Article

Co-temporal Force and Fluorescence
Measurements Reveal a Ribosomal Gear ShiftMechanism of Translation Regulation by
Structured mRNAs
Graphical Abstract

Authors

Cy3•EF-G
binding
hairpinRibosome
Extension

Extension

Cotemporal force & fluorescence measurements
1codonhairpinopening
EF-G arrivaland departure
Time

Ribosomes shift into a slow gear to unwind
strong mRNA hairpins
Weak hairpin
Fast path (90%)

Strong hairpin
Fast path (50%)

Slow path (10%)

Slow path (50%)

Highlightsd

The mRNA hairpin is opened after EF-G binding and before
30S head reverse rotationdEF-G-catalyzed translocation is not affected by mRNA
secondary structure stabilitydRibosomes operate in two alternative (fast and slow) gears
during translationdIncreased hairpin stability increases the flux through theslow gear

Desai et al., 2019, Molecular Cell 75, 1007–1019
September 5, 2019 ª 2019 Elsevier Inc.
https://doi.org/10.1016/j.molcel.2019.07.024

Varsha P. Desai, Filipp Frank,
Antony Lee, ..., Harry F. Noller,
Ignacio Tinoco, Jr., Carlos Bustamante

Correspondence
carlosb@berkeley.edu

In Brief
Desai et al. used optical tweezers withsingle-molecule fluorescence detection
to show that mRNA hairpin opening andtranslocation by the ribosome occur
concurrently. Moreover, they unravel anallosteric gear shift mechanism oftranslation regulation in which ribosomes
switch into an alternative, slower kinetic
pathway in response to strong mRNA
hairpins.

Molecular Cell

ArticleCo-temporal Force and Fluorescence Measurements
Reveal a Ribosomal Gear Shift Mechanismof Translation Regulation by Structured mRNAs
Varsha P. Desai,1,2 ,9 Filipp Frank,2 ,9 Antony Lee,2 ,3 Maurizio Righini,1,2 ,4 Laura Lancaster,5 Harry F. Noller,5Ignacio Tinoco, Jr.,1 and Carlos Bustamante1,2 ,3 ,4 ,6 ,7,8 ,10,*
1Department of Chemistry, University of California, Berkeley, Berkeley, CA 94720, USA
2Jason L. Choy Laboratory of Single-Molecule Biophysics, University of California, Berkeley, Berkeley, CA 94720, USA
3Department of Physics, University of California, Berkeley, Berkeley, CA 94720, USA
4California Institute for Quantitative Biosciences, QB3, University of California, Berkeley, Berkeley, CA 94720, USA
5Department of Molecular, Cell, and Developmental Biology and Center for Molecular Biology of RNA, University of California, Santa Cruz,
Santa Cruz, CA 95064, USA
6Howard Hughes Medical Institute, University of California, Berkeley, Berkeley, CA 94720, USA
7Department of Molecular and Cell Biology, University of California, Berkeley, Berkeley, CA 94720, USA
8Kavli Energy Nanoscience Institute, University of California, Berkeley, Berkeley, CA 94720, USA
9These authors contributed equally
10Lead Contact*Correspondence: carlosb@berkeley.edu
https://doi.org/10.1016/j.molcel.2019.07.024

SUMMARY

The movement of ribosomes on mRNA is often interrupted by secondary structures that present mechanical barriers and play a central role in translation
regulation. We investigate how ribosomes couple
their internal conformational changes with the activityof translocation factor EF-G to unwind mRNA
secondary structures using high-resolution opticaltweezers with single-molecule fluorescence capability. We find that hairpin opening occurs during
EF-G-catalyzed translocation and is driven by the forward rotation of the small subunit head. Modulating
the magnitude of the hairpin barrier by force showsthat ribosomes respond to strong barriers by shifting
their operation to an alternative 7-fold-slower kinetic
pathway prior to translocation. Shifting into a slow
gear results from an allosteric switch in the ribosomethat may allow it to exploit thermal fluctuations to
overcome mechanical barriers. Finally, we observethat ribosomes occasionally open the hairpin in two
successive sub-codon steps, revealing a previously
unobserved translocation intermediate.

INTRODUCTION
Ribosomes are macromolecular machines responsible for thesynthesis of proteins in all cells. During translation initiation, ribosomes assemble on an mRNA and subsequently enter the elongation phase (Schmeing and Ramakrishnan, 2009). Here, they
move along the mRNA in steps of one codon (3 nt) to decode
its message and correspondingly add an amino acid to thegrowing polypeptide chain (Schmeing and Ramakrishnan,2009). This mechanical movement of the ribosome on themRNA is called translocation and is catalyzed by the guanosine
triphosphatase (GTPase) activity of a trans-acting elongation
factor, EF-G (Rodnina et al., 1997). Translocation is also accompanied by large-scale conformational changes within the ribosome. After peptide bond formation, the small (30S) and the large(50S) ribosomal subunits rotate with respect to each other spontaneously, resulting in the translocation of the acceptor ends ofthe tRNAs in the 50S subunit (Cornish et al., 2008; Frank andAgrawal, 2000; Moazed and Noller, 1989b; Sharma et al.,2016; Zhang et al., 2009). During this process, however, thecodon of the mRNA and the anticodon of the tRNAs that residein a cleft between the head and the body domains of the 30S
subunit have not yet moved. Several lines of evidence indicatethat subsequent forward and/or reverse rotation of the 30S
head completes this movement, resulting in the net translocationof the mRNA relative to the ribosome by one codon (Guo andNoller, 2012; Ratje et al., 2010; Zhou et al., 2014).
During translocation, ribosomes often encounter mechanicalbarriers due to structures such as hairpins and pseudoknots
adopted locally by the mRNA (Ding et al., 2014; Katz and Burge,
2003; Kertesz et al., 2010; Li et al., 2012; Mustoe et al., 2018).
Crystal structures have revealed that the entry tunnel of the ribosome can only accommodate single-stranded RNA (Yusupova
et al., 2001). Biochemical studies have shown that while ribosomes can intrinsically unwind mRNA secondary structures during translocation (Takyar et al., 2005), the rate of translation is
reduced in front of such barriers (Qu et al., 2011; Wen et al.,2008); as such, these barriers are thought to play a key regulatoryrole in cotranslational protein folding (Chaney and Morris, 1978;
Guisez et al., 1993; Watts et al., 2009), mRNA localization (Chartrand et al., 2002; Young and Andrews, 1996), and protein abundance (Duan et al., 2003; Nackley et al., 2006), and, in combination with additional cis elements, they can induce ribosomal

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Molecular Cell 75, 1007–1019, September 5, 2019frameshifting, leading to the synthesis of alternative protein
products (Jacks et al., 1988; Tsuchihashi, 1991). Despite astrong effect of mRNA secondary structures on translation, anumber of questions remain as to how ribosomes couple theirhelicase activity with their translocation. Specifically, when aremRNA secondary structures opened during a given translation
cycle? How is the activity of EF-G coupled to strand opening
and translocation? Finally, how are the internal conformationalchanges of the ribosome involved in barrier crossing?
Positively charged amino acid residues located at the mRNA
entry tunnel are known to play a crucial role in the destabilizationof secondary structures prior to translocation (Qu et al., 2011).
Mutating these residues does not affect translocation along a single-stranded mRNA but prevents progress on a double-stranded
RNA template, indicating that the unwinding of secondary structures at the entry tunnel may occur prior to translocation (Takyar
et al., 2005). Alternatively, unwinding could occur concomitantlywith EF-G binding, whereby the free energy gained upon bindingis used to destabilize the hairpin. Finally, unwinding could occur
after EF-G binding and concomitantly with mRNA translocationby the forces generated during the forward or the reverse rotationof the 30S head domain (Liu et al., 2014).
Moreover, the mechanism by which the translation rate ismodified by these barriers has been debated. Previous singlemolecule optical tweezers studies postulate that mRNA hairpins
should directly and selectively reduce the rate of EF-G-catalyzed
translocation (Qu et al., 2011), whereas single-molecule fluorescence studies have proposed that mRNA secondary structures
allosterically modify the ribosome to delay E-site tRNA release,
thereby affecting parts of the translation cycle not related totranslocation (Chen et al., 2013a). A limitation of these measurements is that they only monitor either the mechanical or the fluorescence coordinate and therefore cannot directly correlate
motions within the ribosome and/or the activity of trans-acting
factors such as EF-G to hairpin opening. Such questions arealso broadly applicable to many molecular motors that couple
the activity of protein or chemical factors to their movement
along a substrate (Bustamante et al., 2011).
Therefore, in this study, we use a recently developed high-resolution optical tweezers with single-molecule fluorescencecapability (‘‘fleezers’’) (Comstock et al., 2011) to directly measure
hairpin opening while simultaneously visualizing the binding andrelease of EF-G. We find that a downstream hairpin, irrespectiveof its stability, is always only opened after EF-G binding, unequivocally establishing that translocation and hairpin opening
are temporally coupled. Moreover, using antibiotic perturbation
experiments, we show that the unwinding of secondary structures results from the force generated upon forward 30S headrotation. Then, we increase the stability of the downstream barrier by decreasing the force applied to the hairpin and discoverthat strong hairpins only marginally reduce the rate of translocation; instead, they act as allosteric switches that globally slow
down the translation cycle, biasing ribosomes into a kineticallyaltered 7-fold-slower pathway. Finally, the use of two orthogonal—mechanical and fluorescence—channels to monitor
translation enabled us to unravel a novel unwinding intermediatein which each hairpin opening event occurs via two successive
sub-codon steps while EF-G remains bound. These findings uncover fundamental insights on the mechanism of mRNA unwinding and translocation by the ribosome.
RESULTS
Ribosomes Open mRNA Hairpins during Translocation
To test whether unwinding occurs prior to, concomitant with, orafter EF-G binding, we perform an optical tweezers assay wherein
we simultaneously measure mRNA hairpin opening in parallel with
EF-G arrival and release, using high-resolution optical tweezers
with single-molecule fluorescence capability (Comstock et al.,2011; Wen et al., 2008) (Figures 1A and 1B). This assay uses anmRNA hairpin with repeating valine codons that is tethered byits 50 and 30 ends to two polystyrene beads, each held in an opticaltrap (Figures 1A and S1). A ribosome is stalled at the base of thehairpin by the omission of Val-tRNAVal. After capturing the stalledribosome complex on the beads, translation is restarted by supplying a mixture containing tRNAVal, valyl-tRNA synthetase, valine,
Cy3-labeled EF-G, EF-Tu, ATP, and GTP.
This scheme enables us to measure translocation in real timethrough two channels: (1) the optical tweezers channel, to follow
the stepwise opening of the hairpin (Figure 1C), and (2) the fluorescence channel, to monitor the binding of fluorescently labeled
EF-G (Figure 1D). In this experiment, each extension step corresponds to the unwinding of exactly one codon (Figure 1C). The
waiting time between two consecutive codon steps is called a

Figure 1. Ribosome Opens an mRNA Hairpin after EF-G Binding(A) Experimental setup for simultaneous measurement of hairpin opening and EF-G binding using a time-shared optical trap with single-molecule confocal
imaging, fleezers. The time resolution of the optical tweezers channel is 7.5 ms and that of the fluorescence channel is 10 ms. The assembly of ribosome-stalled
complexes is outlined in Figure S1.
(B) Possible scenarios of when hairpin unwinding occurs with respect to EF-G binding. Case (i): the hairpin is spontaneously opened prior to EF-G binding due tothe destabilization energy contributed by the ribosome (Qu et al., 2011). Case (ii): the hairpin is opened concomitant with EF-G binding. Here, either EF-G binding
itself induces a conformational change (e.g., the forward or reverse movement of the 30S head) that results in the opening of the hairpin, or the 30S head rotatesback and forth as a Brownian ratchet, and only the binding of EF-G effectively rectifies the position of the 30S head, leading to hairpin opening. Case (iii): thehairpin is opened a variable amount of time after EF-G binding. Opening could result from events such as EF-G , GTP tight binding, GTP hydrolysis, Pi release,
EF-G , GDP release, or internal conformational changes of the ribosome.
(C and D) An example of a single-ribosome fleezers trajectory along an mRNA hairpin held under a high external force of 13–16 pN with 10 nM Cy3-labeled-EF-G.
(C) Optical tweezers channel: ribosome opens the hairpin in one-codon steps ( = 6 nt hairpin opened), separated by dwells, t dwell . The data were recorded at
133 Hz and displayed at 13 Hz. The yellow box shows a magnified event. (D) Fluorescence channel: each spike in fluorescence corresponds to the binding of an
EF-G. The yellow box shows a magnified event. The data were recorded at 100 Hz and displayed at 10 Hz. Additional magnified events are shown in Figure S2.
(E) Summary of average t dwell (gray) and average EF-G residence times before unwinding (t unwinding , green) and after unwinding (trelease , green) for a weak hairpin
held under an external force of 13–16 pN (n = 55 events, 9 molecules). The error bars represent SEMs.

Molecular Cell 75, 1007–1019, September 5, 2019 1009

‘‘dwell,’’ during which the mRNA is stationary on the ribosome
and a number of chemical events occur, including the additionof one amino acid to the nascent polypeptide chain. Likewise,
each spike in the fluorescence channel corresponds to the binding and dissociation of a single EF-G molecule (Figure 1D).
Strikingly, at each codon, we observe a strict order of events:
hairpin unwinding occurs always after EF-G arrival; similarly, therelease of EF-G occurs always after the unwinding event (yellow
box in Figures 1C, 1D, and S2). The total time that EF-G is boundto the ribosome is the sum of the time between EF-G binding andthe unwinding of the hairpin, tunwinding , and the time between unwinding and EF-G release, trelease (Figure 1E). On average,t unwinding is 0.25 ± 0.07 s and trelease is 0.39 ± 0.1 s, when thehairpin is held at high force (>13 pN) (Figures 1E and S2; distributions of tunwinding and t release are shown in Figures 4D and S6A,
respectively). Therefore, unwinding occurs only after EF-G binding and not before by the independent helicase action at themRNA entry site; neither does it occur simultaneously with EFG arrival via the free energy gained upon binding. As the mRNA
entry tunnel can only accommodate single-stranded RNA (Yusupova et al., 2001), hairpin opening must then occur after EF-G
binding and concomitant with mRNA translocation.
As mentioned above, the translocation cycle begins after peptide bond formation, when the 30S and 50S subunits rotate withrespect to each other, resulting in the movement of the acceptor
ends of the tRNAs in the 50S subunit by one codon. Then, after
EF-G binding, two coordinated conformational changes occur
within the ribosome that result in the movement of the tRNAs
and the mRNA in the 30S subunit (Guo and Noller, 2012; Mohan
et al., 2014; Ratje et al., 2010; Zhou et al., 2014). First, the 30S
head rotates forward, moving the tRNAs and the mRNA relative
to the 30S body by one codon. Second, the 30S head detaches
from the tRNAs and the mRNA and moves back into the nonrotated position. As a result, the entire ribosome moves itself
relative to the mRNA by one codon and completes a translocation cycle.
To distinguish whether forward or reverse 30S head rotation
results in hairpin opening, we performed experiments with antibiotic fusidic acid (Figure 2). Previous studies have shown thatwhile fusidic acid binds to an early translocation intermediate,its kinetic effect on forward 30S head rotation and subsequenttRNA-mRNA movement is small. Conversely, fusidic acid greatly
reduces the rate of late translocation events such as reverse 30S
head rotation, E-site tRNA release, and EF-G dissociation (Belardinelli and Rodnina, 2017; Borg et al., 2015; Ramrath et al., 2013;
Wasserman et al., 2016). Therefore, by measuring whether
t unwinding or t release increases in the presence of fusidic acid, we
can determine the relative timing of 30S head rotation and hairpin
opening (Figure 2A). We find that tunwinding is unchanged (0.21 ±0.1 s), but t release increases dramatically (!10-fold) to 3.81 ±0.84 s in the presence of fusidic acid (Figures 2B–2D). These results establish that hairpin opening occurs before 30S head
reverse rotation and that the helicase and translocase activitiesof the ribosome are tightly coupled and occur simultaneously.
Strong mRNA Hairpins Reduce Translation Rate
Next, we asked how the ribosome responds to the presence of astronger mRNA hairpin. Does the rate of hairpin unwinding

1010 Molecular Cell 75, 1007–1019, September 5, 2019

decrease upon the increased strength of the hairpin, or are otherparts of the translation cycle modified by the downstream
hairpin? In our assay, we can modulate the strength of the barrier
by applying force to destabilize the junction (Bustamante et al.,
2004; Qu et al., 2011; Tinoco and Bustamante, 2002) (Figure 3A).
A key advantage of using force rather than GC content to alter
the hairpin stability is that the mRNA coding sequence and identity of the tRNAs remain invariant, so that steps in the translation
cycle unrelated to mechanical movement, such as decoding,remain unaffected.
We measure the average residence time of the ribosome ateach codon (tdwell ) as a function of force applied to destabilize
the hairpin junction (Figure 3B). At saturating concentrations of
EF-G (10 mM), we find that ribosomes spend !1.3 s/codon in
front of a weak barrier (high force >13 pN) and !2.5 s/codon in
front of a strong barrier (low force <7 pN; Figure 3B). Hence,
the rate of translation decreases 2-fold from !0.8 codon/s to!0.4 codon/s as the strength of the downstream barrier increases. The translation rate measured at high force in our single-molecule experiments is slightly slower than the 3–10
codon/s reported in in vitro translation assays performed inbulk, possibly due to temperature effects as our measurements
are made at room temperature as opposed to 37"C (Holtkamp
et al., 2015; Pavlov and Ehrenberg, 1996).
The Effect of a Strong Hairpin on Translation Rate Is Not
Restricted to the Unwinding Step
What change in the single-ribosome trajectories accounts forthis 2-fold decrease in translation rate observed with increased
strength of the barrier? We found earlier that the dwell duration
per codon (t dwell ) increases by on average !1 s at low force,when the junction is more stable (Figure 3B). Intuitively, we would
expect that because EF-G-catalyzed translocation is the forcesensitive step, its lengthening should account for the full dwell
time increase observed in front of a barrier. Therefore, since
the hairpin is opened only after EF-G binding, we expect thatthe residence time of EF-G before unwinding (t unwinding ) would
correspondingly increase by !1 s/codon at low force in ourassay, possibly due to a slower rate of forward head rotation(Figures 3C, 3D, and S4).
Surprisingly, the application of low force increases tunwinding byonly !0.35 s on average, while t release remains statistically invariant(Figure 3E). Hence, our results directly indicate that an additional
kinetic event either prior to EF-G binding or after EF-G release becomes rate limiting and accounts for the lengthening of the totaldwell time by the increased strength of the junction barrier. Thisis unlikely to be A-site tRNA binding or peptide bond formation,
which have been shown to be insensitive to downstream mRNA
secondary structures (Chen et al., 2013a, 2014; Kim et al., 2014).
We can also exclude photobleaching artifacts of the bound EF-G
since we determine the average bleaching times from fusidic
acid measurements to be at least 5 s, which are 5-fold longer
than the total residence time of EF-G on the ribosome (Figure 2D).
Translation Occurs via Two Parallel Pathways
Further analysis of the single ribosome trajectories showed thatfitting of the distribution of total dwell time (tdwell ) requires a minimum of two exponentials (Figures 4A and S5) according toAFigure 2. Hairpin Opening Occurs during
Forward 30S Head Rotation(A) Diagram of predicted EF-G residence times in thepresence of antibiotic fusidic acid for unwinding resulting from either forward or reverse 30S headrotation.
(B and C) An example of a single-ribosome fleezers
trajectory in the presence of 50–200 mM fusidic acidand 10 nM Cy3-EF-G. The hairpin is held under ahigh external force of 13–16 pN (B). (C) Opticaltweezers channel: data were recorded at 133 Hz anddisplayed at 13 Hz. (D) Fluorescence channel: data
were recorded at 100 Hz and displayed at 10 Hz.
Additional magnified events are shown in Figure S3.
(D) Summary of average EF-G residence times
before and after unwinding show that trelease increases dramatically in the presence of fusidic acid(n = 25 events, 13 molecules). The error bars
represent SEMs.

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PðtÞ = f1 ,k1 ,e%k1 , t + f2 ,k2 ,e%k2 , t
where PðtÞ is the probability of observing a dwell of duration t, k1
and k2 are the respective rates of the two exponential distributions, f1 is the fraction of events that a given ribosome translateswith rate k1 , and f2 is the fraction of events that it translates with
rate k2 . The need for two exponentials to describe the distribution of dwell times indicates that the kinetic mechanism of theribosome elongation cycle is not a simple linear scheme butthat it bifurcates into two alternative pathways with rates k1
and k2 .
Significantly, the stability of the hairpin junction determines thefraction of time that the ribosome chooses one or the other path.
At the high force of 15 pN, during which the mRNA hairpin is highly destabilized, we observe that !90% of the translation eventsoccur at the rate k1 (!1.1 codon/s), while the remaining eventsoccur through a slow pathway with the rate k2 (!0.2 codon/s)(Figure 4B, rates k1 and k2 ; and Figure 4C, fraction of eventswith rate k2 ). Notice that the rate k2 is !6-fold slower than k1 ;hence, we define k1 hkfast path andk2 hkslow path . As the force applied to thehairpin is reduced and its stability increases, the values of kfast path andkslow path remain relatively constant (Figure 4B); however, the fraction of translation
events that enter the slow pathway increases to 30%–50% as the force decreases below 7 pN (Figure 4C). These
slow events do not arise from a subset of
impaired ribosomes (static dispersion)
since the biphasic translation rate isobserved for any individual ribosome trajectory, indicating that the same ribosome
can switch between the two pathways(dynamic dispersion) (Figure S5). Thus,
the presence of a strong barrier biases theribosomes into the slower pathway withminimal alteration of the pathway rates
and accounts for the 2-fold decrease in the average translation
rate observed in these conditions.
Pathway Identity Is Maintained through EF-G-Catalyzed
Hairpin Opening
Next, we asked whether the translation kinetic pathway bifurcation occurs prior to EF-G binding through hairpin unwinding or after unwinding during EF-G release and resetting of the ribosome(see Data S1). The distribution of lifetimes of t unwinding (the residence time of EF-G before unwinding) is also best fit by a double
exponential (Figure 4D) with fast and slow rates that are also notsignificantly force sensitive (Figure 4E). At high and low force,
fast
slow
is !24/s and kunwindingis !2/s (Figure 4E). Significantly,
kunwinding
however, the fraction of unwinding events displaying slow kislow) increases from !15% at high force (>12 pN)
netics (kunwinding
to !40% at low force (<9 pN; Figure 4F).
Notice that both kdwell (Figure 4B) and kunwinding (Figure 4E) aredescribed by a mixture of two exponentials, but, more importantly, that the fraction of events going through the slow branch

Molecular Cell 75, 1007–1019, September 5, 2019 1011

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increase in approximately the same proportions for both distributions (kdwell and kunwinding ) as a function of force (Figures 4C
and 4F). These results indicate that the translation pathway bifurcates prior to EF-G binding and the ribosome remains in the fastand slow branches through EF-G binding and subsequent
hairpin unwinding. Consistently, analysis of t release reveals thatthe distribution of this parameter is insensitive to the strengthof the hairpin (Figure S6), indicating that the two pathways
have converged during the unwinding events.
It is important, however, to note that the rates of unwinding
fast
slow
!24/s and kunwinding!2/s; Figure 4E) are much faster(kunwinding
than the overall rates of the fast and slow pathways (kfast path
!0.9 codons/s; kslow path !0.15 codons/s; Figure 4B). This observation implies that there must exist an earlier kinetic transition(denoted below as kintermediate ) in both the fast and slow pathways
that is rate limiting for each of the respective branches.

Kinetic Scheme for Translation through mRNA Hairpins
From the above results, we conclude that the translation pathway
bifurcates prior to EF-G binding and converges after EF-G-catalyzed hairpin unwinding (Figure 4G; Data S1). Our data require
three sequential steps to describe the kinetics of translation
through mRNA secondary structures. First, there is a ‘‘hairpin
sensor’’ step that occurs rapidly and irreversibly, which gives riseto the bifurcation of the translation pathway into a ‘‘fast’’ and a
fast
slow
and ksensor, respectively. The ratio‘‘slow’’ branch with rates ksensor
of these rates is force sensitive and is equal to the fraction of events

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Molecular Cell 75, 1007–1019, September 5, 2019

Figure 3. Force Dependence of Translation
Rates(A) Schematic showing the effect of force on thestability of the hairpin junction. High applied force
results in a weaker hairpin junction.
(B) Average time per codon increases at low forces
as the hairpin presents a stronger barrier (n = 889
events from 24 molecules at 15 pN, n = 776 eventsfrom 19 molecules at 10 pN, n = 256 events from 8
molecules at 5 pN, and n = 551 events from 19
molecules at 3 pN). The concentration of EF-G is
10 mM. Error bars represent SEM.
(C and D) An example of a single-ribosome fleezers
trajectory with 10 nM Cy3-EF-G, and hairpin is held
under a low external force (5–7 pN). (C) Opticaltweezers channel: data were recorded at 133 Hz anddisplayed at 13 Hz. (D) Fluorescence channel: data
were recorded at 100 Hz and displayed at 10 Hz.
Additional magnified events are shown in Figure S4.
(E) Summary of average t dwell (gray) and average EFG residence times before unwinding (t unwinding , green)
and after unwinding (trelease , green) for a strong
hairpin held under a low external force of 5–7 pN(n = 62 events, 14 molecules). The error bars represent SEMs. When compared to data collected at highforce (Figure 1E), tdwell increases by !1 s at low force,but t unwinding only increases by !0.3 s and treleaseremains statistically invariant.

going througheach pathway—i.e.,
fast
slow
ksensor=ksensor= ffast path =fslow path . Second,an intermediate kinetic step occurs in both
fast
slow
and kintermediate). This step is insensitive topathways (kintermediate
force and determines the rates of translation through the two paths(Figure 4B). These two sequential ‘‘sensor’’ and ‘‘intermediate’’
steps are necessary to explain the force-dependent population
shift and the force-independent rates of the two pathways, respectively. In the third step, the ribosome opens the hairpin in the presfast
slow
and kunwinding, neither of which isence of EF-G via rates kunwinding
rate limiting in the respective pathways. The two pathways
converge after the unwinding event, EF-G is released, and the ribosome is reset for another round of elongation. Taken together,
these results indicate that the presence of a strong barrier at the
RNA junction allosterically switches the ribosome prior to EF-G
binding from a fast to a slow overall translation speed. We also
tested other possible kinetic schemes, in particular one in which
the ribosome could be stuck in an inactive conformation prior to
EF-G binding, and found that this scheme does not satisfy all of
the data obtained here (see Data S1 for details).
Detection of Translocation Intermediates
In the presence of a strong barrier, we occasionally observe that
unwinding, instead of taking place in a single transition, occurs in
two (smaller than one codon) steps, a process accompanied byunusually long EF-G binding durations (Figures 5A–5F). The firstsub-step covers on average 42% ± 17% of the total step sizeand occurs soon after EF-G binding. In the majority of events,
the dwell time in the intermediate state lasts up to a few seconds,
and EF-G release is observed only after the second step of unwinding has occurred (Figures 5A, 5B, and 5D–5F). However,

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Figure 4. Ribosome Translates through a Hairpin via Two Parallel Pathways that Bifurcate before EF-G Binding and Converge after Hairpin
Opening(A) Cumulative density of t dwell , the time ribosome spends at each codon, is best fit by a mixture of two exponentials given by CdfðtÞ = 1 % ffast path ,e%kfast path ,t % fslow path ,e%kslow path ,t .
(B) Summary of fast and slow pathway rates, kfast path and kslow path , obtained from fits to the cumulative distribution of t dwell at forces 3 pN (n = 551 events, 19
molecules), 5 pN (n = 256 events, 8 molecules), 10 pN (n = 776 events, 19 molecules), and 15 pN (n = 889 events, 24 molecules). The error bars represent 95%
confidence intervals. The concentration of EF-G is 10 mM. kfast path and kslow path remain constant at !0.92 codon/s and !0.15 codon/s, respectively, at various
forces.
(C) Fraction of events going through the slow pathway, fslow path , as obtained from fitting shown in (A). fslow path increases from !10% at high force (>10 pN) to!30%–50% at low force (<7 pN). The error bars represent 95% confidence intervals.
(D) Cumulative density of t unwinding , the time between EF-G binding and hairpin opening, is best fit by a mixture of two exponentials given by CdfðtÞ = 1 %
fast
slow
fast
slow
funwinding,e%kunwinding ,t % funwinding,e%kunwinding ,t .
fast
slow
(E) Summary of bi-exponential rates kunwinding
and kunwinding
obtained from fits to the cumulative distribution of t unwinding at forces 5–8 pN (n = 62 events, 14
molecules), 7–9 pN (n = 46 events, 10 molecules), 12–13 pN (n = 80 events, 20 molecules), and 14–17 pN (n = 53 events, 22 molecules). These experiments were
performed in passive mode—in other words, the force is maintained in a particular regime rather than being held constant. The concentration of EF-G is 10 nM.
The error bars represent 95% confidence intervals.
slow(F) Fraction of events going through the slow unwinding pathway, funwinding, increase from !15% at high force (>12 pN) to !40% at low force (<9 pN). The error
bars represent 95% confidence intervals.
(G) Proposed kinetic scheme: the ribosome ‘‘senses’’ the hairpin barrier and irreversibly switches into either a fast state (green) or a slow state (red) via rates
fast
slow
fast
slow
ksensoror ksensor, respectively. The ratio ksensor=ksensor
is force sensitive and determines the fraction of translation events that go through either pathway, as
fast
slow
shown in (C) and (F). Then, the ribosome in either the fast or the slow state must undergo an intermediate transition (kintermediateor kintermediate) that becomes rate
limiting at low force and determines the overall rates shown in (B). It is possible that this intermediate transition is the rate of EF-G binding (see Discussion).
(legend continued on next page)

Molecular Cell 75, 1007–1019, September 5, 2019 1013

in a few events (e.g., the one occurring at !0.5 s in Figure 5C) the
first half-step is reversible, and the full step occurs only at !3.5 s
via the intermediate. In this particular case, there is a potential
EF-G binding event at !3.5 s, when the full step occurs, but
due to the low intensity of the signal, we cannot conclusively
determine the presence of EF-G. It is important to note that theuse of labeled EF-G in these experiments and the lengtheningof its dwell time on the ribosome clearly indicate that the substeps are not merely a manifestation of hairpin dynamics.
In contrast, we sometimes observe large-scale excursions(!2 codons) of the ribosome along the mRNA (Figure 5G). Thesemovements closely resemble those previously described during
frame shifting along a slippery sequence (Yan et al., 2015). During these excursions, several EF-G molecules are seen to bindand unbind with prolonged dwell times on the ribosome (seeEF-G events marked by a star in Figure 5G). Thus, in the presence of a strong mRNA junction, the ribosome appears to switch
into a diffusive mode over the extent of two codons, during which
multiple EF-Gs are seen to bind and unbind.
Mechanism of Translocation through mRNA Secondary
Structures
The above results lead us to propose a mechanism in which theribosome can operate in either of two distinct kinetic modes or‘‘gears,’’ both of which are competent to bind EF-G and translocate. After peptide bond formation, two major conformationalchanges are known to occur in the ribosome before translocation: (1) inter-subunit rotation and (2) partial forward 30S headrotation (Figure 6, step 2). In this state, the 30S and 50S subunits
spontaneously rotate !6"–8" with respect to each other,concomitant with the formation of tRNA hybrid states (Frankand Agrawal, 2000; Moazed and Noller, 1989b; Zhang et al.,2009). Structural evidence shows that this state is accompanied
by an intermediate degree of rotation (5"–7") of the 30S head
domain with respect to the 30S body, along an axis orthogonal
to that of the inter-subunit rotation, before EF-G binding (Mohan
et al., 2014). In fact, deconvolution of förster resonance energy
transfer (FRET) measurement data from donor and acceptor positions at S13 and L33 or at S6 and at L9, respectively, reveal asimilar mobility in the 30S head domain before EF-G arrival(Belardinelli et al., 2016). This partial head rotation is distinct
from the full !22" forward head rotation that occurs after EF-G
binding that is accompanied by a partial reverse rotation of the
30S body (Guo and Noller, 2012; Ratje et al., 2010; Zhou et al.,2014). As shown here, the full forward head rotation results inthe opening of the hairpin junction at the entry port (Figure 6,
step 3).
Consequently, as the ribosome senses the hairpin prior to
EF-G binding, we propose that the intermediate head rotationthat occurs during inter-subunit rotation could operate as thesensor that switches the ribosome between the fast versus
slow gears and that leads to the subsequent rate-limiting stepof each pathway. Even though the detailed structural intermediates involved during the slow gear path remain unknown, since
the hairpin junction abuts the interface of the 30S head and the
30S body, we favor a model in which two distinct head rotation
modes result in the fast and slow gears of ribosome operation.
We propose that the 30S head ‘‘senses’’ the downstream junction during its initial rotation (Figure 6, step 2) and switches into
an altered conformation in response to a strong barrier. Several
structures of the ribosome trapped using antibiotics or non-hydrolyzable analogs of EF-G , GTP that block translocation,
show this partial degree of head rotation (!6"), hinting at the possibility that such conformational change gives rise to the low gear
(Brilot et al., 2013; Chen et al., 2013c; Mohan et al., 2014; Ratje
et al., 2010; Svidritskiy et al., 2014; Tourigny et al., 2013; Zhang
et al., 2016).
DISCUSSION
Many cellular processes are carried out by sophisticated molecular machines that move in a precise manner to produce forces,
torques, and displacements. These machines operate throughcycles that couple many chemical events such as internal
conformational changes, nucleotide binding and hydrolysis,
release of catalysis products, and binding and unbinding oftrans-acting factors to their mechanical task (Bustamante
et al., 2011). How these various chemical and mechanical steps
are coordinated in the operational cycle of these machines remains a central question in biophysics. Although the advent of
single-molecule methods has greatly improved the ability to
extract mechanistic details from the individual molecular trajectories, the challenge is compounded by the fact that, more often
than not, these measurements are one dimensional, tracking asingle event of the cycle.
Optical tweezers experiments, in which forces and displacements can be detected with millisecond resolution, have provided unprecedented details about the mechanical operationof such machines (Moffitt et al., 2008). In this way, the steppingof nucleic acid motors, such as polymerases, helicases, andtranslocases, has been resolved at the single base pair level(Chemla, 2010). Likewise, torque generation by topoisomerases,
DNA translocases, and flagellar motors has been measured
directly and has provided significant mechanistic insights into
their operation (Moffitt et al., 2006; Ryu et al., 2000). However,these mechanical measurements are blind to the concomitantchemical events of the operational cycles of the machines. Alternatively, the use of single-molecule fluorescence methods haveenabled researchers to follow, for example, in real time, some of
the conformational changes of these machines during their operation, but at the expense of disregarding their progress along themechanical coordinate (Joo et al., 2008).
The recent development of a new generation of optical tweezers instruments, endowed with single-molecule fluorescence

fast
slow
Then, the ribosome unwinds the hairpin via rates kunwinding
and kunwinding
shown in (E). Finally, EF-G is released in a similar fashion in either pathway, suggestingthat the bifurcated pathways converged upon unwinding (Figure S6). An illustration of the fleezers trajectory for both the fast and slow pathways isshown above and below the kinetic scheme. Notice that while t unwinding increases in the slow pathway, it does not increase enough to account for the total
increase in tdwell .

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A

B

C

capability (referred to here as fleezers [fluorescence opticaltweezers]), has opened the possibility of monitoring molecularmachine trajectories along two (and, in principle, several) orthogonal reaction coordinates in a co-temporal manner (Comstock
et al., 2011; Hohng et al., 2007; Lang et al., 2004; van Mameren
et al., 2008). Hence, these instruments make it possible to simultaneously record, correlate, and causally relate chemical events(e.g., binding of ligands, conformational changes) with the corresponding mechanical events (force or torque generation anddisplacement). Here, we have used a fleezers instrument(Comstock et al., 2011; Whitley et al., 2017) to investigate thecoupling of ribosome translocation to the binding and activityof elongation factor EF-G and to uncover how downstream secondary structures in the mRNA template regulate this process.

D

E

F

Mechanism of Helicase and Translocase Activities of the
Ribosome
During protein synthesis, ribosomes translocate along the mRNAin one-codon steps. This movement is often impeded by structural elements in the downstream mRNA such as hairpins andpseudoknots that are situated at the entry pore on the small
30S subunit. Previous optical tweezers studies have proposedthat ribosomes unwind mRNA secondary structures via twomodes of active helicase activity: one in which the positively
charged amino acid residues at the surface of the mRNA entry
tunnel destabilize helical regions prior to EF-G binding, and theother in which the forces generated during EF-G-catalyzed
translocation actively open the secondary structures (Qu et al.,2011). Here, by co-temporally monitoring both the mechanicaland the fluorescence coordinates of a ribosome translating on
an mRNA hairpin, we find that junction opening always occurs
a variable length of time after EF-G binding, unequivocally establishing that the helicase and translocase activities of the ribosome occur simultaneously (Figure 1). This observation therefore
indicates that the surface of the mRNA entry tunnel is not responsible for the opening of secondary structure prior to translocation(Qu et al., 2011), but it is still possible and likely that these positively charged amino acids contribute to the destabilization ofdownstream hairpins.
Direct measurements of the one-codon displacement of themRNA on the ribosome have shown that ribosomes are capableof generating forces that can unwind downstream secondary
structures during translocation (Liu et al., 2014). Such forces
could be exerted during either the forward or the reverse rotationof the 30S head domain, which results in the coordinated movement of the mRNA codons and the tRNA anticodons in the 30S
subunit. By using an antibiotic to stall the reverse 30S head rotation, we can identify the forward 30S head rotation, which ensues
after EF-G binding, as the likely candidate for the force generation step of the ribosome that results in hairpin opening (Figure 2).
Consistent with this finding, a previous single-molecule study
has reported a small rotation of EF-G with respect to the ribosome after GTP hydrolysis that could exert force to ‘‘unlock’’
the ribosome and drive translocation (Chen et al., 2016). In thiscase, unlocking corresponds to a ribosome intermediate inwhich the 30S head has fully rotated forward, whereas the 30S
body has moved back toward its non-rotated state. Furthermore, structures of the ribosome captured in intermediate states

G

Figure 5. Novel Ribosome-Dependent Hairpin Dynamics(A–F) Examples of single-ribosome fleezers trajectories in the presence of
10 nM Cy3-labeled-EF-G, where the unwinding step is split into two substeps that add up to one codon. Notice that EF-G remains bound until theentire step is completed. For comparison, a second unwinding step that
occurs in a single transition is shown in (A). The sub-steps are highlighted
with red arrows.
(G) An example of a single-ribosome fleezers trajectory in the presence of
10 nM Cy3-labeled-EF-G showing a region of hairpin hopping where several
EF-G molecules unproductively bind for long timescales (stars).

Molecular Cell 75, 1007–1019, September 5, 2019 1015

Figure 6. Model for Tight Coupling between Helicase and Translocase Activity of the Ribosome
At the beginning of each elongation cycle, the ribosome decodes the mRNA codon in the A-site and transfers the peptide chain from the P-site tRNA to the A-sitetRNA (step 1). Then, two major conformational changes occur in the ribosome before EF-G binding: inter-subunit rotation and partial 30S head rotation (step 2).
We propose that the ribosome could sense the presence of an mRNA hairpin during partial head rotation (step 2) to shift into a slow gear. After EF-G binding, thehead further rotates forward to a full 22" swivel as the body domain rotates back along the inter-subunit rotation axis (step 3). Our results demonstrate that
downstream mRNA hairpins are opened during this step. Finally, EF-G is released, and the ribosome is reset for another round of elongation (step 4). The 50S
subunit is shown in blue, the 30S body domain is shown in yellow, and the 30S head domain is shown in green. Inter-subunit rotation axis is shown in gray and thehead-rotation axis is shown in brown. The magnitude of rotation is represented by the size of the arrow.

of translocation also support that hairpin opening must occur
prior to or during forward 30S head rotation as the mRNA entry
tunnel cannot accommodate double-stranded RNA in the 30S
head-rotated state (Zhou et al., 2014). Overall, our results show
a tight temporal coupling between EF-G binding, internal dynamics of the ribosome, and mRNA hairpin opening.
This coupling is maintained even in cases in which the presence of a mechanical barrier of an mRNA hairpin junction forces
the ribosome to move in sub-codon steps; in these cases, wefind that the EF-G residence time is prolonged until the full codontranslocation has been completed (Figure 5). We speculate thatthese sub-codon steps could correspond to the head trapped inintermediate states of rotation, as have been observed structurally in vacant ribosomes or in the presence of EF-G analogs (Mohan et al., 2014; Pulk and Cate, 2013). We suggest that such astrict coordination between EF-G and the mechanical movementof the ribosome could be necessary to ensure that the reading
frame of the mRNA is always maintained.
Along the same lines, the !2 codon diffusive movement of theribosome at low force is accompanied by several futile EF-G
binding events (Figure 5G). As the mRNA hairpin used in theseexperiments consists of repeating valine codons, such largescale movement of the ribosome along the mRNA could arise
from the re-pairing of peptidyl tRNA to the neighboring valine codons. This hypothesis is consistent with a previous observationthat similar slips in the reading frame occur during frame shifting
only when the slippery sequence is positioned at the decodingsite of the ribosome (Yan et al., 2015). Moreover, EF-G binding
does not seem to be the rate-limiting event to exit this diffusive
state of the ribosome, as several EF-G molecules are seen tobind and unbind. Rather, given that we do not provide aminoacyl

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Molecular Cell 75, 1007–1019, September 5, 2019

tRNAs for alternative reading frame codons, it is possible that thearrival of EF-G must coincide with the ribosome re-registering in
the correct reading frame to allow the ribosome to exit this stateand continue translation.
Translation Regulation by Secondary Structures
The stability of downstream mRNA secondary structures has
been shown to modulate the rate of translation (Chen et al.,2013a, 2014; Qu et al., 2011; Wen et al., 2008). Here, we haveused force to modulate the strength of the mRNA barrier in frontof the ribosome to investigate how the magnitude of this barrier
affects the translation cycle. Until now, the naive expectation has
been that the resulting reduced translation rate is due to theslowing down of the mechanical step of mRNA unwinding before
a strong barrier (Qu et al., 2011). However, we find that thehairpin opening step of the ribosome catalyzed by EF-G is only
marginally affected in the presence of stronger hairpins (Figure 3).
Instead, the ribosome responds to stronger barriers by shifting
its operation !50% of the time into an alternative kinetic pathway(i.e., a slower gear, Figure 4).
We postulate that this shift into an alternative slower pathway
in front of strong mechanical barriers occurs prior to EF-G binding. A previous study showed that a frameshifting stimulatingstem loop destabilizes the inter-subunit rotated or hybrid stateof the ribosome and biases the ribosome toward the non-rotatedor classical state (Kim et al., 2014). EF-G binding to such a destabilized hybrid state could be impaired and possibly result in theslower pathway here. Consistent with this interpretation, in
another study in which a frameshifting stimulating pseudoknot
is present at the mRNA entry port, a moderate inhibitory effecton EF-G binding was observed (Caliskan et al., 2014).

However, the presence of downstream barriers in the context
of frameshifting also significantly slowed down events after EF-G
binding such as unlocking of the ribosome (Kim et al., 2014) andlate translocation events such as 30S head reverse rotation,
E-site tRNA release, and EF-G dissociation (Caliskan et al.,2014; Chen et al., 2013a; Kim et al., 2014). While we do observethat in the presence of an mRNA hairpin, the ribosome can takean alternative slower pathway to unwind the hairpin after EF-G
slow
in Figures 4D–4F), thereby slowing down thebinding (kunwinding
unlocking of the ribosome, we do not observe any delay in
EF-G release as a function of the strength of the barrier (Figure S6). The latter result is in contrast to previous studies but
could be rationalized because these late-stage translocationtransitions are likely specific to the presence of a slippery
sequence during frameshifting, which is not present in our study.
In fact, several studies have now shown that the slipping of thetRNA-mRNA complex occurs during the reverse 30S head rotation (Caliskan et al., 2014; Chen et al., 2013a, 2014; Kim et al.,2014). Consistently, structural data suggest that the paddles
that hold the mRNA reading frame in position disengage only
during reverse 30S head rotation (Zhou et al., 2013), allowing
for the uncoupled movement of the reading frame and subsequent frameshifting.
Moreover, the identity of the secondary structure could also
play an important role in determining which parts of the translation cycle are slowed down. For example, pseudoknots are mechanically stronger barriers than mRNA hairpins and have beenshown to result in a tilted conformation of the 30S head domain
during reverse 30S head rotation (Ramrath et al., 2012). It is
possible that either such a tilted conformation favors frameshifting or that the ribosome must frameshift to relieve this tilted stateand proceed with translation normally. It is also important to notethat in frameshifting conditions, the ribosome is not translating
through an mRNA structural barrier and only encounters thejunction upon approaching the slippery sequence; therefore,
many frameshifting studies may not have captured the bifurcated pathway that we observe prior to EF-G binding. In the
future, it would be very interesting to pursue the current experiments in frameshifting conditions, and we predict that a strong
effect on t release and downstream resetting of the ribosome
would be observed.
Although the structural details of the ribosome conformation
resulting in the slow gear have yet to be established, we speculate that the 30S head domain plays an important role in shifting
between the two distinct gears in the presence of an mRNA
structural barrier (Figure 6). Several studies have shown that ribosomes take alternate translation pathways in response to awide variety of challenging conditions such as in the absenceof GTP hydrolysis (on EF-G; Belardinelli et al., 2016), in the starvation of aminoacyl tRNAs during frameshifting (Caliskan et al.,2017), and in the presence of antibiotics (Peske et al., 2004).
Whether the mechanism of the switch into the slower pathway
is the same for all of these conditions has yet to be determined.
Finally, shifting into a slower gear to unwind secondary structures could be thermodynamically more favorable as the ribosome could take advantage of the thermal fluctuations of thejunction, thus minimizing energy dissipation. A recent study
has shown that the amount of energy dissipated in a non-equilib-

rium process can be significantly reduced if the process slows
down in those areas of the potential energy surface where there
is greater friction (Sivak and Crooks, 2016). This prediction has
been experimentally confirmed in a recent single-molecule study
that uses coincidentally a DNA hairpin (Tafoya et al., 2019); it was
speculated that the high thermodynamic efficiency displayed bymolecular machines could be explained if these have evolved toswitch into a slower gear in regions of their potential energy landscape that are associated with high dissipation, such as mRNA
hairpins.
STAR+METHODS
Detailed methods are provided in the online version of this paper
and include the following:d KEY RESOURCES TABLE
d LEAD CONTACT AND MATERIALS AVAILABILITY
d EXPERIMENTAL MODEL AND SUBJECT DETAILSB Source Organism
d METHOD DETAILSB Construction of mRNA Hairpin Plasmid
B mRNA Hairpin Synthesis
B dsDNA Handle Synthesis
B Protein Purification and Labeling
B Translation Buffer
B Preparation of Ribosome Initiation Complexes
B Deacylation of Total tRNAs
B Charging of DYK tRNAs
B Preparation of Stalling Mix
B Preparation of Ribosome Stalled Complexes
B Assembly of Ribosome Stalled Complexes on Strepta-

vidin Beads
d QUANTIFICATION AND STATISTICAL ANALYSIS

B Hidden Markov Model Step Finding Analysis (Non-fluorescence Ribosome Trajectories)
of Transitions for Fluorescence Ribosome Trajectories
B Bi-exponential Distribution Fittingd DATA AND CODE AVAILABILITY
B Identification

SUPPLEMENTAL INFORMATION
Supplemental Information can be found online at https://doi.org/10.1016/j.
molcel.2019.07.024.
ACKNOWLEDGMENTS
Dr. Ignacio Tinoco Jr. passed away on November 15, 2016. We thank Thy MinhThy Pham and William John van Patten for their help in the preparation of reagents; Charles E. Wickersham, Prof. Matthew Comstock, and Prof. Taepjik
Ha for their help in building the instrument; and Lisa Alexander, Liang Meng
Wee, Maya Sen, and Shannon Yan for useful discussions. This research was
supported by the Howard Hughes Medical Institute (instrumentation and ribosome biochemistry); by NIH grants R01GM071552 and R01GM032543 (fluorescence experiments); and the Nanomachine program (KC1203), funded by
the Office of Basic Energy Sciences of the U.S. Department of Energy (DOE)
contract no. DE-AC02-05CH11231 (data analysis algorithms). F.F. acknowledges support from an EMBO postdoctoral fellowship and M.R. acknowledges support from the Damon Runyon Cancer Research Foundation (DRG2096-11).

Molecular Cell 75, 1007–1019, September 5, 2019 1017

AUTHOR CONTRIBUTIONS
V.P.D., F.F., M.R., L.L., H.F.N., I.T., and C.B. designed the study. V.P.D., F.F.,
and L.L. prepared all of the reagents. M.R. assembled the instrument. V.P.D.,
F.F., and M.R. collected the data. A.L. and V.P.D. designed and implemented
the data analysis procedures. V.P.D., A.L., and F.F. analyzed the data. V.P.D.,
F.F., A.L., L.L., H.F.N., and C.B. wrote the manuscript.
DECLARATION OF INTERESTS
The authors declare no competing interests.
Received: February 15, 2019
Revised: June 12, 2019
Accepted: July 15, 2019
Published: August 27, 2019
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Molecular Cell 75, 1007–1019, September 5, 2019 1019

STAR+METHODSKEY RESOURCES TABLEREAGENT or RESOURCESOURCEIDENTIFIER

Agilent Technologies

Cat # 200152

Bacterial and Virus Strains
SURE 2 Supercompetent Cells
Chemicals, Peptides, and Recombinant Proteins
Ascorbic Acid

Sigma-Aldrich

Cat # A92902

ATP solution

GE Healthcare

Cat # 45-001-34

BsaI-HF

NEB

Cat # R3535

Catalase

EMD Millipore

CAS # 9001-05-2

Chloroform

Sigma-Aldrich

Cat # C2432

Cy3 Maleimide Mono-Reactive

GE Lifesciences

Cat # PA23031

Cyclooctatetraene

Sigma-Aldrich

Cat # 138924

EcoR1-HF

NEB

Cat # R3101S

Glucose Oxidase from Aspergillus Niger

Sigma-Aldrich

Cat # G2133

GTP solution

GE Healthcare

Cat # 45-001-345

KpnI-HF

NEB

Cat # R3142S

L-Ascorbic Acid

Sigma-Aldrich

Cat # A92902

Phenol solution

Sigma-Aldrich

Cat # P4557

Phenol:Chloroform:Isoamyl Alcohol

Sigma-Aldrich

Cat # P2069

p-Nitrobenzyl alcohol

Sigma-Aldrich

Cat # N6251

RNaseOUT

Invitrogen

Cat # 10777019

Superase,InInvitrogen

Cat # AM2694

TroloxSigma-Aldrich

Cat # 238813

Valine tRNA

Sigma-Aldrich

Cat # R2645

Invitrogen

Cat # AM1333

50 handle forward primer: /5Biosg/GGGCGACACGGAAATGTTGAATAC

IDT

N/A

50 handle reverse primer: 50 -GTCCATATGTATATCTCC
TTC(revT or /iSpC3/)GAGCGCTTGTTTCGGCGTGG-30
anneals to: 50 - GAAGGAGAUAUACAUAUGGAC %30

IDT

N/A

30 handle forward primer: 30 GCACATGTCTTGCGTTACTTAA-50 -50 -GAACGCAATGCGTCTGGGCGC-30 anneals to: 50 - CGUGUACAGAACGCAAUGAAUU-30

IDT

N/A

30 handle reverse primer: /5Biosg/TTTTTCTAAATACATTCAAATATGTATCCG

IDT

N/A

gBlock1: TAATACGACTCACTATAGGGCGAATTGGGTACCAGTGGCTGAGGCTTAACTAGTTCTAGAAATAATTTTGTTTAACTTTAAGAAGGAGATATACATATGGACTACAAGGATGACGATGACAAGAAGGTGGTCGTGGTAGTAGTGGTTGTTGTG
GTAGTCGTTGTTGTGGTGGTCGTAGTCGTGGT
CGTGGTGGTTGTTGTAGTTGTTGTTGTTGTTGT
AGTGGTTGTGGTAGTGGTAGTCGTGGTGGTTG
TTGTAGTGGTTGTTGTAGTGGTGGTGGGCCAC
GCGGCGAAAGCCTCATTGGTCTCGTTTTGC
GCGC

IDT

N/A

Critical Commercial Assays
MEGAscript! T7 Kit
Oligonucleotides(Continued on next page)e1 Molecular Cell 75, 1007–1019.e1–e5, September 5, 2019

Continued
REAGENT or RESOURCESOURCEIDENTIFIER

gBlock2:TATATATTTTCGAGACCAATGAGGCT
TTCGCCGCGTGGCCCACCACCACTACAACAACCACTACAACAACCACCACGACTACCACTACCACAACCACTACAACAACAACAACAACTACAACAACCACCACGACCACGACTACGACCACCACAACAACGACTACCACAACAACCACTACTACCACGACCACCGTGTACAGAACGCAATGAATTCGCGCGC

IDT

N/A

Stratagene

N/A

Cython(Behnel et al., 2011)

N/A

MATLAB

Mathworks

N/A

Econo-pac 10DG Desalting Column

Bio-radCat # 7322010

HiPrep 16/60 Sephacryl S-300 HR Column

GE Healthcare

Cat # 17-1167-01

HiTrap Q HP Column

GE Healthcare

Cat # 17115401

HisTrap HP Column

GE Healthcare

Cat # 17524801

Recombinant DNA
pBluescript SK+
Software and Algorithms

Other

illustra MicroSpin G-25 Column

GE Healthcare

Cat # 27-5325-01

Streptavidin Coated Beads (0.84 mm diameter)

Spherotech

Cat # SVP-08-10

TSKgel Phenyl-5PW Column, Glass

Tosoh Bioscience GmbHP/N # 0008804

LEAD CONTACT AND MATERIALS AVAILABILITY
Further information and requests for resources should be directed to and will be fulfilled by the Lead Contact, Carlos Bustamante(carlosb@berkeley.edu).
EXPERIMENTAL MODEL AND SUBJECT DETAILS
Source Organism
Plasmid for mRNA hairpin was cloned in SURE 2 Supercompetent cells. All other plasmids were cloned in Escherichia coli DH5a cells.
All proteins were purified from Escherichia coli BL21 (DE3) cells. Ribosomes were harvested from Escherichia coli MRE600 cells.
METHOD DETAILS
Construction of mRNA Hairpin Plasmid
To generate the DNA sequence containing the complete mRNA used in the experiments two gBlocks (IDT), each containing one armof the long hairpin structure, were ordered. gBlock1 (covering the 50 side of the hairpin sequence) was digested with KpnI and BsaI,
whereas gBlock2 (covering the 30 side of the hairpin sequence) was digested with BsaI and EcoRI. BsaI digestion generated the
UUUU tetraloop that caps the long hairpin structure. The resulting DNA pieces were inserted into the KpnI and EcoRI sites of pBluescript SK+. This generated the following sequence (sequence contributed by pBluescript SK+ including the T7 promoter as well as
KpnI and EcoRI sites is underlined):
TAATACGACTCACTATAGGGCGAATTGGGTACCAGTGGCTGAGGCTTAACTAGTTCTAGAAATAATTTTGTTTAACTTTAAGAAGGAGATATACATATGGACTACAAGGATGACGATGACAAGAAGGTGGTCGTGGTAGTAGTGGTTGTTGTGGTAGTCGTTGTTGTG
GTGGTCGTAGTCGTGGTCGTGGTGGTTGTTGTAGTTGTTGTTGTTGTTGTAGTGGTTGTGGTAGTGGTAGTCGTGGTGGTTGTT
GTAGTGGTTGTTGTAGTGGTGGTGGGCCACGCGGCGAAAGCCTCATTGGTCTCGTTTTCGAGACCAATGAGGCTTTCGCCGC
GTGGCCCACCACCACTACAACAACCACTACAACAACCACCACGACTACCACTACCACAACCACTACAACAACAACAACAACTACAACAACCACCACGACCACGACTACGACCACCACAACAACGACTACCACAACAACCACTACTACCACGACCACCGTGTACAGAACGCAATGAATTC
The resulting plasmid was then linearized by digestion with EcoRI to generate the template for in vitro transcription.

Molecular Cell 75, 1007–1019.e1–e5, September 5, 2019 e2

mRNA Hairpin SynthesismRNA hairpin was made by in vitro transcription of the plasmid linearized with EcoRI using MEGAscript T7 Transcription Kit. 0.5 mg oftemplate was used per 20 mL of reaction and transcription was carried out at 37"C for 4 hours. mRNA was then purified via a phenolchloroform extraction followed by ethanol precipitation and a MicroSpin G25 column. mRNA was stored at %20"C.
dsDNA Handle Synthesis
The mRNA hairpin was tethered between two beads via a 50 handle and 30 handle as described in Figure S1. Both the handles are!2.5 kb in length and were generated by PCR using modified primers. The 50 handle forward and the 30 handle reverse primers werelabeled with biotin at their respective 50 ends through which these handles are attached to streptavidin coated polystyrene beads.
The 50 handle reverse primer has an inverted base that efficiently terminates Phusion DNA polymerase to generate a 21 nts overhangthat anneals to the 50 end of the mRNA hairpin. The 30 handle forward primer has 18 inverted bases at its 50 end that efficiently terminates either Phusion or Taq polymerase and results in an 18 nts overhang that anneals to the 30 end of the mRNA hairpin.
Protein Purification and Labeling
The ribosomes were harvested and purified from Escherichia coli MRE600 cells (Moazed and Noller, 1989a) and S-100 enzymes were
purified as described previously (Traub et al., 1981). Initiation factors (Lancaster and Noller, 2005), wild-type EF-G (Wilson and Noller,
1998) and EF-Tu (Boon et al., 1992) were purified via the engineered His-tag using a HisTrap column followed by anion exchange
chromatography with a HiTrap Q column. Plasmid for Val-RS was obtained from Prof. Susan Marqusee and purified via an engineered His-tag. Plasmid for S73C EF-G was obtained from Prof. Joseph Puglisi. To fluorescently label EF-G (Chen et al., 2013b),
S73C EF-G was dialyzed into a labeling buffer (20 mM Tris,HCl pH 7.5, 100 mM NaCl and 0.1 mM TCEP) and 6x molar excess of
Cy3-monomaleimide was added. Protein + dye solution was rotated gently in a nutator at room temperature for 2 hours and thenat 4"C over night. Free Cy3 was removed by passing the labeled protein through three 10DG desalting columns and then througha sephacryl S300 size exclusion column. Labeled EF-G was separated from unlabeled EF-G through hydrophobic interactions chromatography with a TSKgel Phenyl 5-PW column. All proteins were stored in 25 mM Tris,HCl pH 7.5, 60 mM NH4Cl, 10 mM MgCl2 and5 mM b-Me unless stated otherwise. All proteins were flash frozen in liquid nitrogen and stored in %80"C.
Translation Buffer
The in vitro translation buffer is composed of 40 mM HEPES,KOH pH 7.5, 60 mM NH4Cl, 10 mM Mg(OAc)2, and 6 mM 2-mercaptoethanol. Additionally, either 10 mM sodium azide (for non-fluorescence experiments) or 12.5 mM ascorbic acid (for fluorescence
experiments) was added to the translation buffer to prolong tether lifetimes in optical tweezers.
Preparation of Ribosome Initiation Complexes
Initiation complexes were assembled in bulk by mixing 2 mM of 70S ribosomes, 0.1 mM mRNA, 1.75 mM charged f-Met tRNA, 1 mM
GTP and 1 mM of each initiation factor (IF1, 2 and 3) in translation buffer and incubating at 37"C for 15 min. They were flash frozen in
2 mL aliquots; each aliquot contains 0.2 pmol of mRNA and 4 pmol of 70S ribosomes.
Deacylation of Total tRNAs
Total tRNAs were dissolved in ultra-pure water to a concentration of !1 U/mL and stored in 50 mL aliquots at %80"C. To deacylate andunfold tRNAs, 300 mL of 45 mM Tris,HCl pH 8.0 was added to a 50 mL aliquot of total tRNA and incubated at 85"C for 2 min and thenroom temperature for 10 min. 6.25 mL of 1 M MgCl2 was added and the reaction was incubated at 37"C for 30 min to allow the tRNAs
to refold. Then, 30uL of 3M KOAc pH 5.3 was added and the reaction was placed on ice till cold. Finally, the deacylated tRNAs were
purified via two phenol extractions and one chloroform extraction followed by ethanol precipitation. The deacylated tRNAs were
resuspended in 10mM KOAc pH 5.3 and stored in %80"C.
Charging of DYK tRNAs
25 U of deacylated total tRNAs were charged using 20 mL of DEAE purified S100 extract in a total volume of 200 mL with a buffer
composed of 50 mM HEPES,KOH pH 7.5, 50 mM KCl, 10 mM MgCl2, 5 mM DTT, 250 mM of each amino acid (D, Y, and K) and
4 mM ATP. This reaction was incubated at 37"C for 20 min. Then, 20 mL of 3M KOAc was added and the reaction was placed on
ice till cold. Finally, two phenol extractions and one chloroform extraction were performed followed by ethanol precipitation. The
charged DYK tRNAs were resuspended in 10mM KOAc pH 5.3 and stored in %80"C.
Preparation of Stalling Mix
A 50 mL reaction of 1 mM GTP, 1 mM ATP, 24 mM EF-Tu, 4 mM EF-G and 1 mL of Superase,In was made in translation buffer andincubated at 37"C for 15 min. 6.4 U of charged DYK tRNAs were added to the reaction and further incubated at 37"C for 5 min.
The reaction was then placed on ice until cold and diluted with 350 mL of translation buffer containing 1 mM ATP and 1mM GTP.
This stalling mix was flash frozen in 10 mL aliquots and stored in %80"C.

e3

Molecular Cell 75, 1007–1019.e1–e5, September 5, 2019

Preparation of Ribosome Stalled Complexes
One 10 mL aliquot of stalling mix was added to one 2 mL aliquot of ribosome initiation complexes and the resulting sample was incubated at 37"C for 7 min. 1 mL of 250 nM 50 handle was added to the reaction and further incubated at 37"C for 2 min and then at room
temperature for 5 min. The sample was kept on ice for the rest of the day. A fresh sample of stalled complexes was made every day.
Assembly of Ribosome Stalled Complexes on Streptavidin Beads
20 mL of 0.1% solution of streptavidin coated polystyrene beads was made in translation buffer and vortexed at high speed for 30 min.
To make the ‘sample beads’, 2 mL of the ribosome stalled complexes were deposited on 0.75 mL of 0.1% streptavidin coated polystyrene beads (0.84 mm diameter) and incubated at room temperature for 5 min. ‘Handle beads’ were made by mixing 1 mL of 0.1%
streptavidin coated polystyrene beads with 1 mL of 50 nM 30 handle. To form a tether, a ‘sample bead’ with ribosome-mRNA-50 handle
complex was held in an optical trap and brought close to a ‘handle bead’ with the 30 handle held in the other optical trap. Upon the
hybridization of the 30 overhang of the 30 handle bead with the 30 side of the mRNA, a tether was formed (Figure S1A). We then unfold
the partially translated mRNA hairpin to verify that the ribosome is accurately stalled with the first valine codon in the P-site (Figure S1B). Then, the tethered stalled ribosome is held at either a constant force with force feedback (for experiments without fluorescence) or a semi-passive constant force with a range of 13-16 pN for high force or 5-7 pN for low force (experiments with fluorescence), and translation is restarted by the addition of a factor mix. The factor mix, composed of 1 mM ATP, 1 mM GTP, 400 mM
valine amino acid, 2 mM Val-RS, 2 mM valine tRNA, 4 mM EF-Tu, and variable amount of EF-G (dependent on the experimental condition), is incubated at 37"C for 20 min to pre-charge valine tRNA. When, Cy3-EF-G was used, a cocktail composed of 4 mM Trolox,
1 mM 1,3,5,7-cyclooctatetraene, 1 mM p-nitrobenzyl alcohol, 0.8% (w/v) glucose, 300 mg/ml glucose oxidase, 40 mg/ml catalase and12.5 mM ascorbic acid was added to the factor mix to prolong fluorophore and tether lifetimes. Catalase was pre-incubated with
RNase-Out to suppress RNase activity. Factor mix was made fresh every 2 hours due to pH change caused by the activity glucose
oxidase and catalase.
QUANTIFICATION AND STATISTICAL ANALYSIS
Hidden Markov Model Step Finding Analysis (Non-fluorescence Ribosome Trajectories)
An automated algorithm was developed to detect one-codon steps while being robust against the main source of spurious noise, i.e.,
non-cooperative hairpin unzipping (such as the one seen in Figure S5A, middle trace around time = 20 s). We model the stepping
process as follows: at any given time t, the ribosome may be at any codon position c in the hairpin (1 %c%cmax = 50); and somemore base pairs (nR0) may additionally be open at the base of the hairpin. For practicality, we assume n < nmax = 10 (this assumption
is discussed below). Assuming that the initial tether extension is d and the size of one base single-stranded RNA is s, the tether extension at any given time is d + 2s3ð3c + nÞ where, ð3c + nÞ gives the total number of base pairs open. For now, we will assume that d ands are known, fixed values. The measurement noise can be approximated as normally distributed with a standard deviation s, which isadded to the actual tether extension.
At any given time point, one of four transitions may occur:1) the ribosome advances by one codon (c /c + 1); this is associated by a decrease of n by 3 – if at least that many bases wereopened – (n/maxðn % 3; 0Þ),2) the ribosome does not advance (c/c), but the hairpin opens by one base (n /n + 1),3) the ribosome does not advance (c/c), but the hairpin closes by one base – if at least one base is open (n/maxðn % 1; 0Þ),4) neither c nor n change.
Let pfwd , popen , pclose , and 1 %pfwd % popen % pclose be the respective probabilities of the four events.
Overall, this setup fully describes a hidden Markov model with cmax 3nmax states (each indexed by a pair ðc;nÞ), with the structure of
the transition matrix implied by the transitions listed above. The parameters pfwd , popen , pclose , and s can fitted using the standard
Baum-Welch algorithm (Rabiner, 1989). Note that the transition matrix, of size ðcmax nmax ; cmax nmax Þ, is actually sparse —there are
only 4cmax nmax nonzero entries—; this important observation allows us to speed up the summations in our custom Cython (Behnel
et al., 2011) Baum-Welch implementation, both in the forward-backward step and the update step, by skipping most summands.
In practice, it is difficult to a priori obtain d or s with high accuracy. Instead, we start with approximate ranges of estimates for them(d from the approximate position at the start of the trace, s from a freely-jointed chain estimate). We then repeat the above HMM fit
with various values of d and s sampled in their respective ranges, and pick the ðd; sÞ pair for which the HMM fit yields the highest final
likelihood.
The output of the procedure is thus a ðd; sÞ pair and a HMM fit, which includes in particular a gt ðc; nÞ matrix (in the notation used by(Rabiner, 1989)) indicating the probability, at any time point, that the system is at a state ðc;nÞ. In practice, we do not care about the
number of additional open base pairs; we thus marginalize gt over n (i.e., for each value of c, sum over all values of n) to obtain theprobability gt ðcÞ that the ribosome is at codon c at time t. We then select for each time t the codon c with the maximum probability.
However, there remain two issues with our approach. First, the HMM as described above yields, in practice, a clearly unsatisfactory fit, where the value of the codon position is always underestimated, and all fluctuations in the signal are assigned to fluctuations

Molecular Cell 75, 1007–1019.e1–e5, September 5, 2019 e4

in the number of additional open base pairs. This arises because changes in n have better resolution than changes in c and can goboth forward and backward; it is thus easier for the algorithm to adjust n than c). To avoid this effect, we force popen =pclose (at every
iteration of the Baum-Welch algorithm) to a fixed ratio, estimated from the thermodynamic stability of the hairpin as computed bymfold (adjusting for the destabilization from the tension and the additional ribosome destabilization) (Qu et al., 2011; Zuker, 2003).
In practice, this ratio is small (!0.1) and suitably penalizes excessive fluctuations in n.
Second, note that our estimate of c may not be equally good at every time point. In particular, when the number of additional open
base pairs is much more than one codon, any reasonable knowledge of the actual ribosome position is essentially lost (this is thereason why we could assume n < nmax : there is little information in the time intervals where nR10 anyways). Such time intervalsP
are characterized by the quantity gt ðc; n < 3Þ = n˛f 0;1;2g gt ðc; nÞ being relatively small; in other words, there is a large probability
that there is more than one codon worth’s of additional open base pairs. We chose to set a cutoff on this value: we excluded any
region where gt ðc; n < 3Þ < 1=2. Additionally, if additional base pairs are opened, we discard downstream steps depending on thenumber of base pairs opened. If n%3, we discard the current and the following step. If 4 %n%6, we discard the current and thefollowing two steps. If 7 %n%9, we discard the current and the following three steps. Finally, if n > 9, we discard the current andthe next four steps.
All high force data (force R 10 pN) was downsampled to 133 Hz and all low force data (force % 5 pN) was downsampled to 66 Hz,
except data collected at 5 pN with 30 nM EF-G, where data had to be downsampled to 44 Hz for convergence of the fit. At high force,at least one to two points were required to establish a dwell and at low force five to seven points were required due to a decrease insignal to noise at low force.
Identification of Transitions for Fluorescence Ribosome Trajectories
Fluorescence and tweezers data were truncated to ± 5 s around the event of interest. Transitions in the data were fitted using thefollowing two algorithms:
1) A classic two-state hidden Markov model (using hmmlearn, https://hmmlearn.readthedocs.io/en/0.2.0/).
2) The Pruned Exact Linear Time (PELT) segmentation method (Killick et al., 2012) (we used a custom Cython (Behnel et al., 2011)
implementation of the algorithm), which efficiently finds, for a given time series, the best least-square fit to it by a stepwise constant function with a given number of steps. More accurately, PELT finds the best fit for a given per-step penalty; we used adichotomic search to find the penalty that yielded a given number of steps.
Bi-exponential Distribution Fitting
All fits were made to cumulative distributions using MATLAB’s inbuilt fit function. Bayesian information criterion was used to establish
when a bi-exponential fit was better than a single exponential fit. Several initialization conditions were tested to ensure convergenceof fits. All conditions converged to a single result despite different initialization condition except data collected at 5 pN with 10 mM
EF-G. We constrain this fit by using the same kslow path as that obtained from 5 pN data at 1 mM EF-G since we know that kslow path
is independent of EF-G concentration (Table S1). Fits were also performed by an orthogonal method, Maximum likelihood estimation,and the resulting fits were within error of current fits and no trends were changed. For 10 mM EF-G, we obtained n = 889 events from
24 molecules at 15 pN, n = 776 events from 19 molecules at 10 pN, n = 256 events from 8 molecules at 5 pN and n = 551 events from
19 molecules at 3 pN. For 1 mM EF-G, we obtained n = 747 events from 23 molecules at 15 pN and n = 403 events from 13 molecules
at 5 pN. For 100 nM EF-G, we obtained n = 732 events from 21 molecules at 15 pN and n = 509 events from 18 molecules at 5 pN. For
30 nM EF-G, we obtained n = 333 events from 9 molecules at 15 pN and n = 304 events from 11 molecules at 5 pN. For fluorescencemeasurements, we obtained n = 55 events from 9 molecules at 13-16 pN and n = 62 events from 14 molecules at 5-7 pN.
DATA AND CODE AVAILABILITY
The published article includes all relevant data generated or analyzed during this study. Additional data and codes are available fromthe corresponding author on request.

e5

Molecular Cell 75, 1007–1019.e1–e5, September 5, 2019

Molecular Cell, Volume 75

Supplemental Information

Co-temporal Force and Fluorescence Measurements
Reveal a Ribosomal Gear Shift Mechanismof Translation Regulation by Structured mRNAs
Varsha P. Desai, Filipp Frank, Antony Lee, Maurizio Righini, Laura Lancaster, Harry F.
Noller, Ignacio Tinoco Jr., and Carlos Bustamante

A

V50 mRNA:

AUG
M

Pre-stalling sequence
DYKDDDDKK

150 bp hairpin encoding 50 Valines Short residual hairpin
VVVV...
GHAAKASLVS

180bp

180bp

AUG

{

Traps brought close
to form a tether

streptavidin
coated

18 nts ssDNA overhang with sequence Handle beadcomplementary to the
3’end of the mRNA hairpin

Sample beadB

biotin3’

5’
streptavidin
coatedbiotinBulk assembly3’

biotinstreptavidin
coatedHandle beadSingle-molecule tethering of assembled ribosomes

180bp

22

Add 5’handle
and then deposit on
streptavidin beads

180bp

5’
5’ AUG

AUG

Formation of 18 bp
RNA/DNA hybrid
results in a tether

Sample bead

Force (pN)biotin{5’
streptavidin
coatedbiotin

3’

AUG

3’

20
18
16

hairpin no ribosome

14
2700
2800
2900
Trap position (nm)

180bp

180bp

5’

Force (pN)

22

Initiate
Ribosomes

3’5’

biotin

E P A

3’

20
18
16

CANNOTFORM
TETHER

14

12
2700
2800
2900
Trap position (nm)

E P A

22
Force (pN)

Stall
Ribosomes
172bp

172bp
5’
5’

biotin

3’
E P A

18
16

hairpin withstalled ribosome

14
2700
2800
2900
Trap position (nm)

3’
E P A

C

22
Flow intranslation mix

Confirm final postionof the ribosome

Force (pN)Fish for a correct tether
with stalled ribosome

20

5’
30bp

biotin
E P A

3’

20
18
16
14

residual
hairpin

12
2700
2800
2900
Trap position (nm)Figure S1. Assembly and tethering of ribosome stalled complexes, Related to Figure 1

Figure S1. Assembly and tethering of ribosome stalled complexes, Related to Figure 1(contd.)(A) Tethering geometry: a 2.5 kb double-stranded DNA handle with biotin on one end and 20 nts
of single-stranded DNA on the other end is annealed to complementary 20 nts at the 5’ side of anmRNA hairpin and is deposited on streptavidin coated beads – this assembly is called the sample
bead. A different 2.5 kb double-stranded DNA handle with biotin on one end and 18 nts of singlestranded DNA overhang that is complementary to the 3’ side of the mRNA on the other end isdeposited on streptavidin coated beads – this assembly is called the handle bead. A sample and ahandle bead are caught in two optical traps respectively and as they are brought together, theformation of an RNA/DNA hybrid between the 18 nts of ssDNA on the handle bead and the 3’
end of the mRNA hairpin results in a ‘tether’ in our optical tweezers setup.
(B) Sample preparation overview: Left, bulk assembly of ribosome-stalled complexes: First,
ribosomes are initiated on the mRNA upstream from the hairpin. Then, by supplying amino acids
D, Y and K, the ribosomes are stalled when the first valine codon reaches the P-site of theribosome. At this point, the foot-print of the ribosome results in the opening of 8 base-pairs of thehairpin. This bulk stalling strategy results in three types of complexes: 1) mRNA only – mRNA
that harbors no ribosome, 2) initiation complexes – mRNA where the ribosome initiatedsuccessfully but did not move past the start codon to stall at the valine codon, and 3) stalled
complexes – mRNA with ribosome correctly stalled with the valine codon in the P-site. Then, 5’
handle with 20 nts of single-stranded DNA that is complementary to 20 nts of the 5’ side of anmRNA hairpin is added. Right, Tethering of the three complexes formed in bulk and theirunfolding signatures: the 5’ handle can bind only to the mRNA only and stalled complexes. In thecase of mRNA only, there is no ribosome and hence the hairpin is fully closed and results in fourrips in the force-extension curves. As opposed to that, in the stalled complexes, the foot-print of
the ribosome has partially opened the hairpin (by 8 bps) and results in three rips with the first rip
displaying a characteristic shoulder. These unfolding signatures allow us to correctly identify aribosome-stalled complex on the tweezers.
(C) The unfolding signature of the short residual hairpin remaining at the end of translation isshown. If the ribosome stalled before the residual hairpin, the unfolding signature can be used toaccurately determine the exact stopping position of the ribosome.

Distance (codon)
Photon Rate (kHz)τunwinding = 0.2 s

τunwinding = 0.0225 s τunwinding = 0.0675 s

τunwinding = 0.045 s

τunwinding = 0.195 s

τunwinding = 0.025 s

τrelease = 0.01 s

τrelease = 0.0675 sτrelease = 0.225 sτrelease = 0.395 sτrelease = 0.245 sτrelease = 0.0325 s

1

1

1

1

1

1

0

0

0

0

0

0

6

8

5

7

4

6

6

8

3

5

5

6

2

4

10

8

8

7
6

6

5
4

4
0

0.2

0.4

0.6

0

0.1

0.2

1

0

0.2

3

0
Time (s)

12
7

10

4

4
0.2

0.4

0 0.2 0.4 0.6 0.8

2

0

0.2

0.4

Figure S2. Zoomed-in translocation events when hairpin is held under high force, Related to
Figure 1
Top panel: optical tweezers channel – 133 Hz data (shown in grey) is fit as described in Methods,
fit results are shown as solid black lines; Bottom panel: fluorescence channel – 100 Hz data (shownin grey) is fit as described in Methods, fit results are shown as solid green lines. Concentration of
Cy3-labeled-EF-G is 10 nM.

Time (s)τunwinding = 0.0075 s

τunwinding = 0.005 s

τunwinding = 0.06 sτrelease = 7.3775 sτrelease = 0.7925 sτrelease = 7.015 s

τrelease = 2.65 s

1

1

1

1

0

0

0

0

6

8
7
6

5

8

4

6

5

3

4

2
0

B

2

4

6

8

10

0.4

0

1
0.8
0.6
0.4
0.2
0

0

0.5

1

Tunwinding (s)

1.5

8
7
6
5
4

4

Cummulative Density

Cummulative Density

τunwinding = 0.0425 s

2

1.2 0
Time (s)

0.8

2

4

6

1
0.8
0.6
0.4
0.2
0

8

Cummulative Density

Photon Rate (kHz)Distance (codon)

A

0

5

10

Trelease (s)

15

20

10

3

1

0

2

3

4

10
T total (s)

15

20

1
0.8
0.6
0.4
0.2
0

0

5

Figure S3. Hairpin opening and EF-G binding and release measurements in the presence ofantibiotic fusidic acid, Related to Figure 2(A) Hairpin held at high force (> 13pN, weak hairpin) in the presence of 200 µM fusidic acid and
10 nM Cy3-labeled-EF-G. Top panel: optical tweezers channel – 133 Hz data (shown in grey) is
fit as described in Methods, fit results are shown as solid black lines; bottom panel: fluorescencechannel – 100 Hz data (shown in grey) is fit as described in Methods, fit results are shown as solid
green lines.
(B) Cumulative density distributions of time between EF-G binding and unwinding (!"#$%#&%#' ),time between unwinding and EF-G release (!()*)+,) ) and the total time EF-G is bound to theribosome (!-.-+* = !"#$%#&%#' + !()*)+,) ). Black open circles represent data collected with 200 µM
fusidic acid (n = 25 events, 13 molecules) and grey filled circle represent data without fusidic acid(n = 55 events, 9 molecules). Blue lines are fits to the cumulative distribution summarized in Table
S1.

Distance (codon)
Photon Rate (kHz)τunwinding = 0.08 s

τunwinding = 0.405 s

τunwinding = 0.03 s

τunwinding = 0.0525 s

τunwinding = 0.015 s

τunwinding = 0.625 sτrelease = 0.47 s

τrelease = 0.025 s

τrelease = 0.1 sτrelease = 0.1375 sτrelease = 0.015 s

τrelease = 0.035 s

1

1

1

1

1

1

0

0

0

0

0

0

10

10

12

5

10

4

8

3

9

8

8

8

6

6

8
6
6

6

4
0

0.4

0.8

4

4

7
0

0.2

0.4

0.6

2

0

0.1

0.2

2
0.3 0
Time (s)

0.1

0.2

0.3

2
0

0.04

0.08

0

0.4

0.8

Figure S4. Zoomed-in translocation events when hairpin is held under low force, Related to
Figure 3
Top panel: optical tweezers channel – 133 Hz data (shown in grey) is fit as described in Methods,
fit results are shown as solid black lines; Bottom panel: fluorescence channel – 100 Hz data (shownin grey) is fit as described in Methods, fit results are shown as solid green lines. Concentration of
Cy3-labeled-EF-G is 10 nM.

Figure S5. Bi-exponential distribution is shown in individual traces, Related to Figure 4

Figure S5. Bi-exponential distribution is shown in individual traces, Related to Figure 4(contd.)(A) Top panel: examples of single-ribosome optical tweezers trajectories of hairpin opening at 10µM EF-G (high force > 13 pN, weak hairpin). Black lines represent HMM fit for each step. Notethat the HMM fit is capable of ignoring reversible fluctuations at the hairpin base (middle trace,
long dwell around t = 20 s). Bottom panel: cumulative density distribution of the respective traces.
Single exponential (blue line) and bi-exponential (red line) fits are shown.
(B) Same as (A) but at low force < 7 pN, strong hairpin. Notice the difference in fit quality between
single and bi-exponential fit (blue and red lines respectively) in the cumulative density
distributions.

Figure S6. 01232452 follows a biphasic distribution that is insensitive to force, Related to Figure
4(A) Cumulative density distribution of !()*)+,) at high force (> 13pN, weak hairpin, n = 55 eventsfrom 9 molecules) and low force (< 7 pN, strong hairpin, n = 62 events, 14 molecules) in presenceof 10 nM Cy3-labeled-EF-G.
(B) Average residence times of EF-G before and after hairpin opening. Errors represent SEM.
(C) Results of fits to !()*)+,) at high force (> 13pN, weak hairpin) and low force (< 7 pN, strong
hairpin). Errors represent 95% confidence intervals.

Table S1. Summary of bi-exponential fits at various forces and EF-G concentrations, Related
to Figures 1-4
6&$)**
Force (pN)

[EF-G]

67 (s-1)

87 (%)

69 (s-1)

89 (%)

1510 µM

1.1 ± 0.03

91 ± 20.02 ± 0.06

9±2

15

1 µM

1.19 ± 0.03

84 ± 20.23 ± 0.03

16 ± 215

100 nM

0.66 ± 0.03

96 ± 10.04 ± 0.02

4±1

15

30 nM

0.53 ± 0.01

100

x

x

10

10 µM

0.75 ± 0.01

97 ± 10.05 ± 0.04

3±1

5

10 µM

0.71 ± 0.03

69 ± 20.19 ± 0.01

31 ± 25

1 µM

0.61 ± 0.03

56 ± 40.17 ± 0.01

44 ± 4

5

100 nM

0.41 ± 0.01

76 ± 20.08 ± 0.01

24 ± 2530 nM0.37 ± 0.13

29 ± 290.18 ± 0.03

71 ± 29310 µM

1.1 ± 0.04

51 ± 10.16 ± 0.01

49 ± 1

6"#$%#&%#' ([EF-G] = 10 nM)
Force (pN)

Antibiotic

67 (s-1)

87 (%)

69 (s-1)

89 (%)

14 – 17x

21.13 ± 4.80

92 ± 121.00 ± 3.82

8 ± 12

12 – 14x

27.53 ± 4.70

76 ± 81.89 ± 1.34

24 ± 87–9

x35.48 ± 10.38

61 ± 12

3.17 ± 1.57

39 ± 125–8

x12.7 ± 1.45

62 ± 40.89 ± 0.17

38 ± 413 – 15

Fusidic acid25.4 ± 15.4

96 ± 29

0.23 ± 0.03

4 ± 29

Data S1. Alternate kinetic schemes, Related to Figure 4
The time the ribosome spends at a particular codon before moving to the next codon is defined as!&$)**

τdwell

Codon n + 1
Codon nCodon n – 1
The simplest model to describe the movement of ribosome from codon : to codon : + 1 can be
described as:

kdwell
Codon nCodon n+1

In this case, the movement of ribosome at each codon is given by a single rate limiting event
6&$)** . The probability distribution of observing a dwell of time = would be given by:>(=) = 6&$)** ∙ B CDEFGHH ∙I
The cumulative probability of the above distribution is given by:
K(=) = 1 − B CDEFGHH ∙I
While this distribution fits ~90% of the data at high force, it does not fit data at low force (Figure
4A).
The data instead fits very well to a mixture of two exponential distribution given by:>(=) = 87 ∙ 67 ∙ B CDM ∙ I + 89 ∙ 69 ∙ B CDN ∙ I
where 67 and 69 are the rates of the two exponential distributions, 87 and 89 are the relative
populations of the two exponential distributions, and 87 + 89 = 1
The cumulative probability of the above distribution is given by:
K(=) = 1 − 87 ∙ B CDM ∙ I − 89 ∙ B CDN ∙ I
The dwell time distribution fits a mixture of exponentials (bi-exponential) at both high and low
forces. The result of the fit gives 67 , 69 and 89 (Figures 4B and 4C). We have 87 = 1 − 89 .

For a bi-exponential distribution, a simple scheme where the ribosome goes into an off-pathway
inactive state in presence of a barrier could be proposed:

kdwell
Codon nCodon n+1

kinactive kactive

inactive

Codon nIn order to solve this scheme exactly, let A be the probability of being in the active state at codonn and I the probability of being in the inactive state.
The following system of equation holds:OP= −(6dwell + 6inactive )P + 6active [
O=

(1)

O[
= −6active [ + 6inactive P
O=(2)Then Equation (1) can be rearranged as:[=

OP (6dwell + 6inactive )+P
6active O=
6active1

and then be plugged into Equation (2) to yield:
O9 P
OP+ (6dwell + 6inactive + 6active )+ 6dwell 6active P = 0
9O=O=This is a second order differential equation and the eigenvalues are:]^/C =−(6dwell + 6inactive + 6active ) ± a(6dwell + 6inactive + 6active )9 − 46dwell 6active
2and A and I are linear combinations of exp (]^ =) and exp(]C =).

At = = 0, [ = 0; thus[(=) = g(exp(]^ =) − exp(]C =))and then, from Eq (2), we have,P(=) =

1O[
6active]^ + 6active]C + 6active+[ = ghexp(]^ =) −exp(]C =)i
6inactive O= 6inactive
6inactive
6inactive

The probability of being either in the active or the inactive state at codon n, which is also the
distribution for the dwell time at codon n, is thus
P(=) + [(=) = g h]^ + 6active + 6inactive]C + 6active + 6inactiveexp(]^ =) −exp(]C =)i
6inactive
6inactivei.e., it is biexponentially distributed. Importantly, in this model, the ratio between the
populations of the two exponentials, −(]^ + 6active + 6inactive )/(]C + 6active + 6inactive ), is notindependent of the ratio between the rates of the two exponentials, ]^ /]C : it is generally notpossible to change the ratio between the exponentials’ populations without changing theobserved exponential rates. This is not consistent with our measurements, which show that achange in force causes a change in the population ratio without modifying the rates.
Therefore, we need a scheme where the pathway populations are determined independently ofand prior to the rate-limiting event in the two pathways:

fast

ksensorCodon n

kfast
int

slow

ksensorslow

kint

Branching
ratio(gives rise topopulations seenin Figure 4C)

Rate-limiting
step(gives rise torates seenin Figure 4B)

Now, this bifurcation could occur a) prior to EF-G binding or after EF-G release from previous
cycle, b) prior to EF-G binding through unwinding, or c) after unwinding through EF-G release:a) Pathway bifurcation and resetting occurs before EF-G binding or after EF-G release from previous cycle
Fast

fast

ksensorfast

kintermediate
Codon nCodon n

τunwinding

τrelease
Codon n+1

Codon n+1

kunwinding
slow

slow

ksensorkintermediate

Slowb) Pathway bifurcation occurs before EF-G binding and resets upon hairpin openingτunwinding

fast

kintermediate
Fast

fast

ksensor

Fast

Codon nτrelease

fast

kunwinding
Codon n+1

slow

ksensor

Slow

Slow

slow

kunwinding

slow

kintermediatec) Pathway bifurcation and resetting occurs after hairpin openingτunwinding

τrelease
fast

krelease
Codon nFast

Codon n+1

fast

kintermediate
Codon n+1

kunwindingslow

krelease

slow

Slow

kintermediate

Codon n+1

One possible way to distinguish these schemes is by looking at the distribution of !"#$%#&%#' ,time from EF-G binding to unwinding (Figure 4D). In a), neither !"#$%#&%#' or !()*)+,) would
have a bi-exponential distribution where the populations of the two distributions are forcesensitive. In b), !"#$%#&%#' would be described by a bi-exponential where the populations are
sensitive to force in a manner similar to that observed for !&$)** . In c), !()*)+,) would be
described by a bi-exponential where the populations are sensitive to force in a manner similar tothat observed for !&$)** . In our experiments, we observe that the distribution !"#$%#&%#' is also
best described by a mixture of two exponentials where the rates are force-insensitive (Figure 4E)
but the pathway populations are force-dependent (Figure 4F). Moreover, !()*)+,) , time fromunwinding to EF-G release (Figure S6) is insensitive to force and therefore we propose that thebifurcated pathway has reset prior to release and therefore favor model b).