (A43a and A43b), and the amplitude of the fast phase
in our rate curves does not appear to be so highly dependent on NTP
concentration, as if the biphasic rate curves may have distinct
meanings in the E. coli and H. sapiens systems.
It appears to us that, although homologous, E. coli and
H. sapiens RNAPs may have distinct features in the details
of their catalytic mechanisms, and that many more experiments in both
systems will be required to fully understand the functioning and
regulation of these dynamic molecular machines. For instance, E. coli RNAP may utilize an allosteric mechanism, whereas H. sapiens RNAP II does not appear to require allosteric control,
unless the substrate NTP-induced translocation mechanism that we
propose can be described as "allostery." Generally, allostery
requires distinct binding sites for substrate and effector.
The Substrate NTP-induced Translocation Model--
Our kinetic
model indicates that incoming NTP substrates pair to their cognate DNA
bases before the resulting base pair is translocated into the RNAP II
active site. We therefore inspected the S. cerevisiae RNAP
II EC structure (3) to determine whether such an NTP loading mechanism
is likely or possible. In Fig. 5, we show
how substrate NTPs can pre-load through the RNAP II main channel to
induce translocation. In the S. cerevisiae RNAP II EC
structure, the n + 1, n + 2, and n + 3 DNA template bases are single-stranded and apparently accessible to
base pair with incoming NTPs loaded through the main channel (Fig.
5A) (3). The non-template DNA strand is disordered in the
structure and is therefore unlikely to conflict with NTP loading by
this route. Darst and colleagues (48), Kornberg and colleagues (3, 50),
and Yokoyama and colleagues (49) have suggested that NTPs might load
into the active site through the secondary pore, from the opposite
direction than that we propose. In the RNAP II mechanism, however,
observation of induced fit is evidence for loading the n + 1 NTP before a conformational change that precedes chemistry.
Furthermore, in the TFIIF-stimulated mechanism, the final stage of the
G44 bond addition cycle is also GTP-driven (Fig. 3, D and
B). Therefore, both the n + 1 and the
n + 2 GTPs appear to be loaded prior to rate-limiting
conformational steps, occurring both before and after G44
synthesis.
An advantage to tracking two bonds is that at least one of the two
NTP-driven steps we observe in G44 synthesis, specifically the step
from G44
As noted by Kornberg and colleagues (3, 50), in structures of bacterial
Thermus aquaticus RNAP, the bridge
Modeling NTP loading indicates the importance of conserved S. cerevisiae Rpb1 bridge helix residues
Ile825-Thr831, particularly S. cerevisiae Rpb1 Ala828 and Thr831
(H. sapiens Rpb1 Ala851 and Thr854;
E. coli
It appears that stalled RNAP II ECs may tend to revert to the
pre-translocated state, as observed previously with T7 RNAP (53).
Inspection of the S. cerevisiae RNAP II EC structure
indicates that the RNA-DNA hybrid might be spring-loaded against the
bridge NTP-induced Translocation Enhances Fidelity--
We argue above
that the unusual dependence of H. sapiens RNAP II on
substrate NTP concentration should perhaps not be attributed to
allostery but rather to NTP-induced translocation of the RNAP II active
site. But, if our view is correct, why then is translocation dependent
on prior binding of the next templated substrate NTP? The images we
present in Fig. 5 indicate an assembly line model in which templated
NTPs pre-load up to three at a time, and translocation is continually
driven by the incoming n + 1 substrate NTP. With each
translocation event, a DNA base becomes single-stranded and is expected
to pair to its cognate NTP. In addition to the efficiency of the
molecular assembly line we describe, we propose that NTP-induced translocation is a mechanism for maintaining fidelity of NMP
incorporation during RNA synthesis.
Fidelity is the selection of the specified, templated n + 1 NTP rather than any of the three incorrect choices. We argue that NTP-induced translocation renders mis-incorporation unlikely because the n + 1 base pair must be maintained throughout its long
passage over the bridge DNAP and RNAP Mechanisms--
In the mechanisms of simple DNAPs,
induced fit has been interpreted as a dNTP-induced structural change
preceding chemistry, to convert the EC from a "relaxed"
to a "taut" conformation, also referred to as
conformational coupling or induced fit (22, 25, 27-29). This change is
proposed to enhance fidelity, because, when a dNTP is mis-loaded, the
conformational change is blocked, and the mis-paired substrate can be
dissociated before mis-incorporation occurs. In this paper, we argue
that H. sapiens RNAP II utilizes a substrate NTP-induced fit
mechanism that involves NTP-driven translocation of the RNA-DNA hybrid
past the RNAP II active site. Our mechanism, however, does not preclude
a relaxed antigen and transcription factor IIF. Because recovery from a stall and
the processive transition from one bond to the next can be highly
NTP-dependent, we conclude that translocation can be driven
by the incoming substrate NTP, a model fully consistent with the RNAP
II elongation complex structure.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
antigen (HDAg) strongly stimulates RNAP II elongation in
vitro (6, 9). HDAg is the sole gene product of the small RNA
genome of hepatitis
virus, which is maintained as a satellite
particle by hepatitis B virus. The role of HDAg in elongation may be
clinically significant because hepatitis
virus often complicates
severe and chronic presentations of human hepatitis B virus infection. The general cellular transcription factor IIF (TFIIF) has been shown to
stimulate RNAP II elongation 5-10-fold in vitro, by
suppressing transcriptional pausing (10-16). The role of TFIIF in
elongation may be of particular importance during the promoter escape
phase of the transcription cycle (17, 18). Here viral HDAg and cellular TFIIF are used as probes of H. sapiens RNAP II elongation.
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-32P]CTP, and 20 µM UTP.
ECs were then washed with 1% Sarkosyl and 0.5 M KCl, to
dissociate initiation, elongation, pausing, and termination factors,
and then re-equilibrated with transcription buffer. To commence the
reaction, C40 ECs were incubated in transcription buffer containing 8 mM MgCl2 for 10-40 min (the time varies
according to the time required for individual sample handling) in the
presence of 12 pmol of TFIIF or 77 pmol of HDAg (functionally
saturating amounts) and 20 µM (initial working
concentration) CTP and UTP (to maintain ECs at C40). Downstream of C40
the sequence is 40CAAAGG45. On the bench top,
an equal sample volume of 200 µM ATP (initial working
concentration 100 µM) was added in transcription buffer. The ATP pulse time was 60 s with HDAg, 30 s with TFIIF, and
120 s in the absence of an elongation factor. Times were optimized for each procedure (23). During the pulse, EC samples were injected into the left sample port of the Kintek Rapid Chemical Quench-Flow (RQF-3) instrument and mixed with GTP added from the right sample port.
Due to subsequent equal volume mixing events, the final working NTP
concentrations are 5 µM CTP, 5 µM UTP, 50 µM ATP and the indicated concentration of GTP. Rate
measurements from 0.002 to 4 s were done using the KinTek
instrument, and longer time points were done on the bench top, all at
25 °C.
1, and the fastest rate on the
slow pathway (A43a·GTP
A43b·GTP) is >20 s
1. A
slow step after chemistry is required to fit the sigmoidal shape of G45
rate curves (Figs. 3B and 4A). The rate of this
slow step is well determined as 15.5 ± 0.5 s
1.
1 (not shown) and is estimated as 1200 s
1 in the model shown in Fig. 4B. The model
independent analysis estimates the slower pathway from A43a as >100
s
1 (not shown). Furthermore, there is a requirement for
loading two substrate GTPs in a single G44 bond addition cycle. The GTP requirement is fulfilled by loading both the n + 1 and
n + 2 GTPs to A43a before translocation. As expected from
the S. cerevisiae RNAP II EC structure, GTPs held primarily
by base pairing are bound weakly (GTP off-rates of 10,000 s
1; the DYNAFIT program would select faster dissociation
rates). Elongation rate data in the presence of TFIIF (Figs.
3D and 4B) require a fast elongation pathway that
dominates at high GTP concentrations and a slow pathway that dominates
at limiting GTP concentrations.
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
G45, including a translocation of the RNA-DNA hybrid and template
DNA.
View larger version (32K):
[in a new window]
Fig. 1.
Running start, two-bond protocol. This
method was necessary to measure rapid rates of RNAP II
elongation.
View larger version (49K):
[in a new window]
Fig. 2.
RNAP II elongation stimulated by HDAg and
TFIIF. A, running start, two-bond protocol.
B, RNAP II elongation in the absence of a stimulatory
factor, GTP = 250 µM. C, RNAP II
elongation stimulated by HDAg, GTP = 250, 10, and 5 µM. D, RNAP II elongation stimulated by TFIIF,
GTP = 250, 10, and 5 µM. 0* indicates no
ATP pulse, no GTP chase. GTP chase times are in seconds.
View larger version (54K):
[in a new window]
Fig. 3.
Regulation of GTP-dependent steps
in G44 and G45 synthesis by HDAg and TFIIF. A, the rate
of G44 synthesis (G44+ % versus time) in the presence of
HDAg. Fraction b (0-27% of total ECs) and fraction
a (27-50% of total ECs) are indicated. B, the rate of
G45 synthesis (G45+ %) in the presence of HDAg. Rate curves for
A and B were drawn according to the kinetic model
shown in Fig. 4A. C, the rate of G44 synthesis in the
presence of TFIIF. Fraction b (0-10% of total ECs) and
fraction a (10-65% of total ECs) are indicated.
D, the rate of G45 synthesis in the presence of TFIIF. Rate
curves for C and D were drawn according to the
kinetic model shown in Fig. 4B. Corresponding residuals are
shown below each graph, indicating the quality of modeling
to the data.
View larger version (24K):
[in a new window]
Fig. 4.
The simplest kinetic mechanisms for HDAg- and
TFIIF-stimulated elongation. A, HDAg-stimulated
mechanism. B, TFIIF-stimulated mechanism. C,
equilibrium sliding model. The dominant pathways at high GTP
concentration are shaded in gray. Black
triangles indicate GTP-dependent rates, with units of
µM 1 s
1. All other rate
constants are in units of s
1. Estimated
Kd for GTP binding are indicated in
boxes. Estimated rate constants are also shown for
GTP-dependent steps because these are the values used in
DYNAFIT simulations. Unidirectional arrows indicate normally
irreversible steps. According to our interpretation, A43a is
pre-translocated; A43b is post-translocated; A43c is paused; G44d may
be a taut conformation of RNAP II that is relaxed in the
slow step after chemistry. The pausing pathway can lead to backtracking
and RNA cleavage. The equilibrium sliding model is not consistent with
RNAP II elongation rate data, which demands alternate
GTP-dependent routes from A43a and A43b, as shown in
A and B.
1 (see under
"Experimental Procedures")). An additional 23% of A43 ECs are more
dependent on GTP substrate to advance rapidly (Fig. 3A,
fraction a; 27-50% of total ECs;
kobs,slow >20 s
1 (see under
"Experimental Procedures")). Because two classes of A43 EC are
detected with different responsiveness to GTP concentration, this is
evidence of induced fit in the RNAP II mechanism. GTP binds to a less
highly poised EC (fraction a) and converts it to a more highly poised
EC (fraction b). In the presence of HDAg, 50% of A43 ECs are initially
paused (Fig. 3A; 50-100% of total ECs;
kforward ~0.05 s
1).
1 (see under
"Experimental Procedures")). These A43 ECs are highly poised to
bind GTP substrate and incorporate GMP. A distinct fraction of A43 EC
(about 55% of total) elongates rapidly at high GTP concentrations but
much more slowly at low GTP concentrations (Fig. 3C,
fraction a; 10-65% of total ECs;
kobs,slow >100 s
1). This fraction
of A43 EC (fraction a) requires prior GTP binding to convert to a
catalytically competent state (fraction b), consistent with a substrate
GTP-induced fit mechanism for RNAP II elongation. The remaining 35% of
A43 ECs (Fig. 3C; 65-100% of total ECs;
kforward ~0.09 s
1) are strongly
paused but eventually can be extended (Fig. 2D). Because
multiple A43 ECs are detected that respond differently to substrate GTP
concentrations, with both HDAg and TFIIF, the RNAP II EC assumes
conformations that must first bind GTP and then be converted to a form
capable of catalyzing phosphodiester bond formation.
1 (Fig. 4A).
A43b is a conformational
change that may equate with the translocation step. A43a requires prior
GTP binding to make this conformational transition rapidly; otherwise,
the transition is slow. A43b is more highly poised on the forward
synthesis pathway and apparently has already undergone this
conformational transition but is yet capable of binding GTP and
engaging in chemistry. Both the A43a and A43b conformational states are
therefore available to bind GTP in the proposed mechanism. The simple
induced fit mechanism (see Fig. 4A) will account very well
for rates of G44 and G45 synthesis by RNAP II stimulated by HDAg (Fig.
3, A and B). The simple induced fit mechanism
will account for G44 synthesis rates by TFIIF (not shown) but not for
the rates of G45 synthesis, which requires GTP dependence at both the
beginning and end of the G44 bond addition cycle.
G45 synthesis. In the mechanism shown in Fig. 4B, both the
n + 1 GTP and the n + 2 GTP pre-load and then,
in turn, drive translocation of the RNA-DNA hybrid past the RNAP II
active site, establishing the conditions for chemistry. Consistent with
the model in Fig. 4B, the data in Fig.
3D demand two forward pathways for chemistry in the presence
of TFIIF: a faster pathway supported by higher GTP concentrations and a
slower pathway that predominates at limiting GTP concentrations. The HDAg-stimulated rate data only require a single pathway after chemistry
for data fitting. Although our model in Fig. 4B
recapitulates the general shapes of rate curves, the lags observed in
the initial appearance of G45 are more highly GTP-dependent
than we can describe with this kinetic model. This result means that
the TFIIF-stimulated mechanism is more highly
GTP-dependent, at this slow, regulated step from G44
G45 (translocation), than can be accounted for by the simulation shown
in Fig. 4B, a model that confers high GTP dependence.
Therefore, in the presence of TFIIF, translocation of the RNA-DNA
hybrid at this position is a slow step in the mechanism that is highly
dependent on the incoming substrate GTP.
G44 and G44
G45
bond addition cycles, and these experimental observations cannot be
reconciled with an equilibrium sliding model.
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
G45
requires translocation, movement of the RNA-DNA hybrid relative to the RNAP II active site must occur during this GTP-driven phase of the bond
addition cycle. By stalling the EC at A43, however, it appears that we
have established a situation in which formation of G44 can include two
GTP-driven translocations. The first translocation is driven by the
incoming substrate GTP for G44 synthesis, and the second translocation
is driven by the incoming GTP substrate for G45 synthesis. That both
the early and ending phases of the G44 bond addition cycle are
GTP-dependent, in the TFIIF-stimulated reaction, fully
establishes that translocation can depend on a GTP-driven
conformational change in the RNAP II mechanism. We suggest that the
GTP-induced step is translocation, rather than another obligatory but
unknown conformational change that must precede or follow
translocation. Recovery from a stall at A43 and processive elongation
from G44
G45 are likely to be different processes, because the G44
G45 transition requires translocation and pyrophosphate release.
Recovery from a stall at A43 is likely to require translocation for a
fraction of ECs but not pyrophosphate release, which is completed
during the stall. Furthermore, these steps are regulated differently by
TFIIF and HDAg.
-helix to partially close the
catalytic site to dNTP binding (see Refs. 25, 27-29, 40-43). In the
RNAP II NTP-induced fit mechanism, the active site remains open to
substrate binding.
-helix is wedged against the RNA-DNA hybrid (see below). In
such a case, when elongation is stalled, the EC reverts primarily to
the pre-translocated state, and the next templated NTP substrate is
required to drive translocation, as observed in our experiment. This
idea is consistent with our observation that the A43a
(pre-translocated) EC has 55% occupancy in the presence of TFIIF and
only 23% occupancy in the presence of HDAg (Figs. 3 and 4). We believe
that HDAg may loosen the RNAP II clamp to facilitate RNA-DNA hybrid
translocation. In such a case, elongation becomes less dependent on the
prior loading of the next templated NTP because the incoming NTP is no
longer required to force the RNA-DNA hybrid away from the bridge
-helix (see below). We do not believe that the RNAP II mechanism is
truly different in the presence of HDAg and TFIIF. Rather, in the
presence of HDAg, the model appears simpler because loosening of the
RNAP II clamp stimulates translocation and accelerates the slow
elongation pathway, so that it can no longer be distinguished from the
rapid pathway. In support of this idea, in both mechanisms, NTP
substrate-induced fit is observed for G44 synthesis, and a slow step is
observed after G44 chemistry that precedes the first appearance of G45. Therefore, the basic RNAP II mechanism is similar in the presence of
TFIIF or HDAg. In the HDAg-stimulated mechanism, a downside to
relieving tension on the RNAP II clamp may be that the geometry of the
active site becomes less optimal for chemistry, decreasing the fastest
elongation rates, and fidelity may be sacrificed. In hepatitis
virus RNA replication and transcription, HDAg is thought to convert
H. sapiens RNAP II from a DNA template-dependent RNAP to an RNA template-dependent RNAP (9, 44-47), and
loosening the RNAP II clamp may facilitate this loss of fidelity in
template selection.
View larger version (53K):
[in a new window]
Fig. 5.
Pre-loading substrate NTPs through the
main channel and substrate NTP-driven translocation are consistent with
the S. cerevisiae RNAP II EC structure.
A, S. cerevisiae RNAP II pre-translocated EC,
showing that the n + 1, n + 2, and n + 3 NTPs can load through the main enzyme channel to pair with their DNA
cognate bases. The left panel is a simplified view of the
stereo image shown in the right two panels. The main channel
is in front; the secondary pore behind the images shown. The
n base pair (n = RNA length) is
gold; template strand DNA is green; RNA is
red; incoming NTP substrates are dark blue
(n + 1) or light blue (n + 2 and
n + 3); the bridge helix is yellow; other protein
is white; Mg2+-A (the active site
Mg2+) is magenta. B, pre-loading
substrate NTPs into the pre-translocated complex. In the
pre-translocated state, the n base pair (gold)
blocks loading of the incoming n + 1 NTP into the active
site. Only relevant protein and side chains are shown
(yellow). The main channel is in front, and the secondary
pore is behind the bridge helix in this view. C and
D, two views of a post-translocated complex. C,
main channel in front and secondary pore behind the bridge helix (as in
B). D, main channel to the right and secondary
pore to the left of the bridge helix, which is viewed end-on.
Translocation of the n base pair opens a pore for loading
the triphosphate of the n + 1 NTP over the bridge -helix
pulling the n + 1 base pair into position to form the next
phosphodiester bond. Colors of atoms is as follows: carbon is
green; oxygen is red; nitrogen is
blue; phosphorus is magenta (small
spheres).
G45, must include a translocation. If either the
n + 1 or n + 2 GTPs pair to template prior to
translocation, these bases must load through the main channel rather
than through the secondary pore, because otherwise base pairing is
impossible. In the S. cerevisiae RNAP II EC structure,
before translocation, the n + 1 DNA base is oriented toward
the main enzyme channel and is not accessible to the secondary pore. We
believe that the secondary pore is normally the route for pyrophosphate
release and for release of improperly loaded NTPs that are transferred into the active site but cannot subsequently be incorporated into RNA.
We suggest the following: 1) substrate NTPs normally enter the active
site through the main channel, in much the same direction of flow as
downstream template DNA; and 2) substrate NTPs are continuously
pre-loaded in bunches of two or three (Fig. 5). Continuous maintenance
of two or three paired NTPs supports efficiency and fidelity of NMP
incorporation, because accurate base pairing is confirmed and
re-confirmed before loading of a base pair into the active site. No
structural barrier is apparent to loading the n + 1, n + 2, and n + 3 NTPs through the main channel or
to pair them to their DNA cognate bases (Fig. 5A). In the
S. cerevisiae RNAP II EC structure (3), the only observed
barrier to moving the n + 1 NTP-DNA base pair from the main
channel into the catalytic center is the n base pair (Fig.
5, A and B, gold), and the barrier is
removed when the RNA-DNA hybrid translocates (Fig. 5, C and D). Once the n base pair is driven into the
post-translocated position, a pore opens for loading the n + 1 NTP-DNA base pair. We detect no steric or electrostatic barriers to
loading the triphosphate of the n + 1 NTP through the
induced pore, and the amino acid residues that constitute the pore are
conserved in multisubunit RNAPs from bacteria to man (50).
-helix (also called
the F helix) may be bent because of contact with the G-loop or
"trigger" loop protruding from below (48, 49, 51). Bending of the
bridge
-helix in the T. aquaticus RNAP structure prompted Gnatt et al. (3) to propose that bending of the bridge helix might be part of the translocation mechanism. Although structural and
cross-linking studies indicate that the bridge helix is capable of
flexure (3, 48, 49, 51), bending of the bridge helix may not be
necessary during each translocation cycle. As indicated in Fig. 5,
passage of the n + 1 NTP triphosphate over the bridge
-helix may induce movement of the n base pair past the
active site Mg2+ and rotate the n + 1 base pair
into the active site for phosphodiester bond formation, without bridge
helix bending. The most extreme conformation of the bridge helix, seen
in T. aquaticus RNAP structures (48, 52), will not easily
accommodate our model for NTP loading, but conformations in the
S. cerevisiae RNAP II and Thermus thermophilus RNAP structures appear completely consistent with our model (3, 49).
The most strongly bent configurations of the bridge helix are expected
to block movement of the n + 1 DNA base into position, because bending the helix would rotate S. cerevisiae Rpb1
Lys830 (H. sapiens Lys853, E. coli
' Lys789) into the space through which
the DNA base must pass. Thus, the original suggestion that NTPs load
through the secondary pore was made based on a structure with a
severely bent bridge helix that may be incompatible with ongoing
transcription (48, 52).
' Ala787 and Thr790),
which protrude most directly into the space through which the triphosphate of the incoming NTP must pass. Conserved charged residues, S. cerevisiae Asp826 and
Lys830, point down from a straight bridge helix (in the
view shown), away from the proposed NTP loading pore, and are not
expected to complicate or facilitate triphosphate passage, unless the
bridge helix bends. A conserved arginine residue, S. cerevisiae Rpb2 Arg512 (H. sapiens Rpb2
Arg499, E. coli
Arg548) may play
a "gatekeeper" role for NTP loading through the main channel.
S. cerevisiae Rpb2 Arg512 appears to be
positioned to interact with the triphosphate of the n + 1 NTP (Fig. 5B) and then switch, in turn, to contact the triphosphate of the incoming n + 2 NTP, as the
n + 1 NTP moves over the bridge helix and the n + 2 NTP makes its approach (Fig. 5, C and D).
-helix as the RNA-DNA hybrid is pressed by the flexible RNAP II clamp (3). Furthermore, the stalled S. cerevisiae RNAP II EC is found primarily in the pre-translocated form with the active site
Mg2+-A atom poised over the previously formed
phosphodiester bond. In our kinetic analysis, a large fraction of A43
complexes initially appear to be in the pre-translocated A43a state,
even though an adequate kinetic model suggests that elongation passed
through the post-translocated A43b state to access the A43a state (Fig. 4, A and B). The n + 1 DNA base is
rotated about 90° away from the active site Mg2+ atom.
Therefore, catalysis requires that the n + 1 DNA base
first swing into position between the RNA-DNA hybrid and the highly conserved bridge
-helix. Our kinetic data support the idea that the incoming NTP normally pairs with its cognate DNA template base
prior to translocation of the RNA-DNA hybrid. Formation of the
n + 1 base pair, therefore, is expected to facilitate its rotation into the active site pocket. We suggest that base pairing and
movement of the n + 1 NTP over the bridge
-helix provides a major driving force to induce translocation of the RNA-DNA hybrid during rapid catalysis.
-helix (Fig. 5, A-D). To
incorporate AMP for GMP, for instance, would require maintenance of an
unstable dC-ATP base pair throughout movement into the active site.
Most likely, substrate NTP-induced translocation developed as a
mechanism to ensure fidelity of RNA synthesis, because this mechanism
increases efficiency and accuracy by allowing pre-loading and
pre-aligning of multiple NTPs. By using this mechanism, fidelity can be
maintained for ATP, GTP, CTP, and UTP substrates without strict
chemical recognition of individual bases in the RNAP II active site,
which would compromise the flexibility that RNAPs require to utilize four chemically distinct substrates. Very tight binding of substrate is
not required for specificity by RNAPs because recognition of each NTP
substrate is driven primarily by accurate base pairing, through the
NTP-induced translocation mechanism. Incorrectly selected n + 1 NTPs do not remain paired long enough to support translocation, while correctly pairing the specified n + 1 NTP forces the
RNAP into the only conformation capable of forming a phosphodiester bond. Adequate kinetic models indicate that GTP affinity may vary significantly over the course of the reaction (Fig. 4, A and
B). In the TFIIF-stimulated reaction, modeling indicates
that GTP affinity might be highest (estimated as Kd
~6 µM) to the fully translocated substrate-binding
site, as expected from the RNAP II EC structure and the NTP-induced
translocation model, in which NTPs initially load primarily by base
pairing before loading into the RNAP II active site. Because
polymerases utilize four chemically distinct substrates, DNAPs and
RNAPs are characterized by reaction mechanisms that maintain high
fidelity with relatively low substrate affinity. The NTP-induced
translocation model shows one way this biological requirement can be achieved.
taut conformational change that
tightens the active site just prior to chemistry, and we deem such a
conformational change likely. For instance, because translocation by
RNAP II involves a slow, substrate NTP-induced conformational change,
this implies that loosening of the RNAP II clamp and/or a
taut
relaxed RNAP II conversion may accompany NTP-driven translocation. If this is the case, a relaxed
taut conversion must also precede chemistry. For RNAP II,
pyrophosphate release may follow and depend upon a taut
relaxed conversion after chemistry.
![]() |
ACKNOWLEDGEMENTS |
---|
We are grateful to Dorothy Erie for sharing ideas and data prior to publication. We thank Yue Li for help with the operation and calibration of the KinTek RQF-3.
![]() |
FOOTNOTES |
---|
* This work was supported in part by a grant from the National Institutes of Health (to Z. F. B.).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.
§ Both authors contributed equally to this work.
¶ Supported by the Michigan State University Development Initiative in Gene Expression in Development and Disease.
§§ Recipient of support from the Agricultural Experiment Station at Michigan State University. To whom correspondence should be addressed. Tel.: 517-353-0859; Fax: 517-353-9334; E-mail: Burton@msu.edu.
Published, JBC Papers in Press, March 13, 2003, DOI 10.1074/jbc.M301103200
2 D. Erie, personal communication.
![]() |
ABBREVIATIONS |
---|
The abbreviations used are:
RNAP II, RNA
polymerase II;
EC, elongation complex;
HDAg, hepatitis antigen;
TFIIF, transcription factor IIF;
DNAP, DNA polymerase.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Johnson, K. A. (1992) Enzymes 20, 1-61 |
2. | Johnson, K. A. (1995) Methods Enzymol. 249, 38-61[Medline] [Order article via Infotrieve] |
3. |
Gnatt, A. L.,
Cramer, P.,
Fu, J.,
Bushnell, D. A.,
and Kornberg, R. D.
(2001)
Science
292,
1876-1882 |
4. | Conaway, J. W., Shilatifard, A., Dvir, A., and Conaway, R. C. (2000) Trends Biochem. Sci. 25, 375-380[CrossRef][Medline] [Order article via Infotrieve] |
5. |
Shilatifard, A.
(1998)
FASEB J.
12,
1437-1446 |
6. | Yamaguchi, Y., Delehouzee, S., and Handa, H. (2002) Microbes Infect. 4, 1169-1175[CrossRef][Medline] [Order article via Infotrieve] |
7. | Kim, D. K., Yamaguchi, Y., Wada, T., and Handa, H. (2001) Mol. Cells 11, 267-274[Medline] [Order article via Infotrieve] |
8. |
Krogan, N. J.,
Kim, M.,
Ahn, S. H.,
Zhong, G.,
Kobor, M. S.,
Cagney, G.,
Emili, A.,
Shilatifard, A.,
Buratowski, S.,
and Greenblatt, J. F.
(2002)
Mol. Cell. Biol.
22,
6979-6992 |
9. |
Yamaguchi, Y.,
Filipovska, J.,
Yano, K.,
Furuya, A.,
Inukai, N.,
Narita, T.,
Wada, T.,
Sugimoto, S.,
Konarska, M. M.,
and Handa, H.
(2001)
Science
293,
124-127 |
10. |
Renner, D. B.,
Yamaguchi, Y.,
Wada, T.,
Handa, H.,
and Price, D. H.
(2001)
J. Biol. Chem.
276,
42601-42609 |
11. |
Tan, S.,
Aso, T.,
Conaway, R. C.,
and Conaway, J. W.
(1994)
J. Biol. Chem.
269,
25684-25691 |
12. |
Lei, L.,
Ren, D.,
and Burton, Z. F.
(1999)
Mol. Cell. Biol.
19,
8372-8382 |
13. | Bengal, E., Flores, O., Krauskopf, A., Reinberg, D., and Aloni, Y. (1991) Mol. Cell. Biol. 11, 1195-1206[Medline] [Order article via Infotrieve] |
14. |
Izban, M. G.,
and Luse, D. S.
(1992)
J. Biol. Chem.
267,
13647-13655 |
15. | Price, D. H., Sluder, A. E., and Greenleaf, A. L. (1989) Mol. Cell. Biol. 9, 1465-1475[Medline] [Order article via Infotrieve] |
16. |
Funk, J. D.,
Nedialkov, Y. A.,
Xu, D.,
and Burton, Z. F.
(2002)
J. Biol. Chem.
277,
46998-47003 |
17. |
Yan, Q.,
Moreland, R. J.,
Conaway, J. W.,
and Conaway, R. C.
(1999)
J. Biol. Chem.
274,
35668-35675 |
18. | Dvir, A. (2002) Biochim. Biophys. Acta 1577, 208-223[Medline] [Order article via Infotrieve] |
19. | Shapiro, D. J., Sharp, P. A., Wahli, W. W., and Keller, M. J. (1988) DNA (New York) 7, 47-55[Medline] [Order article via Infotrieve] |
20. | Wang, B. Q., Kostrub, C. F., Finkelstein, A., and Burton, Z. F. (1993) Protein Expression Purif. 4, 207-214[CrossRef][Medline] [Order article via Infotrieve] |
21. | Wang, B. Q., Lei, L., and Burton, Z. F. (1994) Protein Expression Purif. 5, 476-485[CrossRef][Medline] [Order article via Infotrieve] |
22. | Johnson, K. A. (1993) Annu. Rev. Biochem. 62, 685-713[CrossRef][Medline] [Order article via Infotrieve] |
23. | Nedialkov, Y. A., Gong, X. Q., Yamaguchi, Y., Handa, H., and Burton, Z. F. (2003) Methods Enzymol., in press |
24. | Foster, J. E., Holmes, S. F., and Erie, D. A. (2001) Cell 106, 243-252[Medline] [Order article via Infotrieve] |
25. | Dunlap, C. A., and Tsai, M. D. (2002) Biochemistry 41, 11226-11235[CrossRef][Medline] [Order article via Infotrieve] |
26. | Kuzmic, P. (1996) Anal. Biochem. 237, 260-273[CrossRef][Medline] [Order article via Infotrieve] |
27. | Patel, S. S., Wong, I., and Johnson, K. A. (1991) Biochemistry 30, 511-525[Medline] [Order article via Infotrieve] |
28. | Wong, I., Patel, S. S., and Johnson, K. A. (1991) Biochemistry 30, 526-537[Medline] [Order article via Infotrieve] |
29. | Showalter, A. K., and Tsai, M. D. (2002) Biochemistry 41, 10571-10576[CrossRef][Medline] [Order article via Infotrieve] |
30. | Guthold, M., and Erie, D. A. (2001) Chembiochem 2, 167-170[CrossRef][Medline] [Order article via Infotrieve] |
31. | Guajardo, R., Lopez, P., Dreyfus, M., and Sousa, R. (1998) J. Mol. Biol. 281, 777-792[CrossRef][Medline] [Order article via Infotrieve] |
32. | Guajardo, R., and Sousa, R. (1997) J. Mol. Biol. 265, 8-19[CrossRef][Medline] [Order article via Infotrieve] |
33. |
Forde, N. R.,
Izhaky, D.,
Woodcock, G. R.,
Wuite, G. J.,
and Bustamante, C.
(2002)
Proc. Natl. Acad. Sci. U. S. A.
99,
11682-11687 |
34. |
Wang, M. D.,
Schnitzer, M. J.,
Yin, H.,
Landick, R.,
Gelles, J.,
and Block, S. M.
(1998)
Science
282,
902-907 |
35. | Wang, H. Y., Elston, T., Mogilner, A., and Oster, G. (1998) Biophys. J. 74, 1186-1202[Abstract] |
36. | Yin, H., Wang, M. D., Svoboda, K., Landick, R., Block, S. M., and Gelles, J. (1995) Science 270, 1653-1657[Abstract] |
37. |
von Hippel, P. H.
(1998)
Science
281,
660-665 |
38. |
Julicher, F.,
and Bruinsma, R.
(1998)
Biophys. J.
74,
1169-1185 |
39. |
Adelman, K.,
La Porta, A.,
Santangelo, T. J.,
Lis, J. T.,
Roberts, J. W.,
and Wang, M. D.
(2002)
Proc. Natl. Acad. Sci. U. S. A.
99,
13538-13543 |
40. |
Li, Y.,
Korolev, S.,
and Waksman, G.
(1998)
EMBO J.
17,
7514-7525 |
41. |
Li, Y.,
and Waksman, G.
(2001)
Protein Sci.
10,
1225-1233 |
42. | Doublie, S., Tabor, S., Long, A. M., Richardson, C. C., and Ellenberger, T. (1998) Nature 391, 251-258[CrossRef][Medline] [Order article via Infotrieve] |
43. | Arndt, J. W., Gong, W., Zhong, X., Showalter, A. K., Liu, J., Dunlap, C. A., Lin, Z., Paxson, C., Tsai, M. D., and Chan, M. K. (2001) Biochemistry 40, 5368-5375[CrossRef][Medline] [Order article via Infotrieve] |
44. | Fu, T. B., and Taylor, J. (1993) J. Virol. 67, 6965-6972[Abstract] |
45. |
Chang, J.,
and Taylor, J.
(2002)
EMBO J.
21,
157-164 |
46. |
Macnaughton, T. B.,
Shi, S. T.,
Modahl, L. E.,
and Lai, M. M.
(2002)
J. Virol.
76,
3920-3927 |
47. |
Modahl, L. E.,
Macnaughton, T. B.,
Zhu, N.,
Johnson, D. L.,
and Lai, M. M.
(2000)
Mol. Cell. Biol.
20,
6030-6039 |
48. | Zhang, G., Campbell, E. A., Minakhin, L., Richter, C., Severinov, K., and Darst, S. A. (1999) Cell 98, 811-824[Medline] [Order article via Infotrieve] |
49. | Vassylyev, D. G., Sekine, S., Laptenko, O., Lee, J., Vassylyeva, M. N., Borukhov, S., and Yokoyama, S. (2002) Nature 417, 712-719[CrossRef][Medline] [Order article via Infotrieve] |
50. |
Cramer, P.,
Bushnell, D. A.,
and Kornberg, R. D.
(2001)
Science
292,
1863-1876 |
51. | Epshtein, V., Mustaev, A., Markovtsov, V., Bereshchenko, O., Nikiforov, V., and Goldfarb, A. (2002) Mol. Cell 10, 623-634[Medline] [Order article via Infotrieve] |
52. | Korzheva, N., Mustaev, A., Nudler, E., Nikiforov, V., and Goldfarb, A. (1998) Cold Spring Harbor Symp. Quant. Biol. 63, 337-345[Medline] [Order article via Infotrieve] |
53. | Huang, J., and Sousa, R. (2000) J. Mol. Biol. 303, 347-358[CrossRef][Medline] [Order article via Infotrieve] |