copyright by jeffrey philip potratz 2012
TRANSCRIPT
Copyright
by
Jeffrey Philip Potratz
2012
The Dissertation Committee for Jeffrey Philip Potratz certifies that this is the
approved version of the following dissertation:
Local and Global Investigations into DEAD-box Protein Function
Committee:
Rick Russell, Supervisor
Kenneth Johnson
Alan Lambowitz
Scott Stevens
Jessie Zhang
Local and Global Investigations into DEAD-box Protein Function
by
Jeffrey Philip Potratz, B.S.
Dissertation
Presented to the Faculty of the Graduate School of
The University of Texas at Austin
in Partial Fulfillment
of the Requirements
for the Degree of
Doctor of Philosophy
The University of Texas at Austin
May 2012
Dedication
To my wife and parents
Acknowledgements
My time at UT has been positively influenced by many individuals that have
helped make this dissertation possible. I would like to thank my advisor, Rick Russell, for
his exceptional patience and clear, effective teaching and communication style. His
logical thinking and attention to detail set an excellent example for his mentees to follow.
Dr. Alan Lambowitz deserves my gratitude for the collaborative scientific efforts
in which I was included and for the generous offerings of his laboratory supplies. I thank
Dr. Kenneth Johnson for teaching me the foundation of enzyme kinetics. Dr. Scott
Stevens and Dr. Jessie Zhang, along with the aforementioned faculty, are my committee
members for whom I would like to show appreciation for their willingness to serve on my
committee and their scientific input.
My fellow lab mates in the Russell lab are also to be acknowledged. The former
members of Cindy Chen, Hari Bhaskaran, Amanda Chadee, and Yaqi Wan made me feel
welcome in the lab and kindly taught me lab protocols. In addition, Pilar Tijerina was a
great source of knowledge for me when I joined the lab. Current lab members Brian
Cannon, Woongsoon Choi, Inga Jarmoskaite, David Mitchell, and Cynthia Pan are
appreciated for fostering a relaxed and comfortable environment in which to work and for
their willingness to help me mature as a scientist through scientific discussions and
critiques.
Special thanks to my mother and father who supported my decision to venture
down to Texas from Wisconsin and to my wife who encourages me, supports me daily,
and is the reason I have called Texas home for the past five years.
vi
Local and Global Investigations into DEAD-box Protein Function
Jeffrey Philip Potratz, Ph.D.
The University of Texas at Austin, 2012
Supervisor: Rick Russell
Numerous essential cellular processes, such as gene regulation and tRNA
processing, are carried out by structured RNAs. While in vitro most RNAs become
kinetically trapped in non-functional misfolded states that render them inactive on a
biologically-relevant time scale, RNAs folding in vivo do not share this same outcome.
RNAs do indeed misfold in the cell; however, chaperone proteins promote escape from
these non-native states and foster folding to functional conformations. DEAD-box
proteins are ATP-dependent RNA chaperone proteins that function by disrupting
structure, which can facilitate structural conversions. Here, studies with both local and
global focuses are used to uncover mechanistic features of DEAD-box proteins CYT-19
and Mss116p. Both of these proteins are general RNA chaperones as they each have the
ability to facilitate proper folding of multiple structured RNAs.
The first study probes how DEAD-box proteins interact with a simple duplex
substrate. Separating the strands of a duplex is an ATP-dependent process and is central
to structural disruption by DEAD-box proteins. Here, how ATP is utilized during duplex
separation is monitored by comparing ATP hydrolysis rates with strand separation rates.
Results indicate that one ATP molecule is sufficient for complete separation of a 6-11
base pair RNA duplex. Under some conditions, ATP binding in the absence of hydrolysis
is sufficient for duplex separation.
vii
Next, focus is shifted to a more global perspective as the function of Mss116p is
probed in the folding of a cognate group II intron substrate, aI5γ, under near-
physiological conditions. Three catalytically-active constructs of aI5γ are used and
catalysis serves as a proxy for folding. Folding of all constructs is promoted by the
presence of Mss116p and ATP. In vitro and in vivo results indicate that a local unfolding
event is promoted by Mss116p, stimulating formation of the native state. Lastly,
orthogonal methods that probe physical features of RNA are used to provide insight into
the structural intermediates with which Mss116p acts.
viii
Table of Contents
List of Tables ........................................................................................................ xii
List of Figures ...................................................................................................... xiii
List of Abbreviations ........................................................................................... xvi
Chapter 1: RNA folding: A Problem and a Solution .............................................1
1.1 Importance of non-coding RNA ...............................................................1
1.2 The folding problem .................................................................................1
1.3 Does misfolding happen in the cell? .........................................................3
1.4 Chaperones assist proper folding ..............................................................3
1.5 Dead-box proteins are RNA chaperones ...................................................5
1.5.1 Structure ........................................................................................5
1.5.2 Not traditional helicases ................................................................5
1.5.3 Mechanism and functions .............................................................6
1.5.4 Specific and general chaperones ...................................................7
1.5.4.1 Specific chaperones ..........................................................8
1.5.4.2 General chaperones ...........................................................8
1.6 Two model general RNA chaperones .......................................................9
1.6.1 CYT-19 .........................................................................................9
1.6.2 Mss116p ......................................................................................11
1.7 Research plan: local and global methods ...............................................12
Chapter 2: DEAD-box proteins can completely separate an RNA duplex using a
single ATP ....................................................................................................19
2.1 Introduction .............................................................................................19
2.2 Materials and methods ............................................................................21
2.2.1 Materials .....................................................................................21
2.2.2 Determination of RNA and nucleotide concentrations ...............22
2.2.3 RNA strand separation ................................................................22
2.2.4 ATP hydrolysis ...........................................................................23
ix
2.2.5 Analysis of strand separation ......................................................23
2.2.6 Stimulation of strand separation by bound ATP .........................25
2.2.7 Derivation of an equation relating the ATP utilization value to the
ATP concentration ......................................................................26
2.2.8 Analysis of data...........................................................................29
2.2.9 Simulations .................................................................................30
2.2.10 Full equation, including intrinsic dissociation of the duplex ....31
2.3 Results .....................................................................................................32
2.3.1 Enhancement of strand separation by bound ATP without hydrolysis
.....................................................................................................34
2.3.2 Increased ATP requirement for longer or more stable duplexes 36
2.3.3 Similar ATP utilization by other DEAD-box proteins ...............37
2.4 Discussion ...............................................................................................38
2.4.1 Strand separation depends on ATP binding, not hydrolysis .......38
2.4.2 What is the role of ATP hydrolysis? ...........................................40
2.4.3 Implications for physiological activities .....................................41
2.5 Footnotes .................................................................................................42
Chapter 3: RNA catalysis as a probe for chaperone activity of DEAD-box helicases
.......................................................................................................................58
3.1 Catalytic activity as a probe of RNA folding..........................................58
3.1.1 Catalytic activity distinguishes the native state from all other
conformations .............................................................................58
3.1.2 Catalytic activity can be used to study chaperone-assisted folding59
3.2 Self-splicing as a readout for native state formation ..............................60
3.2.1 Interpreting chaperone-promoted changes in observed splicing rate
.....................................................................................................60
3.2.2 Potential complications ...............................................................61
3.3 Substrate cleavage as a readout for native state formation .....................63
3.3.1 Setting up a discontinuous assay: folding and catalysis stages...63
3.3.2 Interpreting results from the catalysis stage ................................65
3.3.3 Using the discontinuous assay to probe chaperone-assisted folding
.....................................................................................................67
x
3.4 Other applications of the discontinuous assay ........................................68
3.4.1 Unfolding native structure ..........................................................69
3.4.2 Integrating results with other methods ........................................69
Chapter 4: ATP-dependent roles of the DEAD-box protein Mss116p in group II
intron splicing in vitro and in vivo ................................................................76
4.1 Introduction .............................................................................................76
4.2 Materials and methods ............................................................................81
4.2.1 Recombinant Plasmids ................................................................81
4.2.2 RNA preparation .........................................................................82
4.2.3 Preparation of Mss116p ..............................................................82
4.2.4 Splicing reactions ........................................................................82
4.2.5 Discontinuous catalytic activity assay for D135 RNA folding ...84
4.2.6 S. cerevisiae Northern hybridizations and immunoblotting .......85
4.3 Results .....................................................................................................86
4.3.1 Splicing of LE and SE constructs and Mss116p acceleration .....86
4.3.2 Mutants that are deficient in RNA-unwinding activity ...............88
4.3.3 Two-stage, discontinuous catalytic activity assay for folding of D135
RNA ............................................................................................89
4.3.4 Acceleration of D135 folding by Mss116p .................................92
4.3.5 The role of ATP in acceleration of D135 folding by Mss116p ..93
4.3.6 Disruption of native D135 ribozyme by Mss116p ......................95
4.3.7 Mss116p-promoted splicing of aI5 in vivo ................................96
4.4 Discussion ...............................................................................................98
4.4.1 Requirement for ATP binding and hydrolysis and effects of exon
length on Mss116p-mediated splicing in vitro............................98
4.4.2 The SAT/AAA mutant is compromised for splicing SE and LE
constructs ..................................................................................100
4.4.3 The roles of ATP in Mss116p-promoted intron folding ...........101
4.4.4 Disruption of the native D135 ribozyme ..................................102
4.4.5 Requirement for ATP binding and hydrolysis by Mss116p in vivo103
4.5 Conclusions and implications ...............................................................104
xi
4.6 Footnote ................................................................................................105
Chapter 5: Rapid structure formation within the aI5γ group II intron ................128
5.1 Introduction ...........................................................................................128
5.2 Materials and Methods ..........................................................................130
5.2.1 RNA preparation .......................................................................130
5.2.2 Footprinting data acquisition and analysis ................................130
5.2.3 SAXS data acquisition and analysis .........................................132
5.3 Results ...................................................................................................133
5.3.1 The unfolded state of aI5γ is readily accessible to DMS modification
...................................................................................................133
5.3.2 Addition of Mg2+
results in rapid formation of structural elements
...................................................................................................134
5.3.3 Native state is significantly more protected than early folding time
point state ..................................................................................134
5.3.4 Addition of Mg2+
gives rapid formation of structure in D135
ribozyme ...................................................................................135
5.3.5 SAXS data reveal compaction of D135 ribozyme ....................136
5.3.6 Time-resolved data indicate early compaction .........................137
5.4 Discussion .............................................................................................137
5.5 Footnote ................................................................................................138
Appendix ..............................................................................................................149
A.1 Preliminary DMS footprinting of D135 ribozyme using capillary
electrophoresis ...................................................................................149
Bibliography ........................................................................................................152
xii
List of Tables
Table 2.1: ATP utilization for CYT-19-mediated separation of the 6-bp P1 duplex
...........................................................................................................51
Table 2.2: Dependence of ATP utilization by CYT-19 on Mg2+
concentration and
duplex length .....................................................................................52
Table 2.3: Dependence of strand separation and ATPase rates on duplex length53
Table 2.4: RNA strand separation and ATP hydrolysis rate for duplexes composed
solely of canonical Watson-Crick base pairs ....................................54
Table 2.5: Temperature dependence of ATP utilization by CYT-19 during strand
separation ..........................................................................................55
Table 2.6: ATP utilization by the DEAD-box proteins Mss116p and Ded1p ...56
Table 2.7: Range of conditions for measurements of ATP utilization ...............57
Table 5.1: Nucleotides modified by DMS. ......................................................148
xiii
List of Figures
Figure 1.1: RNA folding pathway .......................................................................14
Figure 1.2: DEAD-box proteins are involved in all aspects of RNA metabolism15
Figure 1.3: Helicase core of DEAD-box proteins and ancillary domains ...........16
Figure 1.4: Structure of Mss116p ........................................................................17
Figure 1.5: Duplex unwinding by a traditional helicase and a DEAD-box protein18
Figure 2.1: ATP hydrolysis and RNA strand separation by CYT-19..................44
Figure 2.2: Effects of a double-stranded extension on RNA strand separation and
ATPase activity .................................................................................45
Figure 2.3: Strand separation by CYT-19 is independent of ‘chase’ CCCUCUA5
concentration (1–10 µM) ..................................................................46
Figure 2.4: ATP-independent strand separation by CYT-19 ...............................47
Figure 2.5: ATP hydrolyzed by CYT-19 per separation event of the 6-base-pair P1
duplex with low Mg2+
concentration (2 mM) ...................................48
Figure 2.6: Model for duplex separation by DEAD-box proteins .......................49
Figure 2.7: Duplex constructs used for measurements of ATP utilization during
unwinding (top, constructs 1-6) and for control experiments (bottom,
constructs 7-9) ...................................................................................50
Figure 3.1: Group I and group II introns .............................................................71
Figure 3.2: Self-splicing constructs .....................................................................72
Figure 3.3: The discontinuous assay ...................................................................73
Figure 3.4: Examples of catalytic reactions with the D135 and Azoarcus ribozymes
...........................................................................................................74
xiv
Figure 3.5: The discontinuous assay with the D135 ribozyme and the DEAD-box
helicase Mss116p ..............................................................................75
Figure 4.1: RNA constructs ...............................................................................106
Figure 4.2: Self-splicing of the LE and SE constructs ......................................107
Figure 4.3: Self-splicing of the LE construct (panel A) and SE construct (panel B) at
different temperatures .....................................................................108
Figure 4.4: Mss116p-stimulated splicing of the LE and SE constructs ............109
Figure 4.5: Mss116p-stimulated splicing of LE and SE constructs ..................111
Figure 4.6: Stimulation of splicing by the SAT/AAA mutant of Mss116p.......112
Figure 4.7: Mss116p motif I mutants K158A and K158R in splicing reactions of the
LE and SE constructs (panels A and B, respectively) ....................113
Figure 4.8: Two-stage catalytic activity assay to monitor folding of D135 ribozyme
.........................................................................................................114
Figure 4.9: Testing of 30 °C, 100 mM Mg2+
for conditions of stage 2 in the
discontinuous catalytic activity assay .............................................116
Figure 4.10: Prefolded and nonfolded reactions for monitoring D135 RNA folding at
42 °C ...............................................................................................117
Figure 4.11: Mss116p accelerates native folding of the D135 ribozyme ............118
Figure 4.12: Progress curves of D135 ribozyme folding ....................................120
Figure 4.13: Proteolysis of Mss116p after incubation with D135 RNA in the absence
of ATP .............................................................................................121
Figure 4.14: Proteinase K digestion of Mss116p in experiment shown in Figure 4.13
.........................................................................................................123
Figure 4.15: Proteolysis of Mss116p after incubation with D135 RNA .............124
Figure 4.16: Unfolding of native D135 RNA by Mss116p .................................126
xv
Figure 4.17: Northern hybridization and correlated immunoblot comparing the ability
of wild-type and mutant Mss116p to promote splicing of aI5 in vivo127
Figure 5.1: Representative DMS footprinting gel .............................................139
Figure 5.2: Many DMS accessible nucleotides in the unfolded state ................140
Figure 5.3: Nucleotides protected and exposed in the native and early folding time
point states ......................................................................................141
Figure 5.4: DMS footprinting results for domain IV ........................................142
Figure 5.5: DMS footprinting profiles of full-length aI5γ ................................144
Figure 5.6: Kratky plots of static samples reveal different compaction peak heights
.........................................................................................................145
Figure 5.7: Time-resolved SAXS data indicate an early compaction event......146
Figure A.1: DMS footprinting profile of the D135 ribozyme ............................150
xvi
List of Abbreviations
AMP-PNP- adenosine 5´-(,-imido) triphosphate
bp(s)- base pair(s)
EDTA- ethylenedinitrilotetraacetic acid
MOPS- 3-(N-morpholino)propanesulfonic acid
RNP- RNA-protein complex
SF1- helicase superfamily 1
SF2- helicase superfamily 2
DTT- dithiothreitol
LE construct- self-splicing construct of the aI5γ intron with long 5 and 3 exons of 293
and 321 nucleotides, respectively
Mss116p SAT/AAA- mutant of Mss116p with S305A and T307A substitutions in motif
III
mt- mitochondrial
nt(s)- nucleotide(s)
PVDF- polyvinylidene fluoride
SDS- sodium dodecyl sulfate
SE construct- self-splicing construct of the aI5γ intron with short 5 and 3 exons of 28
and 15 nucleotides, respectively
Tris- tris(hydroxymethyl)aminomethane
WT- wild-type
ssRNA- single-stranded RNA
tRNA- transfer RNA
rRNA- ribosomal RNA
1
Chapter 1: RNA folding: A Problem and a Solution
1.1 IMPORTANCE OF NON-CODING RNA
RNA has long been known as the middle man in the process of transforming a
DNA sequence into a functional protein. However, through the years the importance of
non-coding RNA has been recognized and, in addition to serving as a template for protein
production, RNA is involved in many other vital cellular processes (1-4). For example,
RNA composes a large percentage of the ribosome which carries out the cellular
production of protein. Additionally, riboswitches are RNA molecules that change
structure upon binding small ligands and are able to regulate gene expression (5, 6).
The cellular roles attributable to RNA greatly increased with the discovery of
catalytic RNA in the early 1980’s by Thomas Cech and Sidney Altman (7, 8). This
finding revealed that RNA can fulfill enzymatic roles in the cell. These catalytic RNA
molecules include group I and group II introns which self-splice out of precursor
messenger RNA. The processing of tRNA molecules is also carried out by catalytic RNA
found in RNaseP, a complex composed of RNA and protein whose RNA component is a
ribozyme and is responsible for catalytic and enzymatic activity (8).
1.2 THE FOLDING PROBLEM
In order for any non-coding RNA to perform its function, it must first fold to a
correct three-dimensional structure. RNA folding follows a progressive, hierarchical
model in which the correct conformation is achieved through a pathway of intermediate
states that have increasing stability. Because secondary structural elements form
extremely rapidly and can persist in the absence of tertiary interactions, molecules with
secondary structure are considered the first intermediates in the folding pathway. Next,
these pre-formed secondary structures interact to form tertiary contacts that result in a
2
further increase in molecular stability and a correctly-folded molecule (9-12) (Figure
1.1). This correct final structure is commonly referred to as the native state. Interestingly,
one RNA sequence can fold into two structurally-distinct conformations that are both
native, as in the case of riboswitches (1). In addition, it has also been suggested that the
native state is not necessarily one particular conformation with minimal global free
energy but is rather composed of an ensemble of native conformations that have deep
local minima that interconvert rather slowly (13). While a pathway of forming the native
state through interactions by pre-formed secondary structural elements is logical, it is not
without obstacles. RNA contains intrinsic properties which complicate folding to the
native state and result in most RNAs studied in vitro becoming trapped in a misfolded
state(s) (14).
The hierarchical folding of RNA, combined with the limited variety of its primary
sequence, leads to trapped, misfolded RNA conformations. RNA consists of only four
standard bases, which endow it with minimal information content. When an RNA
molecule hundreds of nucleotides in length forms secondary structure, the limited
information content of its bases permits a multitude of different combinations of duplex
elements to form. Some of these duplex elements will form quickly even if they are not
present in the native state of the RNA. These duplexes form under kinetic control and are
not found in the minimal free energy conformation of the native state (15). Clearly, these
non-native duplex elements must be disrupted before the RNA can attain its native
conformation. However, RNA duplexes can persist on the time scale of hours or days
even in isolation, making escape from non-native structures and subsequent folding to the
native state quite slow. In addition, tertiary interactions can form that reinforce the
stability of the non-native duplexes (16). Lastly, incorrect tertiary contacts can form or
incorrect strand orientations can occur that would also necessitate correction and lead to
3
slow folding to the native state (17). Therefore, on a biological time scale, the folding
process is often not completed successfully and leaves the RNA trapped in one or more
misfolded states, which prevents the RNA from functioning (15).
1.3 DOES MISFOLDING HAPPEN IN THE CELL?
The RNA folding problem has been well documented in vitro and has been shown
to impact nearly every RNA studied (14). However, in vitro misfolding does not prove
that RNA is prone to misfolding in its cellular environment. There are several rather
apparent differences between the environments of a test tube and the cell in which RNA
folds physiologically. In the test tube, RNA starts as a generally linear molecule in the
absence of cations and forms secondary and tertiary structure upon the addition of
monovalent cations and divalent cations, respectively. These cations reduce the
electrostatic repulsion of the nucleotide phosphate groups, allowing the RNA strand to
interact with itself. Conversely, within the cellular environment the folding process does
not need to wait for the addition of cations and occurs as the RNA is still being
transcribed. It is influenced by events such as pausing of the RNA polymerase and single
stranded binding proteins interacting with the nascent transcript (18). While these factors
undoubtedly influence the folding pathway of RNA, there is compelling evidence that
RNA misfolding occurs within the cell, as described further below.
1.4 CHAPERONES ASSIST PROPER FOLDING
Certainly, if most cellular RNA was trapped in misfolded, inactive conformations,
the cell could not function. Subsequently, most cellular RNA must have a way to escape
misfolded states if they become trapped. Chaperone proteins provide an escape pathway.
They allow RNA to escape misfolded conformations by promoting the disruption of
structural elements which in turn promotes structural conversions that lead to proper
4
folding (19, 20). However, one could make the argument that within the cellular
environment RNA simply avoids misfolding and chaperone proteins are not strictly
necessary. Refuting this argument is the fact that at least one chaperone protein is
required for practically every process performed by structured RNA, (21-24) indicating
that RNA is prone to misfolding inside the cell just as it is outside the cellular
environment and needs assistance to fold properly.
RNA chaperone proteins function distinctly from specific RNA-binding proteins,
even though both proteins ultimately assist the RNA in reaching its native state (19, 25).
Specific RNA-binding proteins, such as CYT-18 and CBP2, function by promoting
formation of correct structural elements or by stabilizing the native structure once
independently formed (25). After either function, these proteins remain bound to the
native state. Without the continued presence of the protein, the RNA would revert back to
a non-functional structure. Conversely, chaperones interact with the RNA transiently,
exert their influence on the folding pathway, and are then dispensable (25). They
accelerate structural transitions between alternative conformations thereby disrupting
misfolded states and giving the RNA another opportunity to fold correctly (15, 26, 27).
Importantly, chaperone proteins have been shown to facilitate RNA folding both in vitro
and in vivo (18, 25, 28-31).
In addition to escaping misfolded conformations, some RNAs have multiple
conformationally-distinct native states which must be populated temporally. The small
nuclear RNAs found in the spliceosome are examples of RNAs that populate multiple
native, functional states during their existence (32, 33). As in the case with misfolded
conformations, RNA chaperone proteins can promote structural disruptions to facilitate
transitions that lead to interconversion of one native structure into another (20).
5
1.5 DEAD-BOX PROTEINS ARE RNA CHAPERONES
One group of proteins that act as RNA chaperones are DEAD-box proteins. They
belong to helicase superfamily 2 (SF2) and make up its largest class (21, 34). There are
38 DEAD-box proteins in humans while Saccharomyces cerevisiae has 25 (21).
Additionally, these proteins are prevalent in all forms of life and are ubiquitous in all
stages of RNA metabolism (22) (Figure 1.2). Among their diverse biochemical activities
is the ability to elicit conformational changes in RNA through ATP-dependent separation
of RNA duplexes.
1.5.1 Structure
The minimal helicase core of all DEAD-box proteins consists of two domains that
are similar in appearance to RecA and are connected by a flexible linker (21, 34-36). The
core may be flanked on either or both sides with non-conserved ancillary domains
(Figure 1.3). Within the helicase core are eleven conserved motifs, including domain II
(D-E-A-D) for which the group is named. These motifs align themselves on the interface
of the two domains when ligands are present (12, 37-42) (Figure 1.4). DEAD-box
proteins cooperatively bind an ATP molecule or analog and a single stranded RNA
segment (43-47). Motifs I, II, III, V, VI, and Q are positioned near the ATP moiety and
help bind and regulate it while motifs Ia, Ib, GG, IV, QxxR, and V bind the single
stranded RNA.
1.5.2 Not traditional helicases
DEAD-box proteins are not traditional helicases, which bind their substrates via a
single-stranded overhang and use ATP hydrolysis to translocate along one strand of
nucleic acid with a specific polarity, either 3’-5’ or 5’-3’, displacing the other strand
along the way (48, 49) (Figure 1.5). This helicase mechanism is quite useful in separating
6
long, continuous duplex segments such as those found in both host and viral genomes.
However, applying this mechanism to chaperone RNA folding would be
counterproductive, as all the helices formed in a structured RNA would be disrupted,
regardless of whether they were native or non-native. This action would require the RNA
to essentially start folding again from a linear molecule.
Rather, DEAD-box proteins are unique ‘helicases’ as they appear to lack
translocation in their strand separation mechanism. The lack of translocation and
concurrent lack of processivity is made apparent by their inability to unwind duplexes
that are longer than 20-25 base pairs (50, 51). In addition, while single-stranded
overhangs can increase the efficiency of duplex separation by DEAD-box proteins, (52,
53) there is no polarity preference for these overhang regions, again indicating no
processivity (51, 54-56). Furthermore, the inclusion of single-stranded overhangs is not
strictly required for duplex separation (57). The unique duplex separation strategy
utilized by DEAD-box proteins is called local strand separation (58) and allows them to
bind directly to one strand of an RNA duplex, absent of any single-strand overhangs, and
efficiently disrupt short duplex elements (59). The duplex lengths disrupted by DEAD-
box proteins are generally less than ten base pairs long, lengths typically found in
structured non-coding RNAs (20, 60). This mechanism of structural disruption allows the
chaperone to function on local areas of the RNA without necessarily disrupting the entire
structure.
1.5.3 Mechanism and functions
The mechanism of a DEAD-box protein is strongly tied to the use of ATP, as
DEAD-box proteins are ATP-dependent strand separators (21). The protein undergoes
changes in ssRNA binding affinity throughout the ATPase cycle. Initially, binding of
7
ATP and ssRNA is cooperative and induces a large conformational change as the two
domains of the helicase core form a ‘closed’ conformation around the nucleotide and
RNA (43-47). In the ATP and/or ADP-Pi state, the protein has a high affinity for the
ssRNA, while after hydrolysis and release of Pi, the protein has a low affinity for the
ssRNA (44, 61, 62). This cycle of ssRNA affinity changes allows DEAD-box proteins to
tightly bind one strand of a duplex in the presence of ATP and/or ADP-Pi. The interaction
with one of the duplex strands induces a conformation in which a duplex cannot exist and
thus promotes duplex dissociation (38). Once the weak binding state of ssRNA is
promoted through Pi release, the protein releases the single strand and is available to
interact with another duplex element.
DEAD-box proteins use this general property of tight binding to ssRNA to
perform multiple functions. In addition to duplex separation activity, DEAD-box proteins
disrupt RNP complexes (63-69). Disruption of RNP complexes is thought to proceed
similarly to duplex disruption as the protein binds tightly to an RNA strand, which leads
to dissociation of the RNA from the protein in the RNP complex. Yet another function
DEAD-box proteins can perform via tight binding to ssRNA is highlighted in the exon
junction complex. In the EJC, the DEAD-box protein eIF4AIII is bound to ssRNA and
acts as a clamp to stabilize the complex. The protein is stably bound to the RNA because
the EJC holds eIF4AIII in the ‘closed’ conformation, which prevents the release of Pi. In
the absence of Pi release, the protein remains in the state which retains high affinity for
ssRNA and cannot let go (70).
1.5.4 Specific and general chaperones
DEAD-box proteins associate with all the cellular processes of RNA from
transcription to decay, and often the same protein is involved in multiple processes (22)
8
(Figure 1.2). Some DEAD-box proteins find their particular substrates with the help of
non-conserved ancillary domains, which flank the helicase core on the N and/or C-
terminus (71, 72) (Figure 1.3). Others use interactions mediated through the helicase core
to find their substrates (40, 73-76). DEAD-box proteins fall into two categories when
grouped according to their substrates, either specific or general. They can either interact
specifically with a target substrate or are capable of interacting with and chaperoning
multiple substrates.
1.5.4.1 Specific chaperones
The DEAD-box protein DbpA from Escherichia coli and its ortholog YxiN from
Bacillus subtilis are examples of chaperones that are directed to a specific substrate, the
23S rRNA. Both DbpA and YxiN interact with it through ancillary C-terminal domains
(77, 78). More descriptively, the C-terminal domain forms an RNA recognition motif and
binds to a secondary structure feature in the 23S rRNA termed hairpin 92 (79, 80). This
additional RNA binding site outside of the helicase core tethers the chaperone in place
and allows the helicase core to bind nearby duplexes and separate them (79) (Figure 1.5).
While the tethering interaction is specific in this case, a similar tethering mechanism
appears to function with general chaperones.
1.5.4.2 General chaperones
There are significantly more distinct RNA sequences in the cell than chaperone
proteins. Further, each RNA can presumably adopt more than one misfolded structure or
contact, so that the number of misfolded structures that may require assistance from
chaperones is very large. Therefore, it logically follows that some DEAD-box proteins
function on multiple substrates non-specifically (20, 26, 30, 81, 82). These proteins are
referred to as general RNA chaperones. Similar to how specific chaperones have been
9
shown to use ancillary domains to anchor near target substrates, general chaperones also
appear to tether near substrates via ancillary domains (53, 83, 84) (Figure 1.5). Ded1p is
an example of a general RNA chaperone found in S. cerevisiae and thought to function in
translation initiation (85-87). A possible role suggested for Ded1p is removing secondary
structures present in mRNA that prohibit the ribosome from reaching the start codon (88,
89). Disrupting structural elements of mRNAs to allow ribosome binding is a process that
inherently would require the protein to act non-specifically because each RNA sequence
would naturally be unique as it encodes for a different protein.
1.6 TWO MODEL GENERAL RNA CHAPERONES
Two DEAD-box proteins that exhibit general chaperone activity have become
model general RNA chaperones. The first is CYT-19 from Neurospora crassa and the
second is Mss116p from S. cerevisiae. Both function physiologically to assist folding of
introns and appear to use an ancillary domain containing many basic residues, referred to
as the ‘basic tail’ or the basic ‘C-tail’, to bind non-specifically to their substrates (84)
(Figure 1.3).
1.6.1 CYT-19
A significant event in the history of DEAD-box protein research occurred in 2002
when CYT-19 was shown genetically and biochemically to function in the folding of
three cognate mitochondrial group I introns (28). While in vivo RNA chaperone activity
had been demonstrated earlier using the nucleocapsid protein and StpA, (90) this was the
first study in which a DEAD-box protein functioned as a chaperone in vivo to disrupt
misfolded structure and promote proper folding of physiologically relevant RNA. In
addition, the finding that one protein could function on three different group I introns
highlighted the general nature of its activity. The generalness of action was demonstrated
10
further when it was shown that CYT-19 can be expressed in S. cerevisiae and at least
partially suppress all the defects in splicing that result from loss of the cognate DEAD-
box protein Mss116p (91). Therefore, not only is CYT-19 able to promote group I intron
folding, but it can promote non-cognate folding of both group I and group II introns
which have different overall architectures.
CYT-19 contains an approximately 50 amino acid ‘C-tail’ that is highly basic and
is a common feature in various sub-families of DEAD-box proteins (92) (Figure 1.3).
This basic C-tail appears to be responsible for non-specific tethering of CYT-19 to its
numerous substrates. The positively charged tail binds to the negative charge on the
phosphate backbone of nucleic acid and tethers the helicase core near adjacent duplexes
(84). Comparison of wild type CYT-19 and a mutant missing the C-tail reveals that the
strand separation activity of the mutant is not enhanced by the presence of a nucleic acid
extension adjacent to the substrate duplex, whereas the same extension enhances strand
separation activity of the wild type protein. This and additional results suggested a model
in which the C-tail can tether the protein to the extension and position the protein in close
proximity to the substrate duplex (93).
CYT-19 has been shown to resolve a misfolded state found in the group I intron
ribozyme derived from Tetrahymena thermophila, (31) further demonstrating its non-
specific activity as a chaperone. Importantly, the misfolded species of the ribozyme is
well characterized, is almost as compact as the native state, and contains the same
secondary and tertiary interactions characteristic of the native state (17, 94). Observing
how the chaperone interacts with the similar misfolded and native states allows features
of its mechanism to be dissected. For instance, does CYT-19 recognize and act on the
misfolded state only or does it also act on the native state? Because the native state and
misfolded state of the ribozyme are so comparable, it is hard to imagine how CYT-19
11
might interact specifically with the misfolded state. In fact, CYT-19 is able to disrupt the
native structure under destabilizing conditions (26). This disruption of native structure
indicates that CYT-19 is so non-specific that it cannot differentiate between the native
state and a misfolded state. Rather, it ‘blindly’ acts on the RNA, with the misfolded
structure being disrupted with greater efficiency because of its lower stability compared
to the native structure, and does not influence the refolding of the disrupted structure.
Therefore, CYT-19 promotes net formation of the native state from a population of
misfolded molecules by kinetically redistributing the RNA and allowing it further
chances to fold to the more stable native state, where it is not productively acted on (26).
While the study referenced above indicates that general chaperones cannot
discriminate between native and misfolded structural features, the native state may indeed
be protected from disruption. CYT-19 appears unable to separate a duplex element of the
Tetrahymena ribozyme when it is involved in a tertiary contact within the core of the
ribozyme (95). This protection offered by a tertiary contact may represent a general
strategy to protect native structure. As is the case for the Tetrahymena ribozyme, native
states may be more tightly packed than misfolded states which may bias chaperone
proteins to productively function predominantly on misfolded states because their
structural elements are more exposed. The misfolded states may contain structural
features that are exposed enough and unstable enough to be disrupted while the native
states may contain structural elements that are protected by tertiary contacts and/or are
too stable to be disrupted.
1.6.2 Mss116p
Mss116p is a general chaperone and physiologically functions in splicing of all
introns found in the mitochondrial genome of S. cerevisiae (91). There are nine group I
12
introns and four group II introns in the mitochondrial genome that have varying overall
architectures. The ability of one protein to help fold RNAs which have varying
architectures displays the non-specific nature of Mss116p activity. Like CYT-19,
Mss116p contains a basic C-tail which may be involved in tethering the chaperone to its
substrates (84).
The crystal structure of Mss116p was solved in 2009 and is very reminiscent of
other solved structures of DEAD-box proteins complexed with an ATP analogue and
ssRNA, with one notable exception (37) (Figure 1.4). The ssRNA is bent twice in this
structure compared to only one bend seen in the previous structures (12, 37-42). This
‘extra’ bend, which results in the RNA being ‘crimped’, is caused by the positioning of
the C-terminal extension.
1.7 RESEARCH PLAN: LOCAL AND GLOBAL METHODS
The overarching goal of my research is to uncover key facets to how DEAD-box
proteins function. The studies used to probe the mechanism start with a narrow focus and
then broaden out to gain a more comprehensive perspective. In the first study, the action
of chaperones is probed through the use of the most basic substrate, a duplex element (see
Chapter 2). The study quantitatively dissects how DEAD-box proteins use ATP to disrupt
this simple secondary structural element. Several general chaperones, including CYT-19
and Mss116p, from different organisms are used on a target substrate derived from the
Tetrahymena ribozyme. In this way, the results are unbiased by any physiological context
and provide a simple look at how many ATPs are required to separate duplex elements.
The second project uses catalytically active RNA (see Chapter 3) and takes a
more global look at how proper RNA folding is promoted through chaperone activity.
The study dissects the role of Mss116p in the folding pathway of the cognate group II
13
intron aI5γ (see Chapter 4). In this way, physiologically relevant chaperone activity is
observed. Lastly, methods used to probe for structural features of RNA, DMS
footprinting and small angle x-ray scattering (SAXS), are used to identify physical
characteristics of intermediate conformations found along the aI5γ folding pathway (see
Chapter 5).
14
Figure 1.1: RNA folding pathway
RNA folding follows a progressive pathway filled with intermediates of increasing
stability. Secondary structure forms first kinetically, followed by modular assembly into
tertiary contacts and a final folded conformation. This figure originally published in (12)
by Landes Bioscience.
15
Figure 1.2: DEAD-box proteins are involved in all aspects of RNA metabolism
Yeast and human ortholog DEAD-box proteins (green circles) are shown connected to
the RNA processes (white boxes) in which they participate. Other SF2 family members
are also depicted (red, light blue, dark blue, and yellow colored circles). This figure
originally published in (22) by Elsevier.
16
Figure 1.3: Helicase core of DEAD-box proteins and ancillary domains
The minimal helicase core shared by all DEAD-box proteins is labeled as ‘DEAD-box’
and shown in light blue. The ‘C-tail’ for CYT-19, Mss116p, and Ded1p is labeled BT for
basic tail and is dark blue. Additional ancillary domains are depicted with various colors
and labeled as follows: RRM, RNA-recognition motif; Di, dimerization domain; RBD,
non-conserved RNA-binding domain; αHR, α-helical region; SMN, survival of motor
neurons protein; SF-1, steroidogenic factor 1 protein; RS arginine-serine-rich domain;
RGG, arginine-glycine-glycine repeats; IQ, calmodulin-binding domain. This figure
originally published in (82). Reproduced with permission by The Royal Society of
Chemistry.
17
Figure 1.4: Structure of Mss116p
(A) Cylindrical representation of the Mss116p sequence. Regions found in the crystal
structure are shown as gray cylinder while regions not found in the crystal structure are
black lines. The abbreviations below the cylinder are as follows: NTE, N-terminal
extension; Domain 1 and 2, the helicase core; CTE, C-terminal extension; BT, basic tail
or ‘C-tail.’ The conserved motifs are highlighted with different colors and labeled above
the cylinder. (B) The crystal structure (37) with domain 1 in blue, domain 2 in green, C-
terminal extension in gray, ssRNA in magenta, and AMP-PNP in orange. C) The crystal
structure (37) in the same orientation as panel B but with the individual motifs now
colored as in panel A. Note the ‘crimp’ in the ssRNA as it is bent twice. This figure
originally published in (96) by Wiley.
18
Figure 1.5: Duplex unwinding by a traditional helicase and a DEAD-box protein
(A) A traditional helicase binds to a single-stranded extension (red) and uses ATP
hydrolysis to translocate directionally through a duplex region (black) and separate the
strands. This model helicase translocates 3’-5’. (B) A DEAD-box ‘helicase’ binds to the
RNA duplex (black) directly with its helicase core (large orange circle) and uses ATP to
separate it with minimal if any translocation. This illustration shows a DEAD-box protein
which possesses an ancillary domain (smaller orange circle) that can bind a region
adjacent to the substrate duplex, depicted here as an adjacent duplex (red), and tether the
helicase core near substrate duplex regions The ancillary domain could bind specifically
or non-specifically to the structure adjacent to the substrate duplex and could be a ‘C-tail’
domain, an RRM domain, or other motifs. This figure originally published in (12) by
Landes Bioscience.
19
The text below and following figures of chapter 2 were originally published by National
Academy of Sciences. Copyright © by the National Academy of Sciences 2008:
Chen, Y*., Potratz, J.P*., Tijerina, P. Del Campo, M., Lambowitz, A.M., Russell, R .
(2008) DEAD-box proteins can completely separate an RNA duplex using a single ATP.
Proc Natl Acad Sci U S A 105(51):20203-20208.
*Chen, Y. and Potratz, J.P. are co-first authors.
Mark Del Campo from the Alan M. Lambowitz lab purified proteins.
Yingfeng Chen and Pilar Tijerina from the Rick Russell lab helped collect data.
Rick Russell, Yingfeng Chen, and Alan Lambowitz helped analyze the data and wrote the
paper.
Chapter 2: DEAD-box proteins can completely separate an RNA
duplex using a single ATP
2.1 INTRODUCTION
To gain insight into the mechanism of CYT-19 activity, we took advantage of the
observation that it can use its non-specific chaperone activity to efficiently separate the
six-base-pair helix termed P1, formed between group I introns and their 5´-exon-intron
junction, and that this unwinding efficiency (kcat/KM) is enhanced by two orders of
magnitude when the duplex is covalently linked to the ribozyme compared to the same
duplex in solution (31). Then, using simple constructs based on group I intron structure,
we found that the activity was also enhanced by simple extensions to the helix.
Interestingly, a single-stranded extension gave a smaller enhancement than a double-
stranded flanking region, and both gave smaller enhancements than the highly structured
intact group I intron.
Whereas conventional DNA and RNA helicases commonly require a 5- or 3-
single-stranded region, which serves as a starting point for translocation into and through
the duplex, results above suggested that the increased activity arose instead from an
20
additional and distinct interaction of CYT-19 with the RNA. Further, the enhancement
under subsaturating conditions (kcat/KM) suggested that this interaction is maintained in
the transition state for strand separation. Additional work showed that the enhancement is
nearly eliminated by deletion of 49 amino acids from the highly basic C-terminus of
CYT-19, whereas strand separation of a duplex that lacks an extension is essentially
unaffected, most simply suggesting that this additional interaction is mediated by the C-
terminal region (93).
Together, these findings led to a model in which interactions with adjacent RNA
structure tether DEAD-box proteins in proximity to exposed helices or perhaps other
elements of RNA structure, where binding of the core domain and ATP-dependent
conformational changes give strand separation (31, 35, 53). Although early studies
demonstrating that DEAD-box proteins can readily separate duplexes of approximately
one helical turn or less but are essentially inactive for duplexes of two or more turns (56,
97) indicated a lack of processivity, the tethering model suggests a more radical
difference in mechanism from processive helicases. This is because continuous formation
of a tethering interaction during duplex unwinding would most simply suggest the
absence of any translocation during the unwinding process.
Strong independent support for essential features of this model has come from
studies in the Jankowsky lab using an elegant set of model duplex substrates. First, they
demonstrated conclusively that a flanking sequence can enhance activity for DEAD-box
proteins without serving as a starting point for translocation by showing that a single-
stranded segment can still give activation if it is not linked to the target duplex but is
instead bound through biotin-mediated interactions with the protein streptavidin (53).
Second, they showed that even model duplexes with an RNA segment flanked on both
sides by DNA can be efficiently separated by DEAD-box proteins, whereas the same
21
proteins are not active on fully DNA substrates, indicating that strand separation can be
initiated internally, without translocation (98). The main features of the model are also
indicated for the E. coli DEAD-box protein DbpA by studies from Uhlenbeck and
colleagues, with the important difference that this protein uses an ancillary domain to
recognize a particular structure within the large subunit ribosomal RNA rather than
interacting with structured RNA more generally (77-80).
In the current work, we have tested and extended this model by measuring the
number of ATP molecules utilized by CYT-19 and other DEAD-box proteins as they
separate RNA duplexes. In the most extreme form of the model, with strand separation
accomplished in the presence of a continuously-formed tethering interaction, it would be
possible that the complete reaction would be accomplished in a single cycle of ATP-
dependent conformational changes and would therefore give hydrolysis of only one ATP.
Indeed, we obtain this result for duplexes of 6–11 bp, characteristic of helices present in
structured RNAs. Further, under some conditions, a significant fraction of strand-
separation events occur in the absence of any ATP hydrolysis. Nevertheless, these events
are dependent on ATP, indicating that bound ATP favors a protein conformation that
promotes local strand separation even prior to ATP hydrolysis.
2.2 MATERIALS AND METHODS
2.2.1 Materials
Oligonucleotides were purchased from Dharmacon. CCCUCUA5 was 5´-end-
labeled with [-32P]ATP by using T4 polynucleotide kinase and gel-purified (99). CYT-
19, Mss116p, and Ded1p were expressed and purified as described (55, 83). AMP-PNP
was treated to remove any contaminating ATP as described (60).
22
2.2.2 Determination of RNA and nucleotide concentrations
The ATP utilization values are linearly dependent on both the duplex and ATP
concentrations, so it is important that the concentrations be measured carefully. We
determined these concentrations spectrophotometrically using extinction coefficients
(260 nm) calculated from base composition: ATP and AMP-PNP, 1.54 104 M
–1 cm
–1;
CCCUCUA5, 1.3 105 M
–1 cm
–1; CCCUCCA5, 1.2 10
5 M
–1 cm
–1; RNA/DNA hybrid
strand, 3.64 105 M
–1 cm
–1 (for hybrid strands with uridine extensions, values were 3.64
105 M
–1 cm
–1 + 0.99 10
4 M
–1 cm
–1 per uridine). Measurements in the presence of 7 M
urea, which disrupts the secondary structure of the P2 extension (31), gave at most small
increases in absorbance (<5%), so concentrations were calculated directly from
measurements made in water. Each concentration was measured at least twice, and
measured values varied by less than 5%.
2.2.3 RNA strand separation
Unless otherwise indicated, reaction conditions were 25 °C, 50 mM Na-MOPS,
pH 7.0, 10 mM MgCl2, 50 µM ATP-Mg2+
, 50 mM KCl, and 5% glycerol. Reactions were
initiated by adding pre-formed duplex (final concentrations of 0.5 µM RNA/DNA hybrid
oligonucleotide, 0.2 µM CCCUCUA5, and trace 32P-labeled CCCUCUA5) to CYT-19 (2
µM), followed by addition of 1–5 µM unlabeled CCCUCUA5 to give a final duplex
concentration of 0.5 µM. Control reactions showed that varying the concentration of
unlabeled CCCUCUA5 across and beyond this range did not affect the rate constant for
strand separation (Figure 2.3), and thus 5 µM CCCUCUA5 was used typically to increase
the signal for strand separation. At various times, aliquots were quenched by adding 70
mM MgCl2 and 1 mg/ml Proteinase K, and then loaded on a 20% nondenaturing
polyacrylamide gel run at 5 ºC. We confirmed that the quench solution was effective, as
CYT-19 did not promote strand separation under the quench conditions (data not shown).
23
Gels were dried, visualized with a phosphorimager (GE Healthcare), and quantitated with
ImageQuant 5.2 (GE Healthcare). Time courses were fit by a single exponential equation
(Kaleidagraph, Synergy Software). Rate constants were converted to steady-state rates by
multiplying by the duplex concentration, 0.5 µM. A control experiment in which
unlabeled chase CCUCUA5 was added before labeled CCCUCUA5, such that CYT-19-
mediated strand separation of the unlabeled P1 duplex was monitored by detecting the
formation of labeled duplex, gave the same rate constant within error (data not shown).
Thus, the presence of the 5’-phosphoryl group on the radiolabeled CCCUCUA5 does not
affect the rate of CYT-19-mediated strand separation.
2.2.4 ATP hydrolysis
Conditions were as above except that reactions included trace [-32P]ATP instead
of 32P-labeled CCCUCUA5. Unlabeled CCCUCUA5 was 1 µM (see above). Aliquots
were quenched with 100 mM EDTA, applied to a polyethyleneimine (PEI) cellulose thin-
layer chromatography plate, developed in 1 M formic acid, 0.5 M LiCl, and quantitated
as above.
2.2.5 Analysis of strand separation
To determine the number of ATP molecules hydrolyzed per stand separation
event, we compared steady-state rates of ATP hydrolysis and strand separation. We
measured the steady-state ATPase rate directly, but for strand separation we measured the
displacement of radiolabeled CCCUCUA5 in a pulse-chase experiment. Thus, it was
necessary to convert the rate constant obtained from a fit by an exponential equation to a
steady-state rate. This conversion was achieved by multiplying the rate constant by the
duplex concentration, typically 0.5 µM (the concentration of the limiting species, the
RNA/DNA hybrid strand). This simple conversion is possible because, although we
24
monitored dissociation only of the labeled CCCUCUA5, the unlabeled CCCUCUA5
present in the reaction is expected to rapidly re-form the duplex, producing a steady state
in which the duplex is largely formed (100-102). Indeed, at the concentrations of
oligonucleotides used, we confirmed that the duplex forms much faster than it is
separated by CYT-19 (data not shown), indicating that the total duplex concentration is
maintained at a steady-state level close to the concentration of the hybrid strand (0.5
µM). This unlabeled duplex is then subject to multiple rounds of unwinding by CYT-19
(31), even though only the first round – separation of the labeled duplex – is monitored
experimentally. The ‘dilution’ of the labeled duplex by continuous formation of
unlabeled duplex is not expected to affect the measured rate constant for strand
separation because CYT-19 is present at subsaturating concentration for binding to the
duplex. The observation that the ATPase rate remains constant at times long after the first
round of strand separation is completed (Figure 2.1B and C) provides strong support for
the interpretation that the duplex concentration remains relatively constant, as control
experiments indicate that the separated strands would give much lower ATPase rates
(Figure 2.1B).
The fraction of labeled CCCUCUA5 that migrated as the duplex at the start of
reactions was typically 0.6–0.7 both in the presence and absence of CYT-19, as shown in
Figure 2.1C. This incomplete retention appears to be caused principally by dissociation of
a modest fraction of the duplex in the gel rather than incomplete duplex formation in the
reaction, as increases in the concentration of the complementary strand up to 4-fold (2
µM) produced little or no increase in the fraction of labeled CCCUCUA5 retained in the
duplex (data not shown). This small loss of duplex is not expected to affect the measured
values of the rate constants because it is not expected to affect the shape of the strand
separation curves, but only to decrease the amplitudes.
25
2.2.6 Stimulation of strand separation by bound ATP
To explore whether bound ATP stimulates unwinding even when it is not
hydrolyzed, we fit the measured values of ATP utilization during unwinding of the 6-bp
duplex (25 °C, 2 mM Mg2+
) by models that included or excluded a pathway that does not
result in ATP hydrolysis, yet gives more rapid strand separation than the pathway
mediated by nucleotide-free CYT-19. The equations relating ATP utilization value to
ATP concentration were derived from Scheme 1, in which S1 and S2 represent the
separated strands, and all separation events are shown as irreversible because of the
presence in the experiment of an excess of one of the unlabeled strands. The rate constant
kATPADP represents the second-order process of CYT-19•ATP binding to the duplex and
separating it to single strands in a process that results in hydrolysis of the ATP. Similarly,
kATP represents the second order rate constant for CYT-19•ATP binding to the duplex and
separating the strands, but in a process that does not result in ATP hydrolysis. The rate
constant kno nuc reflects the second order process of CYT-19 binding to the duplex and
26
separating the strands without bound ATP, and kintrinsic is the first-order rate constant for
strand separation in the absence of CYT-19.
Scheme 1 also includes the following assumptions. 1) ATP binding by CYT-19 is
assumed to be in rapid equilibrium, as suggested by the relatively high KM value
determined previously in steady-state ATPase assays (28). 2) CYT-19 is assumed to be
subsaturating for duplex binding, as indicated by an increasing rate with increasing CYT-
19 concentration across the range used in this and previous studies (10 nM – 2 µM) (31,
83). 3) Release of the products, both the single-stranded products and ADP and Pi are not
shown explicitly; the second-order rate constants are defined as the rate constant up to
and including the rate-limiting step(s). This treatment does not assume that product
release steps are faster than hydrolysis. If one or both of the product release steps are
rate-limiting and hydrolysis is readily reversible, the overall rate constant shown reflects
the equilibrium for ATP hydrolysis and the rate constant for the rate-limiting product
release step.
2.2.7 Derivation of an equation relating the ATP utilization value to the ATP
concentration
The ATP utilization value relates the rate of duplex-dependent ATP hydrolysis to
the rate of CYT-19-dependent strand separation. To derive an equation that relates the
ATP utilization value to the ATP concentration, we assumed for simplicity that all
duplex-dependent ATP hydrolysis events are coupled to successful strand separation
under the conditions of Figure 2.5 (2 mM Mg2+
, 6-bp duplex), which give less than one
ATP hydrolyzed per duplex separated. This assumption allows the ATP utilization value
to be expressed as the rate of ATP-hydrolysis-dependent strand separation events relative
to the total strand separation rate produced by all pathways, or in other words as the
fraction of strand separation events that are coupled to ATP hydrolysis (eq. 1). It is
27
possible that some ATP hydrolysis is ‘wasted’, as is suggested to occur with longer
duplexes and at higher Mg2+
concentration (see Table 2.2). If so, the fraction of
unwinding events that proceed through the ATP-hydrolysis-dependent pathway would be
overestimated, while the fraction that rely on bound ATP without its hydrolysis would be
underestimated. (The total fraction of ATP-dependent events is measured directly from
the increase in unwinding rate in the presence of ATP). Thus, this analysis provides a
lower limit on the acceleration of strand separation provided by ATP when it is not
hydrolyzed.
Equation (1)
Under the conditions of our experiments, the non-enzymatic dissociation of the
duplex can be omitted from the equation (kintrinsic), as this background rate was subtracted
from the raw data (see eq. 5 below for the form of the final equation that includes non-
enzymatic duplex dissociation). Further, this process is substantially slower than the
CYT-19-promoted processes and does not substantially affect the fitted parameters. Eq. 1
can therefore be simplified by omitting this term and by dividing the numerator and
denominator by [Duplex] to give eq. 2.
Equation (2)
The concentration of ATP-bound CYT-19 can be related to the ATP
concentration by dividing the numerator and denominator by [CYT-19] and substituting
ATP utilization=[Duplex][CYT -19 • ATP]kATPADP
[Duplex][CYT -19 • ATP]kATPADP [Duplex][CYT -19]kno nuc [Duplex][CYT -19 • ATP]kATP [Duplex]kintrinsic
[CYT -19 • ATP]kATPADP
[CYT -19 • ATP]kATPADP [CYT -19]kno nuc [CYT -19 • ATP]kATP
28
terms using the expression for equilibrium ATP binding to give the following (KD is the
equilibrium constant for ATP binding):
Equation (3)
Eq. 3 can be rearranged by combining terms and multiplying the numerator and
denominator by KD/(kATPADP + kATP) to give a form that is analogous to the Michaelis-
Menten equation (eq. 4), describing a hyperbolic dependence of the ATP utilization value
on ATP concentration.
Equation (4)
The prominent features revealed by eq. 4 are: 1) the plateau value of the ATP
utilization value with increasing ATP concentration reflects the ratio of rate constants for
ATP hydrolysis-dependent and hydrolysis-independent pathways involving ATP-bound
CYT-19, and 2) the ATP concentration that gives the half-maximal ATP utilization value
(the KM-like term) is composed of the KD for ATP binding and a factor that reflects the
increase in rate afforded by bound ATP. This results in the half-maximal value being
lower than the KD because the increased activity of CYT-19 upon ATP binding causes
most of the separation events to be mediated by ATP-bound CYT-19 even at ATP
concentrations that give ATP binding to only a relatively small fraction of CYT-19.
[ATP]kATPADP
KD[ATP]kATPADP
KD kno nuc
[ATP]kATP
KD
[ATP]kATPADP
kATPADP kATP
KDkno nuc
kATPADP kATP
[ATP]
29
2.2.8 Analysis of data
The ATP utilization data in Figure 2.5 are shown with fits by the model of eq. 4
and two related models. The solid curve shows a fit with eq. 4, which includes the
possibility of a rate acceleration by bound ATP without its hydrolysis (kATP). The fit
gives a maximal value of 0.5 that is well defined by the data, indicating that ATP is
hydrolyzed in only half of the unwinding events mediated by CYT-19•ATP; i.e. kATP is
approximately equal to kATPADP. Further, if the value of KD for ATP binding is assumed
to be equal to the KM value measured previously in steady-state ATPase assays under
similar conditions (200 µM, ref. (28), it is possible to calculate values of kATP and
kATPADP using eq. 4. The best-fit of the data in Figure 2.5 gives a half-maximal value of
5 µM (the KM-like term), which from eq. 4 is equal to KD (kno nuc/(kATPADP + kATP)).
Using the measured value of kno nuc of 2 105 M
–1 min
–1 (converted from the steady-state
rate of 0.2 µM/min (see section 2.3.1) with 0.5 µM duplex and 2 µM CYT-19) and the
relationship that kATP = kATPADP, the values of kATP and kATPADP are determined to be 4
106 M
–1 min
–1 (see Scheme 2 below). Thus, bound ATP is calculated to give a 40-fold
enhancement overall, 20-fold via each of two pathways that include or exclude its
hydrolysis. It should be noted that this overall enhancement represents a lower limit; we
directly observe the strand separation rate to increase linearly to at least 150 µM ATP,
indicating that KM is at least ~200 µM, but we are unable to measure strand separation
above 150 µM ATP because it becomes too fast to measure with hand pipetting.
Nevertheless, the previous determination of KM for ATP of 200 µM in steady-state
ATPase assays under similar conditions (28) and our measurement of 200 µM for KD of
AMP-PNP by inhibition (Figure 2.4) suggests that the KD for ATP under our conditions
is likely to be close to 200 µM.
30
In contrast to the good fit obtained with the model above, the data are not well-
described by a model that excludes the ATP-dependent, hydrolysis-independent pathway
by omitting kATP from the equation, giving clear systematic deviation from the long-
dashed curve in Figure 2.5. Excluding this pathway has the effect of forcing the ATP
utilization value to plateau at unity with increasing ATP concentration, with the physical
implication that any strand-separation process mediated by ATP-bound CYT-19 gives
hydrolysis of the bound ATP. The data are also not well-described by a model in which
strand separation by ATP-bound CYT-19 is permitted without ATP hydrolysis, but bound
ATP does not accelerate strand separation unless it is hydrolyzed (short dashed curve in
Figure 2.5). In this model, the value of kATP is required to be equal to the value of kno nuc.
To be compatible with our observed data, the ATP utilization value is required to plateau
at a value of 0.9 or larger (the plateau value is kATPADP/(kATPADP + kno nuc)), as we
observed directly that the ATP-dependent unwinding rate constant kATPADP is at least 10-
to 20-fold larger than the rate constant for ATP-independent unwinding (kno nuc).
2.2.9 Simulations
To confirm the utility of eq. 4, we performed simulations using Scheme 2, which
31
is modified from Scheme 1 by including values of the rate constants obtained from fitting
eq. 4 to the data on ATP utilization as described above (Figure 2.5). Our prediction was
that, if eq. 4 accurately described the conversion of the fitting parameters (KM- and kcat-
like parameters) to the values of the relevant rate constants (kno nuc, kATPADP, and kATP), a
simulation using these values for the rate constants should return simulated ATP
utilization values that are consistent with the data and the fitted curve.
Simulations were performed using Kinetic Explorer (Kintek) with the rate
constants shown above and the following initial concentrations: CYT-19, 2 µM; S1, 0.5
µM; S2, 1 µM. The duplex was allowed to form with a rate constant of 109 M
–1 min
–1
(100). At several ATP concentrations, steady-state rates of ATPase activity and strand
separation were recorded. From these rates, ATP utilization values were calculated and
plotted on the data and fits from Figure 2.5. The ATP utilization values calculated from
the simulations were in good agreement with the curve fit to the data (data not shown),
indicating that eq. 4 is valid. Its use returned rate constants that give the observed
behavior.
2.2.10 Full equation, including intrinsic dissociation of the duplex
Eq. 5, shown below, is analogous to eq. 4 but includes non-enzymatic duplex
separation (kintrinsic). Terms including the first-order rate constant kintrinsic are divided by
the total CYT-19 concentration, [CYT-19]t, instead of the equivalent form of multiplying
each of the second-order rate constants by [CYT-19]t, to allow the form of eq. 5 to more
closely resemble the simpler eq. 4. (Eq. 5 was not used in any analysis herein but is
included for completeness. It could be necessary to include this term under conditions
that give little or no stimulation of strand separation by CYT-19 in the absence of
nucleotide, such as Mg2+
concentrations of 10 mM or higher.)
32
Equation (5)
2.3 RESULTS
In designing a duplex substrate for these studies, we took advantage of our
previous results that CYT-19 can efficiently separate the P1 duplex of the Tetrahymena
group I ribozyme, leading to dissociation of the oligonucleotide substrate, and the
separation is much more efficient when P1 is covalently linked to the intron or to another
RNA helix (31). This increased activity allows robust experimental signals for strand
separation and duplex-dependent ATPase activity (ref. (31) and results herein). However,
attachment of the intron or even a second RNA duplex would inextricably complicate the
analysis because the additional RNA would be expected to interact with CYT-19 and
stimulate its ATPase activity, and it would not be possible to determine how much of the
total ATPase activity arose from separation of the P1 duplex.
We therefore generated a construct in which the P1 duplex is formed from an
RNA oligonucleotide (CCCUCUA5) and a hybrid RNA/DNA oligonucleotide, resulting
in P1 being flanked by a DNA duplex (Figure 2.1A). This dsDNA extension gave the
same activation of strand separation activity as an equivalent RNA extension and, as
expected from earlier work (106, 107), an oligonucleotide containing only the DNA
[ATP]kATPADP
kATPADP kATP kintrinsic
[CYT -19]t
KD
kno nuckintrinsic
[CYT -19]t
kATPADP kATP kintrinsic
[CYT -19]t
[ATP]
33
portion did not stimulate ATPase activity, implying that it is not actively unwound by the
helicase core (Figure 2.2).
Using this substrate, we measured ATPase activity by CYT-19 under defined
conditions (10 mM Mg2+
; Figure 2.1B). Nearly all of the ATPase activity arose from
interactions with the duplex under these conditions, as the rate was ~10-fold lower in the
presence of either strand alone (Figure 2.1B). Because the duplex separation reactions
included a small excess of CCCUCUA5 and were performed under subsaturating
conditions, we subtracted from the total the rate in the presence of 1 µM CCCUCUA5,
which approximates its free concentration in reactions including the duplex. After
subtracting this background, we obtained a duplex-dependent rate of 0.74 ± 0.05 µM/min
(Figure 2.1B and Table 2.1).
We then measured P1 duplex separation under the same conditions using a pulse-
chase gel mobility shift assay (Figure 2.1C and Figure 2.3). After subtracting the CYT-
19-independent separation, we obtained a rate constant of 1.4 ± 0.1 min–1
from a fit by a
first-order rate equation (Figure 2.1C and additional replicates not shown). To compare
the rate of strand separation with the steady-state rate of ATPase activity measured
above, we converted the rate constant to a steady-state rate by multiplying it by the
duplex concentration (see Section 2.2.5). This conversion gave a steady-state rate of 0.69
± 0.05 µM/min (Table 2.1). Strikingly, the rates of ATPase activity and strand separation
are the same within error, giving a ratio of 1.1 ± 0.1 ATP molecules hydrolyzed per
duplex separated. This ratio, or ATP utilization value, was the same within error across
the range of experimentally accessible ATP concentrations (5–150 µM, data not shown;
KM = 200 µM, ref. (28) and across the more limited range of accessible CYT-19
concentrations (0.5–2 µM, data not shown, see Section 2.5)*. Thus, under these
conditions (10 mM Mg2+
, 25 °C), a single cycle of ATP-dependent conformational
34
changes apparently gives complete strand separation of this 6-bp duplex. If any of the
base pairs are not disrupted directly by the enzyme during this cycle, they must dissociate
spontaneously.
2.3.1 Enhancement of strand separation by bound ATP without hydrolysis
To explore whether hydrolysis of ATP is uniformly required for strand separation
by DEAD-box proteins, we decreased the Mg2+
concentration, shown previously to
increase the strand separation activity of CYT-19 (31, 83). With 5 mM Mg2+
, the ATP
utilization value remained approximately one (Table 2.1). With lower Mg2+
concentration
(2 mM), however, it decreased to 0.45, implying that about half of the strand separation
events proceeded without ATP hydrolysis. ATP-independent separation of this duplex
has been shown for the related DEAD-box protein Mss116p (60), and we confirmed that,
with 2 mM Mg2+
but not 10 mM Mg2+
, CYT-19 gives detectable strand separation in the
absence of nucleotide (with a rate of 0.2 µM/min; Figure 2.4). This activity may be
achieved by ‘strand capture’, analogous to the RNA chaperone activity of proteins that
are not ATPases (60, 108, 109).
We therefore considered a model in which the ATP utilization value of 0.45
reflected a balance between strand separation mediated by ATP-bound CYT-19, which
would result in ATP hydrolysis, and nucleotide-free CYT-19, which would of course not
give ATP hydrolysis. However, it was not clear that the unwinding rate in the absence of
ATP (0.2 µM/min; above) was large enough to support this model. In order for half of the
separation events in the presence of ATP to be mediated by nucleotide-free CYT-19, the
ATP-independent rate would have to be at least half of the total CYT-19-dependent rate
of 0.97 ± 0.05 µM/min (Table 2.1). Although the difference between the expected and
35
observed values is not large, it raised the possibility that bound ATP stimulates the rate of
strand separation by CYT-19 even when it is not hydrolyzed.
We tested this possibility by systematically varying the ATP concentration under
the same 2 mM Mg2+
conditions (Figure 2.5). If the low ATP utilization value resulted
from activity of nucleotide-free CYT-19, it would increase with increasing ATP
concentration, reflecting the increased fraction of CYT-19 bound to ATP, and would
approach unity with saturating ATP (see Section 2.2.6). Instead, the ATP utilization
reached a plateau value of 0.5-0.6, giving no further increase with ATP concentration,
even as the rate of strand separation increased to a value ≥ 40-fold larger than without
ATP. This behavior indicates that ATP is hydrolyzed only in approximately half of the
strand separation events, even when it is bound, and that bound ATP strongly accelerates
strand separation even when it is not hydrolyzed (≥ 20-fold, see Section 2.5)‡. These
results strongly suggest that ATP elicits or stabilizes a protein conformation that induces
or captures local strand separation events (see below and Figure 2.6).
Interestingly, we found that the enhancement from bound ATP is not mimicked
by the non-hydrolyzable analog AMP-PNP, which gave no acceleration of strand
separation beyond the basal level in the absence of nucleotide (Figure 2.4A). The lack of
activity is not due to a failure to bind CYT-19, because AMP-PNP inhibited ATP
stimulation with a KI of 200 µM (data not shown), 5-fold lower than the concentration
used to test for stimulation. Thus, these results suggest that AMP-PNP binding does not
elicit the same conformation as ATP, a conclusion that is strongly supported by recent
work using a series of nucleotide analogs (110).
36
2.3.2 Increased ATP requirement for longer or more stable duplexes
Regardless of the mechanism of strand separation, it would be expected that more
ATP hydrolysis would be necessary for longer duplexes. An increased ATP requirement
is further suggested for DEAD-box proteins by previous observations that longer
duplexes are separated with greatly reduced rates. However, to our knowledge, the
dependence of duplex length on the ATPase activity of DEAD-box proteins has not been
systematically investigated.
Therefore, we extended the 5´-end of the hybrid strand with uridines to form P1
duplexes of seven, nine, or eleven base pairs (see Figure 2.1A and Figure 2.7). As
expected, the ATP requirement increased with duplex length (Table 2.2). This increase
arose from a decreased strand separation rate, while the ATPase rate remained essentially
constant (Table 2.3). The insensitivity of the ATPase rate suggests a simple model in
which CYT-19 manipulates both the longer and shorter duplexes by using a single cycle
of ATP-dependent conformational changes to induce local strand separation, but for the
longer duplexes a fraction of these events do not lead to complete strand separation,
allowing re-zipping of the duplex after the core domain of CYT-19 dissociates (see
Figure 2.6 and refs (52, 61, 97). These experiments also established 11 base pairs as a
lower limit on the unwinding that can be accomplished using a single ATP, as this duplex
gave an ATP utilization value of 1.2 ± 0.1 (2 mM Mg2+
).
We next increased the duplex stability, without increasing the length, by changing
the natural G•U wobble pair within the P1 duplex to a G-C pair (Figure 2.7). The G•U
pair has been shown previously to destabilize the P1 helix, relative to the G-C, in part by
disrupting base stacking (111, 112). We tested the effects of this base pair in the context
of the 6-bp and 11-bp duplexes. Under standard conditions (10 mM Mg2+
), the ATP
utilization value for the 6 bp duplex increased 8-fold (from 1.1 ± 0.1 to 9 ± 3, Table 2.4),
37
and in the context of the 11-bp duplex it increased 5-fold (from 11 ± 2 to 56 ± 11, see
section 2.5)§ Increases were also observed at lower Mg2+
concentrations, although the
changes were smaller (2–4-fold, Table 2.4). These results underscore the established links
between duplex stability and the efficiency of separation by DEAD-box proteins (59),
and they indicate that the less efficient unwinding for a more stable duplex does not
simply result from slower action by DEAD-box proteins, but is accompanied by an
increase in ATP consumption. This increase would not be expected for a conventional
helicase and, as above, may reflect ATP hydrolysis events that are non-productive
because the core domain of CYT-19 dissociates before strand separation is complete (see
Discussion Section 2.4).
A model involving non-productive ATPase cycles would suggest that conditions
that weaken CYT-19 binding or stabilize the duplex would give increased ATP
requirements. Although changes in experimental conditions invariably give complex
effects with multiple physical origins, changing Mg2+
concentration and temperature
generally conformed to these expectations. With increased Mg2+
concentration, strand
separation rates decreased substantially (Tables 2.1, 2.3, and 2.4) and the ATP utilization
values increased. These changes most likely include contributions from increased duplex
stability (31, 57) and weaker binding by CYT-19 (S. Mohr and A.M.L, unpublished
data). Lower temperatures also gave increased ATP requirements (Table 2.5),
presumably in part from increased duplex stability.
2.3.3 Similar ATP utilization by other DEAD-box proteins
To explore whether the insights obtained for CYT-19 extend to other DEAD-box
proteins, we tested the Saccharomyces cerevisiae proteins Mss116p and Ded1p. Like
CYT-19, these proteins separated a 9-base-pair version of the P1 duplex at low Mg2+
38
concentrations with hydrolysis of at most a single ATP (Table 2.6). Further, both proteins
gave an ATP utilization of less than one at 2 mM Mg2+
, indicating that they, like CYT-
19, are capable of strand separation without ATP hydrolysis. With increasing Mg2+
concentration, the ATP utilization by each protein increased, analogous to the behavior of
CYT-19. Mss116p consistently hydrolyzed less ATP per separation event than did Ded1p
or CYT-19, perhaps reflecting tighter binding of Mss116p to single-stranded RNAs and
intermediates formed during strand separation (55).
2.4 DISCUSSION
By quantitatively comparing rates of ATPase activity and strand separation by the
DEAD-box protein CYT-19 and related proteins, we have measured the number of ATP
molecules hydrolyzed during separation of short helices under a range of solution
conditions and temperatures (summarized in Table 2.7). Although analogous
measurements have been made for several processive SF1 and SF2 helicases (reviewed in
ref. (113), few have been reported for DEAD-box proteins (52), and none for DEAD-box
proteins using short helices that are characteristic of structured RNAs. Our central
conclusions are, first, that DEAD-box proteins can separate short duplexes in a single
cycle of ATP-dependent conformational changes. Second, the process of strand
separation is initiated, and sometimes even completed, while the proteins remain in the
ATP-bound form. As described below, these insights provide critical new mechanistic
information on strand separation by DEAD-box proteins, supporting and extending
models for their action on physiological substrates (Figure 2.6).
2.4.1 Strand separation depends on ATP binding, not hydrolysis
Upon binding an RNA duplex, a conformational change is suggested to result in
tight binding of the protein to one strand of the RNA in a conformation that is
39
incompatible with a duplex (Figure 2.6, top left). This conformational change depends on
ATP binding but not hydrolysis, consistent with prior findings of cooperativity between
binding of ATP and ssRNA (44, 45). It may induce local strand separation or trap a
single-stranded segment that emerges due to transient ‘breathing’ of the duplex. After this
initial separation, which is suggested from structural analysis to be limited to five or six
base pairs (38, 39, 41), additional base pairs can apparently dissociate spontaneously to
allow complete separation of duplexes up to at least nine base pairs in the absence of any
ATP hydrolysis (pathway shaded blue in Figure 2.6). The existence of this pathway is
indicated by the ATP utilization values of less than one.
The conclusion that nucleotide-dependent unwinding does not require hydrolysis
is strongly supported by the finding from Jankowsky and colleagues that the non-
hydrolyzable analog ADP-BeFx also promotes unwinding (110), and our demonstration
that ATP gives this enhancement strongly supports the conclusion that ADP-BeFx
provides a good approximate model for the ATP-bound state of DEAD-box proteins
(110). Both studies also show that the enhancement by ATP is not mimicked by AMP-
PNP. The Jankowsky group reports that unwinding of the longer duplexes in their study
is undetectable with AMP-PNP, consistent with earlier work for several DEAD-box
proteins and indicating that any stimulation by AMP-PNP is much less than that of ATP
(79, 114, 115). Our work extends these results; because CYT-19-mediated separation of
the 6-bp duplex can be monitored in the absence of any nucleotide, we are able to show
that AMP-PNP provides no stimulation relative to this basal level. Whereas the lack of
detectable activity with AMP-PNP has been suggested to indicate a requirement for ATP
hydrolysis, the present results indicate that the defect arises at least in part from
differences between the ATP-bound and AMP-PNP-bound states. A similar suggestion
was made previously for the eIF4A protein from differences between ATP and AMP-
40
PNP in RNA binding and crosslinking experiments (44). These results cast doubt on the
general applicability of AMP-PNP as an analog of ATP for interactions with DEAD-box
proteins. Although AMP-PNP can give tight binding of ssRNA to DEAD-box proteins
(38, 45, 110, 116), this tight complex may not reflect an on-pathway intermediate for
ATP-dependent strand separation. Alternatively, it may be on-pathway but attained with
poor efficiency when starting from a double-stranded RNA and bound AMP-PNP.
2.4.2 What is the role of ATP hydrolysis?
Although some strand separation occurs in the absence of ATP hydrolysis, the
ATPase activities of all three DEAD-box proteins are substantially higher in the presence
of a duplex than with either strand alone (Figure 2.1B and data not shown). Thus,
interactions with the duplex stimulate ATP hydrolysis, suggesting that ATP is sometimes
hydrolyzed during strand separation. This ATP hydrolysis could occur from an
intermediate complex in which both strands remain present, in which case it could give
disruption of additional base pairs, or it could follow complete separation and function
solely to facilitate dissociation of the helicase core from the tightly bound strand to allow
additional cycles of unwinding (Figure 2.6 and ref. (110).
Although the precise role remains an open question, it is tempting to suggest that
ATP can be hydrolyzed prior to complete strand separation, as such premature hydrolysis
could account for the observations by us and others that separation of longer duplexes is
accompanied by hydrolysis of more than one ATP (52). If ATP were hydrolyzed prior to
complete strand separation, premature dissociation of the helicase core could then allow
re-zipping of the duplex and ‘wasted’ ATP hydrolysis (boxed pathway in Figure 2.6).
Supporting this interpretation, the ATPase rate remains constant as the length of the
duplex increases, whereas the strand separation rate decreases (52), suggesting that the
41
steps up to and including ATP hydrolysis are not affected by length, but that the longer
duplexes are then less likely to become completely separated. It should be noted that
alternative models for increased ATP utilization are possible, at least for longer duplexes
where the participation of multiple functional units of protein could be imagined.
However, even a duplex as short as 6-bp, which is almost certainly bound by only one
monomer (38), can have an ATP requirement exceeding unity (Table 2.4). These results
lead us to favor the non-productive cycles shown in Figure 2.6 as a central origin of the
increased ATP requirements.
Notably, both the ATP-hydrolysis-dependent and -independent pathways can give
complete strand separation in a single cycle, yielding ATP utilization values of one or
lower and ruling out a general requirement for multiple cycles of ATP-hydrolysis-
dependent translocation along the duplex (see Section 2.2.11). Importantly, the same
general model could apply to tertiary contact disruption or protein displacement, as the
critical feature is that a single strand of RNA is bound tightly and prevented from
interacting with alternative partners.
2.4.3 Implications for physiological activities
These mechanistic features of DEAD-box proteins are likely to be critical for their
roles in manipulating structured RNAs and RNPs. A tethering interaction, which for
CYT-19 and Mss116p appears to be mediated by the C-terminal domain (83, 92),
positions the helicase core to disrupt nearby RNA structure. For CYT-19 and Mss116p,
this interaction is relatively non-specific, whereas for DEAD-box proteins that function
with a defined RNA or RNP, this interaction is likely to be specific (77, 79). For both
classes, it has been proposed that the tethering interaction may be maintained during local
strand separation (31, 79). Here we have tested this model and provide data in strong
42
support of it. A single cycle of ATP-dependent conformational changes is sufficient to
give complete disruption of short helices, and such a cycle can easily be envisioned to
occur while the tethering interaction is maintained. This continued interaction may allow
DEAD-box proteins to disrupt the same local structure repeatedly, which may be
necessary to resolve misfolded RNAs such as group I introns (26), or to remain poised to
facilitate rearrangements of newly-formed intermediates that would otherwise revert
rapidly to non-productive structures. The same mechanism may also permit DEAD-box
and related proteins that function in such processes as pre-mRNA splicing to mediate
rapid and repeated interconversion of alternative sets of contacts, improving fidelity by
allowing sampling of alternative conformations and intermolecular contacts (33, 64, 117).
2.5 FOOTNOTES
*The CYT-19 concentration can be varied only over a limited range with our
current methods because at low concentrations the ATPase rate becomes too small to
measure the fraction of ATP hydrolyzed, and at high CYT-19 concentrations the strand
separation becomes too fast for hand pipetting. Across the accessible range (0.5–2 µM,
with CYT-19 in 4-fold excess of the duplex), the rate constant for strand separation
increased approximately linearly, indicating that CYT-19 is subsaturating with respect to
the duplex, and the ATP utilization value was unchanged within the expected limits of
uncertainty (data not shown).
‡This value arises from the strand separation rate in the presence of ATP under
conditions that do not favor its hydrolysis. The rate is 2–4 µM/min with 150 µM ATP,
with no indication of saturation (data not shown), and therefore at least 8 µM/min with
saturating ATP. ATP is hydrolyzed in only half of the strand-separation events. Thus, the
pathway that involves bound ATP but not its hydrolysis must give half of the total rate (4
43
µM/min), ≥ 20-fold faster than CYT-19-dependent strand separation in the absence of
ATP.
§It is interesting that the G•U to G-C change increased the ATP requirement
approximately equally as a terminal pair (6-bp duplex) or an internal pair (11-bp duplex),
because G•U is more destabilizing as an internal pair in model duplexes (111). The
substitutions also gave similar decreases in the CYT-19-mediated unwinding rate, ~20-
fold. It is possible that the ‘extra’ destabilization from the internal G•U pair is reduced or
absent under our solution conditions (the previous study used 1 M NaCl). Alternatively or
in addition, the terminal G•U may generate local destabilization and a preferred entry
point for CYT-19, compensating for its smaller intrinsic effect on duplex stability.
44
Figure 2.1: ATP hydrolysis and RNA strand separation by CYT-19
(A) Structure of the duplex substrate, derived from the P1 duplex of the Tetrahymena
group I intron. RNA nucleotides are red and DNA nucleotides are black. The DNA
portion has the equivalent sequence of the P2 helix, which is adjacent to P1 in the natural
intron. (B) P1 duplex-dependent ATP hydrolysis by CYT-19. ATP hydrolysis was
measured in the presence of the P1 duplex by including both oligonucleotides (), or
with the same concentrations of the RNA/DNA oligonucleotide (0.5 µM, ) or the
excess CCCUCUA5 (1 µM, ) alone. (C) Strand separation of the P1 duplex construct in
the presence () or absence () of CYT-19. Excess CCCUCUA5 (5 µM) was present to
prevent re-annealing of the 32P-labeled CCCUCUA5. An equivalent reaction with 1 µM
CCCUCUA5 gave the same rate constant within error (Figure 2.3), but the higher
concentration allowed more precision by increasing the extent of displacement of the
labeled CCCUCUA5. Experimental conditions for panels B and C were 25 °C, pH 7.0, 10
mM Mg2+
, 50 µM ATP-Mg2+
, and 2 µM CYT-19.
45
Figure 2.2: Effects of a double-stranded extension on RNA strand separation and
ATPase activity
(A) Separation of the 6-base-pair P1 duplex. Rate constants are plotted against the CYT-
19 concentration for the P1 duplex alone (construct 7 in Figure 2.7, , 2.6 105 M
–1
min–1
) or with an adjacent helix composed of RNA (◊, 3.2 106 M
–1 min
–1) or DNA (,
4.4 106 M
–1 min
–1) with 2 mM ATP-Mg
2+ (constructs 8 and 1, respectively, in Figure
2.7). The enhancement in strand separation activity conferred by the additional helix, 10–
20-fold relative to the P1 duplex alone, is the same within error whether the additional
helix is composed of DNA or RNA. (Data for constructs 7 and 8 are reprinted from ref.
(31) for comparison. All reactions were performed at 25 °C, pH 7.0, 10 mM Mg2+
). (B)
ATPase activity under identical solution conditions (25 °C, pH 7.0, 10 mM Mg2+
) in the
presence of 50 µM ATP-Mg2+
. Symbols are the same as above. After subtracting the
basal ATP hydrolysis rate, 0.05 µM/min, ATPase rates are 0.02 µM/min for P1 alone
(), 1.6 µM/min for P1 with an activating RNA helix (◊), and 0.6 µM/min for P1 with an
activating DNA helix (). The activating DNA helix gives less enhancement of ATP
hydrolysis by CYT-19 than the activating RNA helix. Further, the increase in ATPase
activity from the activating DNA helix (30-fold) is in the same range as the increase in
strand separation, suggesting that the increased ATPase activity arises from the increased
rate of strand separation of the attached P1 RNA helix, not from direct ATPase
stimulation by the DNA portion. (C) The DNA extension does not itself stimulate ATP
hydrolysis. Progress of ATP hydrolysis is shown in the presence of 0.5 µM of the
standard RNA/DNA chimeric oligonucleotide (present in constructs 1 and 5 of Figure
2.7, ), the same concentration of an oligonucleotide that includes only the DNA portion
(construct 9 in Figure 2.7, ), or CYT-19 alone ().
(Panels A and B are the work of Cindy Chen.)
46
Figure 2.3: Strand separation by CYT-19 is independent of ‘chase’ CCCUCUA5
concentration (1–10 µM)
Progress curves for separation of the 6-bp P1 duplex construct (Figure 2.1A and construct
1 in Figure 2.7) are shown in the presence of 1 µM (, 1.61 min–1
), 2 µM (∆, 1.60 min–1
),
5 µM (, 1.60 min–1
), and 10 µM (, 1.41 min–1
) unlabeled CCCUCUA5 chase. An
equivalent reaction in the absence of CYT-19 is also shown (, 0.08 min–1
). Conditions
for all reactions were 25 °C, pH 7.0, 10 mM Mg2+
, 50 µM ATP-Mg2+
, and 0.5 µM of the
P1 duplex construct. The independence of the rate constant on the concentration of
unlabeled CCCUCUA5 indicates that CCCUCUA5 does not inhibit strand separation at
concentrations ≤10 µM. Thus, 1 µM CCCUCUA5 was used in ATPase measurements to
minimize the ATP hydrolysis arising from interactions of free CCCUCUA5 with CYT-19,
and 5 µM CCCUCUA5 was used in strand separation experiments to maximize the extent
of visible strand separation. Lower concentrations of chase are expected to give smaller
extents of visible strand separation, as shown here, because a substantial fraction of
labeled material remains in the duplex when equilibrium is reached. These fractions were
somewhat lower than calculated from the relative concentrations of chase CCCUCUA5
and the duplex, apparently because a fraction of the duplex dissociates during gel
electrophoresis (see Section 2.2.5).
(This figure is the work of Cindy Chen.)
47
Figure 2.4: ATP-independent strand separation by CYT-19
(A) Separation of the 6-base-pair P1 duplex with the DNA extension (Figure 2.1A) in the
presence of 2 mM Mg2+
. The rate constants are 0.12 min–1
in the absence of CYT-19 (∆),
0.46 min–1
in the presence of CYT-19 alone (), 0.47 min–1
in the presence of CYT-19
and 1 mM AMP-PNP (), and 2.30 min–1
in the presence of CYT-19 and 50 µM ATP-
Mg2+
(). The CYT-19-dependent strand separation rate in the absence of ATP is
calculated by subtracting the CYT-19-independent rate constant and multiplying by the
duplex concentration: (0.46 min–1
– 0.12 min–1
) 0.5 µM = 0.2 µM/min. These results
indicate that CYT-19 possesses ATP-independent strand separation activity under these
conditions and that AMP-PNP does not enhance this activity. CYT-19 was bound to
AMP-PNP in our experiments, because AMP-PNP inhibits ATP-dependent strand
separation with a Ki value of 200 µM (data not shown), 5-fold lower than the
concentration used here. (B) Separation of the 6-base-pair P1 duplex under standard
conditions with 10 mM Mg2+
. The rate constants were 0.08 min–1
in the absence of CYT-
19 (∆), 0.09 min–1
for CYT-19 alone (), and 1.8 min–1
for CYT-19 with 50 µM ATP-
Mg2+
(). Under these conditions, no significant ATP-independent strand separation by
CYT-19 was detected.
(These figures are the work of Cindy Chen.)
48
Figure 2.5: ATP hydrolyzed by CYT-19 per separation event of the 6-base-pair P1
duplex with low Mg2+
concentration (2 mM)
Open and filled circles show results from two identical experiments. The solid curve
shows the best fit by a model that includes stimulation of unwinding by bound ATP
without hydrolysis (see Section 2.2.6). Dashed curves show best fits to discarded models
in which only free CYT-19 gives ATP hydrolysis-independent strand separation (long
dashes) or in which ATP-bound CYT-19 can give strand separation without hydrolysis,
but with the same efficiency as nucleotide-free CYT-19 (short dashes).
(This figure is the work of Cindy Chen.)
49
Figure 2.6: Model for duplex separation by DEAD-box proteins
Interactions are formed between the ATP-bound helicase core and the RNA (radiolabeled
RNA indicated by an asterisk), and a tethering interaction is formed adjacent to the core
by an ancillary site, as shown, or by an additional protomer (53). Concomitant with or
subsequent to initial binding by the helicase core, a conformation that is dependent on
ATP binding but not hydrolysis induces or captures local strand separation. Complete
strand separation can be achieved without ATP hydrolysis (left, shaded blue) or with
ATP hydrolysis (pathways down and to the right), which accelerates dissociation of the
helicase core and may induce additional strand separation. Premature dissociation of the
helicase core after ATP hydrolysis leads to a futile cycle (counterclockwise within box).
Throughout the entire strand separation process, the tethering interaction may remain
intact, as shown, allowing the protein to perform multiple cycles of structure disruption
on the same RNA without being lost to solution.
(Figure prepared by Cindy Chen.)
50
Figure 2.7: Duplex constructs used for measurements of ATP utilization during
unwinding (top, constructs 1-6) and for control experiments (bottom,
constructs 7-9)
All constructs for ATP utilization measurements included the same DNA sequence
(black). The P1 duplex was increased in length by using different DNA/RNA hybrid
strands that included various numbers of uridine nucleotides at the 5-end as shown
(constructs 2-4 and 6). The all-Watson-Crick duplexes (5-6) were constructed by using
CCCUCCA5 instead of CCCUCUA5.
(Figure prepared by Cindy Chen.)
51
[Mg2+
]
, mM
Total
ATPase rate
Duplex-
dependent
ATPase rate†
Total strand
separation rate
CYT-19-
dependent
strand
separation rate‡
ATP
hydrolyzed per
strand
separation
2 1.10 ± 0.03 0.50 ± 0.03 1.05 ± 0.05 0.97 ± 0.05 0.45 ± 0.04
5 1.25 ± 0.04 1.10 ± 0.05 1.25 ± 0.10 1.20 ± 0.10 0.90 ± 0.09
10 0.80 ± 0.05 0.74 ± 0.05 0.73 ± 0.05 0.69 ± 0.05 1.1 ± 0.1
20 0.13 ± 0.01 0.11 ± 0.01 0.10 ± 0.01 0.05 ± 0.01 2.0 ± 0.4
Table 2.1: ATP utilization for CYT-19-mediated separation of the 6-bp P1 duplex
Reactions were performed at 25 °C, 50 mM Na-MOPS, pH 7.0, 50 µM ATP-Mg2+
, 0.5
µM duplex, 1.2 µM total CCCUCUA5, 2 µM CYT-19. All rates are µM/min. Values are
averages and standard deviations from two to four independent determinations.
†Values are the total ATPase rate minus the ATPase rate in the presence of the
approximate concentration of excess CCCUCUA5 expected to be single-stranded (0.7 – 1
µM). This difference represents the rate of ATPase activity arising from CYT-19
interacting with the duplex. The background ATPase activity was measured in the
presence of 1 µM CCCUCUA5 to allow for the possibility of incomplete duplex
formation, and control reactions established that the background rate depends only
weakly on CCCUCUA5 concentration between 0.7 and 1 µM (<20% at 2 mM Mg2+
and
less than 10% at 10 mM Mg2+
; data not shown).
‡Values are the observed rate constant for strand separation minus the basal strand
separation rate constant in the absence of CYT-19, with the difference multiplied by the
duplex concentration (0.5 µM) to give a steady-state rate.
(This table is the work of Cindy Chen.)
52
[Mg2+
],
mM 6 bp 7 bp 9 bp 11 bp
2 0.45 ± 0.04 0.4 ± 0.1 1.1 ± 0.1 1.2 ± 0.1
5 0.90 ± 0.09 1.3 ± 0.4 1.4 ± 0.4 4.4 ± 0.7
10 1.1 ± 0.1 1.3 ± 0.2 5 ± 2 11 ± 2
20 2.0 ± 0.4 2.0 ± 0.4 14 ± 2 18 ± 2
Table 2.2: Dependence of ATP utilization by CYT-19 on Mg2+
concentration and
duplex length
Values are ATP hydrolyzed per duplex separated (ATP utilization value). Longer
duplexes were derived from that shown in Figure 2.1A by extending the 5´-end of the
RNA/DNA strand with uridine nucleotides. All reactions were performed at 25 °C, 50
mM Na-MOPS, pH 7.0, 50 µM ATP-Mg2+
, 0.5 µM duplex, and 2 µM CYT-19.
(This table is the work of Cindy Chen.)
53
duplex
length
[Mg2+
],
mM
Total ATPase
rate
Duplex-dependent
ATPase rate†
Total strand
separation rate
CYT-19-
dependent strand
separation rate‡
ATP
hydrolyzed per
strand
separation
7 bp¶ 2 1.10 ± 0.05 0.65 ± 0.05 1.8 ± 0.3 1.8 ± 0.4 0.4 ± 0.1
5 1.6 ± 0.3 1.4 ± 0.5 1.2 ± 0.3 1.1 ± 0.3 1.3 ± 0.4
10 0.9 ± 0.2 0.9 ± 0.3 0.58 ± 0.02 0.55 ± 0.04 1.3 ± 0.2
20 0.12 ± 0.03 0.10 ± 0.03 0.07 ± 0.02 0.038 ± 0.003 2.0 ± 0.4
9 bp 2 2.0 ± 0.4 1.7 ± 0.4 1.4 ± 0.2 1.4 ± 0.2 1.1 ± 0.1
5 2.1 ± 1.0 2.1 ± 1.3 1.3 ± 0.4 1.3 ± 0.4 1.4 ± 0.4
10 1.3 ± 0.4 1.2 ± 0.4 0.28 ± 0.07 0.27 ± 0.08 5 ± 2
20 0.30 ± 0.03 0.25 ± 0.10 0.020 ± 0.002 0.014 ± 0.002 14 ± 2
11 bp 2 2.0 ± 0.2 1.7 ± 0.3 1.8 ± 0.5 1.8 ± 0.6 1.2 ± 0.1
5 3.2 ± 0.2 3.00 ± 0.05 0.9 ± 0.4 0.9 ± 0.4 4.4 ± 0.7
10 1.7 ± 0.2 1.6 ± 0.3 0.15 ± 0.05 0.14 ± 0.05 11 ± 2
20 0.15 ± 0.05 0.10 ± 0.03 0.006 ± 0.001 0.005 ± 0.001 18 ± 2
Table 2.3: Dependence of strand separation and ATPase rates on duplex length
Reaction conditions were 25 °C, 50 mM Na-MOPS, pH 7.0, 0.5 µM duplex, 2 µM CYT-
19. Rates are µM/min. Values are averages and standard deviations from at least three
independent determinations. Duplexes are as shown in Figure 2.1A, with longer versions
achieved by extending the 5´-end of the chimeric RNA/DNA oligonucleotide with
uridine nucleotides (constructs 2–4 in Figure 2.7).
†Duplex-dependent ATPase rate is the total ATPase rate minus the ATPase rate in the
presence of CYT-19 and the approximate concentration of CCCUCUA5 expected to be
present as ssRNA (1 µM).
‡CYT-19-dependent strand separation rate is the strand separation rate constant minus
the basal dissociation rate constant in the absence of CYT-19, both multiplied by the
duplex concentration.
¶Although this construct has sequence potential to form a 7-bp duplex (compare
constructs 1 and 2 in Figure 2.7), recent work using the same duplex extension in the
context of the intact Tetrahymena ribozyme suggests that the single-base extension does
not form an additional base pair (111). Consistent with this interpretation, the unwinding
rates are the same within error for this construct and the 6-bp duplex construct that lacks
this single-nucleotide extension (Table 2.1).
(This table is the work of Cindy Chen.)
54
Table 2.4: RNA strand separation and ATP hydrolysis rate for duplexes composed
solely of canonical Watson-Crick base pairs
Reaction conditions were 25 °C, 50 mM Na-MOPS, pH 7.0, 50 µM ATP-Mg2+
, 0.5 µM
duplex, and 2 µM CYT-19. Rates are µM/min. Values represent averages and standard
deviations from two independent determinations.
†The duplex constructs are the same as shown in Figure 2.1A, except that the P1 portion
of the constructs used here includes a G-C base pair instead of the natural G•U pair
shown in Figure 2.1A. The 6-bp and 11-bp duplex constructs are numbered 5 and 6,
respectively, in Figure 2.7.
‡The values shown are the ATP utilization values (ATP hydrolyzed per strand separation,
second-to-right column) divided by the corresponding values for the duplex of the same
length that includes the natural G•U pair instead of a G-C pair.
(This table is the work of Cindy Chen.)
Duplex
length†
[Mg2+
]
, mM
Total
ATPase
rate
Duplex-
dependent
ATPase rate
Total strand
separation rate
CYT-19-
dependent
strand
separation rate
ATP
hydrolyzed per
strand
separation
ATP
utilization
rel. to G•U
duplex‡
6 bp 2
1.6 ±
0.5 1.4 ± 0.4 1.3 ± 0.5 1.3 ± 0.5 1.2 ± 0.3 2.7 ± 0.7
5
2.2 ±
0.8 2.1 ± 0.8 0.9 ± 0.5 0.9 ± 0.5 2.9 ± 0.7 3.2 ± 0.8
10
0.31 ±
0.11 0.28 ± 0.10 0.035 ± 0.01 0.032 ± 0.01 9 ± 3 8.2 ± 2.8
11 bp 2
1.4 ±
0.6 1.3 ± 0.5 0.29 ± 0.11 0.29 ± 0.11 4.3 ± 1.5 3.6 ± 1.3
5
1.7 ±
0.7 1.6 ± 0.7 0.10 ± 0.05 0.10 ± 0.06 17 ± 7 3.9 ± 1.8
10
0.35
±0.06 0.31 ± 0.06 0.006 ± 0.0007 0.006 ± 0.0007 56 ± 11 5.1 ± 1.4
55
T (ºC)
Total
ATPase rate
Duplex-
dependent
ATPase rate
Total strand
separation rate
CYT-19-
dependent
strand
separation rate
ATP hydrolyzed
per strand
separation
15 0.22 0.19 0.006 0.005 38
25 0.49 0.45 0.036 0.033 14
37 0.18 0.13 0.21 0.14 0.93
Table 2.5: Temperature dependence of ATP utilization by CYT-19 during strand
separation
Reaction conditions were 50 mM Na-MOPS, pH 7.0 (at 25 °C), 0.5 µM duplex, 2 µM
CYT-19, 10 mM Mg2+
, and 50 µM ATP-Mg2+
. Rates are µM/min. Experiments
monitored separation of the 6 bp duplex that included a G-C base pair instead of the
natural G•U pair (construct 5 in Figure 2.7). A reaction at 25 °C, whose results are
included here, was performed in parallel with reactions at 15 °C and 37 °C, and gave
similar, but not identical, results to those shown in Table 2.4 from independent,
equivalent experiments.
(This table is the work of Cindy Chen.)
56
[Mg2+
], mM Mss116p Ded1p CYT-19
2 0.5 ± 0.1 0.52 ± 0.09 1.1 ± 0.1
5 0.8 ± 0.3 1.5 ± 0.9 1.4 ± 0.4
10 0.8 ± 0.2 1.9 ± 0.5 5 ± 2
20 2.6 ± 0.5 6 ± 3 14 ± 2
Table 2.6: ATP utilization by the DEAD-box proteins Mss116p and Ded1p
Values are ATP hydrolyzed per duplex separated. Separation of a 9-bp duplex was
monitored (construct 3 in Figure 2.7). Conditions were identical to experiments for CYT-
19 except that lower concentrations of proteins, substrates, and ATP were used to
compensate for increased activity of these proteins. For experiments with Mss116p, the
protein concentration was 100 nM, the duplex concentration was 25 nM, and the ATP
concentration was 0.5–100 µM, depending on the Mg2+
concentration. For experiments
with Ded1p, the protein concentration was 100–500 nM, the duplex concentration was
25–125 nM (maintaining a constant ratio of Ded1p and duplex concentrations), and the
ATP concentration was 1–200 µM. In all cases, the ATP utilization values reported are
saturating with respect to ATP concentration (data not shown). Corresponding values for
CYT-19 are reproduced from Table 2.2 for comparison.
(Pilar Tijerina contributed data used in this table.)
57
Variable Range Other conditions*
CYT-19 concentration 0.5 – 2 µM 10 mM Mg2+
, duplex 1 (6 bp) (not shown)
ATP concentration 1 – 150 µM
5 – 150 µM
2 mM Mg2+
, duplex 1 (6 bp), Figure 2.5
10 mM Mg2+
, duplex 1 (6 bp) (not shown)
All ATP concentrations tested are subsaturating for
CYT-19 binding, based on a previous determination
of KM of 200 µM (28) and our results that strand
separation rate increases with ATP concentration
across the range tested (data not shown).
Mg2+
concentration 2 – 20 mM
2 – 10 mM
Duplexes 1–4 (6–11 bp), Tables 2.1 and 2.3
Duplexes 5 and 6 (6 and 11 bp), Table 2.4
Duplex length 6 – 11 bp G•U background (duplexes 1-4), 2–20 mM Mg2+
,
Tables 2.1 and 2.3
G-C background (duplexes 5 and 6), 2–10 mM Mg2+
,
Table 2.4
Duplex stability G•U or G-C 2, 5, and 10 mM Mg2+
, 6 and 11 bp (duplexes 5 & 6),
Tables 2.1, 2.3, and 2.4
Temperature 15 – 37 °C Duplex 5 (6 bp), 10 mM Mg2+
, Table 2.5
Protein identity CYT-19,
Mss116p
Ded1p
See above
Duplex 3 (9 bp), 2 – 20 mM Mg2+, 100 nM
Mss116p,
0.5 – 100 µM Mg2+
, Table 2.6
Duplex 3 (9 bp), 2 – 20 mM Mg2+, 100 – 500 nM
Ded1p,
1 – 200 µM Mg2+
, Table 2.6
Table 2.7: Range of conditions for measurements of ATP utilization
* Duplex numbering refers to Figure 2.7.
(This table is the work of Cindy Chen.)
58
The text below and following figures of chapter 3 were published originally by Elsevier.
Copyright © by Elsevier Inc. 2012. All rights reserved.
Potratz, J.P. and Russell, R. (2012) RNA catalysis as a probe for chaperone activity of
DEAD-box helicases. Methods in Enzymology. 511, 111-130.
Rick Russell helped organize the paper content.
Chapter 3: RNA catalysis as a probe for chaperone activity of DEAD-
box helicases
3.1 CATALYTIC ACTIVITY AS A PROBE OF RNA FOLDING
To track productive folding of an RNA, a signal must exist that allows the native
state of the RNA to be distinguished from all other conformations. Functional assays
provide a powerful probe, because even folding intermediates with extensive native
structure can be readily distinguished if they are unable to function. Catalytic RNAs are
well suited for this purpose, as their native states can be readily detected by monitoring
chemical conversion of a substrate to its corresponding product.
3.1.1 Catalytic activity distinguishes the native state from all other conformations
The central goal of using catalytic activity to monitor RNA folding is to measure
the fraction of the RNA that is present in the native state unambiguously and
quantitatively. This information can then be combined with other approaches to gain
insights into the structural properties of folding intermediates. In the case of the ribozyme
derived from a group I intron in Tetrahymena thermophila, a misfolded conformation
exists that contains the full set of native secondary and tertiary contacts and is difficult to
distinguish from the native state by physical approaches (17, 118). However, it is
straightforward to distinguish between the two structures in a catalytic activity assay
because the natively folded ribozyme, but not the misfolded ribozyme, can cleave an
59
oligonucleotide substrate in trans. Using this assay, the transition from the misfolded state
to the native state has been measured (17, 119-121).
3.1.2 Catalytic activity can be used to study chaperone-assisted folding
The ability to track native RNA folding allows the influence of chaperone
proteins to be monitored. While this chapter highlights group I and group II introns and
the DEAD-box helicase chaperone proteins, the general techniques can be applied to a
broad range of catalytically active RNAs and chaperone proteins. Indeed, RNA catalytic
activity of the hammerhead ribozyme was used nearly two decades ago to study
chaperone activities of the HIV NC protein (122, 123), and group I introns have been
used to monitor in vitro and in vivo chaperone activities of a variety of RNA binding
proteins (90, 124-126).
Group I and group II introns are mobile genetic elements that catalyze their own
excision from precursor RNA via two transesterification splicing reactions (Figure 3.1A).
In order for these splicing events to occur, the RNA must fold to an active three-
dimensional conformation. Introns can be converted to ribozymes with constructs that
lack the exons (Figure 3.1B,C). Ribozymes of group I and group II introns cleave
oligonucleotide substrates that include a 5´ splice site in reactions that mimic the first step
of splicing (99, 127-131).
Group I and group II introns and their corresponding ribozyme constructs are well
suited for studying chaperone-assisted RNA folding because they have extensive
networks of secondary and tertiary structures (Figure 3.1B,C), and RNAs from both
groups have been shown to fold slowly and to accumulate intermediates (132-140). These
RNAs are sufficiently complex to include diverse sets of intermediates and corresponding
kinetic barriers during folding, allowing detailed probing of the abilities of chaperones to
60
assist in overcoming these barriers, while remaining simple enough to deeply probe the
folding processes and pathways.
3.2 SELF-SPLICING AS A READOUT FOR NATIVE STATE FORMATION
A straightforward way of measuring native folding is to follow self-processing of
intron-containing constructs after folding is initiated by addition of Mg2+
. The reaction
can be followed in the format of a continuous assay, in which folding and the catalytic
steps take place concurrently and in the same reaction. If the rate of native RNA structure
formation is slower than the subsequent catalytic steps, the observed splicing rate
provides a good measure of the rate of folding to the native state.
It is common practice to body label the RNA to visualize all the products of intron
splicing. This can be accomplished by including a radiolabeled nucleotide, commonly [α-
32P] UTP, to label nucleotides throughout the RNA (30, 91, 141). The reaction is then
monitored by denaturing polyacrylamide gel electrophoresis, which allows the unspliced
precursor to be separated from the splicing products (Figure 3.2A). The loss of precursor
RNA over time is quantitated to indicate the observed rate of splicing, and in turn folding
to the native state (30, 81, 142) (Figure 3.2B).
3.2.1 Interpreting chaperone-promoted changes in observed splicing rate
A simple reaction scheme for intron folding and splicing is shown below and is
instructive for understanding how a chaperone protein affects the splicing process.
Scheme 1
61
To identify whether an RNA helicase influences folding, the observed rate
constant (kobserved) is compared in the absence and presence of multiple protein
concentrations and in the presence and absence of ATP. If the helicase increases the rate
of folding to the native state (kfold), most likely by accelerating resolution of one or more
intermediates (I), this increase will lead to an increase in kobserved if the splicing rate
constant (ksplice) is larger than the folding rate constant. For introns with robust self-
splicing activity this condition is generally met because large RNAs usually fold slowly
in vitro (143, 144).
The observed rate constant is typically plotted as a function of protein
concentration to determine the efficiency with which the chaperone promotes folding
(Figure 3.2C). The observed rate constant may increase with protein concentration at low
concentrations and then level or begin to decrease at higher protein concentrations
(Figure 3.2C). Possible physical sources for this inhibition are trapping of intermediate
conformations in a protein-bound state (shown in Scheme 1 as a single intermediate (I)
for simplicity) and chaperone-induced unfolding of the native state. Although unfolding
of native RNAs may be physiologically important for RNAs that must cycle between
functional structures, this inhibition complicates analysis of the effects of the chaperone
on the folding process toward the native state. Thus it is most straightforward to interpret
the data in terms of protein-accelerated folding at relatively low protein concentrations
where these additional effects are minimized (Figure 3.2C).
3.2.2 Potential complications
Since the chaperone is present during the splicing reaction, it could affect the
catalytic steps of splicing (ksplice) in addition to the folding steps. If the catalytic steps
(ksplice) are rate-limiting, influencing them will alter the overall rate constant (kobserved)
62
even if the protein does not affect the rate of native structure formation (kfold). However,
this will not be an issue provided that folding remains slower than splicing. Group I
introns allow the rate-limiting step to be determined. A folding incubation is performed in
the absence of the exogenous guanosine cofactor and then guanosine is added to permit
splicing. If this reaction gives a larger splicing rate than the reaction in which guanosine
was present continuously, it indicates that folding is rate limiting.
A second issue concerns the multi-step nature of the splicing process. Although it
is shown as a single step for simplicity in scheme 1, some group I and group II introns
display rapid, reversible first steps of splicing, giving accumulation of intermediates in a
fast phase that is followed by slower completion of the second step and formation of
products (145-148). If native folding is slower than the reversible first step but faster than
the second step, the precursor will be lost with an initial rate constant that reflects
folding, as in the simple case with a more rapid second step. A slow step will result in
additional loss of precursor. This loss could mistakenly be assigned to an additional
folding pathway that suggests slower folding of the precursor, whereas it actually reflects
completion of the second splicing step. This behavior can be identified by the appearance
and subsequent disappearance of the intermediates.
A final issue concerns the choice of RNA constructs. Although emphasis is
typically on the intron for folding, exon length and composition can cause substantial
effects (142, 149, 150). The exons may misfold themselves and/or stabilize structure
within the intron, and either of these effects may be important biologically. Care must be
taken when comparing results from constructs that differ in the properties of the exons.
63
3.3 SUBSTRATE CLEAVAGE AS A READOUT FOR NATIVE STATE FORMATION
Many of the potential complications associated with using self-splicing constructs
in continuous assays can be avoided by using ribozyme versions. Lacking exons,
ribozyme constructs allow interpretation of all folding processes to reflect the intron
domains. Further, although ribozyme versions can be used in continuous assays, the fact
that they cleave oligonucleotide substrates in trans makes them well suited for
discontinuous assays, in which folding and catalytic activity are separated into two
discrete stages (Figure 3.3A). The main advantage of this separation is the ability to
directly assess the fraction of native ribozyme during folding. This allows productive
folding to be dissected and unfolding of the native structure to be probed, (see Section
3.4.1, below). In addition, the discontinuous assay permits the use of folding conditions
that do not support robust catalysis, an option not available using a continuous assay.
3.3.1 Setting up a discontinuous assay: folding and catalysis stages
The discontinuous assay is composed of two stages, the folding stage (stage 1)
and the catalysis (or cleavage) stage (stage 2) (Figure 3.3A). The folding stage contains
the ribozyme alone or with the chaperone under conditions desired for chaperone-assisted
RNA folding. To prevent catalytic activity at this stage, the oligonucleotide substrate is
omitted. In the catalysis stage, conditions are changed such that further folding to the
native state is blocked, and radiolabeled oligonucleotide substrate is added, typically in
small excess of the ribozyme (2-3-fold over the ribozyme). The cleavage reaction is
allowed to proceed for various times (t1), and the substrate and product from each time
point are separated on a denaturing polyacrylamide gel (Figure 3.3B). The fraction of
product, normalized by the substrate and ribozyme concentrations, is plotted against
cleavage time (t2) (Figure 3.3B,C). Most commonly, the cleavage stage produces a burst
of product formation, with the amplitude reflecting the fraction of ribozyme in the native
64
state (see Section 3.3.2, below). This fraction increases as a function of time spent in the
folding stage (t1), giving a rate constant for native folding of the ribozyme (Figure 3.3D).
After initial experiments have been performed and the cleavage rate constant is known, a
single time point in the cleavage reaction (stage 2) may be sufficient to determine the
burst amplitude (see Figure 3.4A). This timepoint should be chosen after completion of
the burst phase, but before significant contribution from subsequent turnovers is seen.
A key advantage of the discontinuous assay is that any set of conditions can be
used for the folding stage. However, setting up the catalysis stage requires care, as
conditions must support enzymatic activity while blocking further native folding. This is
because ribozyme that reached the native state during stage 2 would produce cleavage
products and cause the calculated fraction of native ribozyme to be artificially high.
The ability of the catalysis stage to prohibit folding is probed by comparing
cleavage reactions from a ribozyme that has been prefolded to the native state and a
ribozyme that is transferred directly to the catalysis stage, omitting the folding step. (In
practice, testing for suitable conditions for the catalysis stage can be undertaken
simultaneously with learning how to prefold ribozyme to the native state, as described
below.) The burst amplitude of product from ribozyme that skipped the folding stage
should be much less than that from the prefolded ribozyme, indicating little formation of
native ribozyme during the cleavage reaction. It may not be possible to completely block
native folding in stage 2, but a relatively small fraction of native ribozyme produced in
this stage can be accounted for by normalization (142).
Although optimal conditions for stage 2 are likely to be different for different
catalytic RNAs, some general guidelines are applicable. A study using a ribozyme
engineered from the group II intron aI5γ from S. cerevisiae, D135 ribozyme (Figure
3.1C), used a high pH in the catalysis stage to accelerate cleavage, and high Mg2+
65
concentration (100 mM) and low temperature (15 C ) to enhance the arrest of folding
(Figure 3.4A) (see Section 4.3.3) (142). Cleavage is not particularly fast under these
conditions (10-2
min-1
). However, slow cleavage is acceptable for the catalysis stage in a
discontinuous assay as long as cleavage is significantly faster than folding. In addition,
proteinase K is included at the catalysis stage to ensure that protein transferred from the
folding stage has no effect on substrate cleavage.
To compare a ribozyme prefolded to the native state with an unfolded ribozyme, it
must be known how to fold the ribozyme to the native state. Establishing how to fold the
ribozyme to the native state can be undertaken concurrently with exploring conditions to
use for the catalysis stage. Folding of a ribozyme (typically by adding Mg2+
) is initiated
and aliquots are removed at different times from the folding stage. These aliquots are
transferred to stage 2 conditions in the presence of a small excess of substrate. The
product burst amplitudes are plotted against folding time. When the burst amplitude no
longer increases as a function of folding time, the ribozyme has been folded to the native
state as fully as it can be under that set of conditions. Care must be taken to avoid
misinterpreting a slower phase of folding as a plateau indicating complete folding (Figure
3.4B).
3.3.2 Interpreting results from the catalysis stage
To make optimal use of the discontinuous assay, it is critical to interpret the burst
amplitude quantitatively in order to determine the fraction of native ribozyme. Selected
examples from work involving chaperones will be covered below, and a more thorough
guide to interpreting results from catalytic rate measurements was published last year
(151).
66
The interpretation of the burst amplitudes depends on the relative rate constants of
different steps in the catalytic cycle, which is shown in Scheme 2.
Scheme 2
In general, the cleavage stage is performed under multiple turnover conditions,
and the fraction of product is normalized by the substrate and ribozyme concentrations,
giving the ratio or product to ribozyme. This ratio is plotted against cleavage time (t2)
(Figure 3.3B,C). The reaction products are bound by base-pairing, with or without
additional tertiary contacts, and for many ribozymes their dissociation rate constants
(kP5´release and kP3´release) are small and limit the rates of subsequent turnovers. In all, there
are three possible regimes: (1) both products are released quickly relative to the catalytic
step (kcleavage); (2) both are released slowly; (3) one is released quickly and the other
slowly (151). Regimes 2 and 3 result in bursts of product formation, and the amplitude of
this burst can be used to calculate the fraction of native ribozyme, as described in two
examples.
The D135 ribozyme was used in a catalytic reaction with three-fold excess
substrate. When performed with a ribozyme prefolded to the native state, the cleavage
stage resulted in a burst of product with an amplitude approximately equal to one
turnover of the ribozyme (Figure 3.4A) (see Section 4.3.3) (142). This is consistent with
regime 3, in which there is a burst of product formation with a rate constant equal to the
cleavage rate (kcleavage) and a subsequent linear phase dictated by the slow release of one
of the products. Because the 5´ product of the oligonucleotide substrate forms twelve
67
Watson-Crick base pairs with the ribozyme, product release might be slow. Under this
reaction regime, the amplitude of the burst phase is equal to the fraction of D135
ribozyme folded to the native state.
The ribozyme engineered from a group I intron from Azoarcus evansii (Figure
3.1B) was used in a catalytic reaction with two-fold excess substrate (136). When the
ribozyme was prefolded to the native state, the cleavage stage resulted in a burst of
product with an amplitude approximately equal to half of the ribozyme concentration
(Figure 3.4B). This is consistent with regime 2, in which the slow release of both
products allows equilibrium between the cleavage (kcleavage) and ligation (kligation) to be
reached. The burst amplitude is smaller than the fraction of native ribozyme and must be
corrected by the value of the internal equilibrium to reveal the fraction of native
ribozyme.
3.3.3 Using the discontinuous assay to probe chaperone-assisted folding
When optimal conditions for stage 2 are established and the relationship of the
burst amplitude to the fraction of native ribozyme is understood, the discontinuous assay
can provide important insights into chaperone-assisted RNA folding. The most
straightforward experiment is to compare the folding reaction in the presence of various
concentrations of chaperone, plotting the fraction of native ribozyme against time (t1)
(Figure 3.3D). A chaperone may increase the rate of native state formation (31, 136), an
effect that most likely arises from accelerated resolution of one or more kinetically-
trapped intermediates. This interpretation is particularly clear for RNAs that are known to
misfold (31, 134, 136, 152). Further confirmation can be obtained by allowing the RNA
to misfold first and then adding the chaperone (31, 136).
68
It is possible for a chaperone to increase the fraction of ribozyme that reaches the
native state rapidly without having a significant impact on the observed rate constant
(Figure 3.5A,B) (142). This effect may arise from an influence of the chaperone early in
folding, which decreases the probability of misfolding at later folding steps.
Alternatively, resolution of an intermediate can become sufficiently fast so that this
pathway is then indistinguishable from pathways that avoid the intermediate.
Analogous to effects on self-splicing constructs, higher protein concentrations can
inhibit folding of ribozymes to the native state (Figure 3.5C). The physical processes
responsible for inhibition are presumably the same for ribozymes and self-splicing
constructs, but the discontinuous assay allows the origins of the inhibition to be
distinguished. For self-splicing constructs, inhibition by trapping protein-bound non-
functional intermediates and by unfolding native RNA both lead to a decrease in the
observed splicing rate (Figure 3.2C). In a discontinuous assay unfolding of native
ribozyme decreases the endpoint of the folding curve (Figure 3.5C). Trapping of folding
intermediates, without unfolding of natively-folded RNA, would result in a decrease in
the rate of native ribozyme formation but not a decrease in the endpoint.
3.4 OTHER APPLICATIONS OF THE DISCONTINUOUS ASSAY
The discontinuous assay is amenable to a diverse set of experiments. It can be
used to monitor a decrease in the native ribozyme, and an additional stage can be
included to probe the role of ATP in chaperone-mediated folding (see Section 4.3.5) (55,
142). Further, the progress of native ribozyme formation obtained from the assay can also
complement insight from other powerful physical approaches, and thus provide a more
complete understanding of the action of RNA chaperones.
69
3.4.1 Unfolding native structure
Many chaperones function non-specifically and are capable of disrupting the
native states of RNAs as well as misfolded states (26). For probing the mechanisms of
chaperone activity in structure disruptions, it can be very useful to monitor the native
state because it is relatively homogeneous and the structure may be known. In contrast,
folding intermediates may be heterogeneous and their structure poorly defined.
To monitor loss of the native ribozyme, the ribozyme is first pre-folded to the
native state, and then the chaperone protein is added. Even for general chaperones, the
level of activity may be reduced for the native structure because it is typically highly
stable, so it may be necessary to lower the Mg2+
concentration and/or to use relatively
high protein concentrations to detect net unfolding. A decrease in the fraction of native
ribozyme over time indicates that the chaperone protein has mediated at least partial
unfolding of the native ribozyme, giving intermediates that do not readily refold to the
native state upon transfer to the stage 2 conditions. To ensure that the loss of native
ribozyme reflects reversible unfolding, the protein should be proteolyzed and the
ribozyme again folded to the native state (Figure 3.5D and see Section 4.3.6) (142).
3.4.2 Integrating results with other methods
The materials and methods required to employ the use of RNA catalytic activity
are standard for many laboratories, and the ease of implementing this method makes it a
convenient tool for obtaining a kinetic view of the fraction of native ribozyme. Results
can be highly complementary to those from methods that provide physical information on
folding intermediates (see Chapter 5).
Three physical probes that have been used extensively for RNA folding studies
are chemical footprinting, small angle X-ray scattering (SAXS), and single-molecule
FRET. Time-resolved chemical footprinting can be performed with several different
70
probes and provides a highly specific view of nucleotides that are engaged in secondary
or tertiary contacts during a folding process (153-158). Hydroxyl radical footprinting has
been particularly valuable for probing structured intermediates and elucidating folding
pathways (159-161). The orthogonal information provided by catalytic activity
measurements – how much of the ribozyme is in the native state – is tremendously
valuable because it can be used to place constraints on the folding pathways modeled
from footprinting data (161, 162). SAXS provides rich information on the overall size
and shape of RNA as it folds, which is highly complementary to footprinting (163, 164),
and again coordinated activity measurements under the same conditions can assist greatly
in constraining physical descriptions of intermediates (136, 161-163, 165). Last, single-
molecule FRET experiments are uniquely powerful for detecting and characterizing
intermediates that do not accumulate in bulk experiments (140, 166-170), and the
concurrent detection of catalytic activity can be used to tremendous advantage for
distinguishing the native state from folding intermediates (167-169, 171).
71
Figure 3.1: Group I and group II introns
(A) Self-splicing reactions. Left panel, Group I intron splicing reaction. An exogenous
guanosine attacks the 5´ splice site in the first step, and the 5´ exon attacks the 3´ splice
site in the second step. Right panel, Group II intron splicing reaction. A bulged adenosine
near the 3´ end of the intron attacks the 5´ splice site, generating a lariat intermediate. The
5´ exon then attacks the 3´ splice site to ligate the exons together. (B) Secondary structure
of the Azoarcus group I intron ribozyme. The nine-nucleotide substrate is lowercase. The
cleavage site is indicated by a thin arrow, and two tertiary contacts are indicated by thick
arrows. (C) Secondary structure of the aI5γ group II intron from S. cerevisiae (131). The
domains shown in black are present in the D135 ribozyme. Tertiary interactions are
indicated by Greek letters. Interaction sites between exon and intron sequences are
indicated by the abbreviations IBS and EBS (Intron Binding Site and Exon Binding Site,
respectively). The 24-nt substrate is shown at the right and the cleavage site is indicated
by an arrow.
72
Figure 3.2: Self-splicing constructs
(A) Denaturing gel showing splicing products for a group II intron: (from top to bottom)
lariat intron, unspliced precursor, linear intron, and spliced exons. (B) Simulated plot of a
splicing reaction showing the fraction of precursor as a function of time. The simulated
data are fit by a single exponential curve to obtain rate constants for splicing in the
presence (circles) and absence (diamonds) of protein. (C) Simulated plot showing how
the observed splicing rate varies with protein concentration. The rising linear portion of
the data (circles) is fit with a line to obtain a second order rate constant for chaperone-
accelerated folding. The plateau and decrease in rate (diamonds in gray area) reflect
inhibition by the chaperone at higher concentrations (see Section 3.2.1).
73
Figure 3.3: The discontinuous assay
(A) Reaction schematic. The ribozyme folds in the first stage, and then it cleaves the
oligonucleotide substrate (S) in the second stage under reaction conditions that block
further native folding. (B) Denaturing gel showing the results of cleavage of a 5´ labeled
oligonucleotide substrate by native ribozyme. (C) Simulated plot of multiple cleavage
reactions representing different folding times prior to the cleavage reaction. The curves
with larger bursts represent longer folding times, giving greater accumulation of native
ribozyme. The amplitude values are shown with filled symbols. (D) Simulated plot
showing the fraction of native ribozyme (fN) plotted as a function of folding time (t1).
The burst amplitudes from simulated cleavage reactions in panel C are plotted as a
function of folding time. In this simulated scenario, the folding progress can be fit by a
single exponential function, giving a single rate constant for native state formation.
74
Figure 3.4: Examples of catalytic reactions with the D135 and Azoarcus ribozymes
(A) Identifying conditions that block native folding in the catalysis stage of the
discontinuous assay. Comparing burst amplitudes resulting from both D135 ribozyme
placed into stage 2 conditions with (circles) or without (triangles) a prior incubation in
Mg2+
-containing buffer to allow prefolding to the native state. The burst is much smaller
without the preincubation, indicating that these conditions for stage 2 (pH 8.0, 100 mM
Mg2+
, 500 mM KCl, 15 °C) effectively block folding. The solid circle indicates how a
single time point can be sufficient to determine the fraction of native ribozyme if the
kinetics of cleavage are known. (B) Ribozyme prefolding to the native state. The burst
amplitudes from cleavage reactions of the Azoarcus ribozyme are smaller than they
would be for stoichiometric product formation, and further work showed that this results
from an equilibrium between substrate cleavage and ligation (136). Note also that the
bursts from ribozyme prefolded at 37 °C and 10 mM Mg2+
for 15-45 min are identical,
suggesting that 15 minutes is sufficient for complete folding (see Section 4.1). Panel A
reprinted from (142) with permission from Elsevier. Panel B adapted from (136). This
research was originally published in Journal of Biological Chemistry. Selma Sinan,
Xiaoyan Yuan, and Rick Russell. The Azoarcus Group I Intron Ribozyme Misfolds and
Is Accelerated for Refolding by ATP-dependent RNA Chaperone Proteins. JBC. 2011;
286:37304-37312. © the American Society for Biochemistry and Molecular Biology
75
Figure 3.5: The discontinuous assay with the D135 ribozyme and the DEAD-box
helicase Mss116p
(A) Cleavage time courses initiated with ribozyme folded in the presence of Mss116p for
the times indicated. The increase of the burst amplitude with folding time (t1) indicates
productive folding to the native state. (B) Comparison of ribozyme folding in the
presence (diamonds) and absence (circles) of Mss116p. The protein changes the folding
profile from multi-phasic to a single exponential phase. (C) High Mss116p concentrations
inhibit the native conformation. At 500 nM (diamonds) and 1000 nM (inverted triangles)
Mss116p, the fraction of native ribozyme at steady state is lower than in the presence of
lower protein concentrations (gray symbols). (D) The native conformation of D135 can
be unfolded by Mss116p. Ribozyme prefolded to the native state (black/gray solid circle)
is divided into reaction tubes containing a high concentration of Mss116p (black circles)
or no Mss116p (gray circles). After approximately 10 min, the protein is proteolyzed and
the ribozyme is treated to allow it to refold to the native state (black and gray diamonds),
demonstrating that the loss of native ribozyme arises from unfolding and not an
irreversible process. Panels A-D adapted from (142) with permission from Elsevier.
76
The text below and following figures of chapter 4 were originally published by Elsevier.
Copyright © by Elsevier 2011.
Potratz JP, Del Campo M, Wolf RZ, Lambowitz AM, & Russell R (2011) ATP-
Dependent Roles of the DEAD-Box Protein Mss116p in Group II Intron Splicing In Vitro
and In Vivo. J Mol Biol 411(3):661-679.
Mark Del Campo performed experiments with proteins and self-splicing constructs.
Rachel Wolf performed the in vivo experiments.
Mark and Rachel are both in the Alan. M. Lambowitz lab.
Rick Russell and Alan M. Lambowitz helped analyze the data and write the paper.
Chapter 4: ATP-dependent roles of the DEAD-box protein Mss116p in
group II intron splicing in vitro and in vivo
4.1 INTRODUCTION
Autocatalytic group I and group II introns have proven to be valuable model
systems for understanding RNA folding, structure, and function (14, 172-175). The
mitochondrial (mt) genome of S. cerevisiae encodes nine group I introns and four group
II introns, all of which require the DEAD-box protein Mss116p for efficient splicing in
vivo (91). These introns differ substantially in their structural features and global
architectures, suggesting that the roles played by Mss116p in their splicing reflect non-
specific interactions with RNA. Further, other DEAD-box proteins are able to mitigate
the defects from loss of functional Mss116p (30, 55, 81, 176). These proteins include the
cytoplasmic S. cerevisiae protein Ded1p, which functions in cellular compartments that
lack group I and II introns and therefore does not function naturally in folding of these
introns.
The mechanism by which DEAD-box proteins promote group I intron splicing
has been analyzed biochemically. Mss116p and its Neurospora crassa homolog CYT-19
77
interact with cognate and non-cognate group I introns and accelerate conformational
transitions, including those from kinetically-trapped, misfolded intermediates to the
native states (26, 28, 31, 55, 177). These observations and the established propensity of
group I introns to misfold (14, 20, 178) have led to models in which the principal
function of DEAD-box proteins in group I intron folding is to disrupt structure non-
specifically, allowing misfolded intermediates additional opportunities to fold to the
native state (20, 25, 28, 81, 143).
Analogous models have been proposed for Mss116p and CYT-19 in group II
intron folding (30, 81, 82, 91, 175). Group II introns consist of six domains with a
complex set of local and long-range tertiary contacts that generate a functional structure
with an active site for splicing, which is in many cases stabilized by the binding of
specific proteins (175, 179-181). To understand how Mss116p functions, attention has
focused on the yeast mtDNA group II introns that are its natural substrates. Two of these
introns, aI1 and aI2, are closely related group IIA introns and encode maturase proteins,
which are required for structural stabilization during RNA splicing. In the absence of
Mss116p in vivo, unspliced precursor RNA accumulates in a complex with the maturase,
suggesting that Mss116p can function at a step after stable maturase binding,
hypothesized to be the resolution of kinetic traps (91). The other two yeast mt group II
introns, aI5 and bI1, are small subgroup IIB introns that do not encode maturases, and
their splicing is accelerated by Mss116p and CYT-19 in vivo and under near-
physiological conditions in vitro (30, 55, 91, 176). Although the specific folding steps
accelerated by Mss116p remain to be established, biochemical studies showed that
Mss116p functions on bI1 as an RNA chaperone, as it promotes ATP-dependent
formation of an active intron structure and is then dispensable for activity (30).
78
The mechanisms of acceleration for the remaining yeast group II intron, aI5,
have been the subject of debate. This intron was the first for which self-splicing via lariat
formation was demonstrated (182) and thereafter has been used as a model for RNA
folding and catalysis (174). Early studies used a shortened version of the intron termed
D135, which lacks exons and some intron domains and functions as a ribozyme by
cleaving an RNA oligonucleotide substrate, and were done at elevated temperature and
high ion concentrations (42 °C, 100 mM Mg2+
, 500 mM KCl). Under these conditions,
hydroxyl radical footprinting and catalytic activity measurements suggested concerted
folding in 1-2 min (0.6 min–1
) (183, 184). The denaturant urea did not accelerate folding,
consistent with the absence of a rate-limiting kinetic trap.(183)
In contrast, aI5 splicing is accelerated by Mss116p and other DEAD-box
proteins in vitro under near-physiological conditions (30 °C, 8 mM Mg2+
, 100 mM KCl)
(30, 55, 60, 176). Under these conditions, a native gel shift assay showed that folding of a
D1356 ribozyme derivative to compact species is slower and more complex, giving as
many as four kinetic phases with rate constants spanning at least three orders of
magnitude (>1 min–1
to <10–3
min–1
) (137, 138). The multiple phases suggest multiple
pathways and rate-limiting steps, and the time scale of hours for major populations would
be surprising for intramolecular diffusive processes. Nevertheless, the compaction rates
of the slow, dominant phases are unaffected by urea, suggesting that if there are rate-
limiting kinetic traps, they do not require substantial regions of buried RNA to become
exposed to solvent in the transition state ensemble (137).
Starting with the conclusion that the folding of aI5γ is not rate-limited by the
resolution of kinetic traps, Solem et al. suggested that Mss116p and other DEAD-box
proteins promote splicing of aI5 without unwinding RNA by binding and stabilizing an
on-pathway intermediate required for intron compaction (176). A central piece of
79
evidence was the behavior of an Mss116p mutant in which the SAT sequence of motif III
is mutated to AAA (abbreviated SAT/AAA) (185). This mutant was reported to promote
in vitro splicing of aI5 -wild-type efficiency but to be inactive in unwinding
of moderately-long RNA duplexes (12–18 bp) (176). A subsequent analysis, however,
revealed that this mutant retains residual ATP-dependent unwinding activity for shorter
RNA duplexes (6–10 bp), of a length more representative of the helices in group II
introns, and that the decrease in unwinding efficiency can be as large as the decrease in
splicing activity under the same solution conditions (8–28-fold) (60). Thus, this mutant
did not provide experimental support for the suggestion that Mss116p promotes aI5γ
splicing without unwinding RNA (60).
The experiments above used a standard construct of aI5, which includes the 887-
nt intron and relatively long exons (~300 nt, termed LE construct). Recently, the same
authors compared splicing of the LE construct and a construct containing short exons of
<30 nt (SE). They concluded that Mss116p functions on the LE construct as an ATP-
dependent RNA chaperone to resolve kinetic traps involving exons but promotes splicing
of the SE construct by binding and stabilizing an on-pathway intermediate (149). A key
basis for this conclusion was the finding that although the SAT/AAA mutant was
compromised in promoting splicing of the LE construct, it appeared to be as efficient as
wild-type Mss116p for the SE construct. Other work showed that Mss116p accelerates
compaction of an RNA that consists only of aI5 domain I (DI), (138) suggesting that this
step is accelerated during folding of the SE construct (138, 176). However, acceleration
of DI compaction is ATP-independent, whereas acceleration of splicing for the SE
construct, like the LE construct, requires ATP. This difference suggests either that one or
more other folding steps for the SE construct require ATP-dependent RNA unwinding or
that ATP hydrolysis is required to accelerate dissociation of Mss116p following an ATP-
80
independent role (138, 149). Finally, recent single molecule studies with the D135
ribozyme revealed that Mss116p-promoted folding involves an initial ATP-independent
step, presumably compaction of DI, and at least one later ATP-dependent step (140). The
authors favored the interpretation that this later step involves dissociation of bound
Mss116p to permit further RNA folding but left open the possibility of ATP-dependent
RNA unwinding to resolve a kinetic trap.
To delineate the roles of Mss116p in aI5 splicing, here we compare Mss116p-
promoted folding of three constructs, all under the same near-physiological conditions. In
addition to the standard LE construct, with 5 and 3 exons of 293 and 321 nt,
respectively, (186) we use a version with greatly shortened exons of 28 and 15 nts (SE
construct), (138, 149, 150) and the D135 ribozyme (Figure 4.1) (131). We show that the
ability of Mss116p to promote folding is ATP-dependent for all of the constructs, and
SAT/AAA and other mutants that are deficient in RNA-unwinding activity are deficient
in promoting splicing of both the SE and LE constructs. Catalytic activity measurements
with the D135 ribozyme and additional splicing assays with the SE and LE constructs
indicate that a major role of ATP is to promote the disruption of RNA structure by
Mss116p. Last, the relative abilities of Mss116p mutants to support ATP-dependent
splicing and RNA unwinding in vitro correlate well with their abilities to support aI5γ
splicing in vivo in a strain that lacks all other mt group I and group II introns. Together
our results indicate that the physiological function of Mss116p in aI5 splicing includes a
critical role for ATP-dependent RNA unwinding to resolve inactive structures.
81
4.2 MATERIALS AND METHODS
4.2.1 Recombinant Plasmids
pMAL-Mss116p, used to express Mss116p from E. coli for biochemical studies,
contains the Mss116p coding sequence (codons 37–664) with an in-frame N-terminal
MalE fusion cloned downstream of a tac promoter in the expression vector pMAL-c2t
(55, 81). pHRH197, used to express Mss116p in S. cerevisiae for genetic analysis,
contains the MSS116 gene with its endogenous promoter cloned in the centromere-
containing (CEN) plasmid vector pRS416 (92). Sequences encoding Mss116p mutations
(K158A, K158R, S305A/T307A) were introduced into these plasmids by Quikchange
mutagenesis (Stratagene). pJD20 encodes the long-exon (LE) construct downstream of a
phage T7 promoter in Bluescribe (186). The LE construct is a 1501-nt precursor RNA
that includes a 293-nt 5 exon, the 887-nt aI5 intron, and a 321-nt 3 exon. The 5 exon
consists of 20 nt derived from the vector (plus three guanosines from the T7 promoter)
and 270 nt of COX1 5-exon sequence, and the 3 exon consists of 291 nt of COX1 exon
sequence followed by 30 nt of vector sequence. Plasmid pUC19::aI5-SE, which encodes
the SE construct, was created by PCR amplification of pJD20 using the primers 5-
TAATACGACTCACTATAGGGACTTACTACGTGGTGGGAC-3 and 5-
TTGATAATACATAGTATCCCGATAGGTAGACC-3, which added a T7 promoter
(underlined). The resulting PCR product was re-amplified with the primers 5-
GCCCcatatgTAATACGACTCACTATAGGG and 5-
GGGCaagcttAATACATAGTATCCCGATAGG to add NdeI and HindIII sites
(lowercase), and cloned between the corresponding sites of pUC19. The SE construct is a
930-nt precursor RNA that includes a 28-nt 5 exon, the 887-nt aI5 intron, and a 15-nt 3
exon. The 5 exon consists of three guanosines from the T7 promoter and 25 nt of COX1
82
5-exon sequence, and the 3 exon consists of 11 nt of COX1 exon sequence followed by
4 nt of vector sequence. pQL71 encodes the D135 ribozyme (see Figure 4.1B) (183).
4.2.2 RNA preparation
The LE and SE RNA constructs were transcribed in vitro from HindIII-digested
plasmids by T7 RNA polymerase in the presence of [α-32P]-UTP (Perkin Elmer) using a
Megascript kit (Ambion). RNA was isolated by phenol-chloroform extraction and size
exclusion chromatography using two consecutive G-50 columns. D135 RNA was
transcribed in vitro from HindIII-digested pQL71 using T7 RNAP and purified via an
RNeasy column (Qiagen). The RNA oligonucleotide substrate for D135
(CGUGGUGGGACAUUUUCGAGCGGU) was 5-end-labeled with [α-32P]-ATP
(Perkin Elmer) by using T4 polynucleotide kinase (New England Biolabs). RNA
concentrations were determined by specific activity of the [α-32P]-UTP precursor or
spectrophotometrically using extinction coefficients at 260 nm of 1.76 107 M
–1 cm
–1 for
the LE construct, 1.17 107 M
–1 cm
–1 for the SE construct, 5.86 10
6 M
–1 cm
–1 for the
D135 construct, and 2.36 105 M
–1 cm
–1 for the D135 substrate oligonucleotide.
4.2.3 Preparation of Mss116p
Wild-type and mutant versions of Mss116p were expressed and purified as
described (55, 60). After purification, protein was dialyzed overnight against storage
buffer solution (20 mM Tris-Cl, pH 7.5, 500 mM KCl, 1 mM EDTA, 1 mM DTT, 50%
glycerol), flash frozen, and stored at –80 °C.
4.2.4 Splicing reactions
Splicing reactions were performed in a thermal cycler in 20 or 50 µl (50 mM Na-
MOPS, pH 7.0, 100 mM KCl, 8 mM MgCl2, and 5% glycerol). The RNA concentration
was 20 nM for reactions shown in Figure 4.2 and 4.3, and it was 1 nM for reactions in the
83
presence of Mss116p (Figure 4.4, 4.6, and 4.7). Control reactions at the lower
concentration in the absence of Mss116p gave rate constants within 2-fold of those at the
higher concentration (Figure 4.4A and data not shown). When indicated, reactions also
included 1 mM ATP, added as a stoichiometric complex with Mg2+
. RNA was heated
briefly in the absence of Mg2+
(92 °C, 1-2 min), cooled rapidly, and then splicing
reactions were initiated by adding RNA to a pre-incubated tube containing splicing buffer
solution or by adding RNA and then MgCl2. Portions of reactions were quenched at
various times by adding 4 µl of 100 mM EDTA to 2 µl of reactions in Figure 4.2 and 4.3
or by adding 5 µl of 50 mM EDTA, 0.1% SDS, 1 mg/ml proteinase K to 3 µl of reactions
in Figure 4.4, 4.5, 4.6, and 4.7. Splicing products were separated on a denaturing 4%
polyacrylamide gel and quantified using a phosphorimager and ImageQuant TL (GE
Healthcare). Rate constants were obtained by fitting the time-dependent decrease in
precursor RNA, relative to reaction products, either by an exponential decay or by a line
for slow reactions of the LE construct in the absence of Mss116p (Kaleidagraph, Synergy
Software). In the latter case the rate constants were inferred from the initial splicing rate.
Unless otherwise indicated, rate constants are reported as the average and standard error
from 2-4 independent determinations. In the presence of higher concentrations of
Mss116p, some splicing reactions included a lag phase. The lag was included in the
analysis by allowing the y-intercept of a single exponential equation to vary. Inclusion or
exclusion of the lag phase did not significantly affect the rate constant for the slower,
major phase of splicing. The origin of the lag is unclear, but its appearance at higher,
inhibitory Mss116p concentrations suggests that it originates from a molecular process
associated with inhibition by Mss116p.
84
4.2.5 Discontinuous catalytic activity assay for D135 RNA folding
The discontinuous activity assay for monitoring D135 ribozyme folding consisted
of two stages. In stage 1, D135 ribozyme (50 nM) was allowed to fold for various times.
The ribozyme was first denatured at 90 °C for 1 min in buffer solution containing 50 mM
Na-MOPS, pH 7.0, and then folding was initiated by addition of D135 to a stage 1
reaction (30 °C, 50 mM Na-MOPS, pH 7.0, 100 mM KCl, and 8 mM MgCl2 unless
otherwise indicated). ATP was also present (1 mM) when indicated, added as a
stoichiometric complex with Mg2+
. In stage 2, substrate cleavage was monitored by
diluting portions of the stage 1 folding reaction 5-fold into a solution with higher Mg2+
concentration and pH and lower temperature (15 °C, 500 mM KCl, 100 mM MgCl2, 80
mM HEPES, pH 8.1, 1 mg/ml proteinase K). This solution also included the radiolabeled
oligonucleotide substrate (30 nM, 3-fold in excess of D135 ribozyme after dilution).
Time points (2 µl) were quenched by adding 4 µl of 100 mM EDTA, and the substrate
and product were separated in a denaturing 20% polyacrylamide gel.
Cleavage time courses were fit by an exponential phase followed by a linear
phase, with the amplitude of the exponential phase reflecting the fraction of native D135
ribozyme (134). After early experiments established the rate constants of the fast and
slow phases, the burst amplitude was determined from a single time point at 300 min
(119). This time was chosen because it allows completion of the fast phase while
minimizing the contribution from the slow phase. Burst amplitudes were normalized and
scaled to reflect the fraction of the population that was native at the time of transfer to
stage 2. First, the raw burst amplitude was corrected by subtracting an amount
representing the fraction of unfolded molecules that fold rapidly in stage 2 (~0.15 under
standard conditions). This value was then divided by the value reflecting full native
folding, determined in reactions with pre-folded ribozyme, after subtracting an equivalent
85
amount (e.g. 1 – 0.15 = 0.85). Folding rate constants are reported as the average and
standard error from at least two independent determinations.
4.2.6 S. cerevisiae Northern hybridizations and immunoblotting
The S. cerevisiae wild-type strain 161/U7-aI5γ is a derivative of 161-U7 MATa
ade1 lys1 ura3, in which the mtDNA contains a single intron aI5 (91). In the isogenic
strain mss116Δ-aI5, the MSS116 gene was replaced by a kanr cassette and the resident
mtDNA was replaced by cytoduction with mtDNA containing only aI5 (91). Wild-type
and mutant versions of Mss116p with a C-terminal myc tag were expressed from the
CEN plasmid pHRH197, which carries a ura3 marker (92). Transformants of the
mss116Δ-aI5 strain containing the CEN-plasmids were selected by plating on 2% agar
plates containing 2% dextrose and Hartwell’s complete (HC) media (1X YNB solution,
1X HC dropout 6 amino acid solution, plus 1% each of lysine, tryptophan, and histidine;
0.1% adenine; and 2% leucine) lacking uracil (187).
For Northern hybridizations, cells were grown at 30 °C to O.D.600 of 1.0 – 1.6 in
Hartwell’s Complete liquid medium containing 2% raffinose and lacking uracil. RNA
was isolated as previously described (92). Samples containing 1.0 µg of RNA were
denatured by incubating with 5.6% glyoxal in 50% DMSO, 0.1 M NaPO4 at 65°C for 15
min and electrophoresed in a 1.5% agarose gel with RNA-grade 1X TAE (40 mM Tris-
acetate, pH 8, 1 mM EDTA) at 25°C. The gels were blotted onto a nylon membrane
(Hybond-XL, GE Healthcare) overnight, hybridized with a 5′-end-labeled DNA
oligonucleotide probe complementary to COX1 exon 6
(GAATAATGATAATAGTGCAAATGAATGAACC), and scanned with a
phosphorimager.
86
For immunoblotting, cells were grown as above, and proteins were precipitated
with trichloroacetic acid as described, (188) except that the resulting protein pellets were
washed once with 0.5 ml of 1 M Tris base before being resuspended in 150 µl of SDS-
PAGE sample buffer. Samples (~60 µg of protein measured by O.D.280) were run in a
pre-cast 4-20% polyacrylamide gradient gel (BioRad) with 0.1% SDS in the running
buffer and transferred to a Sequi-BlotTM PVDF membrane (BioRad) using a BioRad
Criterion blotter apparatus. The blot was probed with anti-Mss116p guinea pig primary
antibody (1:5,000 dilution), (185) developed using an ECL Plus Western Blotting-kit (GE
Healthcare), and imaged using Kodak Biomax XAR film. To confirm equal loading of
protein samples, the PVDF membrane was stripped of antibodies using RestoreTM Plus
Western Blot Stripping Buffer (ThermoScientific) and stained with AuroDye Forte (GE
Healthcare) following the manufacturer’s directions.
4.3 RESULTS
4.3.1 Splicing of LE and SE constructs and Mss116p acceleration
To probe how exons influence the mechanisms of Mss116p-promoted folding of
the aI5γ intron, we first compared self-splicing of the LE and SE constructs. Under near-
physiological conditions in vitro, splicing was considerably faster for the SE construct,
consistent with a recent report using essentially the same constructs and conditions
(Figure 4.2A and Figure 4.3) (149). At 30 °C, the rate constant for the SE construct
determined from the disappearance of precursor and appearance of lariat and linear intron
was 1.4 (± 0.5) 10–3
min–1
, whereas splicing of the LE precursor was 80-fold slower
(1.8 (± 0.5) 10–5
min–1
). Both constructs spliced faster at higher temperatures, with the
LE construct displaying a greater temperature dependence (Figure 4.2B). Thus, one or
both long exons slow splicing and add to the enthalpic barrier, consistent with previous
87
findings that exon sequences can strongly affect aI5γ (150). Figure 4.2B also
shows that the splicing rates of both constructs level and begin to decrease with
increasing temperature, suggesting that the active structures are unstable at higher
temperatures. It is notable that this transition occurs at higher temperature for the LE
construct, suggesting that the longer exons stabilize the native structure in addition to
slowing its formation.
We then measured Mss116p-accelerated splicing of the SE construct. Consistent
with a recent report, (149) the concentrations of Mss116p used previously for the LE
construct (60, 81) gave strong inhibition with the SE construct, and we therefore used
lower concentrations of Mss116p and RNA (1 nM RNA). For comparison, we performed
experiments with the LE construct under these conditions. Splicing of both constructs
was stimulated by low concentrations of Mss116p, just slightly in excess of RNA, in the
presence of ATP (Figure 4.4A and Figure 4.5). In general, the time courses were well
described by single rate constants, with endpoints of >90% for reactions that reached
completion within the observation time (Figure 4.4B). Both constructs required ATP for
maximal stimulation, although SE construct splicing was weakly stimulated in the
absence of ATP (Figure 4.4B). Analogous ATP-independent stimulation was observed
previously for group I intron splicing by Mss116p (55) and at a very low level for the
aI5γ LE construct by the E. coli DEAD-box protein SrmB (81).
As a quantitative benchmark for comparison, we measured the Mss116p
concentration dependences (Figure 4.4C). In earlier work with higher RNA and protein
concentrations, acceleration of the LE construct by Mss116p and other DEAD-box
proteins gave sigmoidal dependences, suggesting roles for multiple protomers, (81)
although the upward curvature was more obvious for the other proteins and had
previously gone undetected for Mss116p (60). Here, across the limited range of
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stimulatory protein concentrations, the dependence was approximately linear, giving a
second-order rate constant of 4.7 106 M
–1 min
–1. Although the possibility of a higher-
order dependence cannot be excluded, for simplicity we use the linear dependence for
comparison with the SE construct and Mss116p mutants (see Section 4.5).*
We found that Mss116p stimulates splicing of the SE construct at least as
efficiently as it stimulates splicing of the LE construct (>4.2 106 M
–1 min
–1, Figure
4.4C) (60). Only a lower limit could be determined for the SE construct because even at
the lowest attainable protein concentration, the splicing rate was not readily distinguished
from the maximal value. This plateau for the SE construct results from strong inhibition
of splicing by Mss116p at concentrations as low as 4 nM, considerably lower than for the
LE construct (Figure 4.4B,C and Figure 4.5) (60). Together, these results indicate that the
long exons cause or exacerbate slow folding, Mss116p stimulates splicing of both the LE
and SE constructs, and that maximal Mss116p stimulation of both constructs requires
ATP.
4.3.2 Mutants that are deficient in RNA-unwinding activity
To explore further the properties of Mss116p required for acceleration of aI5γ
splicing, we tested three mutants compromised in ATP-dependent RNA unwinding for
stimulation of splicing of the LE and SE constructs. The SAT/AAA mutant gave an
efficiency of 4.2 105 M
–1 min
–1 for the LE construct (Figure 4.6), 10-fold lower than
wild-type Mss116p (WT) and within the range of relative rate constants found previously
(60). For the SE construct, the efficiency for the SAT/AAA mutant was 8.9 105 M
–1
min–1
, at least 4- to 5-fold lower than WT (a minimum estimate because, as noted above,
we could only determine a lower limit for WT). Thus, the decrease in efficiency for the
SAT/AAA mutant with the SE construct is comparable and could be even larger than for
89
the LE construct. Acceleration of the SE construct by Mss116p SAT/AAA is ATP-
dependent, with little or no ATP-independent acceleration up to at least 10 nM protein
(Figure 4.6B). Additionally, two mutants in motif I, K158A and K158R, which are
defective in ATP binding and hydrolysis and in RNA unwinding, (176) failed to stimulate
splicing of either the LE or SE construct at any concentration tested (≤15 nM; Figure
4.7).
These results show that mutations that decrease RNA-unwinding activity
commensurately reduce the ability of Mss116p to stimulate splicing of both the SE and
LE constructs. Our conclusion that the SAT/AAA mutant is similarly deficient in splicing
of both constructs disagrees with the conclusion but not the data from the earlier work
(149) (see Discussion Section 4.4). The similar requirements for the SE and LE
constructs could reflect that Mss116p accelerates a common folding step of these two
constructs or that different folding steps coincidentally give similar dependences on
Mss116p and ATP.
4.3.3 Two-stage, discontinuous catalytic activity assay for folding of D135 RNA
Previously, the folding of the D135 ribozyme was measured by catalytic activity
using a continuous assay, in which folding and substrate cleavage occur simultaneously,
(184) but this assay can give information on the folding rate only under conditions in
which the rate of substrate cleavage exceeds that of folding (e.g. 100 mM Mg2+
at 42 °C)
(183). To follow folding at lower Mg2+
concentrations, where catalytic activity is weaker,
we designed a two-stage, discontinuous activity assay (Figure 4.8A). Folding takes place
in stage 1, and then portions of the folding reaction are transferred at various times into a
second set of conditions (stage 2). This second set of conditions must allow substrate
cleavage by ribozyme that has already folded to the native state at the time of transfer but
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inhibit the remainder of the population from reaching the native state on the time scale of
the cleavage reaction (189).
Based on previous work on group I intron ribozymes, (17, 119, 121) we tested
whether low temperature and high Mg2+
concentration would provide suitable conditions
for stage 2. When D135 was prefolded to the native state under established conditions (42
°C, 100 mM Mg2+
, 20 min), then transferred to 15 °C with 100 mM Mg2+
, and a small
excess of substrate was added, an initial phase of substrate cleavage gave a rate constant
of 8.7 (± 0.4) 10–3
min–1
and an amplitude approximately equal to one turnover of the
ribozyme (Figure 4.8B). Cleavage of the remainder of the substrate was slower,
presumably because at least one of the cleavage products is released slowly from the
ribozyme and limits subsequent turnovers. The 5 portion of the substrate base pairs with
exon-binding site (EBS) 1 and 2 of the ribozyme and is therefore a strong candidate for
slow release.
Although a cleavage rate constant of ~9 10–3
min–1
indicates a rather slow
reaction at 15 °C, it is within the expected range considering that the reaction proceeds at
1 min–1
under the same solution conditions at 42 °C. We reasoned that 15 °C and 100
mM Mg2+
could be suitable for stage 2 if folding to the native state under these
conditions is even slower than substrate cleavage by the native ribozyme. Indeed, we
found that ribozyme that was transferred directly from buffer solution in the absence of
Mg2+
into stage 2 gave substantially less product formation (Figure 4.8B, termed
‘nonfolded’ control), indicating that most of the ribozyme did not fold to the native state
on the time scale of the cleavage reaction. The low temperature of stage 2 was critical, as
a parallel experiment in which stage 2 was 30 °C gave minimal difference between the
prefolded and nonfolded reactions (Figure 4.9). Nevertheless, even at 15 °C a small
amount of product (15%) appeared in the nonfolded control with roughly the same rate
91
constant as product in the prefolded reaction (Figure 4.8B). This product formation most
likely reflects a subpopulation of the ribozyme that was able to fold rapidly to the native
state in stage 2. In subsequent experiments, we normalized the data to account for this
subpopulation (see Section 4.2.5).
We next measured the kinetics of native folding in stage 1 by incubating the D135
ribozyme for various times and then transferring portions into stage 2 and measuring the
burst amplitude from time courses of substrate cleavage (see Figure 4.8A). As an initial
control, we monitored folding under conditions similar to those used previously in the
continuous activity assay (42 °C, 100 mM Mg2+
). As expected, this experiment gave
essentially the same result as the continuous assay, with complete folding to the native
state occurring with a rate constant of 0.5 ± 0.1 min–1
(Figure 4.8C) (183). As indicated
above, data were normalized by the fraction of ribozyme that reached the native state
rapidly upon transfer from buffer solution in the absence of Mg2+
, which was larger when
the ribozyme was transferred from 42 °C (Figure 4.10).
We next measured folding of D135 at the near-physiological conditions used for
the LE and SE constructs. Under these conditions, folding was slower and more complex,
with at least two phases (Figure 4.8D). A minor fast phase gave a rate constant of 1.0 (±
0.2) min–1
, and a second phase gave a rate constant of 1.4 (± 0.6) 10–3
min–1
with an
apparent endpoint of ~50-60% native ribozyme. These results are crudely consistent with
those from a published gel-shift assay used to measure compaction of the related D1356
ribozyme under similar conditions, which gave two observed phases (0.16 min–1
and
0.006 min–1
), an unresolved fast phase (>1 min–1
), and a slower phase (~10–3
min–1
,
complete in 24 h) (138). It is not clear whether the apparent endpoint in our experiment
reflects equilibration of native and non-native forms or whether an unobserved third
phase would give still more native RNA at longer folding times. Regardless, the results
92
show that under the near-physiological conditions, D135 folds in a complex, multi-phasic
process that differs from the single-phase folding at higher temperature and Mg2+
concentrations (183, 184, 190).
4.3.4 Acceleration of D135 folding by Mss116p
We next used the discontinuous activity assay to probe the effects of Mss116p on
D135 ribozyme folding under near-physiological conditions. Addition of Mss116p and
ATP with Mg2+
gave rapid accumulation of the native ribozyme (Figure 4.11A), reducing
the folding process to a single phase with a rate constant of 0.61 ± 0.07 min–1
and an
apparent endpoint of 0.76 ± 0.04 (Figure 4.11B). While this rate was slightly slower than
that for the minor fast phase without Mss116p, the major effect of Mss116p was to
increase the amplitude of the rapid native state formation. The rate constant did not
change systematically across the accessible range of Mss116p concentrations (Figure
4.12A), preventing determination of a second-order rate constant for acceleration.
However, as observed with the LE and SE constructs, higher Mss116p concentrations
were inhibitory. In contrast to the inhibition of splicing, which gave a decreased rate
constant, here the inhibition gave a decreased endpoint. This result suggested either that
higher concentrations of Mss116p can bind and trap folding intermediates, limiting the
extent of native ribozyme formation, or that Mss116p can unfold the native ribozyme,
generating a steady-state mixture of native and non-native ribozyme.
We then used the endpoint for the single fast phase as a diagnostic tool to
examine the effects of ATP and Mss116p mutations. The enhanced folding by Mss116p
was strongly dependent upon ATP, as 100 nM Mss116p by itself gave only a small
increase in endpoint relative to the fast phase of a reaction without protein (Figure 4.11B,
cf., with Figure 4.8D). As expected from the ATP requirement, the motif I mutants
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K158A and K158R also gave only small increases in endpoint (Figure 4.11C). The
SAT/AAA mutant was partially active, giving higher endpoints than the other mutants
but lower than WT Mss116p, especially at lower protein concentrations (Figure 4.11C
and 4.12B, cf., with Figure 4.11B). Because the rate constants did not depend
systematically on the concentration of Mss116p SAT/AAA, we could not determine
quantitatively whether the mutant is compromised for D135 folding relative to the WT
protein (Figure 4.12). Nevertheless, the lower endpoints suggest either that the mutant
protein is unable to promote folding of a fraction of the ribozyme population or that it
gives a lower steady-state level of native ribozyme.
4.3.5 The role of ATP in acceleration of D135 folding by Mss116p
Next we used the discontinuous assay to probe the role of ATP in Mss116p-
mediated folding. It has been suggested that for D135 and the splicing constructs,
acceleration of the critical folding step by Mss116p is ATP-independent, but ATP is
required to promote dissociation of Mss116p, allowing rapid folding to the native state
(138, 140, 149). The discontinuous activity assay afforded an incisive test of this model,
which predicts that Mss116p would be active in the absence of ATP if an alternative
means were provided to remove Mss116p prior to the determination of catalytic activity.
Thus, we incubated Mss116p with D135 and Mg2+
for various times in the
absence of ATP and then added proteinase K (1 mg/ml) to degrade Mss116p (Figure 4.13
and Figure 4.14). After additional incubation of up to 60 min to allow further folding of
any ribozyme that was ‘poised’ to form the native state following Mss116p dissociation,
we transferred aliquots to stage 2 and measured the fraction of native ribozyme by
activity. The prediction from the model above was that this reaction would give
accumulation of ribozyme that could quickly reach the native state upon removal of
94
Mss116p. However, with 100 nM Mss116p (2-fold excess over D135 ribozyme), removal
of Mss116p by proteolysis did not promote rapid native ribozyme formation (Figure
4.13A, closed symbols) beyond the level in an equivalent reaction without the proteolysis
step (open symbols). Further, inclusion of ADP or the non-hydrolyzable analog AMP-
PNP with Mss116p did not lead to significant native ribozyme formation after proteolysis
(Figure 4.15A). These results most simply suggested that the role of the ATPase cycle is
not solely to accelerate Mss116p dissociation.
We next considered an extension of the model in which the ATP requirement
stems from Mss116p being sequestered at non-productive sites within the RNA, requiring
multiple cycles of binding and release to bind productively. Thus, we increased the
Mss116p concentration so that sufficient protein would be available even after non-
productive sites were filled. With 400 nM Mss116p, 8-fold excess over D135, we
observed a variable increase in native ribozyme upon proteolysis (10-25% of the
population; Figure 4.13B, closed symbols, different colors indicate replicate
experiments), which reached an intermediate level between reactions with Mss116p in
the presence and absence of ATP (compare with Mss116p+ATP level of ~0.8 shown in
Figure 4.11B). As above, inclusion of ADP or AMP-PNP with Mss116p gave results that
were the same within error as those in the absence of added nucleotide (Figure 4.15B).
These results indicate that for a fraction of the ribozyme, ATP-independent activity of
Mss116p and removal by proteolysis is indeed sufficient to promote folding to the native
state. However, much of the ribozyme population remains non-native, most likely
because it forms additional intermediates that require further activity of Mss116p.56 Still
higher concentrations of Mss116p (up to 1200 nM) did not give more native ribozyme
upon proteolysis than observed with 400 nM Mss116p (data not shown).
95
The variable but significant increase in native D135 ribozyme after Mss116p
binding and proteolysis in the absence of ATP prompted us to investigate whether this
might also occur with the splicing constructs. Incubation of 1 nM RNA with 2 nM or 10
nM Mss116p in the absence of nucleotide, or in the presence of ADP or AMP-PNP,
followed by proteolysis gave no detectable splicing for the LE construct and at most a
small increase in splicing rate for the SE construct (0-10%; data not shown). Together the
results suggest that for a subpopulation of the ribozyme, ATP hydrolysis and product
release are necessary only to accelerate Mss116p dissociation and allow continued
folding as proposed (138). Nevertheless, ATP is required for additional steps in
Mss116p-dependent folding of much of the D135 population and most or all of the
populations of the SE and LE constructs.
4.3.6 Disruption of native D135 ribozyme by Mss116p
A strong prediction of models in which Mss116p functions as a general chaperone
is that it would not specifically recognize non-native structure and would therefore be
able to use ATP to unfold the native intron structure. This inherently non-specific
mechanism can nevertheless lead to native ribozyme accumulation if the native state is
more stable than populated misfolded states and therefore unfolded less efficiently or is
biased to form intermediates after protein-induced unfolding that preferentially refold to
the native state (26).
Thus, we used the discontinuous assay to probe for an Mss116p-dependent
decrease in the fraction of native D135 ribozyme (151). We prefolded D135 to the native
state and then added Mss116p at a concentration that inhibited productive D135 folding
(Figure 4.16). At various times thereafter, we transferred portions of the reaction to stage
2 and determined the fraction of native ribozyme by activity. In the presence of Mss116p,
96
the fraction of native ribozyme decreased rapidly, suggesting that Mss116p can unfold
the native D135 ribozyme to give intermediates that do not readily refold to the native
state in stage 2. Although substantial loss of native ribozyme was also observed for
Mss116p in the absence of ATP, the decrease was faster and larger with ATP. Mss116p
was then proteolyzed and the ribozyme was again incubated under conditions known to
give native folding (42 °C, 100 mM Mg2+
, 20 min). The fraction of native ribozyme
returned to near its original value (Figure 4.16, diamonds), confirming that the Mss116p-
dependent decrease was due primarily to disruption of the native structure rather than a
process giving irreversible inactivation.
4.3.7 Mss116p-promoted splicing of aI5 in vivo
Finally, to determine which activities of Mss116p are physiologically relevant for
aI5 splicing, we tested whether the Mss116p mutants can promote splicing of aI5 in
vivo (Figure 4.17). We used a previously developed in vivo splicing assay in which WT
or mutant Mss116p is expressed from a centromere-containing (CEN) plasmid in an
MSS116 deletion strain (mss116) (91). Because Mss116p is required for the synthesis of
mt respiratory components, null mutants cannot grow on non-fermentable carbon sources,
such as glycerol, but grow well on raffinose, a non-repressing fermentable sugar,
enabling their splicing phenotype to be assessed by Northern hybridization. To eliminate
the possibility of indirect effects from defects in splicing of other introns in the COX1
gene, we used isogenic strains containing mtDNAs with the single intron aI5 (91). The
COX1 pre-mRNAs in these strains contain aI5 with long 5 and 3 exons (~1.5 and 0.5
kb, respectively, to the 5 and 3 ends of COX1 mRNA), and thus most closely resemble
the LE construct used in vitro.
97
Figure 4.17A shows a Northern blot comparing the abilities of WT and mutant
versions of Mss116p to promote splicing of aI5 in vivo. The blot was hybridized with a
COX1 exon probe to detect spliced mRNA and unspliced precursor RNA. Because the
COX1 gene contains only aI5, the blot shows two major COX1 transcripts,
corresponding to spliced mRNA and an unspliced precursor RNA, with the ratio of the
bands providing a measure of the splicing efficiency. As expected, the wild-type strain,
which has a functional chromosomal copy of MSS116, spliced aI5 efficiently, yielding a
predominant band corresponding to COX1 mRNA (lane 1), while the mss116 strain
accumulates unspliced precursor RNA (lane 2). Expression of WT Mss116p from the
CEN-plasmid efficiently complemented the splicing defect in the mss116 strain,
restoring aI5 splicing to nearly the level of the wild-type strain (lane 3). By contrast, the
K158A and K158R mutants were unable to promote splicing of aI5 substantially above
the low residual level in the mss116 strain (null phenotype; lanes 4 and 5), although
K158A gave a very small increase that paralleled a small amount of residual ATPase
activity (176). The SAT/AAA mutant gave an intermediate phenotype, with
approximately equal amounts of unspliced precursor and spliced mRNA (lane 6). We
verified by immunoblotting that the mutant proteins were expressed at or near the level of
WT Mss116p (Figure 4.17B,C). Previous experiments using strains with multiple COB
and COX1 introns similarly showed that K158A and other motif I mutants gave a null
phenotype, and that the SAT/AAA mutant gave an intermediate phenotype for splicing of
aI5 and all other mt group I and II introns examined (91, 185, 191). Collectively, the
findings for the motif I mutants show that ATP binding and hydrolysis are essential for
Mss116p-promoted in vivo splicing of all S. cerevisiae mt group I and II introns. Further,
the splicing efficiencies of the motif I and SAT/AAA mutants agree with their activities
in splicing and folding aI5 RNAs in vitro and, as for all other Mss116p mutants
98
examined, correlate with their residual RNA-unwinding activities for appropriately sized
duplexes (60).
4.4 DISCUSSION
Here we used in vitro and in vivo approaches to probe the mechanisms by which
the yeast DEAD-box protein Mss116p promotes splicing of the group II intron aI5γ. Our
results indicate that a major physiological role of Mss116p in splicing this intron, as with
other group I and group II introns, is to use ATP to promote conformational transitions
that require the transient disruption of RNA structure.
4.4.1 Requirement for ATP binding and hydrolysis and effects of exon length on
Mss116p-mediated splicing in vitro
To probe the functional requirements of Mss116p in aI5γ splicing, we tested
mutants that are compromised in ATP binding, hydrolysis, and RNA unwinding, and we
used the SE and LE constructs to test whether the ATP requirement depends on the
lengths of the flanking exons. The experimental design was essentially the same as in a
recent report, (149) and where overlapping experiments were performed, the data are
largely consistent. Thus, the LE construct splices much slower than the SE construct;
both constructs require Mss116p and ATP for accelerated splicing; and with higher
concentrations of Mss116p, the splicing rates level and then decrease. The plateau
between activation and inhibition occurs at a higher Mss116p concentration and a higher
splicing rate for the LE construct than the SE construct. Also consistent with prior work,
the mutants K158A and K158R, which are deficient in ATP binding and hydrolysis, are
unable to stimulate splicing of either construct (176). Similar results were obtained for
other Mss116p and CYT-19 motif I mutants with the LE construct (30, 55).
99
However, key conclusions differ from those in previous work (149). Zingler et al.
compared the rate constants for the LE and SE constructs at Mss116p concentrations near
the plateau regions and concluded that the LE construct is stimulated to a greater extent
by Mss116p, perhaps reflecting Mss116p recruitment by the exons or facilitation of
protein oligomerization. In our view, it is dangerous to interpret these maximum values
as a measure of activation because they reflect the intersection of activation and
inhibition activities and can be influenced by changes in either or both activities. Instead,
we interpreted the slope of the rising portion of the Mss116p concentration dependence,
which reflects the overall reaction efficiency — i.e. the free energy change in going from
free protein and RNA in solution to the rate-limiting transition states — and is the part of
the concentration dependence that is least affected by inhibition. Making this comparison,
we observe that Mss116p stimulates splicing of the SE construct at least as efficiently as
the LE construct. Thus, rather than stimulating the SE construct less strongly, we
conclude that Mss116p inhibits it more strongly.
The slower, more temperature-dependent splicing of the LE construct suggests
that the long exons cause or contribute to kinetic barriers in RNA folding. This
conclusion is consistent with previous work indicating that sequences in the 5 exon
upstream of the intron-binding sequences (IBS1 and IBS2) can have large effects on aI5γ
splicing (150). The inhibition of splicing from exon sequences could reflect the formation
of misfolded exon structures that must be resolved prior to splicing, as demonstrated for
group I introns (192, 193). Alternatively or in addition, the long exons may stabilize
structure within the aI5γ intron, presumably by interacting with the intron, thereby
increasing barrier heights for disruption of both non-native and native structure. Evidence
in support of this possibility comes from the lower, more temperature-dependent splicing
rate of the LE construct in the absence of Mss116p, indicating a higher enthalpic barrier,
100
and from the greater stability of the LE construct. The latter is suggested by the higher
temperature optimum for LE construct splicing and its greater resistance to inhibition by
Mss116p, which presumably reflects disruption of the native intron structure. Our finding
that stimulation of both the SE and LE constructs by Mss116p depends upon ATP and is
affected similarly by Mss116p mutations raises the possibility of common rate-limiting
steps and barriers for these two constructs, which may be hindered by exon stabilization
in the LE construct. Further work will be necessary to define the structures of folding
intermediates for these constructs and the roles played by exons.
4.4.2 The SAT/AAA mutant is compromised for splicing SE and LE constructs
The motif III mutant of Mss116p (SAT/AAA) follows a design used with eIF4A
and other DEAD-box proteins (43, 47, 194) and was originally described in a Ph.D. thesis
from H.R. Huang in Philip Perlman’s lab, where it was classified as a ‘weak allele’ based
on in vivo analysis of translation and splicing (185). This mutant has since been the
subject of substantial study and contributed to physical models of Mss116p function (60,
176). Recently, the Mss116p SAT/AAA mutant was reported to be impaired in splicing
of the aI5γ LE construct but as efficient as WT Mss116p for splicing of the SE construct
(149). Because it is deficient in RNA-unwinding activity, this result was taken as
evidence that RNA unfolding is required for the exons but not within the intron. In
contrast, our data show that Mss116p SAT/AAA is reduced in activity by at least a
comparable amount for the SE construct as for the LE construct. The difference in
conclusions can be understood from inspection of both sets of data. The previous
conclusion that Mss116p SAT/AAA promotes splicing of the SE construct with WT
efficiency came from a comparison of splicing rates with 15 nM protein, which is well
into the inhibitory regime for WT but much closer to the plateau for the SAT/AAA
101
mutant (Figure 2 of ref. (149)). The reduced efficiency of Mss116p SAT/AAA with the
SE construct is visible at lower protein concentrations, both in our data (Figure 4.4 and
4.6) and in Figure S2B of Zingler et al (149). Thus, the SAT/AAA mutant again fails to
provide evidence that Mss116p can promote splicing of aI5γ without unwinding RNA.
4.4.3 The roles of ATP in Mss116p-promoted intron folding
Using a new two-stage, discontinuous catalytic activity assay, we found that
Mss116p accelerates native folding of the D135 ribozyme under near-physiological
conditions, supporting and extending previous studies of Mss116p on folding of aI5γ
(55, 138, 140). We then used the discontinuous assay to gain new insights
into the roles of ATP in Mss116p-mediated folding of aI5γ. It was suggested previously
that Mss116p promotes folding by stabilizing an on-pathway folding intermediate in a
reaction that is inherently ATP-independent, but that ATP is needed to promote release of
Mss116p after this step (149). A key prediction of this model is that removal of Mss116p
by proteolysis after binding in the absence of ATP would allow the ribozyme to fold to
the native state, which could be detected by activity in stage 2 of our assay. In contrast to
this expectation, we found that with two-fold excess Mss116p over D135, proteolysis of
Mss116p does not give additional native ribozyme. However, with 8-fold excess
Mss116p, proteolysis does allow 10-25% of the ribozyme to reach the native state. The
partial recovery indicates that for a subpopulation of the ribozyme, Mss116p accelerates
native folding by promoting a step, presumably compaction of domain I, (138) without
requiring ATP, and it provides experimental support for models in which Mss116p
dissociation is required for productive folding (149). Acceleration of domain I
compaction may arise from transient binding and stabilization of a folding intermediate,
as suggested by the ATP independence and the ability of several basic proteins to
102
promote this step with low efficiency (140, 149, 176). Alternatively or in addition,
Mss116p and other DEAD-box proteins may accelerate this step by disrupting local
structure in a process that does not strictly require ATP, perhaps giving transient
unstacking of coaxially stacked helices to allow bending of an internal loop within D1
(60, 195, 196).
Nevertheless, even under the most favorable conditions, the yield of native D135
ribozyme upon incubation with Mss116p followed by proteolysis is substantially smaller
than in the presence of Mss116p and ATP, indicating that most of the population requires
Mss116p and ATP for at least one additional step. The fraction that is able to avoid this
additional requirement is smaller for the SE construct and undetectable for the LE
construct. The increased dependence on ATP with increasing exon length supports the
hypothesis that the additional ATP-dependent step(s) include localized RNA unwinding,
because the larger size and complexity of these RNAs are expected to lead to folding via
more complex pathways with more trapped intermediates, and likely with intermediates
of greater stability due to contributions from the exons. It is also reasonable to expect that
binding of Mss116p to at least some sites within the intron would be inhibitory
throughout the folding process, and thus it is likely that acceleration of Mss116p
dissociation is an additional role of the ATPase cycle independent of any structural
stabilization.
4.4.4 Disruption of the native D135 ribozyme
Models for DEAD-box proteins as general RNA chaperones postulate that they
disrupt RNA structure non-specifically, generating a kinetic redistribution of folding
intermediates and additional chances for productive folding (26). A corollary is that there
is no absolute mechanism for distinguishing native from non-native structure. In support
103
of this hypothesis, we used the discontinuous activity assay to show that Mss116p can
disrupt the native D135 ribozyme in an ATP-dependent manner. This result parallels
previous findings for a native group I intron ribozyme (26, 60) and indicates that
Mss116p is capable of promoting native folding of aI5γ by disrupting misfolded
intermediates, which are less stable than the native state. Although it was reported that
Mss116p cannot unfold the native structure of a domain I construct of aI5γ, (138)
unfolding may have gone undetected by non-denaturing gel assays if the isolated domain
I refolds rapidly to the native structure or to alternative compact forms that migrate
similarly to the native structure.
4.4.5 Requirement for ATP binding and hydrolysis by Mss116p in vivo
To investigate which activities of Mss116p are required for its in vivo function,
we used yeast strains that lack all mt group I and group II introns except aI5γ (91). The
major COX1 pre-mRNA in these strains contains aI5 with long 5 and 3 exons (~1.5 and
0.5 kb, respectively) and thus most closely resembles the LE construct used to analyze
the effect of Mss116p in vitro. In other yeast strains that contain multiple COX1 introns,
the COX1 pre-mRNAs are expected to be even longer and more heterogeneous,
depending upon the order in which the introns upstream of aI5 are spliced.
Our Northern blot analysis indicates that the motif I mutants are essentially
inactive for aI5γ splicing, with a very low level of residual activity for K158A, and that
Mss116p SAT/AAA is more active than the motif I mutants but nevertheless significantly
compromised relative to WT Mss116p, as expected from its decreased RNA-unwinding
activity (60). These effects mirror those of the same mutations in vitro, most simply
suggesting similar barriers to group II intron folding in vivo and in vitro.
104
Our results are in agreement with previous in vivo studies of Mss116p mutations
in strains containing multiple introns, which showed that K158A and other motif I
mutations give a null phenotype for splicing of aI5 and all other group I and II introns
examined, and that SAT/AAA and other motif III mutations give an intermediate
splicing-defective phenotype for aI5 and all other group I and II introns examined (91,
185, 191). We note in particular that the initial characterization of the SAT/AAA
mutation in a strain with three COX1 introns showed that “this mss116 allele does not
function as well as the wild-type MSS116 on splicing” (185) (in contrast to statements in
the Discussion of Zingler et al (149)). Although another mutant, Q412A, has been
suggested to promote efficient RNA splicing in vivo despite low RNA-unwinding
activity, (149, 191) its unwinding activity was assayed only with a relatively long duplex
(17 bp) and was greater than that of a motif III mutant (T307A) in the same study. Thus,
in our view, the simplest interpretation is that Q412A supports splicing by virtue of its
residual RNA-unwinding activity, which is expected to be higher for shorter duplexes of
the type found in group I and II intron RNAs.
4.5 CONCLUSIONS AND IMPLICATIONS
Together, our results indicate that a critical activity in Mss116-mediated folding
of the aI5γ intron is ATP-dependent RNA-unwinding activity. ATP is required for
maximal stimulation of folding and splicing of all constructs tested, and Mss116p is
capable of disrupting even the most stable global structure of the intron, the native state.
Further, for all Mss116p mutants analyzed to date, the ability to promote splicing of aI5γ
in vitro and in vivo correlates with RNA-unwinding activity with appropriately sized
duplexes, suggesting that this unwinding activity is necessary for stimulation of aI5γ
splicing. Nevertheless, it remains likely that different RNAs and different folding
105
intermediates require different activities and that DEAD-box proteins use multiple
mechanisms to promote RNA folding. The recent advances in understanding the folding
of aI5γ
transitions accelerated by DEAD-box proteins and the detailed mechanisms through
which the accelerations are achieved.
4.6 FOOTNOTE
* The efficiency reported in the current work is approximately 5-fold larger than
previously reported (60, 81). In principle, this difference could be caused by the
decreased RNA and protein concentrations, changes in buffer from Tris to MOPS, small
accompanying changes in counterion identity and concentration, and/or recent
preparations of protein being somewhat more active than earlier preparations. Further
experiments indicate that the difference is not caused by the change in buffer conditions,
which result in a small decrease in activity (2-fold), whereas there may be contributions
from the changes in RNA and protein concentration (≤ 2-fold) and differences in activity
between different protein preparations (data not shown).
106
Figure 4.1: RNA constructs
(A) Long exon (LE) and short exon (SE) splicing constructs. Each intron domain is
shown in a different color and indicated with a label, and exon lengths are indicated. (B)
D135 ribozyme construct. Truncated and intact domains are shown in the same colors as
in panel A, and the intact domains are labeled. As indicated, the ribozyme includes 37 nts
at the 3 end, which are derived from a multiple cloning site (131). The secondary
structures were generated by modifying a diagram from the Comparative RNA Website
(197).
(Figure prepared by Mark Del Campo.)
107
Figure 4.2: Self-splicing of the LE and SE constructs
(A) Gel images from splicing reactions performed at 33 °C. Splicing was monitored as
the fraction of radiolabeled material present as precursor RNA (see Section 4.2.4). Bands
in this and subsequent gel images are precursor (P), excised intron lariat (I-lar), excised
linear intron (I-lin), and ligated exons (E1-E2). (B) Temperature dependences for self-
splicing of the LE (red) and SE (blue) constructs. Rate constants, as determined from
single exponential fits or from initial rates of slower reactions, are plotted on a log scale
against the inverse of temperature (in Kelvin) multiplied by 1000. Reaction conditions
were 50 mM Na-MOPS, pH 7.0 (determined at 25 °C), 8 mM MgCl2, 100 mM KCl
(including a contribution from Mss116p storage buffer), 1 mM ATP-Mg2+
, and 10%
Mss116p storage buffer (see Section 4.2.4). All values shown reflect the average and
standard error from 2–4 independent determinations except those at 41 °C, which reflect
a single determination. Downward arrows indicate that non-specific products were
observed under those conditions, allowing determination of only an upper limit on the
rate constant for splicing. The regions in which splicing rate increased with temperature
gave ΔH values of 57 kcal/mol and 44 kcal/mol for the LE and SE constructs,
respectively.
108
Figure 4.3: Self-splicing of the LE construct (panel A) and SE construct (panel B)
at different temperatures
Reactions were performed in a thermal cycler with a temperature gradient. Temperatures
for each reaction were estimated by linear interpolation from the temperature settings.
109
Figure 4.4: Mss116p-stimulated splicing of the LE and SE constructs
110
(A) Gel images of LE and SE splicing in the presence and absence of Mss116p (10 nM
and 2 nM Mss116p for the LE and SE constructs, respectively). In this and all other
experiments with Mss116p, the total intensity of radiolabeled RNA in the quantified
region of the gel decreased less than 2-fold during the experiment, indicating minimal
RNA degradation. (B) Progress curves of LE and SE splicing in the presence of various
Mss116p concentrations, as indicated by color. Circles show results from reactions in the
presence of 1 mM ATP and triangles indicate reactions without ATP. Points represent the
fraction of labeled material present as precursor as a function of time, and the curves
reflect the best fit by a first-order rate equation. (C) Mss116p concentration dependences
for splicing of the LE and SE constructs in the presence of ATP. For the SE construct,
Mss116p was saturating or nearly saturating even at low concentrations equivalent to that
of the radiolabeled RNA (~1 nM), preventing determination of a second-order rate
constant for activation.
(This figure is the work of Mark Del Campo.)
111
Figure 4.5: Mss116p-stimulated splicing of LE and SE constructs
Using Tris buffer conditions (10 mM Tris-Cl, pH 7.5, 100 mM KCl, 10% glycerol), with
10 nM RNA. (A and B) Gel images of splicing in the presence and absence of Mss116p
for the LE construct (40 nM Mss116p, panel A) and the SE construct (10 nM Mss116p,
panel B). (C) Concentration dependences of Mss116p stimulation for the LE and SE
constructs. Rate constants from this experiment are 1.8 106 M
–1 min
–1 for the LE
construct and ≥1.8 106 M
–1 min
–1 for the SE construct. (D) Concentration dependences
for Mss116p SAT/AAA for splicing of the LE and SE constructs under Tris buffer
conditions. Rate constants are 1.2 105 M
–1 min
–1 for the LE construct and 4.4 10
5 M
–1
min–1
for the SE construct.
(This figure is the work of Mark Del Campo.)
112
Figure 4.6: Stimulation of splicing by the SAT/AAA mutant of Mss116p
(A) Progress curves of splicing of LE (left) and SE (right) constructs in the presence and
absence of the Mss116p SAT/AAA. Circles indicate reactions in the presence of 1 mM
ATP and triangles indicate reactions without ATP. (B) Concentration dependence of
Mss116p SAT/AAA for stimulation of splicing of the LE and SE constructs in the
presence of ATP.
(This figure is the work of Mark Del Campo.)
113
Figure 4.7: Mss116p motif I mutants K158A and K158R in splicing reactions of the
LE and SE constructs (panels A and B, respectively)
All reactions included 1 mM ATP and were performed under standard near-physiological
conditions at 30 °C (50 mM Na-MOPS pH 7.0, 100 mM KCl, 8 mM MgCl2, and 5%
glycerol). The parallel reactions shown in the absence of Mss116p (black x’s) and in the
presence of WT Mss116p (purple diamonds) were performed side-by-side and gave the
same rates within error as equivalent reactions shown in Figure 4.4.
(This figure is the work of Mark Del Campo.)
114
Figure 4.8: Two-stage catalytic activity assay to monitor folding of D135 ribozyme
(A) Reaction schematic depicting standard, near-physiological conditions for stage 1. In
subsequent panels, results are shown plotted against folding time in stage 1 (t1) or
cleavage time in stage 2 (t2). (B) Prefolded and nonfolded controls. Ribozyme was either
prefolded (circles, 42 °C, 100 mM MgCl2, 500 mM KCl, 50 mM Na-MOPS, pH 7.0, 20
min) or transferred to stage 2 directly from buffer solution (triangles, 30 °C, 50 mM Na-
MOPS, pH 7.0). As expected for a single turnover followed by slow product release, the
reaction with prefolded ribozyme gave a stoichiometric burst of product (see Section
4.2.5) and a second phase reflecting subsequent turnovers of the ribozyme. The
nonfolded ribozyme reaction gave less substrate cleavage, with five determinations
giving an average burst corresponding to 15% of the ribozyme population. (C) Progress
115
of D135 RNA folding in stage 1 under non-physiological conditions similar to those used
previously (42 °C, 100 mM Mg2+
, pH 8.1), (183) with the fraction of native ribozyme
determined from the burst amplitude of substrate cleavage in stage 2. (D) Progress of
D135 folding under near-physiological conditions. Burst amplitudes are normalized by
the maximal burst amplitude from prefolded ribozyme, as shown in panel b. Fifteen
determinations gave a fast phase with an amplitude of 0.22 ± 0.02 and a rate constant of
1.0 ± 0.2 min–1
. A slower phase was also present, and six determinations for this phase
gave a rate constant of 1.4 (± 0.6) 10–3
min–1
and a final amplitude of ~0.6.
116
Figure 4.9: Testing of 30 °C, 100 mM Mg2+
for conditions of stage 2 in the
discontinuous catalytic activity assay
The amount of rapid substrate cleavage from ribozyme transferred directly from buffer
solution lacking Mg2+
at 30 °C (triangles, the nonfolded reaction) is not much less than
that from an equivalent amount of ribozyme that was prefolded (42 °C, 100 mM Mg2+
, 20
min) before being transferred to the stage 2 conditions. A comparison of this plot with
Figure 4.8B demonstrates that the lower temperature in that experiment for stage 2, 15
°C, is more effective for blocking folding of D135 on the time scale of substrate cleavage
by the native ribozyme. Thus, the lower temperature was used for all subsequent
experiments.
117
Figure 4.10: Prefolded and nonfolded reactions for monitoring D135 RNA folding at
42 °C
Black symbols and curves show reactions that were transferred from 42 °C into standard
stage 2 reaction conditions (15 °C, 100 mM Mg2+
, 80 mM Na-HEPES, pH 8.1). The
circles show the progress of substrate cleavage for a reaction that was first prefolded (42
°C, 100 mM Mg2+
, 50 mM Na-MOPS, pH 7.0, 20 min) and then transferred, and the
triangles show results from a reaction that was transferred from 42 °C (80 mM Na-
HEPES, pH 8.1) to the stage 2 conditions without prefolding. HEPES buffer was used
here so that the reaction would be identical to previous work (see Section 4.3.3).(184)
The gray symbols show analogous reactions that were transferred from 30 °C (50 mM
Na-MOPS, pH 7.0, ± 100 mM Mg2+
), either with or without prefolding, reprinted from
Figure 4.8B for comparison. The prefolded reactions are essentially the same as expected,
indicating nearly 100% native ribozyme, but the nonfolded reaction transferred from 42
°C gives substantially more cleavage than the equivalent reaction transferred from 30 °C,
despite the essentially identical conditions in stage 2 for the two reactions.
118
Figure 4.11: Mss116p accelerates native folding of the D135 ribozyme
119
(A) Time courses of substrate cleavage in stage 2 after incubation with Mss116p under
near-physiological conditions in stage 1. D135 ribozyme (50 nM) was incubated with 100
nM Mss116p and 1 mM ATP for the indicated times before substrate cleavage was
measured under stage 2 conditions (see Section 4.2.5). Also shown are a ‘nonfolded’
control reaction, in which D135 was added directly to stage 2 conditions, and a prefolded
control, in which D135 was incubated at 42 °C for 20 min in the presence of 100 mM
Mg2+
to form native ribozyme. (B) Progress of D135 folding in stage 1 in the presence of
100 nM Mss116p with 1 mM ATP (circles, kobs = 0.61 ± 0.07 min–1
, amplitude = 0.76 ±
0.04) or without ATP (diamonds, kobs = 0.64 ± 0.07 min–1
, amplitude = 0.28 ± 0.01). The
fraction of native ribozyme was determined by the burst amplitude in stage 2, as
described in Materials and Methods. (C) Progress of D135 folding in stage 1 with
Mss116p mutants in the presence of 1 mM ATP and 100 nM protein. Proteins are
Mss116p SAT/AAA (blue, kobs = 0.5 ± 0.2 min–1
, amplitude = 0.50 ± 0.09), Mss116p
K158A (orange, kobs = 0.7 ± 0.1 min–1
, amplitude = 0.34 ± 0.06), or Mss116p K158R
(red, kobs = 0.7 ± 0.2 min–1
, amplitude = 0.33 ± 0.01).
120
Figure 4.12: Progress curves of D135 ribozyme folding
50 nM D135 ribozyme in the presence of 1 mM ATP and various concentrations of
Mss116p (panel A) or Mss116p SAT/AAA (panel B).
121
Figure 4.13: Proteolysis of Mss116p after incubation with D135 RNA in the absence
of ATP
Mss116p was added in stage 1 at 100 nM (panel A) or 400 nM (panel B) and incubated
for the indicated times under near-physiological conditions. Proteinase K (1 mg/ml) was
122
then added at the indicated times (closed symbols) and incubated for an additional 8 – 60
min to permit further folding before aliquots were transferred to stage 2 and the fraction
of native ribozyme was determined by measuring the substrate cleavage burst amplitude.
The fraction of native ribozyme did not depend on the incubation time after proteinase K
addition (8–60 min), and the symbols show the average values. Open symbols show
equivalent reactions with Mss116p and without nucleotide, to which proteinase K was
not added. Including or omitting 0.5% SDS with proteinase K to ensure removal of
peptide fragments had no significant effect on the results (data not shown). For the
experiments shown in panel B, 0.5% SDS was added immediately after proteinase K.
Results from independent determinations are shown in different colors. It can be seen that
the increase in native ribozyme upon proteinase K treatment is variable and that the
variation is larger between experiments than within experiments.
123
Figure 4.14: Proteinase K digestion of Mss116p in experiment shown in Figure 4.13
When proteinase K is added to Mss116p in the stage 1 conditions (30 °C, 50 mM MOPS,
pH 7.0, 8 mM Mg2+
), Mss116p is undetectable within 30 s (right lane). The second lane
from the right shows an equivalent reaction to which proteinase K was omitted, and the
third lane from the right shows an equivalent amount of proteinase K alone (at dye front).
124
Figure 4.15: Proteolysis of Mss116p after incubation with D135 RNA
Mss116p was present at 100 nM (Panel A) or 400 nM (panel B). Nucleotides were
present, as indicated, at 1 mM. Closed symbols show the fraction of native ribozyme for
125
reactions in which proteinase K (1mg/ml) was added at the indicated time, followed by
additional incubation of 5 – 30 min before transfer to stage 2. The results did not depend
on this incubation time, and the average values are shown. SDS (0.5%) was added
immediately after proteinase K to ensure that peptide fragments were prevented from
interacting with the ribozyme. Open symbols show equivalent reactions to which
proteinase K was not added. Reactions in the absence of nucleotide (circles) were
performed side-by-side for comparison and gave the same results within error as the
equivalent reactions shown in Figure 4.13. Results from two independent experiments are
shown in red and blue.
126
Figure 4.16: Unfolding of native D135 RNA by Mss116p
The ribozyme (1.8 µM) was prefolded at 42 °C, 100 mM Mg2+
and then diluted to 30 °C
and 8 mM Mg2+
prior to addition of Mss116p (1.2 µM Mss116p; 74 nM D135 after
dilution). Reactions included 1 mM ATP (green) no ATP (red) or the same volume of
storage buffer without Mss116p (blue). Reactions were incubated for the indicated times
and the fraction of native ribozyme was determined by transferring aliquots to stage 2 and
measuring substrate cleavage. After 12 min, proteinase K (1 mg/ml) and additional Mg2+
(100 mM) were added and the RNA was again folded to the native state by incubation at
42 °C for 20 min prior to determining the fraction of native ribozyme as above
(diamonds). This refolding step was included to ensure that the decrease in native
ribozyme upon incubation with Mss116p arose from unfolding rather than an irreversible
process such as RNA degradation.
127
Figure 4.17: Northern hybridization and correlated immunoblot comparing the
ability of wild-type and mutant Mss116p to promote splicing of aI5 in
vivo
(A) Northern hybridization. The blot shows whole-cell RNAs (1.0 µg) from the indicated
strains separated in a 1.5% agarose gel and hybridized with a 32P-labeled oligonucleotide
complementary to COX1 exon 6. Lanes: (1) WT 161-aI5 (WT); (2) mss116-aI5
transformed with CEN plasmid pRS416 (empty vector); (3-6) mss116-aI5 transformed
with CEN plasmids expressing Mss116p mutants K158A, K158R, and SAT/AAA. (B)
Immunoblot. The blot shows TCA-precipitated proteins (~60 µg) from the same strains
as in panel (A) separated in a 4-20% polyacrylamide gradient gel and probed with an
anti-Mss116p antibody. (C) Immunoblot stained with AuroDye Forte to confirm equal
loading. The numbers to the left of the gel in (b) and (c) indicate the positions of size
markers (Precision Plus Protein Dual Color Standards; Bio Rad).
(This figure is the work of Rachel Wolf.)
128
Chapter 5: Rapid structure formation within the aI5γ group II intron
5.1 INTRODUCTION
Using catalytically active RNA to probe folding pathways has been an effective
method for obtaining kinetic information on the fraction of RNA in the native state and
for detecting the presence of intermediate, non-catalytic RNA conformations.
Intermediates may be non-catalytic because they are kinetically-trapped misfolded
conformations (134, 136) or because they are more stable than the native state in the
absence of a cofactor (198, 199). However, without the use of orthogonal techniques that
probe for physical information, studies using catalytic activity are unable to reveal
structural information about the folding intermediates and cannot differentiate between
the two types of non-catalytic conformations mentioned above.
Two powerful techniques that probe physical features of RNA are chemical
footprinting and small angle X-ray scattering (SAXS). Chemical footprinting experiments
use exogenous probes to modify the RNA according to its structural features. For
example, DMS footprinting uses dimethyl sulfate to methylate the base-pairing faces of
accessible adenine and cytosine nucleobases. Formations of base-pairing interactions or
tertiary interactions that involve the base-pairing face of the nucleotides reduce the
accessibility of dimethyl sulfate and provide protection from modification. The
modification pattern of accessible and protected nucleotides can be used to obtain
structural information regarding which features are formed in a particular RNA
conformation. Unlike the mostly local structural information provided by DMS
footprinting, SAXS is able to provide a more global view of folding intermediates. SAXS
is a solution-based method that allows the overall size and shape of RNA to be probed.
While the resolution capabilities of SAXS are not as fine as X-ray crystallography,
compaction events during RNA folding can often be detected (94, 163, 165).
129
In this study, knowledge acquired about the aI5γ group II intron folding pathway
through catalytic activity measurements (see Chapter 4) is combined with DMS
footprinting and SAXS data to probe structural features of any folding intermediates that
might be populated. Folding and splicing of the full-length aI5γ construct in near-
physiological conditions (100 mM KCl, 8 mM Mg2+
, 30 °C) are extremely slow with a
rate constant near 10-5
min-1
(see Section 4.3.1) (142). The DEAD-box protein Mss116p
has been shown to stimulate folding and splicing of aI5γ (see Section 4.3.1 and (60, 81,
142). There is evidence that this stimulation occurs because of and is dependent on the
RNA unwinding ability of the protein. For example, a motif III SAT/AAA mutant of
Mss116p retains ATPase and RNA binding activity but has a decreased efficiency for
separating duplexes compared to wild-type Mss116p (60). Correspondingly, this mutant
has a decreased ability to stimulate folding and splicing of aI5γ constructs, indicating that
Mss116p promotes native folding by promoting a local unfolding event (see Section 4.3.2
and (60)). Importantly, this local unfolding event is also indicated in vivo (see Section
4.3.7). Additionally, a ribozyme version of the intron populates the native state to a
greater degree after interaction with Mss116p and ATP than after interaction with
Mss116p alone, again indicating that an ATP-dependent strand separation event promotes
native state formation (see Sections 4.3.4-5). The requirement for a local unfolding event
implies that there is at least one misfolded conformation populated along the folding
pathway.
To probe the physical characteristics of any intermediates populated along the
folding pathway of aI5γ, DMS footprinting and SAXS strategies were utilized. DMS
footprinting was carried out on full-length aI5γ (~1500 nts) and the derived D135
ribozyme (~600 nts) (see Chapter 4, Figure 4.1). Both constructs have nucleotides that are
in duplex and loop regions in the native state become protected from DMS modification
130
soon after the addition of Mg2+
, indicating rapid formation of structural features.
Additionally, initial SAXS data on the D135 ribozyme reveal a degree of global
compaction upon addition of Mg2+
, again indicating rapid formation of structural features
that underlie this compaction.
5.2 MATERIALS AND METHODS
5.2.1 RNA preparation
Group II intron aI5γ was transcribed in vitro from HindIII-digested plasmid by T7
RNA polymerase using a Megascript kit (Ambion). RNA was isolated by phenol-
chloroform extraction and size exclusion chromatography using two consecutive G-50
columns. D135 RNA was transcribed in vitro from HindIII-digested pQL71 using T7
RNAP and purified via an RNeasy column (Qiagen). RNA concentrations were
determined spectrophotometrically using extinction coefficients at 260 nm of 1.76 107
M–1
cm–1
and 5.86 106 M
–1 cm
–1 for aI5γ and D135, respectively.
5.2.2 Footprinting data acquisition and analysis
DMS footprinting reactions were carried out using 0.5 pmol of full-length aI5γ.
The RNA was denatured at 92°C for 1 minute in the absence of Mg2+
and then transferred
to reaction conditions. The states of the RNA probed were ‘unfolded’ (50 mM Na-MOPS
pH 7.0, 100 mM KCl), ‘native’ (50 mM Na-MOPS pH 7.0, 100 mM KCl, 8 mM MgCl2,
100 nM Mss116p, 2.5 hr+, 30 °C), and an ‘early folding time point’ that was initiated
from the unfolded conditions with the addition of Mg2+
(50 mM Na-MOPS pH 7.0, 100
mM KCl, 8 mM MgCl2, 30 °C, < 5 min). In addition, all reactions contained 1 mM ATP-
MgCl2 and 5% glycerol. After the RNA was folded, it was incubated with 1 µL of 70 mM
DMS for 2.5 minutes at 30 °C. A published protocol was followed using P32 labeled
131
primers to obtain cDNA fragments that were separated on 8% polyacrylamide gels and
visualized with a PhosphorImager (154) (Figure 5.1).
SAFA was used to quantify the individual bands in the footprinting gels (200,
201). These bands were then normalized by cDNA full-length products that were
quantified using ImageQuant (GE Healthcare) and multiplied by 1000. In part because
the cDNA products vary in size depending on if aI5γ has spliced or not, an area that
covered both cDNA product lengths was boxed and counted as full-length product. For
the majority of nucleotides in the intron, data from two or more reactions were averaged
and the standard deviations were determined. However, data from only one reaction was
obtained for nucleotides 632,633, and 725-780 for the native state and nucleotides 862-
886 for the early folding time point state. Additionally, nucleotides 789-886 were not
probed in the native state.
The data for unfolded RNA incubated with DMS were compared to data with
unfolded RNA not incubated with DMS to observe which nucleotides were accessible to
DMS. In order for a nucleotide to be considered accessible, the value +DMS must be ≥ 1
larger than the value –DMS, the –DMS value cannot be larger than 1, and the standard
deviation for the +DMS value cannot overlap with the standard deviation for the –DMS
value. In order for a nucleotide to be considered protected, the unfolded+DMS value
must be ≥ 0.5 larger than the value for the native+DMS or early folding time point+DMS
reactions, the standard deviations cannot overlap, and the –DMS reactions cannot be
larger than 1. Certain nucleotides show greater protection in the native state than in the
early folding time point state. These nucleotides are considered protected in the early
folding time point state and have early folding time point+DMS values ≥ 0.4 larger than
values with the native+DMS reactions. Lastly, some nucleotides are not protected but are
made more accessible to DMS after folding. If a nucleotide had a value in the
132
native+DMS reaction ≥ 0.1 larger than the unfolded+DMS reaction, the standard
deviations did not overlap, and the –DMS reactions were less than 1 then that nucleotide
was labeled as being more accessible after folding.
Experiments probing the native state were carried out both with the addition of
0.1 mg/ml proteinase K after aI5γ incubation with Mss116p and without proteinase K
addition. Because minimal differences in DMS profiles were noted and the amount of
nucleotides protected was similar in the presence and absence of proteinase K, the
experiments were treated as identical during analysis. Additionally, the native state was
confirmed to have significantly spliced by comparing the full-length products
corresponding to unspliced and spliced aI5γ generated in the reverse transcription
reaction (Figure 5.1).
5.2.3 SAXS data acquisition and analysis
SAXS data were collected on 0.5 µM or 1 µM of D135 ribozyme at the Advanced
Photon Source beamline 12-ID-C with twenty 1 second exposures for static
measurements and a variable number of 1 second exposures for the time-resolved data at
a sample-detector distance of 2 meters. The states of the RNA probed statically at room
temperature were ‘unfolded’ (50 mM Na-MOPS pH 7.0), ‘native’ (50 mM Na-MOPS pH
7.0, 500 mM KCl, 50 mM MgCl2, 42 °C for 20’), ‘early time point’ (50 mM Na-MOPS
pH 7.0, 100 mM KCl, 8 mM MgCl2, 30 °C for 20-30’), ‘Mss116p +ATP’ (50 mM Na-
MOPS pH 7.0, 100 mM KCl, 8 mM MgCl2, 30 °C for 20-30’, stoichiometric amount of
Mss116p), and ‘Mss116p –ATP’ (50 mM Na-MOPS pH 7.0, 100 mM KCl, 8 mM
MgCl2, 30 °C for 20-30’, stoichiometric amount of Mss116p). All of the samples also
contained 5% glycerol and 1 mM ATP-MgCl2 (except for the Mss116p –ATP sample,
which lacked ATP-Mg2+
, and the unfolded sample, which only contained 50 mM Na-
133
MOPS pH 7.0). The samples were folded to the aforementioned states at the temperatures
indicated and then were put on ice and exposed to the beam at room temperature.
Conversely, the early time point, Mss116p +ATP, and Mss116p –ATP samples were
observed with time dependence at 30 °C.
Data were collected during two separate visits to APS and were qualitatively in
good agreement when comparing the relative peak heights of Kratky plots of different
states from one visit to the next. Analysis of the SAXS data was performed with IGOR-
Pro (WaveMetrics) and ATSAS (version 2.3) software.
5.3 RESULTS
5.3.1 The unfolded state of aI5γ is readily accessible to DMS modification
First, the unfolded state of aI5γ in the presence of 100 mM KCl was probed
(Figure 5.2 and Figure 5.4A). Predictably, many A and C nucleotides that are thought to
reside in loops or not be base paired in the native state were modified by DMS (nts: 27,
38, 41, etc.). In fact, almost every loop region has at least one nucleotide modified by
DMS, indicating a lack of tertiary structure formation. Somewhat surprisingly, but in
agreement with an earlier study (202), many nucleotides that are base paired in the native
state are also accessible to DMS in the presence of 100 mM KCl (nts: 66,67,129, etc.),
indicating a lack of native secondary structure formation. This could result from Mg2+
being necessary in order for the RNA to form certain secondary structural elements (202)
or from non-native secondary structure forming that prohibits the formation of secondary
structure found in the native state. Because full-length aI5γ is being probed, non-native
secondary structure involving interactions between the exon(s) and intron may form,
which could trap the RNA in a misfolded state (60, 138, 142).
134
5.3.2 Addition of Mg2+
results in rapid formation of structural elements
To determine whether any structural features form early in the folding pathway,
changes in DMS modification patterns were probed soon after folding was initiated. The
conformation being investigated is referred to as an early folding time point state. To
reach the early folding time point conformation, the RNA was folded in near-
physiological conditions (100 mM KCl, and 8 mM Mg2+
at 30 °C for 2’) before DMS
was added. Although this folding incubation leads to much less than 1% of the RNA
reaching the native state and splicing (see Chapter 4, Figure 4.3), significantly more
nucleotides were protected from DMS in the early folding time point state than the
unfolded state (Figure 5.3 and Figure 5.4B, green, blue, and orange nts). While many of
the protected nucleotides appear to simply form secondary structure upon the addition of
Mg2+
(nts: 180, 181, 218 etc.), nucleotides in loop regions involved in tertiary contacts
were also protected (nts: 348 in α’ and 617 in µ). Protection of these nucleotides, and
others in loop regions that do not have identified tertiary contacts (nts: 209, 210, 395,
etc.), indicates tertiary structure formation early in the folding pathway. Therefore, upon
the addition of Mg2+
, some secondary and tertiary structural features form in less than
two minutes.
5.3.3 Native state is significantly more protected than early folding time point state
The last conformation probed was the native state, which was formed by
incubating the RNA with Mss116p for > 2.5 hours. The majority of RNA spliced under
these conditions, as confirmed by the different lengths of the cDNA products generated
from unspliced or spliced RNA (Figure 5.1). Predictably, numerous nucleotides that were
not protected in the early folding time point state are now protected in the native state,
indicating further formation of structure unique to the native state (Figure 5.3 and Figure
5.4B, red nts: 27, 51, 63 etc.). Also, some nucleotides that were protected in the early
135
folding time point state are protected to a greater degree in the native state (Figure 5.3
and Figure 5.4B, orange nts: 187, 191, 195, etc.). Additionally, two nucleotides are made
more accessible to DMS in the native state compared to the unfolded state (Figure 5.3
and Figure 5.4B, magenta nts: 420 and 734). This information leads to a model in which
the native state contains more structural elements than the early folding time point state
and further stabilizes some of the structures that are already formed in the early folding
time point state. Further, the native state has undergone splicing, which may give rise to
some of the changes in DMS modification between the early folding time point and the
native states.
5.3.4 Addition of Mg2+
gives rapid formation of structure in D135 ribozyme
A global comparison of the footprinting profiles for the unfolded, early folding
time point, and native state are shown in Figure 5.5A,B while Table 5.1 indicates all of
the nucleotides that were modified by DMS in full-length aI5γ. One of the most readily
apparent observations from the data with full-length aI5γ is that the early folding time
point state has many more nucleotides protected from DMS modification compared to the
unfolded state, indicating that significant structural features have formed soon after the
addition of Mg2+
(see Figure 5.5A and compare the unfolded profile (black) with the
early folding time point profile (blue)). Correspondingly, preliminary footprinting data
using the derived D135 ribozyme also reveals formation of structural elements soon after
the addition of Mg2+
(see Appendix and Figure A.1). Moreover, many of the structures
that are indicated to have formed in the D135 ribozyme are also formed in the full-length
aI5γ.
136
5.3.5 SAXS data reveal compaction of D135 ribozyme
The full-length aI5γ construct did not provide useful SAXS data at small angles.
In addition, the combined length of the exons is over two thirds the length of the intron
domains and would make data analysis complex. Therefore, the D135 ribozyme was used
to gather SAXS data. The kinetic information on the fraction of natively-folded
molecules of the D135 ribozyme revealed by the discontinuous assay (see Section 4.3 and
(142) can be combined with SAXS data to evaluate global structural features of the native
state and any intermediates that may be populated. First, the SAXS profile for the
unfolded (see Section 5.5)* structure (50 mM Na-MOPS pH 7) was compared to the
profile for D135 ribozyme that was prefolded to the native state‡ (50 mM Na-MOPS pH
7, 500 mM KCl, 50 mM Mg2+
42 °C for 20’). Observing the different profiles on a
Kratky plot plainly reveals the large compaction of the native state relative to the
unfolded state. The native state profile shows a clear, sharp peak indicative of a compact
conformation while the unfolded state profile is relatively flat and has no peak (Figure
5.6, black and gray traces). Analogously, the compaction of D135 ribozymes that have
undergone different prefolding incubations can be probed. States such as an early time
point (50 mM Na-MOPS pH 7.0, 100 mM KCl, 8 mM Mg2+
30 °C for 20’), Mss116p –
ATP (50 mM Na-MOPS pH 7.0, 100 mM KCl, 8 mM Mg2+
, Mss116p, 30 °C for 20’),
and Mss116p +ATP (50 mM Na-MOPS pH 7.0, 100 mM KCl, 8 mM Mg2+
, Mss116p, 1
mM ATP-Mg2+
, 30 °C for 20’) were probed. Previous catalytic activity measurements
showed that these incubations give ~30%, ~40%, and ~80%, respectively, of the
ribozyme folded to the native state. (see Sections 4.3.3-5 and note that here the raw, non-
normalized fraction of active D135 molecules inferred from the discontinuous assay is
used) The height of the peak in the Kratky plot increases as the fraction of native
ribozyme increases (Figure 5.6).
137
5.3.6 Time-resolved data indicate early compaction
In an attempt to follow compaction from the unfolded state to the states
mentioned above, a time-resolved mixing setup was used. RNA and buffer were mixed
with folding conditions to initiate the reaction and data were collected. First, folding to
the early time point state was observed as RNA was folded at 30 °C in 100 mM KCl and
8 mM Mg2+
for ~4 minutes, an incubation that produces ~30% native ribozyme. A Kratky
plot revealed that there was no systematic increase in peak height as a function of time
(Figure 5.7A), which would have indicated compaction during the monitored incubation
time. Rather, even the earliest time point after mixing (~3 sec) showed significant
compaction of the ribozyme compared to the unfolded state. Analogously, both the
Mss116p –ATP and Mss116p +ATP reactions indicated that no significant compaction
occurs after the initial compaction that takes place in the dead time of the experiment
(Figure 5.7B,C). It appears that the compactness possessed by these three states after 20-
30 minutes is essentially complete ~3 seconds after mixing with the folding solution
conditions (Figure 5.7, compare thick lines to thin lines).
5.4 DISCUSSION
Both the DMS footprinting data and SAXS data reveal significant formation of
structure for aI5γ constructs soon after the addition of Mg2+
. The footprinting data
illustrate that large amounts of secondary structure and some tertiary structure have
formed while the SAXS data indicate that significant global compaction of the RNA also
occurs within minutes of Mg2+
addition. Moreover, the SAXS data show that the majority
of compaction appears to be complete after 3 sec, as no significant further compaction
was observed beyond the initial compaction seen after the dead time of the experiment.
While this study did not uncover any specific misfolded conformations of aI5γ,
misfolded RNA conformations do not necessarily strictly differ in overall appearance or
138
structural features when compared to native states. Rather, they can mimic the native
state quite closely in some cases (17). Additionally, it is still possible that a multitude of
misfolded intermediate structures exist. These structures would not strictly give rise to a
distinct DMS footprinting profile because of their inherent heterogeneity but would
almost certainly be more compact than the unfolded conformation. As for the conclusions
about the group II intron aI5γ folding pathway derived from this study, it appears that
significant structural formation occurs early in the folding pathway upon the addition of
Mg2+
.
5.5 FOOTNOTE
* The unfolded state of the D135 ribozyme does not contain KCl, as opposed to
the unfolded state of aI5γ, which contains 100 mM KCl. The unfolded state for D135
ribozyme lacks KCl because a discontinuous activity assay revealed that a significant
portion of the ribozyme folds productively in the presence of KCl (data not shown).
‡ A control reaction using the discontinuous assay revealed that the same amount
of D135 ribozyme folds to the native state whether 50 or 100 mM Mg2+
is in the folding
reaction (data not shown).
139
Figure 5.1: Representative DMS footprinting gel
A DMS footprinting gel is shown above. The red box indicates the two different full-
length cDNA products resulting from reactions in which the intron was spliced with
Mss116p. The shorter cDNA product being more prevalent than the longer cDNA
product indicates that the majority of the intron is spliced and the reverse transcription
reaction terminates at the 5’ splice site most often. Lanes 1 and 2 are probing the native
state while lanes 3,4, and 5 are probing the unfolded state. The gel shows from
approximately nucleotide number 600 (bottom of gel) through the 5’ end.
140
Figure 5.2: Many DMS accessible nucleotides in the unfolded state
Full-length aI5γ was subjected to DMS footprinting. Shown on the secondary structure
diagram are the A and C nucleotides that were accessible to DMS modification (black) or
were not accessible (gray) when the RNA was in an unfolded state. Note that even many
nucleotides involved in secondary structures in the native state are accessible here,
indicating that they lack defined secondary structure in these conditions. Additionally, as
expected, many nucleotides in loops or involved in tertiary contacts in the native state are
accessible to DMS in these conditions. Known tertiary contacts are noted with Greek
letters.
141
Figure 5.3: Nucleotides protected and exposed in the native and early folding time
point states
Nucleotides that were protected from DMS modification or made more accessible, as
compared to the unfolded state, are shown. Nucleotides that are protected to a similar
degree in both the early time point and native states are green while those that are
protected in both states but are protected significantly more in the native state are shown
in orange. Protections only observed in the native state are red while those only protected
in the early time point state are blue. Therefore, together the green, orange, and blue
nucleotides represent the protections observed in the early time point state. In addition,
nucleotides that were more accessible to DMS in the native state compared to the
unfolded state are shown in magenta. Nucleotides shown in black were not protected in
any state. Domains V and VI were not probed in the native state due to a primer issue and
were not included in the analysis for the native state (gray box). See Figure 5.5A,B and
Table 5.1 for the raw footprinting data and a list of all the nucleotides modified by DMS.
142
Figure 5.4: DMS footprinting results for domain IV
Panel A shows the DMS accessible nucleotides in black while panel B shows protections
and enhancements to DMS modification by comparing the unfolded state to the other
states. Color scheme is the same as in Figure 5.3. The gray box indicates nucleotides that
were not probed in the native state.
143
Nucleotide #
144
Nucleotide #
Figure 5.5: DMS footprinting profiles of full-length aI5γ
The normalized signal intensity values for A and C residues for the unfolded, early
folding time point, and native states are shown. Panel A shows the profiles in the
presence of DMS and panel B shows the profiles in the absence of DMS.
145
Figure 5.6: Kratky plots of static samples reveal different compaction peak heights
D135 ribozyme samples (0.5 µM) were folded for approximately twenty minutes under
specified conditions and then put on ice until exposed to X-rays at room temperature. The
states probed are native (black), Mss116p +ATP (green), Mss116p –ATP (red), early
time point (blue), and unfolded (gray). These states correspond to 100%, 80%, 40%,
30%, and 15% native ribozyme, according to a discontinuous activity assay (142).
146
Figure 5.7: Time-resolved SAXS data indicate an early compaction event
Time-resolved data (thin lines in all panels) are compared to static data (thick lines in all
panels) acquired on the (A) early time point state, (B) Mss116p –ATP state, and (C)
Mss116p +ATP state. The time-resolved data are collected beginning ~3 seconds after
mixing D135 ribozyme with folding conditions and continue for ~4 minutes. Time-
resolved data are grouped together in colors from earliest to latest time points. Thin red
lines represent times from ~3-15 seconds after initiating folding. Thin green lines
represent times from ~15-30 seconds. Thin yellow lines represent times from ~40-130
147
seconds. Thin blue lines represent times from ~130-220 seconds. Note that while the time
dependent signal fluctuates, there is no systematic increase in peak height as a function of
time. By comparing the first time points of the time-resolved data (red thin lines) to the
static data (thick lines), it appears that the RNA is as compact after ~3 seconds as it is
after twenty minutes. All the data were collected using 1 µM D135 ribozyme except for
the unfolded state, which used 0.5 µM ribozyme with the signal normalized to account
for this difference.
(Time-dependent data acquired by Woongsoon Choi.)
148
Nucleotide reaction with DMS Total Number Color Coding in Figures
accessible 156 Black
27, 38, 41, 51, 63, 66, 67, 71, 77, 82, 92, 93, 94, 100, 101, 105, 114, 115, 117, 121, 125, 126, 127, 128,
129, 130, 133, 148, 152, 157, 159, 169, 176, 177, 180, 181, 187, 188, 191, 193, 195, 196, 198, 204, 205,
208, 209, 210, 214, 215, 217, 218, 219, 220, 225, 227, 228, 229, 230, 233, 237, 238, 240, 241, 242, 245,
251, 252, 253, 254, 259, 262, 264, 269, 272, 289, 290, 293, 307, 309, 310, 331, 332, 333, 341, 348, 355,
362, 365, 368, 382, 383, 389, 395, 403, 406, 410, 411, 412, 413, 415, 416, 418, 419, 441, 442, 444, 445,
447, 449, 463, 482, 496, 561, 585, 586, 589, 594, 598, 599, 600, 604, 605, 617, 627, 629, 630, 639, 644,
645, 646, 648, 651, 652, 653, 661, 668, 729, 745, 753, 757, 758, 762, 802, 803, 804, 806, 830, 832, 851,
860, 861, 868, 869, 876, 886
early folding time point protections 71 Blue, Green
129, 169, 180, 181, 187, 188, 191, 193, 195, 204, 205, 209, 210, 218, 219, 220, 229, 230, 233, 237, 238,
240, 241, 242, 307, 309, 310, 332, 348, 368, 389, 395, 403, 406, 410, 411, 413, 415, 416, 441, 442, 444,
445, 463, 482, 561, 585, 586, 589, 594, 599, 600, 604, 617, 627, 644, 645, 646, 652, 668, 729, 753, 757,
758, 762, 802, 803, 804, 806, 876, 886
native protections 96 Red, Green, Orange
27, 51, 63, 66, 67, 71, 77, 92, 100, 101, 105, 117, 121, 130, 133, 148, 159, 169, 180, 181, 187, 188, 191,
193, 195, 196, 198, 204, 205, 208, 209, 210, 214, 215, 217, 218, 219, 220, 227, 228, 229, 230, 233, 237,
242, 245, 259, 264, 307, 309, 310, 333, 341, 348, 382, 389, 395, 403, 406, 410, 411, 413, 415, 416, 418,
419, 441, 463, 482, 585, 586, 589, 594, 598, 599, 600, 604, 605, 617, 627, 630, 639, 644, 645, 646, 648,
651, 652, 653, 661, 668, 729, 745, 757, 758, 762
native greater protections 23 Orange
169, 187, 191, 195, 204, 205, 218, 219, 220, 229, 230, 233, 242, 348, 395, 413, 585, 586, 599, 600, 604,
617, 762
native more accessible 2 Magenta
420, 734
Table 5.1: Nucleotides modified by DMS.
All the nucleotides that were modified by DMS are included. (See Section 5.2.2 for how
these assignments were determined.)
149
Appendix
A.1 PRELIMINARY DMS FOOTPRINTING OF D135 RIBOZYME USING CAPILLARY
ELECTROPHORESIS
DMS footprinting reactions were carried out using 5 pmol of D135 ribozyme. The
RNA was denatured at 92°C for 1 minute in the absence of Mg2+
and then transferred to
reaction conditions. The states of the RNA probed were ‘unfolded’ (50 mM Na-MOPS
pH 7.0), and an ‘early folding time point’ that was initiated from the unfolded conditions
with the addition of Mg2+
(50 mM Na-MOPS pH 7.0, 100 mM KCl, 8 mM MgCl2, ~30
°C for < 2.5 min). After the RNA was folded, it was incubated with 1 µL of 420 mM
DMS for 30 seconds at ~30 °C. A published protocol was followed using Cy5 labeled
primers to obtain fluorescent cDNA fragments that were separated on a Beckman CEQ
instrument (154). The cDNA profiles were analyzed using CAFA software to quantify
the peak area for each nucleotide (203). Nucleotide signals were normalized by the
average peak area.
In order for a nucleotide to be considered accessible, the unfolded value +DMS
must be ≥ 0.2 larger than the value –DMS. Similarly, in order for a nucleotide to be
considered protected the unfolded+DMS value must be ≥ 0.2 larger than the early folding
time point value+DMS.
Despite a number of differences between the footprinting studies conducted on
the full-length aI5γ and the D135 ribozyme (different constructs, different DMS
incubation times, different concentrations of RNA probed, different normalization
procedures for nucleotide signal, and different methods of determining protections), the
general conclusion of structure formation soon after the addition of Mg2+
is maintained.
150
~Nucleotide #
Figure A.1: DMS footprinting profile of the D135 ribozyme
The normalized signal intensities for all the nucleotides of the D135 ribozyme are shown
for the unfolded (black) and early folding time point (blue) states in the presence of
DMS. Highlighted are nucleotides that were accessible in the unfolded state, protected in
151
the early folding time point state, and were also deemed protected in the study using the
full-length aI5γ construct (see Figure 5.3).
152
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