Trigger loop dynamics mediate the balance
between the transcriptional fidelity and
speed of RNA polymerase II
Matthew H. Larsona,1, Jing Zhoub,1, Craig D. Kaplanc,1, Murali Palangatd,2, Roger D. Kornberge,
Robert Landickd, and Steven M. Blocka,b,f,3
University of Wisconsin-Madison, Madison, WI 53706
bDepartment of Applied Physics,
cDepartment of Biochemistry and Biophysics, Texas A&M University, College Station, TX 77843; and
eDepartment of Structural Biology, and
fDepartment of Biology, Stanford University, Stanford,
dDepartment of Biochemistry,
Edited by* E. Peter Geiduschek, University of California, La Jolla, CA, and approved February 28, 2012 (received for review January 18, 2012)
During transcription, RNA polymerase II (RNAPII) must select the
correct nucleotide, catalyze its addition to the growing RNA tran-
script, and move stepwise along the DNA until a gene is fully tran-
scribed.In all kingdomsoflife, transcriptionmustbefinelytunedto
ensure an appropriate balance between fidelity and speed. Here,
we used an optical-trapping assay with high spatiotemporal reso-
lution to probe directly the motion of individual RNAPII molecules
as they pass through each of the enzymaticsteps of transcript elon-
gation. We report direct evidence that the RNAPII trigger loop, an
evolutionarily conserved protein subdomain, serves as a master
regulator of transcription, affecting each of the three main phases
of elongation, namely: substrate selection, translocation, and
catalysis. Global fits to the force-velocity relationships of RNAPII
and its trigger loop mutants support a Brownian ratchet model
for elongation, where the incoming NTP is able to bind in either
the pre- or posttranslocated state, and movement between these
two states is governed by the trigger loop. Comparison of the
kinetics of pausing by WTand mutant RNAPII under conditions that
promote base misincorporation indicate that the trigger loop gov-
erns fidelity in substrate selection and mismatch recognition, and
thereby controls aspects of both transcriptional accuracy and rate.
optical trap ∣ optical tweezers ∣ Pol II
fidelity. RNA polymerase (RNAP) incorporates nucleotides into
nascent RNA chains at rates of 10–70s−1(1, 2), but only inserts
the wrong nucleotide approximately once per 100,000 bases,
on average (3). Although the energetics of base-pairing is funda-
mental to transcription fidelity, the discrimination for correctly
matched nucleotide substrates is kinetically controlled, and
accomplished by active site conformational changes centered on
the trigger loop (TL) (4–7). The TL is an evolutionarily conserved
mobile element that stabilizes substrate NTPs in the active site of
a variety of multisubunit RNA polymerases, including eukaryotic
RNAP I, II, and III, as well as bacterial and archaeal RNAP (8).
In the case of RNAPII, alteration of the TL has important
consequences for activity and fidelity. TL residue His1085, for
example, interacts with the bound NTP substrate, and is fully con-
served among multisubunit RNA polymerases. The importance
of this residue is underscored by the lethality of the H1085A
substitution in yeast (7) and catalytic defects resulting from sub-
stitutions at this position for both the yeast and bacterial enzymes
(7, 9–11). Additionally, a substitution for residue Glu1103 of
the TL has been found not only to promote nucleotide misincor-
poration (6, 7), but also to increase the elongation rate (6, 7, 12),
properties that are jointly consistent with the notion of kinetic
proofreading (13). To explore further the interplay between
elongation rate and transcriptional fidelity, we determined the
effect of E1103G and H1085A substitutions on the nucleotide-
addition cycle (NAC) using single-molecule assays.
he transcription of genetic information stably encoded in
DNA into a transient RNA message occurs with remarkable
A key step in the NAC for all polymerases that remains poorly
characterized is translocation along the DNA. Traditional ensem-
ble techniques do not observe enzyme motions directly, but
instead report the average positions of populations of molecules.
This limitation can be problematic, owing to the intrinsic variabil-
ity in synthesis rates, and for RNAPII, the inherently stochastic
nature of transcriptional pausing (1), which can obscure the
underlying kinetics of translocation. Here, we describe a single-
molecule assay capable of directly observing the motions of indi-
vidual, elongating RNAPII molecules at subnanometer resolution
in real time. Using the assay, we were able to identify and exclude
from analysis transcriptional pauses as short as 1 s, and thereby to
determine the “pause-free” velocity of an individual molecule,
which furnishes an accurate measure of its raw elongation rate.
Our results for the pause-free velocity as a function of applied
force favor a Brownian ratchet model for transcript elongation, in
which the incoming NTPsubstrate is able tobindin either the pre-
or posttranslocated state of RNAPII. By examining the effect of
two TL mutant enzymes—containing either E1103G or H1085A/
E1103G substitutions—on pause-free velocity, we show that the
TL affects NTP binding, translocation and catalysis in the NAC.
Furthermore, the effect of the E1103G substitution on transcrip-
tion provides evidence that the TL promotes fidelity by regulating
nucleotide selection and pausing during elongation, which aids in
the recognition of misincorporation events.
Results and Discussion
Force-Velocity Relationships Distinguish Between Competing Models
for Ordering of Events in the Nucleotide-Addition Cycle. Because
every NAC involves translocation along the DNA template,
the elongation rates of RNA polymerase molecules are sensitive
to modulation by force, which can either assist or hinder trans-
location, depending upon the experimental geometry of the assay
(Fig. 1A). The magnitude of any such modulation, assessed by
determining the force-velocity (F-v) relationship, is affected by
the step size of RNAPII and by its equilibrium between pre-
and posttranslocated states. The F-v relationship is also sensitive
to the temporal ordering of the translocation event with respect
to NTP binding during the NAC when measured as a function of
Author contributions: M.H.L., J.Z., C.D.K., R.D.K., R.L., and S.M.B. designed research; M.H.L.
and J.Z. performed research; C.D.K., M.P., R.D.K., and R.L. contributed new reagents/
analytic tools; M.H.L., J.Z., C.D.K., and M.P. analyzed data; and M.H.L., J.Z., and S.M.B.
wrote the paper.
The authors declare no conflict of interest.
*This Direct Submission article had a prearranged editor.
1M.H.L., J.Z., and C.D.K. contributed equally to this work.
2Present address: Laboratory of Receptor Biology and Gene Expression, National Cancer
Institute, Bethesda, MD 20892.
3To whom correspondence should be addressed. E-mail: email@example.com.
This article contains supporting information online at www.pnas.org/lookup/suppl/
www.pnas.org/cgi/doi/10.1073/pnas.1200939109PNAS Early Edition
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NTP concentration (14), and this placement has been the subject
of some controversy. Kinetic studies have suggested that NTP
binding may precede, and possibly promote, translocation itself
(15–17). Because most substrate-bound RNAPII crystal struc-
tures are in the posttranslocated state, they do not speak to this
issue, but the identification of a nonspecific NTP-binding site in
some of these structures suggests that substrate binding might
precede translocation (18).
We determined the F-v relationships for wild-type (WT) RNA-
PII and two specific TL mutant enzymes, E1103G and H1085A/
E1103G (Fig. 1B) at both saturating and subsaturating NTP levels
(1 mM or 0.1 mM of ATP, CTP, GTP, and UTP, respectively). In
these experiments, we were able to modulate force on the DNA
from 20 pN (assisting load) to −3 pN (hindering load) before
either linkage rupture or irreversible backtracking of RNAPII oc-
curred, respectively. We note that the range of hindering forces is
limited because yeast RNAPII has a higher propensity to back-
track at relatively small hindering loads compared to bacterial
RNAP, consistent with a previous report (19).
Glu1103 lies within a region of the TL that is distal from the
enzyme active site (Fig. 1B), and biochemical data suggest that
E1103G likely affects catalysis via TL dynamics, by biasing the TL
toward a more closed, catalytically active conformation (6, 7).
Previous ensemble measurements have shown that E1103G leads
to an increase in the overall elongation rate (6, 7, 12), which we
confirmed in our single-molecule experiments at saturating NTP
in the presence of 10 pN assisting force (Fig. 1C). We also ac-
quired F-v data in the presence of 100 mM ammonium ion, which
is known to increase elongation rates in vitro to levels closely
resembling transcription rates estimated in vivo (see below)
We considered four different pathways for the NAC to distin-
guish among the competing models for NTP binding and trans-
location (Fig. 2), each of which has been proposed previously for
multisubunit RNA polymerases, based on biochemical or single-
molecule kinetic data (14, 16, 23–28). In Model 1 (Fig.2A), RNA-
PII oscillates back and forth between the pre- and posttranslo-
cated states with thermal motion. That motion is ultimately
rectified in the posttranslocated state by NTP binding (23, 24,27),
after which the NTP is covalently linked into the nascent tran-
script by a condensation reaction catalyzed by RNAPII, and
the NAC is completed with pyrophosphate (PPi) release. In
Model 2 (Fig. 2B), substrate binding occurs prior to translocation,
suggesting that the incoming NTP drives translocation (16), and
the thermal motion between the pre- and posttranslocated states
is rectified by the NTP condensation reaction.
Model 3 (Fig. 2C), where the incoming NTP can bind in either
the pre- or posttranslocated state, is supported by single-molecule
studies of Escherichia coli RNAP (14) and is consistent with stu-
dies of RNAPII that posit the existence of a secondary NTP bind-
ing site, distinct from the primary nucleotide-addition site (18).
(We note that although Model 3 has additional states in the re-
action pathway, fits involve the same number of free parameters,
m ¼ 3, as Models 1 and 2, due to the assumption that NTP bind-
ing to the secondary site is energetically equivalent to active site,
which leads to identical equilibrium constants for NTP binding
pre- and posttranslocation.) Model 4, in which PPirelease can
is inspired by kinetic studies demonstrating that the rate of PPi
release is stimulated by the incoming cognate NTP (26). (Similar
to the assumption made for Model 3, PPirelease is assumed to be
equally likely from either the pre- or posttranslocated state, re-
sulting in a F-v relationship with the same number of free para-
meters as Models 1, 2, and 3.)
Global fits were performed against the models for all the force-
velocity data acquired with wild-type and mutant enzymes: this
arrangement (not to scale). RNAPII (green) is initiated on a nucleic-acid scaffold (dark blue) that includes template and nontemplate DNA strands plus a short
RNA transcript (red). An upstream DNA handle (cyan) and downstream template (black) are ligated to either end of the scaffold. Two polystyrene beads (light
blue) are held in separate optical traps (pink). RNAPII is attached to the smaller bead via a biotin:avidin linkage (yellow and black); the DNA handle is attached
to the larger bead via a digoxigenin:antibody linkage (purple and orange). Direction of transcription is shown (green arrow); in this orientation, tension assists
translocation. (B) Schematic of the RNAPII active site from overlays of two crystal structures, Protein Data Bank (PDB) IDs 2E2H (4) and 1Y1V (41). During
transcription, the TL is proposed to fluctuate between a catalytically active “closed” (magenta, 2E2H) and inactive “open” (light pink, 1Y1V) conformation
(in this case, restrained by the presence of TFIIS; not displayed). During this transition, the TL comes into proximity with the bridge helix (cyan) and the incoming
NTP (orange). The positions of the two TL mutations studied are indicated. (C) Representative records of elongation acquired under 10 pN assisting load for WT
RNAPII (blue) and two TL mutant enzymes, E1103G (red) and H1085A/E1103G (green).
Single-molecule assay and transcription records. (A) Experimental geometry for the DNA-pulling optical-trapping assay, based on a “dumbbell”
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www.pnas.org/cgi/doi/10.1073/pnas.1200939109 Larson et al.
procedure implicitly assumes that enzymes function via the same
reaction pathway, regardless of mutation, albeit with different
rates, and is justified by the goodness of the fit. The best-fit model
(Model 3, Fig. 2C) placed the translocation step at the start of the
NAC, prior to catalysis, with the incoming NTP able to bind
RNAPII in either its pre- or posttranslocated state (reduced
χ2¼ 0.52, ν ¼ 33, N ¼ 45, P-value ¼ 0.99; the three other mod-
els returned P-values < 0.005; Fig. S1). In particular, the F-v
data did not fit an alternative model where NTP binding is re-
quired for forward translocation (16) (Fig. 2B), nor did they sup-
port a model where NTP binds exclusively to the posttranslocated
state (Fig. 2A). The existence of a secondary site for NTP binding,
posited previously in the form of a nontemplated “entry,” or
“E-site” (18), a templated “preinsertion” site (29), or along ex-
posed bases in the downstream DNA template (15), is consistent
with our best-fit model.
Assuming a Boltzmann-type dependence for the load-sensitive
translocation step, the F-v relationship for Model 3 can be ex-
pressed as (for additional description, see ref. 14, SI Text, and
½NTP?f1 þ Kδexpð−Fδ∕kBTÞg þ Kδexpð−Fδ∕kBTÞ; 
where kcatis the rate of nucleotide condensation followed by
PPirelease (these steps have been combined into a single rate;
see SI Text), KDis the NTP dissociation constant, and Kδde-
scribes the equilibrium between pre- and posttranslocated states
(Kδ¼ ½pre?∕½post?). The effective step size, δ, is a parameter cor-
responding to the distance between translocation states; this was
fixed to 1 bp (0.34 nm) based on a previous measurement (14).
We observed a relatively weak effect of force on the pause-free
velocity forthe WT RNAPII (Fig.3A), a dependence that hadnot
been observed previously with RNAPII, presumably due to the
limited range of forces that was explored and the larger experi-
mental uncertainties (19). Based on the fit, WT RNAPII is biased
toward its posttranslocated state (Table 1), indicating that the
translocation step is not rate-limiting for the advance of the
WT enzyme. The apparent KDfor WT RNAPII is somewhat
Model 2: NTP binds before translocation
= 1 bp
Model 1: NTP binds after translocation
= 1 bp
= 1 bp
Model 4: PPi release occurs before or after translocation
= 1 bp
= 1 bp
Model 3: NTP binds before or after translocation
= 1 bp
differs in the placement of the force-dependent translocation step in relation to the NTP-binding, catalysis, and phosphate-release steps. Equilibrium constants
(for reversible steps) and forward rate constants (for essentially irreversible steps) are shown. Groups of states corresponding to the translocation event(s) are
Reaction pathway models of the nucleotide addition cycle. Four variants of the Brownian ratchet model for elongation are presented. Each model
Larson et al. PNAS Early Edition
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higher than that suggested by some previous reports (6); however,
those determinations were carried out for a single NTP species
binding to a given template position, whereas the present value
corresponds to an average of all four NTPs measured over thou-
sands of template positions.
Role of Ammonium in Transcript Elongation. A stimulatory effect of
ammonium on the overall rate of transcription has been pre-
viously reported (20–22), but the mechanism for its action is still
unknown. To assess the role of ammonium on individual steps in
the NAC, we added 100 mM ammonium chloride to the buffer (in
addition to the 130 mM KCl already present) for transcription
reactions of WT RNAPII and its TL mutants. This additional
set of conditions allowed us to discriminate better among the dif-
ferent models for the NAC, as well as to observe transcription by
the H1085A/E1103G mutant enzyme at low NTPs, a condition
which would otherwise produce rates too slow to observe practi-
cally in single-molecule assays due to residual amounts of base-
line drift. Ammonium increased the pause-free velocity by factors
of 1.5 to 4 for WTand mutant RNAPII (Fig. 3 A–C). Based on fits
to Model 3 of the NAC, ammonium ion increases the pause-free
elongation rate by increasing the value of kcatfor the both WTand
mutant RNAPII, leaving KD, and Kδsubstantially unchanged
(Table 1). Furthermore, the maximum pause-free velocities in
the presence of ammonium ion for the WTand E1103G were sta-
tistically indistinguishable, suggesting the existence of a common,
rate-limiting step under conditions where kcatis rapid and an as-
sisting load is applied. Taken together, the results indicate that
ammonium ion primarily alters the rate of the catalytic step of
the nucleotide-addition cycle, and acts independently of trigger
loop changes to E1103 or H1085.
Substitutions in the TL Affect NTP Binding, Catalysis, and Transloca-
tion. The force-velocity dependence of the TL mutant enzymes
(Fig. 3 B and C) confirms that the TL is involved in catalysis
(5, 9, 10), because the E1103G mutation produced a 1.6-fold in-
crease in kcatover the WT (Table 1). We also observed more than
a 2.5-fold decrease in KDcompared to the WT, indicating that
the incoming NTP is more tightly bound to the active site. This
reduction may be attributable to a preferential closure of the
mutant TL around the bound substrate. The increases observed
in both NTP-binding affinity and catalysis were accompanied by
an inhibitory effect on translocation, because the E1103G mutant
enzyme was found to be significantly more biased toward its
pretranslocated state than the WT. A pretranslocational bias is
somewhat counterintuitive, in view of the increased elongation
rate for the mutant enzyme. However, it can be understood be-
cause the TL substitution promotes a net forward translocation
of RNAPII. In the case of E1103G, the combination of tighter
NTP binding and a faster catalytic rate (Table 1) lead to an in-
crease in the elongation rate despite an inhibition of the translo-
cation step. It was previously observed that E1103G displayed a
bias for the pretranslocated state in the absence of the correct
NTP when tested at a single template position, and that this bias
shifted toward the posttranslocated state in the presence of the
correct NTP (6). By contrast, our best-fit model of the NAC,
which supplies uncoupled measures of translocation (Kδ) and
NTP binding (KD) over many different template positions, in-
stead suggests a pretranslocation bias for E1103G, even at high
If the observed properties of the E1103G substitution were at-
tributable to a stabilization of the closed state of the TL (depen-
dent upon a bound, template-matched NTP), which tends to
be more open in the WT, we reasoned that a compensatory de-
stabilization of this closed state might reverse some, or all, of the
mutant enzyme properties. Guided by structural data for the TL
and RNAPII active site (4, 18), we altered a second residue of the
TL, His1085, which, through interaction with NTP substrates,
promotes catalysis and likely favors the closed conformation
of the TL (4, 5, 7, 9–11). H1085A lethality is suppressed by com-
bination with E1103G, but the transcriptional and phenotypic
properties of the double substitution suggest mutual suppression
(11). As anticipated, based on compromise of the critical sub-
strate-interacting sidechain, the H1085A/E1103G double-mutant
enzyme exhibited a significantly slower elongation rate than
either the WTor E1103G single-mutant enzymes, and therefore
returned a reduced value for kcat(Table 1). However, in several
other key respects, the WTcharacteristics were restored. Notably,
the load dependencies of the F-v curves for the H1085A/E1103G
and WTenzymes were similarly flat (Fig. 3C). Moreover, the nu-
cleotide binding affinity (KD) of the double-mutant enzyme was
similar to that of WT, and its translocation equilibrium (Kδ) was
statistically indistinguishable (Table 1). Taken together, these
results suggest that the TL serves to affect not only the rate of
catalysis, as previously found, but also the NTP-binding affinity
and the translocation equilibrium. An inhibitory effect of TL clo-
sure on translocation was recently proposed, based on structural
modeling and computational studies (30), and is consistent with
the proposed “two-ratchet model” for RNAP translocation (27).
In that proposal, the closed state of the TL, which is dependent
mM NTPs, 100 mM NH4
1 mM NTPs
mM NTPs, 100 mM NH4
mM NTPs, 100 mM NH4
1 mM NTPs
mM NTPs, 100 mM NH4
1 mM NTPs
20 151050 -5
Force (pN) Force (pN)
(C) H1085A/E1103G RNAPII at saturating (1 mM) and subsaturating (0.1 mM) NTPs, as well as in the presence of 100 mM ammonium chloride, which was
added to our standard elongation buffer. Global fits to the two-binding site Brownian ratchet model (Model 3, Fig. 2C) are shown (dashed lines).
Force-velocity curves. Average pause-free elongation rates (errors as SEM) measured as functions of force for (A) WT, (B) E1103G, and
Table 1. Trigger loop affects catalysis, NTP binding, and
translocation equilibrium. Global fit parameters (errors as SD)
obtained from modeling of F-v curves in Fig. 3 to the two-binding
site Brownian ratchet model of Fig. 2C.
34 ? 2s−1
77 ? 3s−1
140 ? 16 μM
0.2 ? 0.1
0.34 nm (fixed)
56 ? 2s−1
77 ? 3s−1
49 ? 6 μM
0.8 ? 0.1
0.34 nm (fixed)
6 ? 1s−1
22 ? 2s−1
94 ? 17 μM
0.3 ? 0.1
0.34 nm (fixed)
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www.pnas.org/cgi/doi/10.1073/pnas.1200939109 Larson et al.
upon a match between the incoming NTP and the template
DNA, precludes translocation. However, as the correct NTP
is condensed into the nascent RNA, forward translocation is fa-
cilitated by the opening of the TL, induced by thermal motion,
likely along with a corresponding alteration in the conformation
of the bridge helix (31). The single-molecule data presented
here provide experimental evidence consistent with such TL-
gated translocation. This gating is analogous to models of trans-
location in DNA polymerases, where the reversals of domain
movements coupled to substrate selection and catalysis have been
implicated in the subsequent translocation step, but by and large
have been experimentally inaccessible (32).
Pausing Kinetics Imply a Role for the Trigger Loop in Transcriptional
Fidelity. Because pausing has been implicated in RNAP proof-
reading (33, 34), we examined the characteristics of transcrip-
tional fidelity and pausing behavior in WT RNAPII and its TL
mutants. Misincorporation of certain bases (such as the GTP
analog, ITP) by RNAP results in backtracking along the DNA
template, leading to a relatively long-lived, paused state. The
backtracking motion displaces the 3′-end of the nascent RNA
from the enzyme active site, permitting transcription factors
(such as TFIIS in eukaryotes or Gre factors in bacteria) to acti-
vate cleavage of the most recently incorporated bases, thereby
removing the error and restoring a new 3′ end of RNA in the
active site, allowing transcription to resume (33–35).
The RNAPII pause lifetime distribution could be subdivided
into two categories, the majority of which (∼95%) were “short,”
with an average time of ∼1.1 s, and the remainder of which (∼5%)
were “long” (>15s) (Fig. S3). WT RNAPII entered these long
pauses approximately once every 10 kb at subsaturating NTP
concentrations, a rate that corresponds closely to the frequency
for the incorporation of an incorrect nucleotide substrate, as
measured by ensemble assays in vitro (36). It has been shown
previously that E1103G promotes NTP misincorporation (6, 7).
Consistent with that finding, we observed a slight, but not statis-
tically significant, increase in the long pause density (LPD, num-
ber of long pauses/kb) for the E1103G mutant enzyme over the
WT enzyme at 0.1 mM NTPs (Fig. 4A).
To ascertain whether the increase in long pausing observed for
the RNAPII mutant was caused by an increased rate of mis-
matches arising from a deficiency in substrate selection, we added
1 mM additional nucleotide for each NTP, in turn to subsaturat-
ing NTPs (0.1 mM NTPs), which serves to promote mismatching
of the excess nucleotide by mass action. The LPD for E1103G
increased nearly fivefold upon the addition of excess ATP and
about threefold in the presence of excess CTP or UTP (Fig. 4A).
Only a modest increase in LPD was observed in the presence
of excess GTP. By contrast, the LPD of wild-type RNAPII was
comparatively insensitive to excess NTPs (Fig. 4A) and reached
a minimum in the presence ofadditional GTP.These resultsimply
that the E1103G mutant enzyme differentially undertakes long
pauses dependent on the specific excess NTP. In particular, the
inferred misincorporation events involving GTPare less frequent,
or less able to trigger a long pause, or both. This finding is con-
sistent with a mismatch-specific proofreading mechanism pro-
posed previously (33, 36, 37), and may be attributable to specific
interactions between the TL and the misincorporated 3′ end of
the RNA, which could trigger long pauses.
Inosine triphosphate (ITP), a GTP analog that forms a weak
Watson-Crick pair with cytosine and has a similar binding con-
stant and rate of incorporation into RNA, also inhibits next-
nucleotide-addition rates in a manner similar to that of certain
mismatched base pairs. This property has been exploited pre-
viously toinvestigate postincorporation nucleotide discrimination
and removal by RNAPII (38). At saturating concentration
of NTPs (1 mM) in the absence of ITP, E1103G showed a clear
increase in LPD over WT (Fig. 4B). For the WT RNAPII, the
addition of 200 μM ITP to transcription reactions containing
1 mM NTPs significantly increased the LPD (Fig. 4B), as pre-
viously described for E. coli RNAP (34), which is indicative
of misincorporation-induced pause enhancement. Perhaps sur-
prisingly, the LPD for E1103G decreased in the presence of ITP,
despite the fact that the mutant enzyme is able to incorporate
ITP into transcripts to the same extent as the WT (Fig. S4).
Taken together, these results suggest that the TL promotes tran-
scriptional fidelity in two distinct ways. First, the TL helps to
ensure correct NTP selection from among a pool of mixed sub-
strates. Mutation of the TL (E1103G) generally leads to more
mismatches being added to the transcript. Second, the TL func-
tions in the efficient recognition of miscorporation events by
RNAPII once a selection error has been made, as evidenced by
backtracking (38). The E1103G mutant enzyme also appears to
be deficient in this regard, because the misincorporation of
ITP fails to be recognized, and therefore fails to produce a cor-
responding increase in the proportion of long pauses.
The observation of elongation by individual RNAPII molecules
has provided additional insights into eukaryotic transcription me-
chanisms, and it offers the ability to separate effects on catalysis
from those on translocation, which is critical for the dissection
of the RNAPII mechanism. Based on high-resolution measure-
ments of the load dependence of the elongation rate, we find
support for a biochemical pathway where incoming NTPs can
associate with RNAPII in either its pre- or posttranslocated
state. Both single-molecule and bulk biochemical data, compar-
ing wild-type behavior with substitutions in the trigger loop, sug-
gest that this key subdomain of RNAPII functions not only to
regulate the basic elongation rate, via changes to the transloca-
tion bias and catalytic rate, but also plays a critical role in both
NTP substrate selection and mismatch recognition. In particular,
substrate selection defects were characterized under conditions
of competing levels of matched and mismatched NTPs. The ex-
periments also revealed significant differences in the recognition
of ITP misincorporation by the TL mutant enzyme, implicating
the TL in the promotion of pausing associated with proofreading
repair. We conclude that the sequence of the trigger loop evolved
to strike a critical balance between the opposing constraints of
catalytic prowess and transcriptional fidelity.
Materials and Methods
A detailed description of materials and methods is given in SI Text.
Long pause density (per 10 kb)
Long pause density (per 10 kb)
0.1 mM NTPs
1 mM NTPsATP GTP UTPCTP
1 mM NTPs
0.2 mM ITP
density (long pauses per 10 kb transcribed; errors as SEM) for the WT (blue)
and E1103G mutant enzymes (red) at subsaturating (0.1 mM) NTPS under the
conditions indicated. The long pause density increases for the mutant en-
zyme when a single NTP is present in excess, indicative of a deficiency in sub-
strate selectivity. (B) The inclusion of 200 μM ITP in a saturating NTP buffer
(1 mM NTPs) increases the long pause density for the WT (blue), but decreases
the density for the E1103G mutant enzyme (red), indicative of a failure in
mismatch recognition for ITP incorporation events.
Substrate selection and mismatch recognition. (A) Long pause
Larson et al.PNAS Early Edition
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Transcription Initiation. Biotinylated, 12-subunit Saccharomyces cerevisiae Download full-text
RNAPII (prepared as described in SI Text) was assembled stepwise onto a
DNA:RNA scaffold using a modification of a previous method (39), allowing
us to circumvent the need for initiation factors normally required to assemble
a transcriptional elongation complex. The 2.7 kb upstream and 4.8 kb down-
stream DNA “handles” were ligated to the ends of the template DNA by in-
cubating the RNAPII scaffolded complexes (5 nM) with these DNA fragments
(20 nM each) in elongation buffer (25 mM Hepes-KOH, pH 8.0, 130 mM KCl,
5 mM MgCl2, 1 mM DTT, 0.15 mM EDTA, 5% glycerol and 25 μg acetylated
bovine serum albumin/mL) in the presence of ligase (2 K units) and ATP
(1 mM) at 12 °C for 1 h.
DNA-Pulling Optical-Trapping Assay. Single elongation complexes were teth-
ered between 2 polystyrene beads (0.6 μm and 0.73 μm diameter), each held
in a separate optical trap, forming a “dumbbell.” By labeling either the up-
stream or downstream DNA with digoxigenin, we could apply a controlled
load that either assisted or hindered forward translocation of RNAPII (the
assisting-load version of the assay is depicted in Fig. 1A). Data were acquired
using a dual-beam optical-trapping microscope described previously (14),
and an oxygen-scavenger system was used in buffers to reduce photodamage
Data Collection and Analysis. Records of position data were acquired at 2 kHz
using custom software (written in Labview), filtered at 1 kHz by an 8-pole
lowpass Bessel filter, and analyzed using custom software (written in Igor
Pro). The distance between optically trapped beads was converted into the
DNA contour length (14) and pauses were identified and scored as described
previously (34). Pause-free elongation velocities of single RNAPII molecules
were determined by fitting the velocity distribution to a sum of two Gaus-
sians (one corresponding to pausing, and the other to active elongation) as
previously described (1).
Note Added in Proof. Additional information on materials and methods may
be found in ref. 42.
ACKNOWLEDGMENTS. We thank D. Koslover and V. Schweikhard for help with
data collection, D. Larson for critical reading of the manuscript, and J. Gelles
and members of the Block lab for helpful discussions. This work was sup-
ported by a National Science Foundation Graduate Research Fellowship
(to M.H.L.) andgrantsfrom theNational Institute of General Medical Sciences
(to S.M.B., R.L., and R.D.K.).
1. Neuman KC, Abbondanzieri EA, Landick R, Gelles J, Block SM (2003) Ubiquitous tran-
scriptional pausing is independent of RNA polymerase backtracking. Cell 115:437–447.
2. Darzacq X, et al. (2007) In vivo dynamics of RNA polymerase II transcription. Nat Struct
Mol Biol 14:796–806.
3. Ninio J (1991) Connections between translation, transcription and replication error-
rates. Biochimie 73:1517–1523.
4. Wang D, Bushnell DA, Westover KD, Kaplan CD, Kornberg RD (2006) Structural basis
of transcription: Role of the trigger loop in substrate specificity and catalysis. Cell
5. Vassylyev DG, et al. (2007) Structural basis for substrate loading in bacterial RNA poly-
merase. Nature 448:163–168.
6. Kireeva ML, et al. (2008) Transient reversal of RNA polymerase II active site closing
controls fidelity of transcription elongation. Mol Cell 30:557–566.
7. Kaplan CD, Larsson KM, Kornberg RD (2008) The RNA polymerase II trigger loop func-
tions in substrate selection and is directly targeted by alpha-amanitin. Mol Cell
8. Jokerst RS, Weeks JR, Zehring WA, Greenleaf AL (1989) Analysis of the gene encoding
the largest subunit of RNA polymerase II in Drosophila. Mol Gen Genet 215:266–275.
9. Yuzenkova Y, et al. (2010) Stepwise mechanism for transcription fidelity. BMC
10. Zhang J, Palangat M, Landick R (2010) Role of the RNA polymerase trigger loop in
catalysis and pausing. Nat Struct Mol Biol 17:99–104.
11. Kaplan CD, Jin H, Zhang I, Belyanin A (2012) Dissection of pol II trigger loop function
and pol II-activity dependent control of start site selection in vivo. PLoS Genetics,
12. Malagon F, et al. (2006) Mutations in the Saccharomyces cerevisiae RPB1 gene confer-
ring hypersensitivity to 6-azauracil. Genetics 172:2201–2209.
13. Hopfield JJ (1974) Kinetic proofreading: A new mechanism for reducing errors in bio-
synthetic processes requiring high specificity. Proc Natl Acad Sci USA 71:4135–4139.
14. Abbondanzieri EA, Greenleaf WJ, Shaevitz JW, Landick R, Block SM (2005) Direct
observation of base-pair stepping by RNA polymerase. Nature 438:460–465.
15. Gong XQ, Zhang C, Feig M, Burton ZF (2005) Dynamic error correction and regulation
of downstream bubble opening by human RNA polymerase II. Mol Cell 18:461–470.
16. Nedialkov YA, et al. (2003) NTP-driven translocation by human RNA polymerase II.
J Biol Chem 278:18303–18312.
17. Zhang C, Burton ZF (2004) Transcription factors IIF and IIS and nucleoside triphosphate
substrates as dynamic probes of the human RNA polymerase II mechanism. J Mol Biol
18. Westover KD, Bushnell DA, Kornberg RD (2004) Structural basis of transcription:
Nucleotide selection by rotation in the RNA polymerase II active center. Cell
19. Galburt EA, et al. (2007) Backtracking determines the force sensitivity of RNAP II in a
factor-dependent manner. Nature 446:820–823.
20. Izban MG, Luse DS (1991) Transcription on nucleosomal templates by RNA polymerase
II in vitro: Inhibition of elongation with enhancement of sequence-specific pausing.
Genes Dev 5:683–696.
21. Sluder AE, Price DH, Greenleaf AL (1988) Elongation by Drosophila RNA polymerase II.
Transcription of 3′-extended DNA templates. J Biol Chem 263:9917–9925.
22. Gu W, Reines D (1995) Identification of a decay in transcription potential that results
in elongation factor dependence of RNA polymerase II. J Biol Chem 270:11238–11244.
23. Bai L, Fulbright RM, Wang MD (2007) Mechanochemical kinetics of transcription
elongation. Phys Rev Lett 98:068103.
24. Bai L, Shundrovsky A, Wang MD (2004) Sequence-dependent kinetic model for tran-
scription elongation by RNA polymerase. J Mol Biol 344:335–349.
25. Guajardo R, Sousa R (1997) A model for the mechanism of polymerase translocation.
J Mol Biol 265:8–19.
26. Johnson RS, Strausbauch M, Cooper R, Register JK (2008) Rapid kinetic analysis of tran-
scription elongation by Escherichia coli RNA polymerase. J Mol Biol 381:1106–1113.
27. Bar-Nahum G, et al. (2005) A ratchet mechanism of transcription elongation and its
control. Cell 120:183–193.
28. Erie DA, Kennedy SR (2009) Forks, pincers, and triggers: The tools for nucleotide
incorporation and translocation in multi-subunit RNA polymerases. Curr Opin Struct
29. Temiakov D, et al. (2005) Structural basis of transcription inhibition by antibiotic
streptolydigin. Mol Cell 19:655–666.
30. Feig M, Burton ZF (2010) RNA polymerase II flexibility during translocation from
normal mode analysis. Proteins 78:434–446.
31. Gnatt AL, Cramer P, Fu J, Bushnell DA, Kornberg RD (2001) Structural basis of tran-
scription: An RNA polymerase II elongation complex at 3.3 A resolution. Science
32. GolosovAA, WarrenJJ, Beese LS,Karplus M (2010) Themechanismof the translocation
step in DNA replication by DNA polymerase I: A computer simulation analysis. Struc-
33. Erie DA, Hajiseyedjavadi O, Young MC, von Hippel PH (1993) Multiple RNA polymerase
conformations and GreA: Control of the fidelity of transcription. Science 262:867–873.
34. Shaevitz JW, Abbondanzieri EA, Landick R, Block SM (2003) Backtracking by
single RNA polymerase molecules observed at near-base-pair resolution. Nature
35. Izban MG, Luse DS (1992) The RNA polymerase II ternary complex cleaves the nascent
transcript in a 3′–5′ direction in the presence of elongation factor SII. Genes Dev
36. Sydow JF, et al. (2009) Structural basis of transcription: Mismatch-specific fidelity
mechanisms and paused RNA polymerase II with frayed RNA. Mol Cell 34:710–721.
37. Wang D, et al. (2009) Structural basis of transcription: Backtracked RNA polymerase II
at 3.4 angstrom resolution. Science 324:1203–1206.
38. Thomas MJ, Platas AA, Hawley DK (1998) Transcriptional fidelity and proofreading by
RNA polymerase II. Cell 93:627–637.
39. Kyzer S, Ha KS, Landick R, Palangat M (2007) Direct versus limited-step reconstitution
reveals key features of an RNA hairpin-stabilized paused transcription complex. J Biol
40. Larson MH, Greenleaf WJ, Landick R, Block SM (2008) Applied force reveals mechan-
istic and energetic details of transcription termination. Cell 132:971–982.
41. Kettenberger H, Armache KJ, Cramer P (2004) Complete RNA polymerase II elongation
complex structure and its interactions with NTP and TFIIS. Mol Cell 16:955–965.
42. Palangat M, et al. (2012) Efficient reconstitution of transcription elongation com-
plexes for single-molecule studies of eukaryotic RNA polymerase II. Transcription,
6 of 6
www.pnas.org/cgi/doi/10.1073/pnas.1200939109 Larson et al.