Deprotonation stimulates productive folding in allosteric TRAP hammerhead ribozymes.

Abstract

Hammerhead ribozymes in crystals change conformation in response to deprotonation of the nucleophilic 2' OH, thereby aligning the hydroxyl for in-line displacement at the scissile phosphate. Published data do not address whether deprotonation affects folding in solution. Allosteric hammerhead "TRAPs," when activated by the appropriate oligonucleotide, show the expected log-linear relation between initial cleavage rate and pH. In contrast, attenuated TRAPs shows biphasic kinetics in which a rapid burst is followed by slow cleavage that is nearly independent of pH. Attenuated ribozymes are stimulated by urea at both low and high pH, confirming that rearrangement of secondary structure is rate-limiting for the attenuated ribozymes once they have folded. Plots of burst magnitude versus pH in the absence of urea show a sharp transition around pH 8.3, which is near the kinetic pKa for the cleavage reaction in Mg2+. Raising the pH after folding at pH 7.5 did not activate attenuated ribozymes even when the RNA was incubated at the elevated pH for extended periods prior to addition of Mg2+. In contrast, lowering the pH after folding at pH 9.5 rapidly re-established attenuation. Deprotonation of the ribozyme-substrate complex thus appears to alter the folding landscape such that a metastable "pre-activated" complex forms before the thermodynamically more stable attenuated state can be attained. From the initial partition into active and inactive conformers, we estimate that this deprotonation contributes approximately 1.2 kcal/mol toward stabilization of the active fold at a crucial step during folding of the TRAP. Assuming that the nucleophilic 2' OH is the relevant acid, its deprotonation would thus serve a dual role of favoring productive fold and enhancing the nucleophilicity of this oxygen.

Full text

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Deprotonation Stimulates Productive Folding in
Allosteric TRAP Hammerhead Ribozymes
Vanvimon Saksmerprome and Donald H. Burke*
Department of Chemistry
Indiana University
Bloomington, IN 47405-7102
USA
Hammerhead ribozymes in crystals change conformation in response to
deprotonation of the nucleophilic 20OH, thereby aligning the hydroxyl
for in-line displacement at the scissile phosphate. Published data do not
address whether deprotonation affects folding in solution. Allosteric
hammerhead “TRAPs,” when activated by the appropriate oligonucleo-
tide, show the expected log-linear relation between initial cleavage rate
and pH. In contrast, attenuated TRAPs shows biphasic kinetics in which
a rapid burst is followed by slow cleavage that is nearly independent of
pH. Attenuated ribozymes are stimulated by urea at both low and high
pH, confirming that rearrangement of secondary structure is rate-limiting
for the attenuated ribozymes once they have folded. Plots of burst magni-
tude versus pH in the absence of urea show a sharp transition around pH
8.3, which is near the kinetic pKafor the cleavage reaction in Mg2þ. Raising
the pH after folding at pH 7.5 did not activate attenuated ribozymes even
when the RNA was incubated at the elevated pH for extended periods
prior to addition of Mg2þ. In contrast, lowering the pH after folding at
pH 9.5 rapidly re-established attenuation. Deprotonation of the ribo-
zyme–substrate complex thus appears to alter the folding landscape
such that a metastable “pre-activated” complex forms before the thermo-
dynamically more stable attenuated state can be attained. From the initial
partition into active and inactive conformers, we estimate that this depro-
tonation contributes approximately 1.2 kcal/mol toward stabilization of
the active fold at a crucial step during folding of the TRAP. Assuming
that the nucleophilic 20OH is the relevant acid, its deprotonation would
thus serve a dual role of favoring productive fold and enhancing the
nucleophilicity of this oxygen.
q 2004 Elsevier Ltd. All rights reserved.
Keywords: allostery; hammerhead ribozyme kinetics; RNA folding
landscape; deprotonation; conformational equilibria
*Corresponding author
Introduction
The hammerhead ribozyme is a small RNA
module found in the satellite RNAs of some plant
viroid pathogens1and in the repetitive DNA of
newts, cave crickets and schistosomes.2–4Hammer-
head-like sequences have also been identified in
surveys of genome databases.5In the plant patho-
gens, hammerhead ribozymes catalyze the cis-clea-
vage that separates individual single-strand RNA
genomes following rolling-circle replication, after
which they also catalyze ligation of genomic ter-
mini to yield circularized RNA genomes. Because
cleavage specificity in trans can be readily pro-
grammed throughWatson–Crick
hammerhead ribozymes have received consider-
able attention as gene-inactivation agents.6–8Their
small size also makes them readily amenable to
structuralandmechanistic
mediated catalysis.9,10Several small ribozymes
and many protein ribonucleases utilize similar
chemical strategies in cleaving phosphodiesters
to generate 50hydroxyl and 20,30-cyclic phosphate
termini. Catalysis by hammerhead ribozymes is
strongly influenced by RNA structure, confor-
mational dynamics, binding of divalent metal ions
and proton transfer events. These parameters are
interrelated, and their respective contributions are
base-pairing,
studiesofRNA-
0022-2836/$ - see front matter q 2004 Elsevier Ltd. All rights reserved.
E-mail address of the corresponding author:
dhburke@indiana.edu
Abbreviations used: FRET, fluorescence resonance
energy transfer; TRAP, targeted reversibly attenuated
probes.
doi:10.1016/j.jmb.2004.06.027J. Mol. Biol. (2004) 341, 685–694

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often non-additive. The present work addresses a
heretofore under-appreciated relationship between
protontransferandfolding
ribozymes.
Proton transfer can play essential roles in RNA
structure and catalysis,11and was thought until
recently to be well-understood for the hammer-
head reaction. The logarithm of hammerhead-
catalyzed RNA cleavage rates varies linearly with
pH between pH 5.5 and 8.5. The slope of such
a plot varies considerably with the ribozyme
assayed, but is generally near unity, implying that
a singly deprotonated species participates in the
rate-determining step. The kinetic pKaof the reac-
tion varies with the pKa of the hydrated form of
the divalent metal ion used in the reaction
(approximately 8.0 in the presence of Co2þand 8.5
in Mg2þ), implying that a hydrated metal ion can
participate in the essential deprotonation.12The
rate-determining step could be bond formation
between the scissile phosphate and the adjacent 20
ribose hydroxyl, whose nucleophilicity increases
as much as 100,000-fold upon deprotonation.13On
the other hand, this step could also be a necessary
conformationalchange
requires a deprotonated species, such as alignment
for in-line displacement at the scissile phosphate.
Crystallographic analysis has identified several
conformations that may correspond to structural
intermediates along the reaction pathway of the
hammerhead ribozyme. Interconversion among
conformational states along this path is triggered
by metal ion binding or by deprotonation of the
nucleophilic 20OH. In the ground state, the 20OH
is not oriented for in-line attack on the reactive
phosphate. Cleavage in the crystal was prevented
inthesemolecules by
site ribose with a deoxyribose14or with a 20
O-methylribose.15,16A conformational change inter-
preted as being “early” along the reaction coordi-
nate was captured by freeze-trapping ribozymes
with an unmodified active site ribose in the pre-
sence of MgCl2. The scissile phosphate is moved
approximately 3 A˚in this structure, and the
adjoining ribose and base also move.16Another
conformation in which the 20hydroxyl is more
closely aligned for nucleophilic attack, interpreted
as reflecting a “later” stage on the reaction coordi-
nate, was observed when a kinetic bottleneck was
introduced by replacing the 50OH leaving group
with a 50-O-talo-methyl modification (50ribose
modified from –CH2– to –CH(CH3)–).17A differ-
ent type of bottleneck was produced by engineer-
ing a 50–50linked guanosine at the end of stem 1
(nucleotide 2.6) that base-paired with cytosine 10.4
in stem 2.18This ribozyme is catalytically compe-
tent in solution but not in the crystal, even though
the nucleophilic ribose 20oxygen is less than 2.3 A˚
from the reactive phosphate. A further rigid-body
rotation of stem 1, prevented by the stem 1–stem
2 interaction, is proposed to be necessary to bring
the nucleophilic oxygen into alignment and to
permit cleavage.18Rotation of stem I is also
ofhammerhead
inthe ribozymethat
replacing theactive
predicted
simulations19based on relaxation at 300 K from
the crystal structure of an unmodified hammer-
head with crystallographically identified bound
magnesium ions.16Further rearrangement at the
reactive phosphate is evident in a crystal structure
of the ribozyme–product complex.20Specifically,
the ground state C17–C3 hydrogen bond must be
broken for C17 (immediately 50of the cleavage
site) to loop out and expose the reactive phosphate
to essential nucleotides in domain 1 (the conserved
“CUGA” sequence that forms a uridine turn simi-
lar to that found in transfer RNA14). An NMR
structure of the product complex also shows sub-
stantial changes in the configuration of domain 1
upon cleavage.21
In recent years it has been suggested that depro-
tonation may play an essential role in hammerhead
dynamics. The early conformational change seen in
crystallographic analysis upon addition of Mg2þis
observed independent of pH and can be attributed
to metal binding alone. Similar results have been
noted in solution studies.22–24In contrast, the more
aligned conformation does not accumulate to high
occupancy below pH 8.0 in crystals that contain
the 50-O-talo-methyl modification, and this confor-
mation is not observed at any pH when 20OMe or
20F replaces the nucleophilic 20oxygen.25The
second conformational change therefore appears
to require a deprotonated 20OH at the cleavage
site. Furthermore, catalytically competent ribo-
zymes display biphasic cleavage kinetics in crys-
tals at low pH, with a lag phase that is log-linear
with pH followed by rapid cleavage that is nearly
independent of pH. Even at pH 6.0 the rapid
phase is on the same order as the maximal rate
observed by highly active hammerheads at pH
8.5. A conformational change is proposed to
propagate cooperatively throughout the crystal,
removing a normally rate-limiting conformational
barrier to cleavage. Thus, the pH dependence
normally observed for cleavage by hammerhead
ribozymes in solution may arise from a confor-
mational change that requires deprotonation of
the attacking 20OH,25provided that the crystal
environment does not
constraints.
Allosteric ribozymes are uniquely useful for
studying the factors that drive the formation of
active ribozyme conformations. Folding steps that
might otherwise be invisible can be isolated and
examined, because folding and activity depend on
binding to an effector analyte (such as a small
organic molecule, metal ion, oligonucleotide, pep-
tide or protein26). In the TRAP design (targeted
reversibly attenuated probes†), attenuation, acti-
vation and cleavage specificity are programmed
from solutionmolecular dynamics
introducespurious
†This definition replaces the original definition of
TRAP as targeted ribozyme attenuated probes in order to
acknowledge that aptamers can also be regulated
through TRAP mechanisms.
686
Folding Paths of Deprotonated Hammerhead Ribozymes

Page 3

into separate parts of the RNA.27Two sequence
modules are appended 30to a hammerhead (Figure
1). The distal module (“attenuator”) base-pairs
with the catalytic core and shuts down cleavage of
the RNA target. The proximal module (“anti-
sense”) acts as a flexible tether. Binding of a
“sense” strand to the antisense strand straightens
and stiffens the tether, releasing the attenuator
from the catalytic core and activating cleavage of
substrate RNA. Cleavage of a target RNA (such as
an mRNA for an essential protein) is thus depen-
dent upon the presence of an activator RNA (such
as a tumor-associated mRNA, or viral RNA). The
design has yielded more than 700-fold activation
of cleavageupon addition
oligonucleotide.28This magnitude is easily great
enough to exert a biological effect. Thus, RNA
degradation can be directed to occur in specific
cell types or at specific stages of development.
Kinetic and thermodynamic studies demonstrated
that the relative stabilities of the activation and
attenuating helices are primary determinants of
TRAP kinetics through their effects on pre-cleavage
equilibria.28
Here, we examine the effects of pH and urea on
TRAP cleavage rates. We find that attenuated ribo-
zymes are stimulated by urea at both low and
high pH, consistent with a model in which
rearrangement of base-pairing patterns is rate-
limiting for the attenuated ribozymes.27,28Oligo-
nucleotide-activated TRAPs show the expected
log-linear relation between cleavage rate and pH.
In contrast, the rate of cleavage by attenuated
hammerheads shows almost no dependence on
pH, as could be expected when rearrangement of
base-pairing patterns, rather than chemistry, is
rate-limiting. Unexpectedly, the reaction becomes
markedly biphasic at high pH, with the magnitude
of the burst depending on pH and on the length of
ofsensestrand
the attenuator used. Similar biphasic behavior is
observed for the activated form of the TRAP.
Order-of-additionexperiments
deprotonation of the ribozyme, possibly at the
attacking 20OH, alters the energetic landscape
during the folding process to favor formation of
the active fold without significantly altering the
final equilibrium between activated and repressed
states.
establishthat
Results
pH-dependent biphasic kinetics for
attenuated TRAPs
As a first step in defining the response of attenu-
ated TRAP ribozymes to variation in pH, initial
cleavageratesweremeasured
HH8A.8. This ribozyme is derived from hammer-
head HH829,30and carries both a 20 nucleotide
antisense segmentand
attenuator.27,28In the presence of activating oligo-
ribonucleotide “8A,” the logarithm of the initial
rate of single-turnover cleavage by this species
varied linearly from pH 6.1 to 8.1 with a slope for
log(kobs) versus pH of 0.83. Above pH 8.1 the log of
the initial cleavage rate increased non-linearly,
approaching a plateau (not shown). The activated
TRAP thus responds to pH in a manner that is
similar to previous observations for kinetically
well-behaved ribozymes.13,31
When oligonucleotide 8A is omitted from the
cleavage reaction at pH 7.5 to favor formation of
the attenuated TRAP, single-turnover cleavage
shows biphasic kinetic behavior. A small burst is
completed within a few minutes followed by con-
tinued cleavage at a slower rate.27Unexpectedly,
the biphasic nature of the reaction is much more
forribozyme
an eightnucleotide
Figure 1. Hammerhead TRAP ribo-
zyme design. (A) Schematic represen-
tation of TRAP activation through
bindingofa
oligonucleotide.27,28Green and pink
lines correspond to attenuator and
catalytic core sequences that interact
in the attenuated state. Pairing of a
sense activator strand with the anti-
sense (blue line) activates the ham-
merheadtocleave
Cleavage site in the substrate strand
is indicated by block arrow. (B)
Nucleotide sequences of TRAP ribo-
zymes and activating oligo 8A, fol-
lowing the color scheme used in (A).
Conventional numbering of positions
around the catalytic core is given.
Cleavage substrate is shown in bold
with cleavage site indicated. The
longer attenuator corresponds to that
of HH8A.10, the shorter one to that
of HH8A.8, and HH8A carries no
attenuator.
sense strand
itssubstrate.
Folding Paths of Deprotonated Hammerhead Ribozymes
687

Page 4

pronounced at elevated pH (Figure 2(A)), with
about half of the substrate being cleaved within 15
seconds of adding MgCl2(see inset, Figure 2(A)).
Samples taken just before adding MgCl2(the zero
time point) always showed no evidence of product
formation (Figure 2(B)). During the slow phase
(second phase), cleavage rate was nearly inde-
pendent of pH, ranging from 0.0012 min21to
0.0036 min21over a 100-fold change in hydroxide
concentration. Plotting the log of the fraction
cleaved versus time and extrapolating from the
slow phase back to zero time (Figure 2(A)) reveals
the fraction of the initial ribozyme–substrate
complex that participates in the burst phase. This
fraction shifts abruptly from 5–10% at low pH to
45–51% at high pH, with an apparent pKafor this
transition near 8.3 (Figure 2(D)). The existence of a
burst phase in single-turnover kinetics suggests
that the ribozyme partitions into active and inac-
tive conformations before the reaction is initiated,
and that the two conformations exchange slowly.
The simplest interpretation is that the inactive con-
formation corresponds to a state in which the
attenuator is paired with the core (RAttS), and that
in the active state the attenuator is unpaired and
stem II has reformed (RpS). Addition of MgCl2
converts RpS into the catalytically competent ribo-
zyme, while RAttS must rearrange its base-pairing
patterns in order to form RpS.
It was unexpected that the magnitude of the
burst should vary with pH. Deprotonation of the
ribozyme or substrate appears to alter the distri-
bution among possible pre-cleavage conformations
to favor RpS. To test this hypothesis, cleavage rates
were determined for ribozyme HH8A.10 as a func-
tion of pH. The 10 nt attenuator of this ribozyme
stabilizes the attenuated form relative to the 8 nt
attenuator of HH8A.8.27,28Biphasic kinetics are
again evident at high pH, and the magnitude of
the burst phase again shows a transition near pH
8.3 (Figure 2(C)). The magnitude of the burst is
notably diminished for this ribozyme relative to
that observed for HH8A.8, attributable to greater
stabilization of the “off” state by the longer and
more stable attenuator. Approximately 3% of the
RNA participates in the burst phase at pH 8.1,
increasing to 7% at pH 8.9. Cleavage rates subse-
quent to the burst phase for HH8A.10 appear to
depend more strongly on pH than was observed
for HH8A.8, with logðkobsÞ varying linearly with
pH with a slope of 0.7.
Kinetics in slow second phase is dominated
by refolding
Denaturants
molecules by destabilizing ground state confor-
mations and decreasing the activation energy
required to reach a partially unfolded confor-
mational transition state. To evaluate the effects of
denaturant on TRAP kinetics, cleavage rates were
determined for ribozymes folded in the presence
of urea. For ribozyme HH8A.8, cleavage at pH 7.5
was stimulated fourfold as urea concentrations
increased from 0 M to 2 M (0.002 min21versus
0.008 min21; Figure 3(A)). Higher concentrations
were increasingly disruptive, but did not com-
pletely eliminate cleavage even at 6 M urea. In con-
trast, non-attenuated ribozymes were inhibited by
urea, with half-maximal activity observed between
1 M and 2 M urea both for ribozyme HH8A alone
(lacking attenuator sequences) and for ribozyme
HH8A.8 in the presence of activating oligonucleo-
tide 8A (Figure 3(B)). For each of these ribozymes,
the denaturant is expected to increase the fraction
of time the RNA spends sampling non-ground
state base-pairing arrangements. For the attenuated
ribozyme, this increases the rate of conformational
acceleraterefolding of macro-
Figure 2. pH dependence of substrate cleavage by
HH8A.8andHH8A.10.
uncleaved) versus time display biphasic cleavage by
HH8A.8 in the absence of activator. Diamonds, pH 6.5;
open squares, pH 7.0; triangles, pH 7.5; £ , pH 8.1; aster-
isks, pH 8.5; circles, pH 8.9; filled squares, pH 9.5. Data
points after initial burst were fit to a best-fit line. Insets
here and in (C) show detail of cleavage during first min-
ute. (B) Sample gels showing reaction progress at pH
8.1 (top) and 8.9 (bottom). Time points were taken at 0
second, 15 seconds, 30 seconds, 45 seconds, one minute,
five minutes, ten minutes, 30 minutes, and 60 minutes
(left to right). No cleavage is evident at zero time at
either pH, but the amount cleaved within first 15
seconds is clearly much greater at the higher pH.
(C) Biphasic cleavage by HH8A.10 in the absence of acti-
vator, following same symbols as in (A). (D) Magnitude
of burst phase as determined by extrapolating slow
phase cleavage to zero time plotted as a function of pH
of the reaction for HH8A.8 (diamonds) and HH8A.10
(squares). Transition at pH 8.3 is indicated by a vertical
arrow.
(A)Plots ofln(fraction
688
Folding Paths of Deprotonated Hammerhead Ribozymes

Page 5

re-equilibration from the attenuated ground state
into an active state (RAttS ! RpS). The ground states
of the non-attenuated and activated ribozymes are
already in an active base-pairing arrangement and
are thus inhibited by sampling other base-pairing
arrangements.
Similar effects are observed at pH 8.9, where
urea increases the observed single-turnover clea-
vage rate fourfold from 0.004 min21to 0.017 min21
(Figure 3(C)). High urea concentrations are again
disruptive. Interestingly, at both pH 7.5 and 8.9,
the reaction ceases to be biphasic at elevated urea
concentrations (data at pH 7.5 are shown in Figure
3(D)). This may be due both to slower cleavage
during the burst phase (native conformation par-
tially disrupted by the urea) and to accelerated
cleavage after the burst (more frequent sampling
of alternative conformations by RAttS). The combi-
nation of these two effects would blur the distinc-
tion between the two phases, yielding apparent
monophasic behavior. Unlike the observations at
lower pH, maximal cleavage at pH 8.9 is observed
at 4 M urea.
Raising the pH does not relieve the folding trap
The initial folding partition between RAttS and
RpS determines the magnitude of the initial burst.
To test the relationships among pH, initial folding
partition andsubsequent
HH8A.8 and radiolabeled substrate were folded
together in 50 mM Tris–HCl buffered to pH 7.5.
This RNA was then diluted into an equal volume
of 250 mM Ches-buffered MgCl2 at pH 9.5 to
initiate cleavage at a final pH of approximately
9.4, well above the pH at which the transition is
observed. A relatively small fraction (10%) of the
substrate is cleaved in the burst phase under
these conditions (Figure 4(A), £ ), similar to the
behavior of this ribozyme when both folding and
cleavage are carried out at the lower pH. A low
pH at the time of folding therefore establishes the
partition between the two states of the ribozyme,
even if the cleavage reaction is subsequently
carried out at a higher pH.
refolding,ribozyme
Slow re-equilibration between RAttS and RpS
upon deprotonation could be masked in the
experiments above. To rule out this possibility, the
kinetic stability of the low-pH attenuated form
was evaluated. Hammerhead HH8A.8 was folded
with substrate in dilute Tris–HCl at pH 7.5. This
RNA was made alkaline by the addition of Ches
buffer without added MgCl2, and then incubated
for variable times (t) at the high pH before MgCl2
Figure 4. Examination of refolding upon pH shift.
Cleavage was monitored as a function of time under the
two experimental regimes indicated above the plots and
described in the text. (A) pH was changed after folding,
simultaneously with addition of MgCl2. Triangles, fold
at pH 7.5 and react at pH 7.5; squares, fold at pH 9.5
and react at pH 9.5; £ , fold at pH 7.5 then react at pH
9.4; open circles, fold at pH 9.5 then react at pH 7.8.
(B) pH was shifted from pH 7.5 to 9.4 without added
divalent ions and solution was incubated at elevated pH
for variable time (t) before adding MgCl2 to 10 mM to
initiate cleavage. Values of t: –, 0 minutes; triangles, 10
minutes; squares, 20 minutes; circles, 30 minutes; dia-
monds, 60 minutes; £ , 120 minutes; þ, 240 minutes.
The arrow indicates the burst magnitude when folding
and cleavage are both done at high pH.
Figure 3. Effect of urea on clea-
vage by TRAP ribozymes. (A) Urea
stimulation of HH8A.8 at pH 7.5.
(B)Inhibitionof
HH8A (no attenuator, diamonds)
and of activated HH8A.8 (triangles)
at pH 7.5. (C) Urea stimulation of
HH8A.8 at pH 8.9. (D) Cleavage
kineticbehavior
HH8A.8 at pH 7.5 as a function of
urea concentration: open diamonds,
0 M; open squares, 1 M; open tri-
angles, 2 M; open circles, 3 M; filled
diamonds, 4 M; filled squares, 5 M;
filled triangles, 6 M; filled circles,
7 M.
non-attenuated
ofattenuated
Folding Paths of Deprotonated Hammerhead Ribozymes
689