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Transcript
How the Group I Intron Works: A Case Study of RNA Structure and
Function
James L. Hougland and Joseph A. Piccirilli
Howard Hughes Medical Institute
Departments of Biochemistry & Molecular Biology and Chemistry
University of Chicago
Chicago, Illinois 60637
Marcello Forconi, Jihee Lee, and Daniel Herschlag
Department of Biochemistry
Stanford University
Stanford, California 94305-5307
1
INTRODUCTION
In 1968, Leslie Orgel and Francis Crick wrote back-to-back articles in the
Journal of Molecular Biology, making the same controversial point: that RNA,
because it could adopt structure – at the time tRNA was known to adopt its
famous cloverleaf secondary structure (Fig. 1) – could act functionally as a
catalyst (Crick, 1968; Orgel, 1968). Thereby a solution to the chicken and egg
problem of the origin of life was proposed. Instead of having to co-evolve an
information carrier (such as DNA) and a functional macromolecule (such as
proteins) to copy information from generation to generation, RNA could have
served both roles (Woese, 1967; Crick, 1968; Orgel, 1968). However, the bold
suggestions of Orgel and Crick were largely ignored until 1982, when Cech and
coworkers discovered the self-splicing activity of the group I intron from
Tetrahymena thermophila (Kruger et al., 1982; Cech, 1992). The ability of RNA to
serve as an information carrier is obvious, as it has the same code as DNA and
is even used as such in viruses; RNA’s ability to serve a role analogous to
modern-day proteins was not so obvious. Indeed when this phenomenon was
first encountered, it was disbelieved by many and thereafter viewed as
mysterious.
The difficulty in appreciating RNA as a functional, catalytic molecule
stemmed from both the lack of familiarity with RNA structure [crystal structures
were available only for tRNA (Robertus et al., 1974; Suddath et al., 1974; Giege
et al., 1977; Hingerty et al., 1978; Sussman et al., 1978; Woo et al., 1980)] and
2
from the absence of the protein side chains typically considered as ‘catalytic’.
However, recognizing the ability of a macromolecule distinct from proteins to
adopt globular structures and carry out function has provided a powerful
conceptual counterpoint. The differences between proteins and RNA have
helped us better understand each macromolecule, and the similarities have
helped clarify fundamental features of biological catalysis that extend beyond the
identification of the so-called “catalytic residues”.
In this chapter we use the group I intron to illustrate the interplay between
RNA structure and function. We emphasize the group I intron from Tetrahymena
thermophila, as many of the techniques used in RNA and conceptual
understanding of RNA biochemistry derive from research on this intron. We first
provide an historical overview of the early work on group I RNA structure. We
then turn to the wealth of functional studies with this intron. Finally we place
these functional studies into the context of the recent atomic level structures of
group I introns and articulate future challenges for research in this area.
GROUP I INTRON STRUCTURAL STUDIES
Atomic resolution structures of proteins from x-ray crystallography are now
commonplace and an increasing number of RNA structures are being solved,
including the immense ribosome. Nevertheless, it remains time-consuming and
challenging to solve RNA structures by x-ray and NMR approaches.
Consequently, much structural information about RNA comes from solution
3
structural probing experiments and from phylogeny. The resulting structural data,
while lower in resolution than that obtained by x-ray crystallography, can be used
to guide experiments, to test whether structural features observed in crystals
occur in solution, to correlate functional consequences with changes in structural
features, and to follow structural features associated with kinetic and
thermodynamic studies in solution.
In this section we develop as a case study the historical emergence of
RNA structures derived from phylogenetic and chemical probing studies of the
Tetrahymena group I intron. We hope that this historical perspective is instructive
by introducing techniques, by providing insight into the power and limitations of
these techniques, and by suggesting areas for future development.
Building structural models: Secondary structure from phylogeny and
beyond.
Structure predictions of the group I intron and many other RNAs have
relied on phylogenetic comparisons, initially guided by knowledge of base pairing
rules and later by knowledge of motifs that engage in tertiary interactions (Levitt,
1969; Fox and Woese, 1975; Woese et al., 1980; Branlant et al., 1981; Noller et
al., 1981; Davies et al., 1982; Michel et al., 1982; James et al., 1988; Michel and
Westhof, 1990; Romero and Blackburn, 1991; Gautheret et al., 1995; Lehnert et
al., 1996; Batey et al., 1999; Lilley, 1999; Lee et al., 2003). Phylogenetic
analyses use RNA sequences that show some degree of overall conservation to
identify positions of covariation among residues (Michel and Costa, 1998). For
4
example, if two residues within an RNA sequence are G and C or C and G, but
never G and G or C and C, the residues “covary”. If the covariation is consistent
with base pair formation and if neighboring residues are also consistent with
base pair formation, the presence of a duplex in that region is strongly implied.
Indeed, the original demarcation of the class of introns referred to as “group I”
came from such phylogenetic comparisons, independent of the discovery of the
intron’s catalytic activity (Davies et al., 1982; Michel et al., 1982). Figure 2a
shows one of the two originally proposed secondary structures from phylogenetic
covariation (Michel and Dujon, 1983). Below we describe how these structures
were further developed into models that include secondary and tertiary structure
for the Tetrahymena and other group I introns.
The local and strong nature of RNA helices greatly facilitates our ability to
predict RNA secondary structure from phylogeny and mutagenesis. For proteins,
while residues have differential preferences for α-helix formation, these energetic
preferences are modest, and α-helices are marginally stable or unstable as
isolated entities (Rohl et al., 1996; Fersht, 1999). Further, the energetic “rules” for
β-sheet formation are dominated by non-local side chain interactions (Minor and
Kim, 1994; Smith and Regan, 1995; Smith and Regan, 1997). Thus, secondary
structure predictions for proteins are difficult (Petsko and Ringe, 2004). In
contrast, RNA secondary structure is more stable thermodynamically. Extensive
thermodynamic comparisons of short RNA duplexes (“melting studies”) have led
to comprehensive ‘nearest neighbor’ rules for duplex stability. Because local
interactions dominate the energetics, these rules afford a high predictive ability
5
for duplex stability based only on consideration of the identity of the base pair
and its neighbor to either side (Freier et al., 1986; Mathews et al., 1999).
In 1983, concomitant with publication of a possible secondary structure for
the Tetrahymena group I intron from phylogenetic comparisons (Michel and
Dujon, 1983; Waring et al., 1983), Cech et al. published a secondary structure
model from energy minimization and additional experimental constraints derived
from the intron’s sensitivity to single- and double-strand specific nucleases [Fig.
2b; (Cech et al., 1983)]. This combination of energy minimization and nuclease
mapping predicted a secondary structure that was quite similar overall to the
model derived from phylogeny. The color coding in Figure 2 corresponds to the
complementary helical elements now known to be formed, and the performance
of each of the models in predicting helices is summarized in Table 1.
The most notable difference between the two models (Fig. 2a & 2b) was
the absence of the P3 helical element from the model of Cech et al. (1983). P3 is
a “pseudoknot’ (Pleij et al., 1985), a long-range base pairing interaction that
could not have been found by the traditional energy minimization program for
secondary structure prediction due to sampling constraints in these procedures.
Currently approaches are being developed to overcome this sampling limitation
(Rivas and Eddy, 1999; Xayaphoummine et al., 2003). Additional limitations of
current computational approaches must also be recognized. Whereas the
algorithms contain extensive information about the energetics of duplexes with
Watson-Crick base pairing, much less information about wobble pairs, loops and
bulges is available. Consequently, junctions, loops, mismatches, electrostatics,
6
metal ion binding sites, and tertiary motifs are not included or have incomplete
energy functions. A general and important lesson to be drawn is that one must be
aware of the information used to create a model in order to understand the power
and limits of its predictive value.
Over time, many of the predicted helical regions of the Tetrahymena intron
were tested via mutagenesis and are summarized in Table 1 (Been and Cech,
1986; Burke et al., 1986; Waring et al., 1986; Williamson et al., 1987; Flor et al.,
1989; Michel et al., 1989; Williamson et al., 1989; Burke et al., 1990; Michel et al.,
1990; Suh and Waring, 1990). The logic of the mutagenesis experiments is
simple and parallels that for phylogenetic comparisons. If a G•C base pair is
suspected, mutation of G to C, or C to G, is predicted to be detrimental, whereas
the double mutation, G•C to C•G, is predicted to restore function (or structure).
As is generally the case, no strong conclusion can be made from the disruption
alone –there are many possible explanations for the loss of function. In contrast,
it is improbable that a second mutation would restore function via an indirect
effect; thus, rescue provides strong evidence (though not proof) of a direct
interaction. These experiments provided strong support for many of the helical
assignments inferred from phylogeny and shown in Figure 2.
The P3 helix, suggested in the original phylogenetic secondary structures,
was also tested by mutagenesis and largely confirmed, although modified in the
exact pairing (Williamson et al., 1987). There were also special cases where
implications of the base rescue experiments went beyond simply demonstrating
the importance of the helix. For example, phylogenetic and mutagenetic data
7
provided evidence for helix formation in the nucleotides flanking the C109-G212
base pair, but mutation in the C109-G212 base pair itself did not exhibit rescue,
leading to the suggestion that this base pair participates in tertiary interactions. In
other words, the base pair could not be changed to all other Watson-Crick pairs
because additional interactions are made by the non-base pairing faces (Flor et
al., 1989). This prediction was confirmed by the recent group I intron crystal
structures, which show a tertiary contact between the C109-G212 base-pair and
residue C260 [Tetrahymena numbering; (Adams et al., 2004a; Guo et al., 2004;
Golden et al., 2005)].
Most generally, “rescue” experiments have been extraordinarily powerful
in delineating the relationship between RNA structure and function, providing
information about interactions of individual functional groups and metal ions (see
“GROUP I INTRON FUNCTIONAL STUDIES” and “INTEGRATING
STRUCTURAL AND FUNCTIONAL STUDIES OF GROUP I INTRONS” below).
Moreover, the ability to carry out these types of experiments with populations of
molecules has allowed multiple positions to be probed simultaneously in a single
experiment (Christian and Yarus, 1992; Berzal-Herranz et al., 1993; Gaur and
Krupp, 1993; Conrad et al., 1995; Strobel and Shetty, 1997; Ryder and Strobel,
1999).
Building structural models: Extension to a tertiary structure model.
In 1987 Kim and Cech, starting from the existing secondary structure from
phylogenetic comparisons and mutagenic tests, modeled the three dimensional
8
structure of the Tetrahymena intron (Kim and Cech, 1987); this model is shown in
Figure 3. To guide the modeling, Kim and Cech used information about residue
accessibility from chemical and enzymatic probes, a predilection to place the
most conserved elements together in a catalytic core or center, and a bias for
stacking of helices following observations for tRNA (Robertus et al., 1974;
Suddath et al., 1974). Although the complete structure was not modeled, the Kim
and Cech structure provided the first three-dimensional perspective for a catalytic
RNA. The most novel feature of this structure was the side-by-side packing of
RNA helices, a feature not present in tRNA. Although many of the details have
required revision, including the proposed tertiary interactions, this general feature
of the model and the general disposition of helices in the core were correct.
Retrospectively, the hypothesis of side-by-side packing of helices is an
interesting one. On the one hand, it seems obvious that this or similar packing
would have to occur. After all, enzyme active sites are located in crevices or
cavities, and for RNA to create an active site that behaves analogously, a similar
architecture would be expected (Narlikar and Herschlag, 1997). On the other
hand, it was difficult to envision how RNA could pack efficiently in this manner
given the regular architecture of RNA helices, its limited side chain diversity, and
the expectation that RNA’s negative charge would repel the close approach of
other strands. Moreover, because RNA packing had not been observed
previously, it would have been difficult to distinguish whether this absence was
fundamental or whether the appropriate experiment had yet to be performed.
9
Regarding this latter perspective, the experimental evidence in 1989 for an
“inside” and “outside” of the Tetrahymena intron was greeted with excitement.
Latham and Cech used Fe(II) chelated by EDTA to generate hydroxyl radicals
(HO•) in solution (Latham and Cech, 1989). Tullius and coworkers had provided
evidence that these radicals could indiscriminately react with the RNA backbone
to cause strand scission, an event easily read out by gel electrophoresis (Tullius
and Dombroski, 1985). Latham and Cech simply added Fe•EDTA to solutions of
folded and unfolded intron; they observed regions of protection from cleavage
only when the intron was folded by addition of Mg2+, and these protected regions
were much more extensive than those observed in a tRNA control. Hydroxyl
radical footprinting has since served as a powerful tool in understanding RNA
structure, thermodynamics and folding kinetics (Celander and Cech, 1991; Sclavi
et al., 1998; Ralston et al., 2000; Brenowitz et al., 2002; Takamoto et al., 2004).
At this point it was clear that the intron had a three dimensional structure
and that this structure was critical for function. But questions remained – what
was the structure? How was RNA put together to act as a catalyst? Once again,
phylogeny proved extremely powerful in developing a model for the global
architecture of the intron’s core. In 1969, even before solution of the x-ray
structure of tRNA, Levitt used phylogenetic comparisons to predict long range
base pairs and base triples, thereby developing a tRNA model of remarkable
accuracy (Levitt, 1969). We now describe how Michel, Westhof and coworkers
used phylogeny, mutagenesis, and other functional data, along with common
sense, to develop a remarkable tertiary structure model for the Tetrahymena
10
group I intron (Michel and Westhof, 1990). In the process, they also uncovered
general motifs that allowed modeling of structures for other group I introns and
other RNAs (Michel and Costa, 1998).
Computational work has demonstrated that a small number of the ‘right’
distance constraints can be enormously powerful in distinguishing between
possible three dimensional models (Joseph et al., 2000). Michel and colleagues
recognized that the splicing reaction itself gives a long range distance constraint
because the 3'
-splice site, defined by a universally conserved guanosine residue
at the intron-exon junction, must come together with the 5'
-splice site (see
“GROUP I INTRON FUNCTIONAL STUDIES” below for details). With this insight,
Michel and colleagues reasoned that the binding site for this guanosine must lie
in close proximity to the groups within the intron shown to interact with the 5'
splice site.
The hunt for the guanosine binding site entailed two steps. First, it was
found that most group I introns contain residues that can form base pairs with
residues immediately 5'of the ωG (Burke, 1989; Michel et al., 1989; Michel et al.,
1990). This phylogenetic suggestion was confirmed by compensatory mutations
(Michel et al., 1989; Burke et al., 1990), and the new helical element was called
P9.0 (Fig. 2d). This helical element was likely missed in earlier phylogenetic
analyses because of its short length and the limited number of sequences
available.
As the two residues immediately 5'of the conserved guanosine form one
strand of P9.0, the position of the residues on the other strand now served as a
11
strong clue for locating the G-binding site. Michel and co-workers further
reasoned that the binding site for the universally conserved guanosine residue
would also likely be universally conserved (Michel et al., 1989). Of the two
invariant residues not directly involved in splice site selection, A261 and G264
(base paired to C311 in P7) in the Tetrahymena intron, mutation of G264-C311 to
A264-U311 relaxed discrimination between splicing after guanosine or adenosine,
whereas mutation of A261 while deleterious did not relax this specificity. These
experiments were suggestive of guanosine binding to G264-C311, and allowed a
more direct experimental test to be carried out: the specificity switch, or “rescue”
experiment outlined in Figure 4. If the reactive guanosine were to bind the G264C311 via a base triple (Fig. 4a), then mutation to A264-U311 would decrease
binding of guanosine and increase binding of 2-aminopurine ribonucleoside (Fig.
4b-d). Indeed, this prediction was born out, providing strong evidence for
localization of the G-binding site to the G264-C311 base pair of P7 (see also
“INTEGRATING STRUCTURAL AND FUNCTIONAL STUDIES OF GROUP I
INTRONS” below).
The constraint of the G-binding site and other phylogenetic, mutagenic,
and functional data allowed Michel and Westhof to derive a more accurate and
higher resolution three dimensional model for the intron core than the Kim and
Cech model (Michel and Westhof, 1990). However, progress did not stop there;
recognition of motifs used in both group I and group II introns and additional
phylogenetic comparisons followed by mutagenesis identified long range tertiary
interactions (Jaeger et al., 1991; Jaeger et al., 1994; Murphy and Cech, 1994;
12
Costa and Michel, 1995; Lehnert et al., 1996; Ikawa et al., 2000b). These new
interactions were combined with data on the accessibility of riboses to cleavage
by Fe(II)-EDTA (Latham and Cech, 1989; Celander and Cech, 1991; Heuer et al.,
1991; Murphy and Cech, 1993; Laggerbauer et al., 1994), and to refinements of
the central base triple interactions (Pyle et al., 1992), allowing construction of a
three dimensional model for the entire Tetrahymena intron (Lehnert et al., 1996).
The three dimensional model for the Tetrahymena intron (Fig. 5a)
predicted an overall architecture of the molecule that was later confirmed by xray crystallography [(Adams et al., 2004a; Guo et al., 2004; Golden et al., 2005);
see also “INTEGRATING STRUCTURAL AND FUNCTIONAL STUDIES OF
GROUP I INTRONS” below]. Two sets of coaxially stacked helices make up the
“core” (Fig. 5b, P3-P8, in blue, and P4-P6, in green). These helices are
conserved in all group I introns, whereas the other “peripheral” helices are
conserved only in subclasses of group I introns (Burke, 1988; Cech, 1988; Cech,
1990). As would be expected, the conserved regions make up the active site,
forming a crevice where they come together that allows binding of guanosine and
docking of the P1 duplex that contains the 5'
-splice site (Fig. 5c). The peripheral
helices literally wrap around the core in three dimensions (Fig. 5d & 5e), with
tertiary interactions that connect each of these elements to form a ring around
the core. Because of the recurring motifs, such as tetraloop-tetraloop receptors
and long range base pairing, models for the connectivity of the three dimensional
structures of each of the classes of group I introns are strongly suggested
13
(Jaeger et al., 1993; Costa and Michel, 1995; Lehnert et al., 1996; Westhof et al.,
1996; Batey et al., 1999; Hermann and Patel, 1999).
Perspectives from building structural models.
The odyssey to the group I structural model has provided us with powerful
tools and new insights that can be used to develop models for other RNA
structures and to probe further the structural and functional behavior of the
Tetrahymena intron and other RNAs. Phylogeny can often provide reliable
information on secondary and tertiary structural interactions. Importantly, models
from phylogeny can be tested by mutagenesis rescue experiments. Chemical
and enzymatic structure mapping provides additional constraints, and the
definition of the inside and outside of an RNA molecule by hydroxyl radical
footprinting is especially powerful and has been further applied in studies of
folding thermodynamics and kinetics. A major challenge remains to integrate
these approaches to create a powerful algorithm for solving RNA structures in
solution.
This body of work also helps clarify other substantial challenges that
remain. Although the overall architecture is clear, the energetics that allow this
structure to form cooperatively are not, nor are the interactions or properties that
allow the peripheral elements to enforce formation of active core structure
(Engelhardt et al., 2000). Further, structural models such as that shown in Figure
5a can help in choosing mutations and help guide other experiments, but the
level of information provided by phylogeny and structure mapping is insufficient
14
to provide a detailed mechanistic understanding. As noted above, this limitation
is to be expected as atomic level information from these approaches is far from
comprehensive. For example, while phylogeny identified the tetraloop-tetraloop
receptor motif, the x-ray structure of the P4-P6 domain of the group I intron (Cate
et al., 1996a) revealed a different atomic architecture than had been modeled
(see “INTEGRATING STRUCTURAL AND FUNCTIONAL STUDIES OF GROUP
I INTRONS” below).
In the next section, we examine local interactions and catalytic
mechanisms suggested by studies employing powerful functional approaches. In
the final section we integrate the information from the functional approaches with
the atomic-level structural picture of the entire intron, as recently revealed by xray crystallography.
GROUP I INTRON FUNCTIONAL STUDIES
The group I self-splicing reaction, shown in Figure 6, encompasses many
common features of RNA biochemistry and chemistry (Cech et al., 1981; Kruger
et al., 1982). The intron must fold into its correct native state to be active (Fig. 6,
step 1), uses metal ions in a chemical step, and undergoes conformational
rearrangements that are necessary for function. The 5'
-splice site is recognized
by base pairing to an intron element referred to as the “internal guide sequence”
(IGS), formation of a specific G•U wobble pair, and tertiary interactions that place
the G•U wobble pair in the correct binding register for attack by G (Been and
15
Cech, 1986; Waring et al., 1986; Zaug et al., 1986; Barfod and Cech, 1989;
Doudna et al., 1989; Murphy and Cech, 1989; Pyle and Cech, 1991; Young et al.,
1991; Downs and Cech, 1994; Knitt et al., 1994). The intron binds an exogenous
guanosine cofactor (G) that is used to cleave the RNA strand at the 5'
-splice site
(Fig. 6, step 2). In the first chemical step, the guanosine attacks at the 5'
-splice
site, attaching itself to the 5'
-terminus of the intron while severing the intron’s
covalent connection to the 5'
-exon (Fig. 6, step 3).
Subsequent conformational rearrangements prepare the intron
intermediate for the second chemical step. The newly added guanosine and 5'
end of the intron dissociate from the active site and are replaced by the
guanosine residue that defines the intron’s 3'
-terminus (referred to as “ωG”).
Intron residues immediately upstream of the ωG form base pairs with residues
adjacent to the guanosine binding site to form helical element P9.0, as noted
above, thereby facilitating choice of the correct 3'
-splice site [Fig. 6, step 4;
(Michel et al., 1989; Burke et al., 1990)]. 3'
-Splice site choice is further facilitated
by formation of base pairs between the 5'most residues of the 3'
-exon,
immediately 3'of ωG, and residues of the IGS vacated by the 5'end of intron
(Partono and Lewin, 1990; Suh and Waring, 1990).
The intron is now ready to carry out the second chemical step, which is in
essence the reverse of the first step. The 5'
-exon attacks the 3'
-splice site, with
ωG serving as the leaving group, resulting in ligation of the 5'
- and 3'
-exons (Fig.
6, step 5). Following a dissociation step (Fig. 6, step 6), this reaction yields
ligated exons and free intron. Thus, the intron uses base pairing, helical
16
interactions, tertiary contacts, and other structural elements both to select and
align the 5'and 3'splice sites during the course of the self-splicing.
Despite the important features of RNA structure and function represented
in self-splicing, this reaction has been difficult to study. Uhlenbeck first showed
that the intron can fold into a long-lived inactive conformer in competition with
formation of the native state (Walstrum and Uhlenbeck, 1990), mirroring earlier
studies of tRNA misfolding (Gartland and Sueoka, 1966; Lindahl et al., 1966;
Adams et al., 1967; Ishida and Sueoka, 1968). Indeed, misfolding in vitro is a
common feature of RNA and is thought to arise from the high thermodynamic
and kinetic stability of local secondary structures, and from the ability to form
additional interactions promiscuously, a manifestation of the abundance of
potential hydrogen bonding, stacking and metal ion binding interactions within
RNA (Sigler, 1975; Herschlag, 1995). Further, the commonality of RNA
misfolding coupled with its apparent fundamental physical origins led to the RNA
chaperone hypothesis, which posited the necessity of proteins that act to prevent
and/or resolve misfolded RNAs in vivo (Karpel et al., 1974; Karpel et al., 1982;
Herschlag, 1995). Several of the seminal studies providing strong support for this
hypothesis have been carried out with group I intron systems [(Coetzee et al.,
1994; Mohr et al., 1994; Caprara et al., 1996; Waldsich et al., 2002); reviewed in
(Schroeder et al., 2002; Schroeder et al., 2004)]. Thus, on the one hand the
group I intron has provided an opportunity to explore the conformational behavior
of RNA, but on the other hand this conformational behavior stood in the way of
obtaining molecular understanding of intron function. Indeed, one of the
17
important current challenges in RNA research is to understand the
conformational rearrangements that appear to occur in essentially all RNA
mediated processes, such as pre-mRNA splicing, translation, and protein
trafficking via the signal recognition particle (SRP). The knowledge obtained
about group I introns over the past two decades renders them an attractive
system for future studies to unravel fundamental features of RNA conformational
changes.
A key step that opened up the ability to study group I introns and other
RNA catalysts was the transformation of self-splicing and self-cleaving RNAs into
two-part systems with a separate catalyst and substrate (Fig. 7a)(Uhlenbeck,
1987; Zaug et al., 1988). The conversion to trans-acting ribozymes allowed
favorable renaturation conditions and favorable reaction conditions to be
established through separation of RNA folding and cleavage. Equally as valuable,
such ribozymes allow the concentrations of reactant and catalytic components to
be varied independently and a wide variety of synthetic substrates to be used.
The critical importance of these features is underscored below as we describe
what has been learned from functional studies of the Tetrahymena group I
ribozyme.
Technical advances have also been critical for all aspects of
understanding group I intron structure and function. The ability to transcribe
RNAs from synthetic and plasmid DNA templates allowed the effects of
sequence variation to be probed and allowed large amounts of RNA to be
obtained for functional studies and crystallographic and NMR structural studies
18
(Milligan and Uhlenbeck, 1989; Gurevich, 1996); the ability to carry out solid
phase synthesis of RNA and its analogs provided further chemical control for
functional studies [for reviews see (Letsinger and Mahadevan, 1965; Usman et
al., 1987; Verma and Eckstein, 1998; Muller et al., 2004)]; and the ability to ligate
together transcribed and synthetic RNAs has allowed mutagenesis at the level of
individual functional groups even for RNA catalysts too long to synthesize by
solid phase methods (Moore and Query, 2000; Sherlin et al., 2001). Populationbased screens and selections have also been developed with RNA, allowing
experiments ranging from the selection of new catalysts to the generation of
artificial phylogenies and testing of the roles of individual functional group
substitutions at multiple positions in parallel, as detailed further in
“INTEGRATING STRUCTURAL AND FUNCTIONAL STUDIES OF GROUP I
INTRONS” below [for reviews and representative examples see (Waring, 1989;
Christian and Yarus, 1992; Ekland and Bartel, 1996; Strobel, 1999; Schwans et
al., 2003; Joyce, 2004)]. Finally, the ease of chemical manipulation and wellunderstood base pairing properties of nucleic acids have facilitated numerous
single molecule studies of RNA and protein / nucleic acid systems [for examples
see (Ha et al., 1999; Zhuang et al., 2000; Ha et al., 2002; Bartley et al., 2003;
Tan et al., 2003; Blanchard et al., 2004; Bokinsky et al., 2004; Nahas et al.,
2004)]. Again, while only a subset of technical advances is addressed herein, all
of the functional and structural studies described have relied on such advances.
19
A thermodynamic and kinetic framework for the Tetrahymena group I
ribozyme reaction.
As for any new enzyme, early work on the Tetrahymena ribozyme focused
on establishing assays and reaction conditions (Zaug et al., 1986; Zaug et al.,
1988; Herschlag and Cech, 1990b). As is also common, the initial steady state
kinetic results were largely uninformative, and at times even misleading and
incorrect. The fundamental limitation of steady state kinetics is most simply
described in terms of the information content of the experiment – i.e., the number
of parameters determined relative to the number of variables that need to be
determined to describe the system’s behavior. In this case the information is kcat
and Km (for each substrate), whereas there are typically many more steps in a
reaction scheme, including binding, the chemical step, conformational steps, and
dissociation –this complexity cannot be sorted out from two steady state rate
parameters. Rather, pre-steady state kinetic approaches have been developed in
enzymology to isolate individual reaction steps and dissect reaction mechanisms
(Fierke and Hammes, 1995; Johnson, 1995; Johnson, 1998; Fersht, 1999). The
pre-steady state kinetic analysis of the Tetrahymena group I ribozyme utilized
traditional methodologies, such as rapid mixing and “pulse chase” experiments,
and also utilized new variations of these methods. The resulting kinetic and
thermodynamic framework, outlined in Figure 8, rivals that for the most intensely
studied protein enzymes. Below we describe the framework and some of the
insights derived from it. The reader is referred to the original papers and texts for
information on the presteady state kinetic methodologies used (Herschlag and
20
Cech, 1990b; Herschlag, 1992; Knitt and Herschlag, 1996; Mei and Herschlag,
1996; Narlikar et al., 1997; Narlikar et al., 1999; Karbstein et al., 2002; Karbstein
and Herschlag, 2003).
In Figure 8 “S” is an oligonucleotide that mimics the 5'
-splice site and is
cleaved by guanosine (G) in the first self-splicing step (cf. Fig. 7b). The products
are the dinucleotide GpA and “P”, an analog of the 5'
-exon (which is ligated to the
3'
-exon in the second step of self-splicing). As elaborated below, all reactions
entail attack by a 3'
-OH group, leading to formation and breakage of standard 3'
5'
-phosphodiester RNA bonds. Either ribozyme substrate, S or G, can bind first.
Interestingly, neither binds at the diffusion-controlled limit of ~1010 M-1 min-1, a
rate constant approached for binding of ligands to many protein enzymes (Fersht,
1999). Instead, both ribozyme substrates have been shown to bind by multi-step
processes with considerably slower observed rate constants (108 M-1 min-1 for S
and P; 106 M-1 min-1 for G) (Bevilacqua et al., 1992; Herschlag, 1992; Bartley et
al., 2003; Karbstein and Herschlag, 2003).
For S, an ‘open complex’ forms first, whereby S is held in place solely by
base pairing interactions with the IGS (Fig. 7b); to a reasonable approximation,
the resulting duplex, referred to as the P1 helix, behaves just like a free duplex in
solution, and binding occurs at a rate similar to that for duplex formation between
short oligonucleotides, ~108 M-1 min-1 (Herschlag and Cech, 1990b; Narlikar et al.,
1999). In a second distinct step, the P1 helix docks into tertiary interactions with
the ribozyme’s core (Bevilacqua and Turner, 1991; Pyle and Cech, 1991;
Bevilacqua et al., 1992; Herschlag, 1992; Pyle et al., 1992; Bevilacqua et al.,
21
1994; Pyle et al., 1994; Narlikar et al., 1997; Narlikar et al., 1999; Zhuang et al.,
2000; Bartley et al., 2003). These docking interactions are mediated via 2'
hydroxyl groups along the P1 helix as well as the exocyclic amine of the G•U
wobble pair, which defines the cleavage site in the first step of group I introns.
This exocyclic amine of the cleavage site G•U wobble pair has also been
suggested to play a specific role in organizing the chemical transition state for
ribozyme cleavage, as detailed below (Knitt et al., 1994; Strobel and OrtolevaDonnelly, 1999). Binding of the 5'
-exon analog product, P, follows the same two
step pathway, as it also contains the complementary sequence to the IGS.
One of the unexpected benefits of the discovery of two-step binding is that
the docking step can be viewed and studied as a simplified model for RNA
folding – the progression from secondary to tertiary structure. Dissection of
docking, including experiments at the single molecule level, has led to general
insights into RNA dynamics and thermodynamics (Narlikar and Herschlag, 1996;
Narlikar et al., 1999; Narlikar et al., 2000; Zhuang et al., 2000; Bartley et al.,
2003). One of the remaining mysteries is why docking is so slow, occurring with a
rate constant on the order of 1 s-1, much slower than expected for a
conformational search (Bartley et al., 2003).
Binding of G also occurs in multiple steps, although only one step is
shown in Figure 8 because the steps have not been dissected at the level of
establishing individual rate constants (Karbstein and Herschlag, 2003).
Mechanistic studies suggest that two-step G binding arises because the G-site is
not fully preorganized, and rather must reorganize on the tens of milliseconds
22
timescale to allow binding of G (see “INTEGRATING STRUCTURAL AND
FUNCTIONAL STUDIES OF GROUP I INTRONS” below). This slow binding
may provide an opportunity in splicing for the intron to increase 3'
-splice site
selection specificity via kinetic specificity [see also (Lindner et al., 1999)]. In
splicing, the residues upstream of ωG base pair to form helix P9.0 (Fig. 6 and
discussed above), thereby increasing the residence time of ωG near this site and
the association rate for ωG binding. Whether slow binding was selected to
increase specificity or simply reflects the inherent conformational flexibility of
RNA structure is not known. Nevertheless, these studies uncovered a previously
unrecognized mechanism for enhancing specificity, further underscoring the
value of mechanistic dissection (Karbstein and Herschlag, 2003).
The 5'
-splice site analog, S, and the 5'
-exon analog, P, both dissociate
from the ribozyme more slowly than would be expected based solely on helix
stability. Indeed, this slow dissociation was the original evidence for the
involvement of tertiary interactions in binding and has several implications for
ribozyme function. First, under most conditions where S is subsaturating, the
binding of S is rate-limiting. Additionally, under most conditions where ribozyme
is saturating, product release is rate-limiting. Thus, early steady state kinetic
experiments, instead of probing chemical catalysis by the ribozyme, followed the
physical steps of substrate binding and product release (Herschlag and Cech,
1990b; Herschlag and Cech, 1990c). Second, the slow release of S relative to its
cleavage in the ternary complex (Fig. 7b, k Soff vs kforward) results in low specificity.
Substrate analogs that bind weaker can still be cleaved prior to dissociation from
23
the ribozyme, meaning that ribozyme cleavage is limited by the substrate
association rate. As mismatches do not significantly affect association rates, the
ribozyme cleaves both matched and mismatched substrates efficiently, resulting
in low specificity (Herschlag and Cech, 1990c; Hertel et al., 1996). The
complexity of the relationship between binding affinity and specificity was
important to appreciate in the search for nucleic acid therapeutics that would
recognize specific RNA targets in vivo (Herschlag, 1991), and potentially has
significant implications for specific RNA recognition by small interfering RNA,
microRNAs, and snoRNAs. Finally, the strong binding of the 5'
-exon analog
causes product release to become rate-limiting and slows turnover. While
deleterious for a multiple turnover reaction, this slow release of P makes perfect
sense in the context of the biological self-splicing reaction: after the first chemical
step in self-splicing the 5'
-exon (which P is an analog of) is no longer covalently
attached; dissociation before ligation to the 3'
-exon would abrogate splicing (Fig.
6). Thus, the strong binding likely ensures splicing efficiency by preventing
dissociation of the 5'
-exon (Herschlag and Cech, 1990b; Herschlag and Cech,
1990c).
Once either substrate, S or G, is bound to the ribozyme, the second
substrate binds more strongly – i.e., there is thermodynamic coupling between
the substrates (this also holds for binding of the products, P and GA) (McConnell
et al., 1993; Karbstein et al., 2002). The absence of coupling between P and G
indicated that the reactive phosphoryl group was directly or indirectly involved in
coupling. Subsequent experiments have suggested that a metal ion, referred to
24
as MC (see below), helps mediate coupling by interacting with the phosphoryl
group and the 2'
-hydroxyl of G [Fig. 9c; (Shan and Herschlag, 1999)].
After binding of both substrates to form the E•S•G ternary complex, with S
in the closed complex and coupling between S and G as noted above, at least
one more event must occur prior to the chemical reaction: loss of a proton,
presumably from the attacking 3'
-hydroxyl group of G (Fig. 7c). The interactions
proposed to play roles in accelerating the chemical step are discussed below
(see “Probing the chemical step of the Tetrahymena ribozyme reaction”).
The properties of the reverse reaction (cleavage of GA by P to give G and
S) are analogous to those of the forward reaction, so we will not describe them
further here. The overall reaction equilibrium is near one, as expected for
exchange of one phosphodiester bond for another, and the equilibrium is not
greatly perturbed on the ribozyme (Karbstein et al., 2002). How then does selfsplicing proceed directionally? There are several possible contributions, including
a high cellular concentration of the guanosine nucleophile (in all of its 5'
phosphorylated forms) relative to the concentration of ligated exon and intron
products, self-processing and/or degradation of the free intron, and binding and
sequestration of the spliced exon products in downstream biological processes
(Zaug et al., 1983; Woodson and Cech, 1989). Our knowledge of how RNA
molecules are handled in cellular environments is in its infancy, and the behavior
of catalytic RNAs may provide a tool to help understand RNA behavior in vivo
(Donahue and Fedor, 1997; Brion et al., 1999; Long and Sullenger, 1999; Pichler
and Schroeder, 2002; Yadava et al., 2004).
25
In closing, the kinetic and thermodynamic framework for the Tetrahymena
ribozyme has been central in developing our understanding of the group I intron.
Interpretation of information from substrate analog behavior, ribozymes with sitespecific mutations or chemical changes, and ribozymes with sequences deleted
or added all rely on knowing what steps are being followed and build on previous
characterization of these steps. Even the choice of RNA constructs to use in xray crystallography can be guided by activity measurements in an effort to obtain
uniform populations of molecules to facilitate crystallization. If it is easy to be
mislead about an isolated system such as a group I ribozyme, as illustrated by
the early kinetic studies of this ribozyme, imagine the care and tenacity required
to dissect and understand more complex biological processes such as premRNA splicing by the spliceosome, translation and other aspects RNA
processing! Certainly mechanistic approaches will need to play more and more
central roles in biological investigations.
Revealing catalytic principles through comparison of RNA and protein
enzymes.
In retrospect it should not have been a surprise to find that RNA molecules
can act as catalysts. Although the occurrence of RNA catalysts in Nature was
(arguably) not predictable (Crick, 1968; Orgel, 1968), the ability of RNA to
provide catalysis was. Upon discovery of catalysis by the Tetrahymena group I
intron and the RNase P RNA there was much focus on the absence of the
functionalities and diversity of RNA side chains compared to proteins. For
26
example, RNA lacks groups with solution pKa values near neutrality, as would be
optimally suited for catalysis of proton transfers via general acid and base
catalysis (Jencks, 1987; Narlikar and Herschlag, 1997; Fersht, 1999; Bevilacqua,
2003). While such comparisons are instructive about the make-up and properties
of these distinct biopolymers, ignored from this point of view is the generalization
that protein enzymes use multiple catalytic strategies to accomplish their
enormous rate enhancements. Thus, an alternative catalyst suboptimal in one or
more catalytic strategy could still achieve substantial catalysis via other
strategies. In keeping with this point of view, the initial mystique of Mg2+ as
central to all of RNA catalysis has given way to a more mature view in which
RNA can use multiple catalytic strategies including electrostatic interactions with
Mg2+ ions, general acid-base catalysis, and positioning of reactants and catalytic
groups [reviewed in (Narlikar and Herschlag, 1997; DeRose, 2002; Doudna and
Cech, 2002; Fedor, 2002; Pyle, 2002; Lilley, 2003; Bevilacqua et al., 2004)].
History was to repeat itself in considering DNA as a possible catalyst.
Given the observed differences between RNA molecules with an expanding
richness of structures, versus DNA with its rather pedestrian double strand, much
was made of the special character endowed to RNA by its 2'
-hydroxyl groups.
But ignored in these discussions was the fact that DNA has been selected by
Nature to be ‘boring’ –i.e., to have a regular repeating structure, whereas RNA
has a diverse array of functions. As with the comparison of proteins to RNA, the
importance of RNA’s 2'
-hydroxyls for structure and function does not mean that
one can’t have structure and function without them. But many points are obvious
27
in hindsight, and it took the discovery of catalytic DNAs via in vitro selections to
make scientists comfortable with this notion (Breaker and Joyce, 1995; Cuenoud
and Szostak, 1995; Santoro and Joyce, 1997; Geyer and Sen, 2000; Feldman
and Sen, 2001; Emilsson and Breaker, 2002).
Indeed, cyclodextrins, micelles, and alternative solvents have
demonstrated catalytic ability [for reviews and examples see (Oconnor et al.,
1974a; Oconnor et al., 1974b; Jencks, 1987; Breslow, 1991; Komiyama, 1993;
Breslow, 1994; Tee, 1994; Allohedan and Kirby, 1995; Rathman, 1996)], and
even single stranded nucleic acids can achieve modest catalysis by providing a
base pairing template (Sulston et al., 1968; Chunag et al., 1971; Rosenbaum and
Liu, 2003; Gartner et al., 2004). Thus, the surprise in discovering RNA as a
catalyst arose from a perspective derived from biological dogma at the time –that
all enzymes are proteins, which obscured the perspective from chemistry of
multiple catalytic strategies and catalysts. In general, approaching a problem
from multiple perspectives can deepen understanding and help avoid conceptual
traps.
Indeed, RNA enzymes have provided a vantage point to biological
catalysis that has been instrumental in revealing distinct properties of RNA and
protein molecules and in reinforcing and deepening our understanding of
properties that are fundamental to biological catalysis (Narlikar and Herschlag,
1997). Both protein and RNA enzymes use binding interactions and binding
energy to facilitate catalysis. This most fundamental concept, widely discussed
for protein enzymes, was in fact first directly demonstrated with the hammerhead
28
and Tetrahymena group I ribozymes (Hertel et al., 1997; Narlikar and Herschlag,
1998). Further, the Tetrahymena ribozyme appears to use binding interactions
for positioning of reactants with respect to one another and with respect to active
site functional groups. Additionally, these interactions create destabilization in the
ground state that facilitates catalysis as it is relieved in the transition state
(Narlikar et al., 1995; Narlikar and Herschlag, 1998). These fundamental aspects
of catalysis have been reviewed in depth elsewhere (Narlikar and Herschlag,
1997; Kraut et al., 2003) and are therefore not further elaborated herein.
The final generalization that comes from the world of protein catalysts is
that active sites occur in cavities or crevices. This makes sense from the
standpoint of allowing multiple interactions between catalyst and substrates for
positioning and chemical catalysis. This generality holds for the handful of RNA
enzymes with x-ray structures, with the lone exception of the hammerhead
ribozyme. Some structures of this ribozyme depict its reactive phosphoryl group
projecting into solution away from other conserved residues (Pley et al., 1994;
Scott et al., 1995; Scott et al., 1996); this observation lends further credence to
the substantial functional work suggesting that these x-ray structures do not
reflect the catalyst’s active conformation [(Peracchi et al., 1998; Wang et al.,
1999; De la Pena et al., 2003; Hampel and Burke, 2003; Khvorova et al., 2003;
Canny et al., 2004; Penedo et al., 2004; Heckman et al., 2005); reviewed in
(Uhlenbeck and Blount, 2005)].
Probing the chemical step of the Tetrahymena group I ribozyme reaction.
29
Finding conditions for monitoring the chemical step. As noted above, the first
critical step in studying the mechanism of chemical catalysis by the Tetrahymena
ribozyme was finding conditions under which binding and product release were
not rate-limiting, so that the chemical step could be followed. The known binding
and conformational steps (Fig. 8) are pH independent in the region between pH 6
and 8 (Herschlag and Khosla, 1994; Narlikar et al., 1999; Karbstein and
Herschlag, 2003), and the pH dependencies below and above this range appear
to reflect multiple nucleobase protonations and deprotonations that result in
inactivation of the ribozyme (Knitt and Herschlag, 1996). Conversion of the
ternary complex to products increases in rate log-linearly with pH with a slope of
one, consistent with the loss of one proton prior to the rate-limiting transition state
(Jencks, 1987). Herschlag and Khosla suggested that under these conditions,
the chemical step limits the ribozyme reaction rate and postulated that the
attacking 3'
-oxygen of G might interact with a metal ion (a model that was later
supported, as described below; Fig. 9b) (Herschlag and Khosla, 1994). Therefore,
the simplest model that could account for the observed pH dependence was loss
of the proton from the 3'
-hydroxyl group of G prior to the chemical step (Fig. 7c).
Further support for a rate-limiting chemical step under certain reaction
conditions came from the observation of a thio-effect – i.e., a rate effect from
sulfur substitution of the pro-RP nonbridging phosphoryl oxygen atom at the
cleavage site; the rate decrease of ~3 fold is of the same magnitude as the
intrinsic effect of thio-substitution on the nonenzymatic reactions of phosphate
diesters (Herschlag et al., 1991). Finally, the same pH dependence and thio-
30
effects were observed for substrates that varied in reactivity over 103-fold,
providing no indication of a rapid chemical step masked by a rate-limiting
conformational step (Herschlag and Khosla, 1994). Nevertheless, conformational
steps can also be pH-dependent (Bayfield et al., 2001; Xiong et al., 2001;
Rodnina and Wintermeyer, 2003) so that claims that a log-linear pH dependence
alone is indicative of a chemical rate-limiting step, while reasonable, are overinterpretations of pH-rate data.
Finding transition state interactions: Metal ion rescue and beyond. It is well
known in science (and in life!) that it is much easier to mess something up than to
fix it. In experiments, it is much easier to find factors that give deleterious effects
than factors that enhance a particular activity. Even if discovered, such
enhancements often end up having mundane explanations; for example, the
increased value of kcat for Tetrahymena ribozymes with mutated sequences
arose not because interactions in the chemical step were improved but rather
because rate-limiting product release was rendered faster via a disruption of the
tertiary contacts that strengthen binding (Young et al., 1991).
Because there are many ways to disrupt catalysis upon mutagenesis or
atomic level substitution, such effects are often difficult to interpret. However,
experiments that probe the rescue of an atomic substitution provide an exception,
as the deleterious effect is “rescued” –i.e., the activity is restored by a second
change. Logically this is analogous to second site revertants in genetics and
phylogenetic or mutagenic analysis of base pairing in RNA described above: if a
31
G•C base pair is suspected, mutation of the G to U or the C to A are each
deleterious but mutation of both together to give an U•A base pair will restore or
“rescue” activity. A negative result does not show the absence of a base pair, as
there may be additional interactions or alternative folds that lead to a preference
for a particular base pair identity. In contrast, a positive result can be strongly
suggestive.
Rescue of sulfur incorporation by soft metal addition was first used in
independent experiments of Mildred Cohn and Fritz Eckstein with protein
enzymes (Eckstein, 1970; Burgers and Eckstein, 1979; Cohn et al., 1982; Jaffe
et al., 1982; Eckstein, 1983; Eckstein, 1985). In this approach, a phosphoryl
oxygen atom is replaced with sulfur (Scheme 1). If the oxygen atom interacts with
a Mg2+ ion, function may be compromised because the “soft” sulfur atom does
not interact strongly with a “hard” metal ion like Mg2+ (Pearson, 1963). However,
replacement of the Mg2+ by a softer metal ion such as Mn2+ or Cd2+ can restore
activity, provided that the steric and geometrical differences do not interfere too
greatly with function. Nitrogen substitutions can also be used, as nitrogen is also
a softer metal ligand than oxygen.
Figure 9b shows the three atomic positions within the group I ribozyme
transition state that gave metal ion rescue in early experiments (Piccirilli et al.,
1993; Sjogren et al., 1997; Weinstein et al., 1997). Additionally, thio-substitution
of the pro-SP oxygen atom gave a large deleterious effect of ~104-fold, in contrast
to the 2-3 fold effect for the pro-RP oxygen atom described above (Rajagopal et
al., 1989; Herschlag et al., 1991; Yoshida et al., 1999; Yoshida et al., 2000). The
32
differential effect of thio-substitution of these chemically equivalent atoms
provided strong evidence for an interaction with the pro-SP oxygen atom.
However, rescue could be obtained only after more was understood about the
ribozyme and the metal ion rescue experiments themselves, leading to evidence
for two metal ion interactions with this oxygen atom (see below).
Thus, the early experiments provided evidence for three metal ion
interactions with the chemical transition state, and subsequent experiments
provided evidence for two additional metal ion interactions. But these
experiments could not distinguish how many metal ions were involved in these
five interactions. To address this question, a new rescue approach was
developed that we refer to as Thermodynamic Fingerprint Analysis [TFA; (Shan
and Herschlag, 1999; Shan et al., 1999)]. TFA uses functional assays specifically, metal ion rescue of modified substrates- to determine the affinity of
the rescuing metal ion. The affinity then provides a “fingerprint” characteristic of
that metal ion. If rescue at two different sites gives different fingerprints, then
rescue occurs via two distinct metal ions; if the fingerprints are the same, the
same metal ion could be making both interactions or two metal ions with the
same affinity could be involved. To distinguish between these possibilities the
two modifications can be introduced together and the concentration dependence
of metal ion rescue determined; if rescue still depends on one metal ion,
evidence suggests that a single metal ion makes both interactions, whereas a
steeper dependence on metal ion concentration provides strong evidence for
distinct metal ions making each interaction.
33
While simple in principle, in practice there are several critical controls and
criteria for TFA, and implementation is most powerful in the context of a kinetic
and thermodynamic framework for the reaction of interest, such as that shown in
Figure 8 for the Tetrahymena ribozyme (Shan et al., 1999). Briefly, the criteria
are as follows. The starting state of the system is the ribozyme with its active site
unoccupied; this prevents the different thio (or amino)-substituted substrate
analogs from directly interacting with the rescuing metal ion in the reaction’s
ground state and artifactually causing binding affinity to be different for the
different rescuing metal ions. Also, rescue is presented in terms of “krel”, the rate
constant for the thio-substrate relative to that for an unsubstituted control; this
accounts for most effects from soft metal ions on both reactions that are
unrelated to rescue, and using a high ‘background’ of Mg2+ further minimizes
effects from other sites (Shan et al., 1999). Indeed, the initial conclusion of no
Cd2+ rescue of the SP-thiophosphate lacked this control; once the inhibitory effect
of high concentrations of Cd2+ was uncovered and controlled for, selective rescue
of this sulfur substitution was revealed (Shan et al., 2001). Finally, the same
reaction steps, including the chemical step, must be followed for both the
substituted and normal substrates; careful attention to this control uncovered a
metal ion that facilitates docking but does not interact with the atoms directly
involved in the chemical step (Shan and Herschlag, 2000).
A series of TFA studies has led to a catalytic model for Tetrahymena
ribozyme involving three active site metal ions making five interactions with the
reaction’s transition state (Fig. 9c). Despite the conceptual simplicity of this
34
approach, results from thio-rescue experiments have often been controversial.
Introduction of a sulfur atom and thiophilic metal ion can result in formation of an
interaction not present in the normal reaction, and evidence for this has been
obtained for the hammerhead ribozyme (S. Wang and D. Herschlag, unpublished
results; V. J. DeRose, unpublished results). Indeed, whenever a mutation is
made it is possible that the reaction path is changed, but given the specificity of
enzyme structure and catalysis we expect that these situations will be exceptions,
especially given the difficulty of restoring an interaction. Another reason that thiorescue experiments have been controversial is that they have sometimes been
carried out in RNA systems without complete thermodynamic and kinetic
frameworks for the structural change or reaction being monitored, rendering
interpretation difficult (Christian and Yarus, 1993; Sontheimer et al., 1997; Basu
and Strobel, 1999; Yoshida et al., 1999; Shan and Herschlag, 2000) (J.
Frederiksen and J. A. Piccirilli, in prep.). Finally, rescue experiments are often
complex in practice, rendering it difficult for readers not expert in kinetic
approaches to evaluate the results and conclusions.
Catalytic model for the Tetrahymena ribozyme. The active site interactions
suggested from metal ion rescue and other functional studies are combined to
produce overall model for the reaction’s transition state as depicted in Figure 9c
(Knitt et al., 1994; Strobel and Ortoleva-Donnelly, 1999; Yoshida et al., 2000;
Shan et al., 2001). Consider the reaction in the direction of attack by guanosine,
the equivalent of the first step in self-splicing. A proton has already been lost
35
from the 3'
-hydroxyl of G, presumably facilitated by metal ion B (MB); MB will
lower the pKa of this group, allowing a higher concentration of a stronger
nucleophile to be formed at physiological pH (Jencks, 1987; Fersht, 1999). The
leaving group atom, the 3'
-oxygen of the cleavage site U, develops negative
charge in the transition state as its bond to the transferred phosphoryl group is
broken. This developing negative charge on the 3'
-oxygen is stabilized by metal
ion A (MA) and by a hydrogen bond donated from the neighboring 2'
-hydroxyl
group whose identification is detailed below.
The cleavage site 2'
-hydroxyl interaction with the 3'
-oxygen leaving group
was identified by first recognizing that the high reactivity of substrates containing
this 2'
-hydroxyl group relative to substrates containing 2'
-deoxy or 2'
-fluoro
substitutions suggested that this group acted as a hydrogen bond donor
(Herschlag et al., 1993a). Subsequent work showed that the enhancement from
the 2'
-hydroxyl group occurred when the leaving group 3'atom was oxygen but
not sulfur, providing a functional link between these atoms (Yoshida et al., 2000).
This 2'
-hydroxyl appears to be oriented via the 2'
-hydroxyl group of ribozyme
residue A207, an interaction that connects the substrate 2'
-hydroxyl to the
exocyclic amino group of the G residue of the cleavage site G•U wobble pair.
After recognizing the importance of a G•U wobble pair at the cleavage site
(Doudna et al., 1989; Knitt et al., 1994; Strobel and Cech, 1995; Strobel and
Cech, 1996), it was suggested that a water molecule or ribozyme group played
such a bridging role based on the crystal structure of an isolated duplex
containing a G•U pair that showed such a bridging water molecule (Holbrook et
36
al., 1991; Knitt et al., 1994). Subsequent work by Strobel and colleagues
revealed a functional connection between the A207 2'
-hydroxyl group and the
G•U wobble pair, providing evidence for this specific bridging interaction shown in
Figure 9c (Strobel and Ortoleva-Donnelly, 1999).
In addition to the interactions with the G nucleophile 3'
-oxygen and both
the 3'
-oxygen leaving group and 2'
-hydroxyl of the cleavage site uridine, there
are also interactions with the 2'
-hydroxyl group of the G nucleophile and one of
the non-bridging oxygen atoms of the transferred phosphoryl group. Metal ion C
(MC) bridges the 2'
-hydroxyl of G and the pro-SP nonbridging phosphoryl oxygen
atom. These interactions appear to be involved in the coupled binding observed
between G and the 5'
-splice site analog [Fig. 9c; (Shan and Herschlag, 1999;
Shan and Herschlag, 2002)] and presumably play a role in positioning the two
substrates with respect to one another. As noted above, the use of binding
interactions to position reacting groups is a fundamental strategy common to
both protein and RNA enzymes (Narlikar and Herschlag, 1997). MA also bridges
to the pro-SP nonbridging phosphoryl oxygen atom and may also help position
the substrates for reaction. The interactions of MA and MC with one of the
nonbridging phosphoryl oxygen atoms may also help stabilize charge on that
oxygen atom in the transition state and/or eliminate or lessen an energetic barrier
to reaction from solvent reorganization (Catrina and Hengge, 1999; Grzyska et
al., 2002; Lopez et al., 2002; Gregersen et al., 2004).
Although we lack a quantitative accounting of catalysis by this ribozyme,
or any enzyme, the interactions shown in Figure 9c are in principle sufficient to
37
account for a substantial fraction of the observed ~1013-fold rate enhancement
provided by this ribozyme (Karbstein et al., 2002; Hougland et al., 2004). The
current challenge, in addition to further testing this model, is to identify the
ribozyme interactions that bind and position the metal ions and substrates and,
further, to determine the overall structural and energetic properties of the
ribozyme that establish these interacting groups. Progress in these areas is
discussed in the next section in the context of the recent crystal structures of
group I introns.
COMPARING STRUCTURAL AND FUNCTIONAL STUDIES OF GROUP I
INTRONS
Table 2 lists the structures of group I introns, domains and fragments
solved or modeled from x-ray crystallographic or NMR studies. The recent
solutions of group I intron crystal structures at atomic resolution have given
researchers a new perspective of a molecule that has been ‘felt’ before, through
many functional assays, but never ‘seen’ up close at atomic resolution. The
ability to now experience this molecule with all of our molecular ‘senses’ provides
strong support for much prior functional work, suggests resolutions to previous
ambiguities, and, in some cases, leads to new models.
In this section we compare models for structure and function based on
functional and structural studies. We tour structural features and assess
energetic contributions and mechanistic models. These comparisons help identify
38
the strengths and limitations of each approach and further underscore the
synergy between structural and functional studies, highlighting how the
combination of these approaches will provide unprecedented power to answer
important remaining questions concerning RNA structure and function.
Overview of the group I intron structure.
The recently reported crystal structures of three group I introns at atomic
resolution provide for the first time an opportunity to compare crystallographic
models for multiple members of the same ribozyme family (Adams et al., 2004a;
Guo et al., 2004; Golden et al., 2005). Overlaying the three group I intron crystal
structures reveals a common global architecture (Fig. 10a). The conserved core
helices, P1-P10, P4-P6, and P3-P9, overlay quite well when aligned from
superposition of the base triple formed at the guanosine binding site within P7.
The angle formed between the P4-P6 stack and the P3-P9 stack is nearly
identical in all three introns, and the relative positions of all helices are the same.
Thus, the structural homology of these three group I introns extends to regions
distal from the catalytic center.
The major difference between the group I intron structures is the presence
or absence of auxiliary or so-called ‘peripheral’ domains. These domains, which
distinguish particular group I intron subclasses, sit outside the conserved core
(Michel and Westhof, 1990; Lehnert et al., 1996). Apparently different introns
have come upon different solutions for stabilizing the active site at the
intersection of the P4-P6 and P3-P9 domains. Such different strategies were
39
previously suggested from comparative hydroxyl radical footprinting and
phylogenetic comparisons (Michel and Westhof, 1990; Heuer et al., 1991).
Understanding how these different elements provide stabilization and how group
I introns have evolved to use different RNA elements and proteins for stability
represents a fascinating current challenge.
It is also of interest to compare the x-ray structures to the models derived
from phylogenetic and functional data as described in "GROUP I INTRON
STRUCTURAL STUDIES". In addition to the Tetrahymena intron, other group I
introns have been modeled, including the intron from Azoarcus [(Michel and
Westhof, 1990; Lehnert et al., 1996; Rangan and Woodson, 2003; Rangan et al.,
2004) (http://www-ibmc.u-strasbg.fr/upr9002/westhof/)]. We described the
Michel-Westhof model of the Tetrahymena intron model in Figure 5 above. The
overall architecture and placement of helices matches that of the crystal structure.
Analogous comparisons for the Azoarcus intron lead to the same conclusion (not
shown).
As expected, there are several regions where the Michel-Westhof model
failed to accurately predict local structure and tertiary contacts. As noted above,
there is insufficient phylogenetic and biochemical information to specify all of the
atomic interactions, and molecular modeling is not sufficiently advanced to make
up for this deficit. Further, small errors can compound to give differences in the
overall shape of the molecule, and such effects are presumably responsible for
the differences in the radius of gyration, small angle x-ray scattering profile, and
hydroxyl radical footprinting observed for the Tetrahymena intron compared to
40
predictions based on the Michel-Westof model (Lehnert et al., 1996; Russell et
al., 2000) (R. Das et al., unpubl.).
The differences in specific regions occur where the local structure in
crystallographic model does not adopt a standard helical geometry, such as the
J8/7 linker region, the hinge region of the P4-P6 domain, and the P7 domain,
which diverges strongly from A-form helix geometry in forming the guanosine
binding site. Below we make comparisons at this more local level between the xray structures and functional data acquired from atomic level perturbations.
Overall there is remarkable congruence between the structural and functional
results, and each substantially enhances interpretation and understanding of the
other.
The Tetrahymena intron P4-P6 domain.
The P4-P6 domain contains the conserved P4, P5, and P6 helical regions
and the P5abc extension that is found in the IC1 and IC2 subclasses of group I
introns [Fig. 2d; (Michel and Westhof, 1990)]. Based on phylogenetic
comparisions as described above, Michel and Westhof postulated an intradomain
tertiary interaction between the minor groove of the P6 helix and the GAAA
tetraloop in the P5b extension, a motif referred to as a tetraloop/tetraloop
receptor interaction (Costa and Michel, 1995). Contemporaneously, Murphy and
Cech demonstrated that the P4-P6 domain forms an independently folding unit.
Hydroxyl radical probing of accessible and protected regions in combination with
mutational analysis led to a model in which folding required a long-range
41
interaction between the GAAA tetraloop and the P6a helix. They also showed
that mutation of the conserved residue A186 in the bulge in P5a (Fig. 2d)
disrupted the structure of the P5abc subdomain in isolation and destabilized
folding of the P4-P6 domain (Murphy and Cech, 1993; Murphy and Cech, 1994).
Direct visualization of the P4-P6 domain by electron microscopy revealed
Mg2+-induced compaction, whereas domains mutated so as to extend the P5
helix, through base pairing of the J5/5a region, exhibited a rod-like shape in the
presence of Mg2+ (Wang et al., 1994). In summary, the combination of
phylogenetic, biochemical, mutagenesis, and electron microscopy data predicted
that the P4-P6 domain would contain a sharp bend in the J5/5a region, thereby
allowing the P5abc domain to lie alongside the coaxially stacked P5-P4-P6-P6aP6b helices and make tertiary contacts from its P5b tetraloop and P5a bulge to
the P5-P4-P6-P6a-P6b coaxial stack (Murphy and Cech, 1993; Murphy and Cech,
1994; Wang et al., 1994).
The crystal structure of the P4-P6 domain by Doudna, Cech and
coworkers in 1996 provided an opportunity to compare and contrast models
derived from phylogenetic and prior experimental work with a model derived from
x-ray analysis (Cate et al., 1996b; Cate et al., 1996a). The crystallographic model
generally agreed with overall features of the Michel-Westhof model: the base
pairing was accurately predicted, and the overall structure contained the
predicted sharp bend at J5a/5b and the two predicted regions of tertiary
interactions (Fig. 11). Nevertheless, the atomic details of the bend and of each of
the tertiary interactions were not correctly predicted. This again highlights the
42
importance of recognizing the nature of the information content of an experiment
or analysis. Phylogenetic comparisons can reveal base-base covariation and
sometimes suggest the presence of specific base-base interactions, but
conservation and covariation of groups of residues does not provide information
about the nature of the interactions made by those bases – thus the tetralooptetraloop receptor was not correct at atomic resolution. Additionally, the P4-P6
model provides no information about metal ion binding and interactions involving
the sugar-phosphate backbone – thus the metal ion ‘core’ within the P5abc
subdomain and its interactions mediated by 2'
-hydroxyl groups with P4 were not
identified (Cate et al., 1996b; Cate et al., 1996a).
Interestingly, a combination of site-directed mutagenesis and chemical
footprinting led Cech and coworkers to suggest a contact between G212 in the
P4 helix and the P5a bulge (Flor et al., 1989); residue A183 became exposed to
chemical modification upon mutagenesis of G212. However, the crystal structure
revealed a hydrogen bond between G212 and A184 (Cate et al., 1996b; Cate et
al., 1996a). Thus, the region of interaction was identified, but the loss of
protection at A183 occurred because of disruption of a neighboring interaction,
rather than disruption of an interaction with A183 itself.
Docking of the P1 helix into the intron’s catalytic core.
As described above, the oligonucleotide substrate referred to as “S” in the
ribozyme reaction (Fig. 8) is a 5'
-splice site mimic (see “A thermodynamic and
kinetic framework for the Tetrahymena group I ribozyme reaction” above). This
43
oligonucleotide base pairs to the ribozyme’s IGS to form the P1 helix (Fig. 2d, 7
& 8). To place the substrate’s cleavage site within the active site, the ribozyme
has constructed a binding, or “docking”, site for the P1 helix. The first evidence
for the presence of this site was the strong binding of 5'
-splice site mimics,
relative to the binding expected for simple duplex formation (Herschlag and Cech,
1990a; Herschlag and Cech, 1990b; Herschlag and Cech, 1990c; Pyle et al.,
1990; Bevilacqua and Turner, 1991; Pyle and Cech, 1991; Bevilacqua et al.,
1992; Herschlag, 1992; Pyle et al., 1992; Bevilacqua et al., 1994; Pyle et al.,
1994; Narlikar et al., 1997; Narlikar et al., 1999; Zhuang et al., 2000; Bartley et al.,
2003). Experiments with oligonucleotides containing functional group
modifications and later with ribozymes containing analogous modifications
incorporated into the IGS revealed the importance of specific 2'
-hydroxyl groups
of both strands and of the exocyclic amino group of the conserved G•U wobble
pair that specifies the cleavage site (Herschlag et al., 1993b; Knitt et al., 1994;
Strobel and Cech, 1994; Strobel and Cech, 1995; Strobel and Cech, 1996;
Narlikar et al., 1997; Strobel and Ortoleva-Donnelly, 1999).
Further functional work identified intron functional groups that would
“suppress” the effect of a particular P1 functional group modifications. Such
results, akin to genetic suppression experiments, suggest that the two groups
interact. In principle these interactions can be direct or indirect, but the results
described below suggest that, at least in this system, direct interactions are
responsible in nearly all cases. These experiments are particularly powerful as
44
screens can be carried out on a population of molecules, with subsequent
resolution of the individual variants by gel electrophoresis.
The population-based approach was first used by Christian and Yarus to
identify phosphate oxygen atoms involved in functionally important interactions
(Christian and Yarus, 1992). Phosphorothioates were randomly incorporated into
the intron, via in vitro transcription with a transcription mixture of NTPs doped
with a small amount of an α-thio-NTP, and splicing was allowed to proceed. The
spliced and unspliced RNA could then be separated by denaturing gel
electrophoresis. If a replaced phosphoryl oxygen atom were particularly
important for splicing, then the unspliced population would be overrepresented
with RNAs containing the thio-substitution at that position, relative to RNAs in the
spliced population. The abundance of RNAs containing thio-susbtitutions at each
position can be readily determined in two steps: iodine treatment of radiolabelled
RNA to induce specific cleavage of thio-phosphates followed by a second gel
electrophoresis step. This approach is highly sensitive as it allows side-by-side
comparison of the starting, spliced, and unspliced RNA.
Population-based approaches were then expanded to allow assessment
of the importance of 2'
-hydroxyl groups and, later, of base functional groups
(Gaur and Krupp, 1993; Conrad et al., 1995; Strobel and Shetty, 1997). These
approaches used thio-substitution to ‘mark’ a position that also contained a
second ‘test’ substitution, such as a 2'
-deoxyribose or an N7-deazaadenosine.
Screens with substitutions for each functional group can be carried out to identify
‘important’ positions (i.e., positions that give significant effects under certain
45
conditions), and suppression analysis can be carried out by determining what
positions no longer give deleterious effects when a particular residue or
functional group is changed uniformly in the population. Overall, these
approaches require synthesis of the modified residue, random incorporation via
transcription, and separation of active from less active molecules. Strobel and
coworkers have greatly expanded and expertly utilized the power of this
approach, synthesizing a battery of base analogs and applying them to
numerous ribozyme systems, including group I introns (Ryder and Strobel, 1999;
Strobel, 1999). They refer to the screening and suppression approaches as
NAIM (Nucleotide Analog Interference Mapping) and NAIS (Nucleotide Analog
Interference Suppression), respectively.
We highlight below interactions suggested from functional studies that can
be evaluated via comparison to the recent x-ray structures. Although most of the
functional data on P1 docking has been obtained with the Tetrahymena ribozyme
(Table 3), the partial structure of this RNA lacks the P1 helix (Table 2). However,
there is a high degree of conservation with the Azoarcus intron’s binding site, and
the Azoarcus structure contains a docked P1 duplex, so we use this structure in
comparisons with functional data from the Tetrahymena intron (Table 3). The
more limited functional data with the Azoarcus intron are consistent with that for
the Tetrahymena intron (Strauss-Soukup and Strobel, 2000). The intron regions
that interact with the P1 duplex are in the P4 helix and in J8/7. Specific features
these interactions and the underlying evidence are presented below.
46
Figure 12a shows the conserved splice junction G•U pair in the Azoarcus
intron x-ray structure. The base pair is indeed a wobble pair, as predicted by
functional studies (Doudna et al., 1989; Knitt et al., 1994). Further, functional
data from Strobel and Cech, using inosine instead of guanosine in this base pair,
implicated the exocyclic amino group of G in docking interactions (Fig. 12a & 12
b; G22 in Tetrahymena and G10 in Azoarcus (Strobel and Cech, 1995);
subsequent suppression data identified an interaction with N3 of residue A207
(Strobel et al., 1998), the precise interaction observed in the x-ray structure (Fig.
12a & 12c). Suppression data also implicated the 2'
-hydroxyl group of A207 in
interactions with functional groups on both partners of the G•U wobble pair at the
5'
-splice site – the exocyclic amino group of G22 and the 2'
-hydroxyl group of the
U residue (Strobel and Ortoleva-Donnelly, 1999). These interactions, which
appear to play a role in catalysis (see Fig. 9 and “Probing the chemical step of
the Tetrahymena group I ribozyme reaction” above), were also supported by the
x-ray data (Fig. 12a & 12c). In addition, suppression data linked the 2'
-hydroxyl
group of G22, which contributes to docking, to the 2'
-hydroxyl group and N3 of
residue A114 (Strobel et al., 1998; Soukup et al., 2002); the structure shows
these interactions.
We now turn to residues in the P1 helix upstream from the cleavage site
(Fig. 13a). These residues corresponding to the 5'
-exon sequence are typically
denoted with negative numbers indicating their position relative to the splice site;
i.e., the U of the splice site G•U wobble pair discussed above is U(-1), and
immediately 5'of it in the Tetrahymena 5'
-exon are C(-2) and U(-3) (Fig. 2d).
47
Functional data have implicated tertiary interactions with the 2'
-hydroxyl groups
of both of these residues [Table 3; (Pyle et al., 1992; Narlikar et al., 1997;
Szewczak et al., 1998)]. Functional suppression data from Pyle, Murphy and
Cech, combined with probing of base accessibility by dimethylsulfate (DMS),
implicated an interaction between the U(-3) 2'
-hydroxyl group and residue A302
(Pyle et al., 1992; Narlikar et al., 1997). Based on the ability of U to substitute
partially for A302, and the phylogenetic conservation of A and U at this position,
a hydrogen bond from the 2'
-hydroxyl to N1 of A302 or O4 of U302 was
suggested. The Azoarcus crystal structure indeed shows a hydrogen bond from
N1 of the equivalent A residue. Remarkably, even though the Twort intron has a
U residue instead of A at this position, it still has the corresponding hydrogen
bond to O4 of this residue (Fig. 13b).
In contrast to the excellent agreement between the functional and
structural results for residue U(-3), residue C(-2) does not make the interaction in
the crystal suggested from the functional studies. Suppression data implicated an
interaction between the 2'
-hydroxyl of C(-2) and the exocyclic amino group of
G303 in the Tetrahymena intron (Szewczak et al., 1998). However, this amino
group (at position G169 in the Azoarcus sequence) faces away from C(-2) in the
structure (Fig. 13a). It appears that the exocyclic amino group donates a
hydrogen bond to one of the neighboring phosphoryl oxygen atoms, thereby
positioning the 4'
-ring oxygen atom of the G303/G169 ribose to accept a
hydrogen bond (Fig. 13a). Thus, removal of the exocyclic amino group from
G303 presumably indirectly affects this hydrogen bond (Adams et al., 2004b).
48
At the corresponding position in the Twort x-ray structure, the same 2'
-hydroxyl /
O4'oxygen hydrogen bond appears to be made, but the G residue (G184)
corresponding to G303/G169 in Tetrahymena and Azoarcus is positioned very
differently. The sugar pucker and glycosidic bond are in the 3'
-endo and anti
conformations, respectively, rather than 2'
-endo and syn as in the Azoarcus [Fig.
13a & 13b; (Adams et al., 2004a; Golden et al., 2005)].
These observations underscore a major challenge in RNA research.
Because RNA residues have so many opportunities for hydrogen bonding and
stacking interactions – each base can form at least two hydrogen bonds with
each of the four bases and can hydrogen bond to 2'
-hydroxyl and phosphoryl
groups of any residue– there are many possible interactions within a given
sequence (Sigler, 1975; Herschlag, 1995; Silverman et al., 1999). Thus,
alternative sequences that give different structural interactions may nevertheless
position certain functional groups to allow identical interactions with ligands or
other structural elements. Conversely, individual mutations or functional group
substitutions can result in significant conformational rearrangements.
Finally, we turn to interactions with the IGS residue G25, which base pairs
to C(-4) of the Tetrahymena 5'
-exon. Removal of the 2'
-hydroxyl group of G25
gives a large energetic effect [Table 3; (Strobel and Cech, 1993; Narlikar et al.,
1997)]. The corresponding Azoarcus residue, C13, is part of P2 instead of P1 as
in the Tetrahymena intron. However, functional data suggested that P2 and P1
stack in the intron subclass to which the Azoarcus (but not the Tetrahymena)
intron belongs and may therefore behave like a single helix. Numerous group I
49
introns have a conserved length of 13 for the P1 and P2 helices combined and a
GNRA tetraloop at the end of P2 (where N is any base and R is a purine).
Mutagenic data strongly suggest that this length specifies a favored site for
cleavage due to positioning of the P2/P1 stack via an interaction with the
tetraloop (Michel and Westhof, 1990; Peyman, 1994). The Azoarcus x-ray
structure has verified this stacked arrangement and shows the predicted GNRA
tetraloop interaction with J8/8a. Therefore, the Azoarcus P2 helix may be
considered part of P1, and the Azoarcus residue C13 may be structurally
homologous to G25 of the Tetrahymena intron. Indeed, functional suppression
data for the Tetrahymena intron and the x-ray data for the Azoarcus intron
indicate that a homologous interaction is made, involving the 2’-hydroxyl and N3
of A301 or A167 in Tetrahymena and Azoarcus, respectively [(Szewczak et al.,
1998; Adams et al., 2004b); interactions not shown]. These results strongly imply
a plasticity in global RNA structure. Uncovering how and why aspects of
interactions have remained or changed in different group I introns may provide a
window into the evolution of new motifs for RNA structure and RNA•protein
interactions.
We now consider the overall geometry of the docking site for P1, located
in the J4/5 and in J8/7 (Fig. 2d & 5a). As noted above, the Azoarcus intron x-ray
structure contains a docked P1 duplex whereas the partial Tetrahymena
structure does not. The structures are nearly identical in the region of J4/5 that
interacts with the splice site G•U wobble pair, and the Twort structure, with its
docked P1 helix, is also nearly identical in this region (not shown). In contrast
50
J8/7 differs considerably between the Azoarcus and Tetrahymena introns (Fig.
14a). Changes in the primary sequence at two positions in J8/7 do not appear to
account for this structural difference (see comparison to the Twort ribozyme
below). We speculate that the slow docking of the P1 helix into its tertiary
interactions arises, at least in part, because the J8/7 residues must realign in
order to make their tertiary interactions with P1 (Bartley et al., 2003). Alternatively,
the structural differences of the Tetrahymena J8/7 sequence could arise because
peripheral regions missing in the crystallized construct are necessary to orient
the P1 binding site (Engelhardt et al., 2000). In either case, these structural
differences underscore the dynamic nature of RNA, the need to better
understand the dynamic properties of RNA, and the need to obtain structural
information in multiple states and under multiple conditions.
We also compare J8/7 for the Azoarcus and Twort introns, both having
docked P1 helices (Fig. 14b). The overall geometry of J8/7 of the Azoarcus intron
is highly consistent with that of Twort intron although there are more sequence
differences between the Azoarcus and Twort introns than between the Azoarcus
and Tetrahymena introns. As described above, a G residue conserved between
these introns has a different sugar pucker and glycosidic bond orientation, yet the
ribose of this residue makes the same interaction (Fig. 13, G169 and G184 in the
Azoarcus and Twort introns, respectively).
As we learn more about RNA structure, folding and energetics, it will be
interesting to ascertain how different sequences, secondary and tertiary
structures can be used to create homologous and nonhomologous solutions for
51
molecular recognition and catalytic challenges. The large number and diversity of
group I introns [see "Gutell Lab Comparative RNA Site";
(http://www.rna.icmb.utexas.edu/)] provide an excellent field for such exploration.
For example, specific tertiary interactions in P1 docking may be conserved
among some, but not all group I introns [Table 3; (Strobel and Cech, 1993;
Narlikar et al., 1997; Testa et al., 1997; Disney et al., 2000; Disney et al., 2001)].
The guanosine binding site.
Several distinct guanosine binding models have been proposed based on
functional studies. As described in “GROUP I INTRON STRUCTURAL
STUDIES”, the reactivity of 2-aminopurine with a mutant ribozyme, in which the
G264-C311 base pair was replaced with an A-U base pair, led Michel et al. to
propose guanosine binding through a base triple interaction (Fig. 4), thereby
localizing G binding to the P7 helix (Michel et al., 1989). Although rescue of the
A264-U311 mutant reaction by 2-aminopurine provides strong evidence for a
hydrogen bond between O6 of G264 and N1-H of bound guanosine, it provides
no information about the other proposed hydrogen bond between the exocyclic
amino group of the bound guanosine and N7 of G264 (Fig. 4) (Yarus et al., 1991).
In the sunY intron, DMS methylation at N7 of the equivalent residue to G264
interfered with G-mediated splicing, and bound G protected the N7 from this
methylation (von Ahsen and Noller, 1993). These observations are consistent
with the Michel base triple interaction, but methylation interference experiments
do not probe hydrogen bonds directly.
52
The absence of direct functional support for the interaction led Yarus et al.
to consider alternative G binding modes, in which G interacts with one or both
adenosine residues (A263 and A265) that flank the G264-C311 base pair,
resulting in a more axial orientation (Yarus et al., 1991). These models were
evaluated by measuring the reactivity of guanosine analogs with mutant
ribozymes and by conducting molecular dynamics simulations and energy
minimization. The mutational analysis revealed an apparent keto-amino
complementarity between the C6 substituent of G and the C6 substituent of A265
below the G264-C311 base-pair plane; although the effects were modest, this
result was consistent with one of the proposed axial binding models. In molecular
dynamics simulations conducted using a seven nucleotide fragment of the P7
helix, the base-triple and axial binding modes had distinct outcomes. The base
triple quickly rearranged to a family of conformations that do not have the
interactions with G264. The axial model also markedly rearranges to a more
stable structure, termed "axial III", but retained the critical hydrogen bond with
G264. In this energy-minimized axial III model, A263 is extruded from the P7
helix, possibly forming a base triple interaction with the minor groove face of the
G264-C311 base pair. Phylogeny supported this possibility: A263 is always
unpaired and covaries as A or C with the C262-G312 base pair (Michel et al.,
1989; Gautheret et al., 1995). However, mutational analysis showed that A263
can be mutated without functional consequence (Yarus et al., 1991).
Using a collection of adenosine analogues, Ortoleva-Donnelly et al.
generated NAIM profiles for A263 and A265 (along with other A residues) in the
53
exon ligation reaction (Ortoleva-Donnelly et al., 1998). Consistent with the lack
of mutational sensitivity reported by Yarus et al., the ribozyme appeared to be
widely tolerant to functional group modifications at A263, although the strong
phosphorothioate interference at this site could have obscured additional
interference from the “test” substitution. At the A265 residue, substitution with
analogues that lack the exocylic amine, which would be expected to weaken G
binding in the axial III mode, did not interfere with exon-ligation activity (OrtolevaDonnelly et al., 1998). This lack of interference was considered uninformative
due to the expectation that the high effective molarity of ωG in the exon-ligation
reaction would render the reaction less sensitive to weakened G binding [see
(Mei and Herschlag, 1996)]. Substitution of N6-methyl adenosine at A265
showed strong interference, consistent with the proposed interaction to O6 of the
guanosine substrate. This was viewed as supporting the axial model; however,
as in the case of DMS protection and interference, these experiments do not
provide direct evidence for hydrogen bonding to the guanosine substrate. The
observed interference could arise from disruption of a hydrogen bond to a
different partner than G or from the steric bulk of the methyl group rather than
from disruption of a hydrogen bond.
Kitamura et al. used NMR to study the guanosine binding site, determining
the structure of a 22-nucleotide RNA mimic of the P7/P9.0 region (Kitamura et al.,
2002). In this RNA, the ωG forms a base triple with the G264-C311 base pair with
no observed contacts to A265. A263 bulges out of the P7 helix, leading to a large
helical twist. There was no evidence for base triple interactions with A263 as
54
defined by the axial III model of Yarus. The NMR analysis established the
stereochemical plausibility of G binding in the base triple mode to the P7 major
groove and the propensity for A263 to bulge out of the helix, but could not
establish the nature of G binding to the full intron as the intron context could alter
the conformation of P7 and / or the orientation of bound G. Indeed, changes in
RNA conformation upon extracting fragments from larger structures are common
(Butcher et al., 1997; Butcher et al., 1999; Rupert and Ferre-D'
Amare, 2001;
Kitamura et al., 2002; Zhang and Doudna, 2002; Guo et al., 2004; Sigel et al.,
2004).
The recent crystal structures of the Azoarcus, Tetrahymena, and Twort
group I introns leave little doubt as to the architecture of the guanosine binding
site (Fig. 15a). All three structures depict ωG forming a base-triple with the G264C311 base pair or the homologous base pair in the Azoarcus and Twort
structures as predicted by Michel et al. and later supported by the NMR structure
(Fig. 15a) (Adams et al., 2004a; Guo et al., 2004; Golden et al., 2005). ωG also
stacks between two additional base triple layers (Fig. 15b & 15c). Below the ωG
triple, the A261 base contacts the major groove of the A265-U310 base pair.
Above the ωG triple, A263 is bulged out of the helix to form two hydrogen bonds
to the minor groove site of the G312-C262 base pair (Fig 15b & 15c), and this
base triple results in significant distortion of the P7 helix from A-form geometry.
Thus, the crystallographic models suggest a guanosine binding site composed of
consecutive layers of stacked base triple interactions, providing both hydrogen
bonding and stacking contacts to bind and position the guanosine for splicing.
55
The architecture of the guanosine binding site provides insight into the
possible origins of slow guanosine binding. As described in the previous section,
guanosine binds in at least two steps, with a conformational change following an
initial encounter complex (Karbstein and Herschlag, 2003). It is possible that this
conformational change is related to the distorted P7 helix that forms the specific
pocket for the guanosine nucleophile. Indeed, it is hard to imagine the guanosine
binding site structure being maintained without guanosine present in the site. As
guanosine sits with base triples above and below it, this stack would be prone to
collapse or rearrangement without the central guanosine (Fig. 15b). Perhaps
without bound guanosine, A263 is bulged into solution as in the NMR structure. It
follows from these considerations that, just as J8/7 might realign to make tertiary
interactions with P1, the G-binding site might require conformational changes to
engage guanosine in hydrogen bonding and stacking interactions.
Some twenty years after the finding by Bass and Cech that the exocyclic
amine of the G substrate contributes to binding, its interaction partner in the
ribozyme had not been functionally identified (Bass and Cech, 1984). In contrast,
the majority of functional studies on the group I introns led to models that were
confirmed (and expanded) by the x-ray structures. Why was the model for G
binding derived from functional data not accurate? In hindsight the answer is
clear and multifaceted, but first it should be recognized that the phylogeny and
specificity switch experiments did correctly localize the guanosine binding site to
P7 and identify the central hydrogen bond of the guanosine•G264-C311 base
triple interaction. The axial model was put forward based on functional data with
56
only modest effects and was bolstered by results from molecular dynamics
simulations. Simply put, molecular dynamics simulations of nucleic acids have
been limited by short time scales (leading to a significant dependence of the
starting conformation chosen), the lack of accurate solvent and solvation models,
and incomplete modeling of the electrostatic environment created by the
polyanionic backbone and its surrounding ion atmosphere [for examples and
reviews see (Auffinger and Westhof, 1998; Auffinger and Westhof, 2000;
Beveridge and McConnell, 2000; Cheatham and Kollman, 2000; Giudice and
Lavery, 2002; Norberg and Nilsson, 2002; Cheatham, 2004; Sorin et al., 2005)].
Unfortunately, conclusions from such computational studies are often accepted
without understanding the underlying theoretical or empirical models used and
without testing decisive predictions that follow from these conclusions. For
example, the interaction between the N7 of G264 and the exocyclic amine of G
proposed in the base triple interaction could have been revealed using 7deazaguanosine, which would be predicted to give interference with guanosine
as the substrate but not with inosine. Nevertheless, it would have been difficult to
arrive at a structural model with the P7 helix correctly kinked based on functional
data alone, and the NMR fragment data suggests that this kink is not intrinsic to
the P7 helix but is enforced by the higher order structure of the RNA.
Metal ions in the group I intron catalytic site.
Functional studies described above utilizing metal ion rescue experiments
and thermodynamic fingerprint analysis (TFA) have led to a model in which three
57
metal ions coordinate to four oxygen atoms of the ribozyme substrates, making
five interactions in the chemical transition state [Fig. 9c; (Shan et al., 1999; Shan
et al., 2001)]. An alternative model of metal ion involvement in the group I
ribozyme transition state was suggested based on a recent crystal structure of
the Azoarcus group I intron [Fig. 9d; (Adams et al., 2004a)]. In this model, there
are two metal ions coordinating to the ribozyme substrates. One metal ion is
coordinated to both the 3'
-oxygen leaving group of the cleavage site uridine and
the pro-SP oxygen of the scissile phosphate, the same contacts made by MA in
the functionally based transition state model (Fig. 9c). The other metal ion
contacts the 2'
-hydroxyl group of the guanosine nucleophile, as does MC in the
functional model, but this metal ion is proposed to form an outer-sphere
coordination to the scissile phosphate through an intervening water molecule
rather than a direct inner-sphere contact. MB, the metal implicated by functional
studies to be interacting with the guanosine 3'
-oxygen atom, is not present in this
model nor is there an interaction with the 3'
-oxygen atom of the attacking (or, in
the reverse reaction, leaving) guanosine.
Neither functional nor structural results are infallible. Structures are nearly
always of inactive complexes, with substrates or reactive portions of substrates
missing or modified, and crystallization conditions typically differ from optimal
conditions for activity. Even when substrates are present, the structure can
rearrange, or the resting structure can be different than the active structure. For
example, there may be a conformational change following deprotonation of the
attacking 3'
-oxygen of guanosine, as a result of the change in hydrogen bonding
58
properties and local electrostatic environment, so that the resting structure with
this group protonated would be different from the active structure [see also
(Wang et al., 1999; Uhlenbeck and Blount, 2005)]. Indeed, the three x-ray
structures from Azoarcus, Twort, and Tetrahymena show differences in the
number and specific contacts of the metal ions bound at the active site. This
heterogeneity in metal ion number and location highlights the inherent difficulty in
assigning catalytic metal ion binding sites using structural approaches, a
common issue with metalloenzymes [e.g. (Pingoud and Jeltsch, 2001; Galburt
and Stoddard, 2002)]. While it is possible that the group I variability represents
species-specific differences, it is far more likely that these differences result from
the different crystallization conditions and from the presence of different ligands.
Functional studies, while the only means to directly interrogate the
transient transition state, also have limitations. Above we described criteria,
including an in-depth knowledge of individual reaction steps (Fig. 8), necessary
for firm interpretation of results from TFA and other functional studies (Shan et al.,
1999; Wang et al., 1999; Shan et al., 2001). But even with this rigor, functional
experiments involve comparisons – comparison of a wild type to a mutant or of
one substrate to a modified substrate. Thus, we can only infer properties of the
wild type or normal reaction from differences in behavior that occur when
changes are made.
Consider MB and its interaction as implied by functional studies (Fig. 9c &
9d). The occurrence of a metal ion interaction with the 3'
-oxygen of guanosine
was first suggested from experiments in which this oxygen was converted to
59
sulfur and catalysis was diminished in Mg2+ but partially rescued upon addition of
thiophillic metal ions such as Mn2+ and Cd2+ (Weinstein et al., 1997). Further,
TFA experiments indicated that the affinity of this rescuing metal ion was different
from that of MA or MC, providing strong evidence for a distinct metal ion (Shan et
al., 1999). While these results strongly suggest that the 3'
-sulfur interacts directly
with a Mn2+ ion that is distinct from MA and MC, these results do not show that MB
is present when there is no thio-substitution; the thiophillic metal ion could be
‘recruited’ by the presence of the sulfur, but not normally present in the reaction.
It is also possible that the inner sphere interaction between MC and the pro-SP
phosphoryl oxygen atom, suggested from rescue experiments with sulfur and the
thiophillic Cd2+ ion (Fig. 9c), is an outer sphere interaction when oxygen and Mg2+
are present as suggested by the Azoarcus x-ray structure (Fig. 9d). The larger
size of sulfur than oxygen and the different size and potential coordination states
of Mg2+ and Cd2+ can contribute to such differences.
Figure 9e presents a hybrid model that combines aspects of the models
derived from the functional and structural data. Tests and refinements of these
models will be most effectively carried out through a combination of structural
and functional studies. One means to relate the structural and functional data has
already been initiated. This approach involves functionally identifying intron
groups that are metal ion ligands [(J. Hougland et al., submitted) (J. Lee et al., in
prep.) (Szewczak et al., 2002); see also (Wang et al., 1999)]. In each intron
structure, the putative active site is formed by the convergence of the J4/5, P7,
and J8/7 regions of the respective introns. The introns’ phosphate backbones
60
pack tightly at this interface, with many of these phosphates also identified
through phosphorothioate interference in population-based cleavage activity
screens, suggesting possible metal ion binding sites (Christian and Yarus, 1993;
Strauss-Soukup and Strobel, 2000). Indeed, the deleterious effects of several of
these phosphorothioate substitutions could be ameliorated by the addition of
Mn2+ (Christian and Yarus, 1993). Thus, these negatively charged phosphoryl
oxygen atoms are good candidates for metal ion ligands.
We have extended TFA to assess which rescuing metal ion is perturbed
by individual thiophosphate substitutions within the ribozyme. TFA was carried
out in the context of variant ribozymes containing atomic mutations within the
ribozymes themselves, e.g. site-specific phosphorothioate incorporations within
the ribozyme’s conserved core. As the apparent affinity for each catalytic metal
ion that serves as its “fingerprint” will depend on the specific coordination
environment within that metal ion’s binding site, changing one of the binding
site’s ligands from oxygen to sulfur should result in a shift in apparent binding
affinity for the rescuing soft metal ion, and the effect should be specific for one of
the rescuing metal ions. Such a shift in metal ion rescue profile reflects a linkage
between the modified functional group in the ribozyme substrate and the
ribozyme core through a specific catalytic metal ion, thereby localizing the
binding site for the specific catalytic metal ion that rescues a given substrate
atomic perturbation.
These modified TFA experiments strongly suggest the identity of the three
metal ion ligands shown in the transition state model of Figure 16a. The pro-SP
61
phosphoryl oxygen of residue C262 appears to coordinate to MC (J. Hougland et
al., submitted), and the pro-SP oxygens of residues C208 and A306 both appear
to contact MA [(Szewczak et al., 2002) (J. Lee et al., in prep.) (A. Kravchuk and D.
Herschlag, unpubl.)]. These phosphoryl oxygens are reasonably positioned to
serve as catalytic metal ion ligands within the available group I intron crystal
structures, suggesting that active site configuration within the structural models
bears relevance to the active site configuration in the transition state of ribozymecatalyzed cleavage (Fig. 16b).
Although substantial progress has been made, distinguishing between
active site models such as those in Figure 9c, 9d, and 9e remains a major
challenge. Additional structures will provide insights into the plasticity of the
active site and will also suggest and refine catalytic models. Ligand candidates
for metal ion B (Fig. 9c) and other putative metal ions will need to be probed in
functional assays, as an integral part of testing and refining the catalytic models.
Functionally identified catalytic metal ion ligands provide “anchor points” between
the model of the chemical transition state based on functional studies and the
group I intron’s global structure derived from crystallographic models. Such
anchor points allow integration of biochemical and structural data within a single
model, leading to a deeper understanding of how this ribozyme (as well as other
enzymes, both RNA and protein) use metal ion coordination and other
interactions to achieve their enormous catalytic power and exquisite specificity.
Summary.
62
The most striking generalizations derived from comparisons between the
functional and structural data are the remarkable agreement and the ability of
these data to reinforce one another. For example, putative hydrogen bonds
observed in structures need not be energetically significant, and occasional long
lengths of these hydrogen bonds can render it difficult to decide whether an
interaction actually occurs. Conversely, functional relationships can be direct or
indirect, and atomic resolution structures allow these interactions to be placed
within the context of the overall structure, aiding understanding of how these
interactions are established.
The biggest surprise from the comparisons of global and local group I
structure is the plasticity of RNA structures that carry out a particular function.
This plasticity may have been and may continue to be highly significant for
evolution of new structures and interactions, but will at the same time render
understanding of the interplay between structure, energetics, and function within
RNA a particular challenge.
PERSPECTIVES AND FUTURE CHALLENGES
RNA structure determination and RNA structures. Powerful approaches exist for
obtaining information about RNA structures in solution. Studies with group I
introns and other RNAs have taken advantage of the power of RNA phylogeny,
both natural and artificial, for determining secondary structures and even tertiary
interactions and motifs; probing base accessibility via chemical modification has
63
supplemented and extended conclusions about secondary structure and
sometimes about tertiary structure. Probing backbone accessibility via hydroxyl
radical footprinting has given information about packing in the overall tertiary
structure. Functional data, especially from rescue and suppression experiments,
has provided evidence for specific interactions at the atomic level. In contrast,
there is no indication that current molecular dynamics or other physics-based
computational methods can give reliable structure predictions beyond the
removal of steric clashes from models obtained by other means.
Thus, there exists a complement of solution-based probes for RNA
structure, and the ability of these approaches to make accurate structural
predictions has been strongly validated through group I intron and other RNA
studies. There are many more RNAs and RNA•protein complexes for which
structures are desired, and many of these may have transient conformations that
would be difficult or impossible to obtain in a crystalline state and would therefore
greatly benefit from solution structural determination. However, despite the power
of the current approaches, modeling to give reliable RNA structures is not yet
straightforward. At least part of the problem results from different groups using
different techniques to derive models. An important current challenge is to
integrate, and possibly extend, the current approaches so a group studying a
particular RNA or RNA•protein complex can obtain a structural model in months
rather than years or decades. Such models, even if imperfect, help generate
hypotheses and guide further experiments. The value of such empirical models
notwithstanding, ultimate understanding and predictive power will require
64
physics-based models that use accurate representations of the underlying forces
to derive structural and ‘behaviorial’ models for any RNA of interest, a goal that is
significantly farther off.
The active site and establishing functional RNA structures. Many functional
studies have provided strong evidence for binding and catalytic interactions
within the group I active site. Reassuringly, the resultant models have largely
been supported by structural work, and in the case of the guanosine binding site
competing models were resolved by x-ray crystallographic data. The primary
remaining questions about catalytic interactions focus on the differences from the
functionally and structurally derived models for catalysis: are there three or two
metal ions, and does one of these metal ions contact the 3'
-oxygen atom of the
attacking guanosine? Structure cannot capture the transition state, and the three
crystallized introns give different bound metal ions in different positions.
Functional data rely on atomic level substitutions and can be subject to
rearrangements or recruitment of metal ion interactions. Most powerful will be a
series of structural and functional studies that test predictions from the current
models and, in the process, deepen our understanding of RNA catalysis.
Understanding of RNA catalysis must extend beyond a description of
active site interactions. Energetic features of the interactions also must be
revealed and the structural context of the active site interactions must be
understood. What are the components of the molecule’s overall structure that
position the groups directly contacting the substrates? Why is activity diminished
65
when peripheral RNA domains are deleted, and how do fragments of group I
intron maintain activity (Ikawa et al., 1997; Ikawa et al., 1999; Engelhardt et al.,
2000; Ikawa et al., 2000a; Ikawa et al., 2003)? What are the thermodynamic
features of folding that allow adoption of the active structure, relative to the
ensemble of all possible alternative structures – in other words, how is
cooperativity maintained? To what extent do structures rearrange locally and
globally in response to mutation, changing conditions, and ligand binding?
Answering these questions will provide information necessary for a deep
understanding of the forces and factors responsible for RNA folding and will
provide an important step toward modeling of RNA structure and dynamics.
Conformational changes in RNA biology. Essentially all cellular processes
mediated by RNA involve conformational changes. Perhaps the ability of RNA to
adopt multiple metastable alternative folds [see (Sigler, 1975; Herschlag, 1995)
and references therein] provides an evolutionary incentive for natural selection to
choose RNA for certain jobs that require a controlled progression of
conformational states. Illustrating this point, the ribosome, the spliceosome, the
signal recognition particle (SRP), and telomerase all act through a series of steps
and conformations.
The group I self-splicing intron undergoes several conformational changes.
Conformational changes associated with binding of substrates, P1 docking and
guanosine binding, have been identified. Much is known about the P1 docking
conformational change, and we have suggested that disorder of the J8/7 region
66
may contribute to the slow observed docking [(Bartley et al., 2003) and
discussion herein]. However, the conformational changes involved in shuttling
groups in and out of the active site between the first and second chemical steps
in splicing (Fig. 6) have been less well characterized (Chin and Pyle, 1995;
Chanfreau and Jacquier, 1996; Emerick et al., 1996; Golden and Cech, 1996;
Costa et al., 1997; Gordon et al., 2000; Karbstein et al., 2004). These
conformational steps in group I intron function should provide a tractable system
for unraveling properties of RNA conformational changes, which represents one
of the major current challenges in understanding RNA structure and function.
The folding of RNA to its functional structure can be thought of as a series
of conformational steps. Understanding how RNA adopts its structure is an
important physical challenge, and a necessary step in understanding how
biological systems modify and control folding and conformational processes. We
expect that folding studies that span levels of complexity from structures like the
Tetrahymena intron to simple RNA secondary and tertiary structural units will
reveal dynamic properties of RNA and will provide information about and tests of
the basic physics underlying RNA behavior.
ACKNOWLEDGMENTS
We thank Tom Cech for his original discovery of the self-splicing introns
and all of the researchers in our and other labs who have contributed to
understanding these fascinating molecules over the past two decades. We thank
V. DeRose, A. Kravchuk, and R. Das for permission to cite unpublished results, T.
67
Cech for permission to reproduce figures; and the NIH (GM49243) and HHMI for
supporting research in this area from the Herschlag and Piccirilli labs,
respectively.
68
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91
Figure 1. “Cloverleaf” secondary structure of alanine tRNA from yeast (Holley et
al., 1965).
Figure 2. Historical tour of proposed secondary structures of the Tetrahymena
group I intron: (a) (Michel and Dujon, 1983); (b) (Cech et al., 1983); (c) (Kim and
Cech, 1987); and (d) (Lehnert et al., 1996). Colors and numbering of the basepaired regions (P) reflect the complementary helical elements now known to be
formed. P1 is violet; P2 and P2.1, orange; P3-P7, blue; P4-P6, green; P5abc,
red; P9, cyan; P9.0, dark green. J8/7, the region between P8 and P7, is in pink.
(“J” refers to “joining regions” in the secondary structure; “L” refers to “loops”.)
Solid lines indicate a direct connection in the primary sequence, and arrowheads
are drawn to show the directionality of the chain. The intron is in upper case
letters and the exons are in lower case.
Figure 3. First proposed three dimensional structure of the core of the
Tetrahymena group I intron. From (Kim and Cech, 1987) with permission.
Figure 4. Specificity switch experiments that led to the localization of the
guanosine-binding site (Michel et al., 1989). The wild type base pair (G264-C311)
is shown in (a) and (c) with guanosine or 2-aminopurine ribonucleoside bound,
respectively. The A264-U311mutant is shown in (b) and (d) with guanosine or 2aminopurine ribonucleoside bound, respectively. The green and red boxes
represent the presence and absence of a hydrogen bonding interaction,
respectively.
Figure 5. The three dimensional structural model of the Tetrahymena group I
intron [(Lehnert et al., 1996); http://www-ibmc.u-strasbg.fr/upr9002/westhof/)].
Color-coding and helices match those in Figure 2d. (a) The entire intron. (b) Two
sets of coaxially stacked helices and joining regions (P4-P6 in green, P3-P8 in
blue, and J8/7 in pink) make up the conserved core. (c) The active site is defined
by a crevice at the junction of P4-P6 and P3-P8 where the P1 duplex (violet)
92
docks. (d) & (e) Long-range tertiary interactions (P13, P14, and L9-P5, Fig. 2d)
bring the peripheral helices (P2, P2.1, P5abc, and P9) in contact (d) and
surround the core [in gray in (e)].
Figure 6. Group I intron self-splicing. The intron is shown in black, the 5'
-exon in
red, and the 3'
-exon in blue. The base paired regions P1, P9.0, and P10 are
shown as lines for Watson-Crick base pairs, dots for wobble pairs, and an open
dot for a mismatch (from the Tetrahymena intron). The exogenous guanosine
nucleophile and the intron’s 3'
-terminal guanosine (ωG) are both shown in green.
Figure 7. The Tetrahymena group I ribozyme. (a) Conversion of the Tetrahymena
group I self-splicing intron into a multiple turnover ribozyme through removal of
the 5'
- and 3'
-splice sites (Zaug et al., 1986; Zaug et al., 1988). The resulting
ribozyme catalyzes an endonuclease reaction in trans. (b) The basic catalytic
cycle of the Tetrahymena ribozyme. (c) Expanded view of the conversion
between bound substrate and products [kforward from (b)], with deprotonation of
the guanosine nucleophile’s 3'
-OH group prior to phosphoryl transfer.
Figure 8. Kinetic and thermodynamic framework for the Tetrahymena ribozyme
reaction. CCCUCUA is the oligonucleotide substrate (S) that is cleaved in a
phosphoryl transfer reaction with the 3'
-terminal adenosine tail transferred to the
guanosine nucleophile (G) to yield the products GpA and CCCUCU (P). The
reaction proceeds as described in the text and references (Herschlag and Cech,
1990b; Bevilacqua et al., 1992; Herschlag, 1992; McConnell et al., 1993; Mei and
Herschlag, 1995; Mei and Herschlag, 1996; Narlikar and Herschlag, 1996;
Karbstein et al., 2002; Karbstein and Herschlag, 2003).
Figure 9. Evolution of the transition-state model for the group I ribozyme reaction.
For consistency, Tetrahymena intron numbering is used throughout. At each step,
novel metal ion interactions are denoted by filled red dots, and metal ion contacts
carried over from previous models are shown as open red dots. The initial
93
proposed interaction with the pro-SP oxygen of the scissile phosphate is shown in
blue. References: (a), [(McSwiggen and Cech, 1989; Rajagopal et al., 1989) and
references in the text]; (b), (Piccirilli et al., 1993; Sjogren et al., 1997; Weinstein
et al., 1997); (c), (Knitt et al., 1994; Shan and Herschlag, 1999; Shan et al., 1999;
Strobel and Ortoleva-Donnelly, 1999; Yoshida et al., 2000; Shan et al., 2001); (d),
(Adams et al., 2004a).
Figure 10. Comparison of group I intron structural models. (a) Pairwise
superpositions of the Tetrahymena (red, PDB ID 1X8W, chain A), Azoarcus (blue,
PDB ID 1U6B), and Twort (green, PDB ID 1Y0Q) group I intron crystal structure
models (Adams et al., 2004a; Guo et al., 2004; Golden et al., 2005). The
conserved core helices are noted in block letters and the peripheral domains
unique to particular introns are noted in colors that correspond to that intron’s
color. (b) Overlay of the P3-P9 and P4-P6 domains from the Tetrahymena crystal
structure (red) and the Michel-Westhof model (black) (Lehnert et al., 1996). (c)
Overlay of the Azoarcus intron crystal structure (blue) and its biochemicallyderived model (black) (Rangan et al., 2004).
Figure 11. Overlay of ribbon diagram of P4-P6 domains from structural modeling
by Michel-Westhof [blue; (Michel and Westhof, 1990)] and x-ray crystallography
[red; (Cate et al., 1996a)]. The two regions of direct tertiary contact are
highlighted: One contact between the A-rich bulge (A183-A187) and the minor
groove of P4 (G212, C109-G110) is shown in cyan for the Michel-Westhof model
and in yellow for the x-ray crystal structure. The other contact between the GAAA
tetraloop (G151-A154) and tetraloop receptor (U247-G251, C222-G227) is shown
in gray and green for the Michel-Westhof model and the x-ray crystal structure,
respectively.
Figure 12. Tertiary interactions between the conserved 5'
-splice junction (the
G•U wobble pair) and the A-rich bulge of J4/5. (a) Hydrogen bonding networks
from the x-ray structure of the Azoarcus intron; distances are in Å (Adams et al.,
94
2004a; Guo et al., 2004; Golden et al., 2005). The residue numbers are shown
for the Tetrahymena and Azoarcus introns in black and green, respectively. The
U(-1) position was thymidine, which contains an extra methyl group on the base.
The 2'
-hydroxyl at this position is absent from the crystallized complex and
therefore was modeled. (b) Wobble base pairs for guanosine (G) or inosine (I)
with uridine (U). (c) Atomic interactions from part (a).
Figure 13. Tertiary interactions between the docked P1 helix and J8/7 from the xray structure of the Azoarcus (a) and Twort (b) introns (Adams et al., 2004a; Guo
et al., 2004; Golden et al., 2005). The P1 helix is shown in red (5'
-exon strand)
and orange (IGS strand), with the bases other than the G•U wobble pair omitted
to allow visualization of the backbone contacts to J8/7, which is shown with its
atoms color-coded. Residue numbering is shown for comparison, with
Tetrahymena numbering in black, Azoarcus in green, and Twort in blue.
Figure 14. Comparisons of the overall geometry of the docking site of the P1
helix. (a) Overlay of x-ray structures from the Azoarcus and Tetrahymena introns
(Adams et al., 2004b; Guo et al., 2004). The P1 region of the Azoarcus intron is
in yellow; J4/5, green; ωG, brown; and J8/7, cyan. The corresponding regions for
the Tetrahymena intron are shown with the darker shades. (Note that P1 is not
present in the Tetrahymena structure.) The J4/5 region of the Tetrahymena intron
is in dark green; ωG, black; J8/7, purple. The J8/7 backbones are also shown as
ribbons to guide the eye, in blue for the Azoarcus intron and in red for the
Tehtrahymena intron. Differences in the J8/7 sequences are denoted by
underlined residues. (b) Overlay of x-ray structures from the Azoarcus and Twort
introns (Adams et al., 2004b; Golden et al., 2005). The Azoarcus structure is
represented as in part (a), and the corresponding Twort regions are in darker
shades.
Figure 15. Guanosine binding site from the group I intron x-ray crystal structures
(Adams et al., 2004b; Guo et al., 2004; Golden et al., 2005). (a) Overlay of the
95
base triples between ωG and G264-C311 base pair (Tetrahymena numbering) or
homologs for the Azoarcus (black), Tetrahymena (blue), and Twort (violet) introns.
(b) Side view of guanosine binding site from the x-ray structure of the
Tetrahymena intron (Guo et al., 2004). (c) Secondary structure of the guanosine
binding site with tertiary contacts shown schematically, with colors corresponding
to those in part (b) for comparison. Parts (b) and (c) are reproduced from (Guo et
al., 2004) with permission.
Figure 16. Transition state model for group I introns with identified catalytic metal
ion ligands. (a) Model includes catalytic metal ion ligands identified from
functional assays (J. Hougland et al., submitted) (J. Lee et al., in prep.) (A.
Kravchuk and D. Herschlag, unpubl.) (Szewczak et al., 2002). (b) The three
functionally identified catalytic metal ion ligands as located within the
Tetrahymena, Azoarcus, and Twort crystal structure models (Adams et al.,
2004a; Guo et al., 2004; Golden et al., 2005). The cleavage site uridine and ωG
are black, and the active site residues are colored to match the color scheme
from (a) with the phosphoryl oxygens that serve as catalytic metal ion ligands
depicted as orange spheres. The hydrogen bond between the cleavage site 2'
hydroxyl [of U(-1)] and the 2'
-hydroxyl of residue A207 (or homolog) is shown as
a dashed red line.
96
Figure 1.
97
Figure 2.
(a)
(b)
C
U U
A U
G C 240
A U
C
P6b A GU
A U
C G
U A
G U
A
A
U AA
U
A
A
180
G
G
A
U
C G P6a
C
GU
U
A
G
C A
U
U
A A A G GG
AU G
G
U
C
C
A
220 G U G
C AC
AC
G UUC
C
A
G G A C 200
C
A P5a U
A
A
A
UG
P5c GG U GU 140
C C G P6
CC
UGG
260
U
UCC A A
A C
G
A
U
G
C
160 G U
C
C G A U C AC
A C AA C U G
GG G
A C
C
A
G A P5b
G
A A AG G G C
120
U G
G
P5
A
U G
P4 U
C
U G
U
U
C
G
A
A
AA
AA A
A
A
U
80
C G
G C AU
U
100 U
C
U
P7
G CU
P2.1
GG
UGU
A
UU
P3 CC G
UA
G
GA
A
G
AA
AU AC C A
A U 280 A A
UU
AA
A
A
C
G
G
UU
G
CG G
U
G AA
U
G
CU
60 C U C
U
P8
C
G
A
P2
A
300 U
C AC C U G G U AG C U
A
G
A
U
G
A CG G AC U A U U G A
A
A
U
40
A
A
A
U
G
A
G
G
P1 AG c
5’
U
20
C
cu ag u au c a
GG
G
u
U
G
uc
A C C UU
A P9b
U
A
A
320
C
-1
U
A
C UC
U
U
UC CU
A
A
U
G AG
U
A ACG
GG
C
A
UG AG GU
U
U A
U GG
CA
A
U A G U G P9.1
CU
340
G
U A
C CG
u
400 G C
G
a
AU G
a
C G
A U
C
gg
U A
u
G
U A 360 A
ga
16 nt
A U
G
3’
U
P9.2UG U
U G
A U
U A 380
A U
U G
G C
A G
AA
P5b
A
A
A
P5c
160
U
C U U UG AG A U G G C CU G
C
G G AA A
UG
A
U
180
G
G
U
A AU
A
P5a G C G A
U
A U
A
U
G
G
UC G
U A
C
A
G C
U A
80
C G
A U
C G
C
A
C U
G
A
C G
U
U
A
C G
C
C
C 240
A U
A
A
A
G U
A U
P2.1 A U
A U
A G
A
120 C G 200
C U P6b
U A
U G
A U
U A
G
A G
P5 G U
U
U A
C
G C
C
C G
A U A
60
A
A
C
G
U
U G
G G
U
G
U
U
G C
G
G
A
C AC
U
A
A
A
A
C A
A
A
260
A
C CUG G UA G C U
G
U C G G
G A
A
A
G
A
C U AA G
U
G C
GG A CU AU U G A
100
U
A U
U
CU
A
A C
G CA
C
G A U
P4
G
A
C
C
U
G
C
40
P2
C G
G
C
G
A
P7 A AG
G
C G
U
A G
G 280
A
G
G C
220 U
C
A
G
A
C
U
C
A
A
P9b
G
320
G P8
A
U A U
G G U AA U
G
A
U
C
A
C
C
C A
U C C C U
U
U AG
U
U
300
20
U
A U
G A G G
G G U U
C
A
C
G
A
C
U
C
U
UA
G
G
U
5’
U
A
400 G U A G G G A
A U
U
A
G
P9.1
U
A C
U
A
C
U AG
C
C
U
340
U
U
G G
U
U A
A AA
C G A UG
G A
P9.2
C
A
U
A U U
C
G
U
A
U
A
G
A
U
A
360 A
A
3’
G
U
G
U
C
P9.1a
G G U
A AU G
C UG
U U
A
A
380
C
U
A G G
P9.2a
G
A
C
A
A
C
A G
GG
G G ACU C U
G
140
(d)
(c)
A
A
C
C
A
U
P9.2
380
L9.2
A
A
G G U CG C C
AG U AU AU UG AU U AG U U U
C U AG CG G
400
P9.2a
P5b
A
A
A
GG
G G AC U C U
G
140
P5a
A
A
C
G
A
U
P1
-1
5’
U A
A
A
A
u
c
u
c
u
c
U
U
U
G
G
A
G
G
G
P5c
160
U
C U U UG AG A U G G C CU G
A
UG
A
C
C
A
U
G
A
UC
U
G
C
C
G
A
C
U
A
U
A
G
G
A
120 C
C
A
U
U A
U
U
A U 80
U G
P5 G
U
C
A
G
A
C
A
G
C G
C
C
A
C G
G
G C
C
40 G
A U P2.1
A
C
20
A U
A
A U
A UA
A
C G
U A
G
P2 UA GU
U A
P4 GG
U A
U A
100
U G 60 C G
C
P3
G C
G
U G
A A A A U
A G C A A G A C C G U C A A AU U
U
280
G
G U A G A AG G G
C UGGCU G U
A
A
A
U
180
G
G
U
A AU
A
CG A
U
G
A
C
G
G
A
C
A
U
G
200
G
U
C
C
U
A
A
C
CA
P6a
C
P6b
G
220
P6 C A A
C
UAA
A
GC
G U CC
GU CA ACA G U
A
U AG U UG U C U U
UU G
U AG G
U A
P7 A A C A C G
260
240
AU CAG C
A U U CU UC U C A U A A G A U A UA GU CG GAC C
U
P8 300
P9b U CC
320 C
U
U
G
A
G
G
G
C
U
A
G
C
G
U
G
A
A
A
U
340 G
A
A
P9.1aUG
G
A
U
G
A
G
G
A
G
C
U
C P9.1U
A
G
C 360 A
U
C
U
G
U
A
G
G
U
G
U 380 A
U
C
A
G
C
C
A
G
A
A
C
A
C
A
G
G
C
A
G
U
C
G
AA
U
A
AU
180 G
G
G U A U
G A C A
C
U
C
200 G
G
G
UU
U
C
C
A
C
G
U
U
P5a
P5c
P5
A A A G G
C
G U U C C G
G
L5c
U
A
160 G
A
G
U
U
U
C
A
P5b
C
U U U G G A G U A C U C G ua ag
U
G 400
A
U P9.2
U
A
G
U
U
A
U
A
U P9.2a
G
A
A
98
P14
3’
A
C
C
AC
G
G
G
C
G
U
A A
A
140
20
J4/5
P1
A
P4 GC
A
G
U
C
U
C
A
G
G
G
G
120
G
A
A
A
A
C
C
A
5
A
C
U
G
G
G
P6
G
C
C
A
A
220 G
U
C
C
U
A
A
G
U
C
A
A
C
A
G
A
U C
L2
U
G
260
U
U
A
C
C
U
U
U
G
G
A
G
G
G
UG G A
G
U U C C U C
UCC
AU
320
A
g
A
a
C
u
G
g
A
g
U
a
A
a
A
u
A
u -1
c
u
c
u
c
C A
U
A U G A
U
A
C
U
GC
A
G
P9.0
G
C
U
G
A
U
U
A
C
U
G
G
C
100
A
C
A
A
G
U
A
J8/7
P6a
300
P6b
A
A
40
A
A CG G AC U AU UG A
A
C
U
C
U
U
C
U
U
A U
P13
CA
G
A
C
U
A
P7
A
A
U
G
U
C
G
G
U
C
P3
G
G 280
G
G
A
A
G
A
U
G
P8
P2.1
A
80
A
A
C G G A A A U U U G G C U AC GU
U
C
C
G
U
C
U
U
U
A
A
A
A
A
C
C
U
U
C
U
G G
G
A AUAGA
C A
P2
60
L9.1
P9.1a
3
A
U U
G A A
A
C
G
U
A
G
G
U
A
U
A
G
U
U
G
U
C 240
U
U
U C AC A A
C
G
AG UG
A U
P9a
U G G G A G
U
G G
G
340
P9b
A
A
C
G G AA A
P9.1 360 A
G C G U A U G U U U A A U C A AG
L2.1
Figure 3.
99
Figure 4.
100
Figure 5.
101
Figure 6.
102
Figure 7.
103
Figure 8.
104
Figure 9.
105
Figure 10.
106
Figure 11.
107
Figure 12.
108
Figure 13.
109
Figure 14.
110
Figure 15.
111
Figure 16.
112
Scheme 1.
113
Table 1 Comparison of the ability of different models of the Tetrahymena group I intron to predict the correct secondary structure
and long-range interactions.
Michel and
Dujon (1983)
+++
Davies et al.
(1983)
+++
Cech et al.
(1983)
+
Kim and Cech
(1987)
+++
Lehnert et al.
(1996)
+++
P2
P2.1
+++
++
+++
++
+++
++
+++
++
+++
+++
P3
++
X
X
++
+++
(3)
P4
+++
+++
+++
+++
+++
(4)
P5
P5a
P5b
P5c
++
+
+++
++
++
+
++
+++
++
+
++
+++
++
+
++
+++
+++
+++
+++
+++
P6
P6a
P6b
+++
+++
+++
X
X
++
+++
X
++
+++
+
++
+++
+++
+++
(5), (10)
P7
+
+
++
++
+++
(3), (6)
P8
+++
+++
+++
+++
+++
(5)
P9.0
P9
P9.1
P9.1a
P9.2
O
X
+++
O
++
O
++
+++
+++
++
O
++
+++
+++
++
O
+
+++
+++
++
+++
+++
+++
+++
+++
(7), (8), (9)
P1
114
Tested by
mutagenesis
(1), (2)
(9)
Table 1 (Continued)
Michel and
Dujon (1983)
Davies et al.
(1983)
Cech et al.
(1983)
Kim and Cech
(1987)
Lehnert et al.
(1996)
Tested by
mutagenesis
P10
O
+++
O
O
+++
(7), (10)
P13
P14
L9-P5
J5a/5c-P4
L5b-J6a/6b
O
O
O
O
O
O
O
O
O
O
O
O
O
O
O
O
O
O
O
O
+++
+++
++
++
++
(11)
(11)
(11)
(12)
+++ = correct prediction; ++ = minor discrepancies; + = helix present but with several incorrect base pairs; X = incorrect
prediction; O = helix not present or not investigated
References: (1) (Waring et al., 1986); (2) (Been and Cech, 1986); (3) (Williamson et al., 1987); (4) (Flor et al., 1989); (5)
(Williamson et al., 1989); (6) (Burke et al., 1990); (7) (Michel et al., 1989); (8) (Burke et al., 1990); (9) (Michel and Westhof, 1990);
(10) (Suh and Waring, 1990); (11) (Lehnert et al., 1996); (12) (Costa and Michel, 1995).
115
Table 2 Chronological list of group I intron and fragment structures.
RNA construct
Description
Technique
Resolution
Reference
P1
Hairpin containing a yeast
group I intron P1 sequence
NMR
-
(Allain and
Varani, 1995)
P4-P6
P4-P6 domain of the
Tetrahymena intron
x-ray
2.8 Å
(Cate et al.,
1996a)
P5b
P5b stem loop from the
Tetrahymena intron with
associated Co(III)
NMR
-
(Kieft and
Tinoco, 1997)
P5a bulge
Conserved five-nucleotide
bulge loop from the
Tetrahymena intron
embedded in a 25-nucleotide
hairpin
NMR
-
(Luebke et al.,
1997)
Tetrahymena
P4-P6, P3-P9
247-nucleotide RNA
containing the P4-P6 and P3P9 domains from
theTetrahymena intron
(missing P1, P2-P2.1, and
P9.1-P9.2)
x-ray
5.0 Å
(Golden et al.,
1998)
P5
14-nucleotide hairpin
containing the P5 helix from
the Tetrahymena intron
NMR
-
(Colmenarejo
and Tinoco,
1999)
P4-P6 ( C209
& A210)
Single-site deletion mutants
of the P4-P6 domain of the
the Tetrahymena intron
x-ray
2.3 Å
(Juneau et al.,
2001)
P7/P9.0/G
22-nucleotide model RNA
containing the guanosine
binding site and ωG
NMR
-
(Kitamura et
al., 2002)
Azoarcus group I
intron
Complete Azoarcus group I
intron with the 5'
- and 3'
exons
x-ray
3.1 Å
(Adams et al.,
2004a)
Tetrahymena
P4-P6, P3-P9
P4-P6 and P3-P9 domains
from the Tetrahymena intron
containing 5 stablizing
mutations (missing P1, P2P2.1, and P9.1-P9.2)
Complete Twort group I intron
with the 5'
- and 3'
- exons
x-ray
3.8 Å
(Guo et al.,
2004)
x-ray
3.6 Å
(Golden et al.,
2005)
Twort group I
intron
116
Table 3. Comparison of tertiary interactions upon P1 docking in group I introns identified by biochemical and structural
approaches.
Tetrahymena thermophila
Position
5'-Splice
Site
Analog
5'-Splice
Site
Wobble
Pair
‡
tert
G
b
(kcal/mol)
G
a
(kcal/mol)
Contacts
identified by
functional
c
experiments
G
d
(kcal/mol)
OH
2'
OH - A207 2'
2'
OH
tert
Contacts
identified
e
by NAIM
Contacts
identified
by x-ray
f
structure
G
g
(kcal/mol)
2'
OH
A207 2'
OH
2'
OH -0.3
2'
OH -0.8
tert
tert
G
h
(kcal/mol)
2'
OH
2'
OMe
-4.8
<1.8
2'
OH
C(-2)
2'
OH
2'
OMe
-1.9
-0.5
2'
OH -0.4
2'
OH - G303 NH2
2'
OH -0.7
2'
OH
G303 O4'
2'
OH -0.3
2'
OH -1.0
U(-3)
2'
OH
2'
NH2
2'
OMe
2'
F
-2.7
0.6
<1.5
0.4
2'
OH -0.9
2'
NH2 -0.1
2'
OH - A302 N1
2'
OH -1.5
2'
OH
A302 N1
2'
OH 0.1
2'
OH -1.5
C(-4)
2'
OH
2'
OMe
-0.5
i
-0.6
i
2'
OH
0
X
N/D
N/D
N/D
2'
OH -0.7
2'
OH 0.4
C(-5)
2'
OH
2'
OMe
-0.8
i
-0.7
i
2'
OH
0
X
N/A
N/A
N/A
2'
OH -0.2
2'
OH 0.9
C(-6)
2'
OH
2'
OMe
-0.8
i
-0.9
i
2'
OH
0
X
N/A
N/A
N/A
2'
OH -0.5
2'
OH 1.2
j
GU/GC
-3.0, -3.4
0
Candida
albicans
U(-1)
G•U(-1)
0.1
Pneumocystis carinii
Azoarcus indigens
GU/GC
117
j
-4.3
GU/GC
j
-2.8
Table 3 (Continued).
Tetrahymena thermophila
Position
Internal
Guide
Sequence
a
‡
G
a
(kcal/mol)
tert
G
b
(kcal/mol)
Azoarcus indigens
Contacts
identified by
functional
c
experiments
tert
G
d
(kcal/mol)
Contacts
identified
e
by NAIM
Contacts
identified by xf
ray structure
Pneumocystis carinii
tert
Candida
albicans
G
tert
G
g
(kcal/mol)
(kcal/mol)
G22
2'
OH
2'
OMe
-3.7
0
2'
OH
-3.1
2'
OH -
A114
OH
N3 & 2'
N/D
2'
OH
N2-NH2
2'
OH - A114
OH
N3 & 2'
N/D
N/D
G23
2'
OH
2'
OMe
-1.6
-1.8
2'
OH
-0.8
N2-NH2 - A207
OH
N3 & 2'
N/D
X
N2-NH2 - A207
OH
N3 & 2'
N/D
N/D
A24
2'
OH
-0.5
2'
OH
0.7
X
N/D
X
X
N/D
N/D
G25
2'
OH
2'
OMe
-3.2
0.8
2'
OH
-2.5
2'
OH -
A301
2'
OH & N3
N/D
2'
OH
A301
2'
OH & N3
N/D
N/D
G26
2'
OH
-1.7
2'
OH
-0.8
OH
2'
OH - U300 2'
N/A
N/A
N/A
N/D
N/D
G27
2'
OH
-0.9
2'
OH
0
X
N/A
N/A
N/A
N/D
N/D
‡
2'
OH -
G (kcal/mol) indicates the energetic effect of 2'
-X groups relative to 2'
-H on the overall transition state stabilization (Herschlag et al., 1993b;
Narlikar et al., 1997). See also (Bevilacqua and Turner, 1991; Pyle and Cech, 1991).
b
tert
-X groups relative to 2'
-H upon P1 docking only (Strobel and Cech, 1993; Narlikar et al., 1997). Note that 2'
G is the energetic effect of 2'
hydroxyl and methoxy substituents render duplexes more stable than 2'
-H, so some of the observed overall effects have been corrected for this
stabilization.
c
Tertiary interactions between in P1 and the core of the Tetrahymena thermophila ribozyme identified by functional studies (Pyle et al., 1992;
Ortoleva-Donnelly et al., 1998; Szewczak et al., 1998; Strobel and Ortoleva-Donnelly, 1999). Atoms in contact are indicated by a single underline;
for case in which more contacts are formed, the atoms in contact are distinguished by a single or double underline.
d
tert
G is as defined in b. The energetic effects were calculated in the Azoarcus indigens ribozyme (Strauss-Soukup and Strobel, 2000).
e
Values taken from (Strauss-Soukup and Strobel, 2000).
f
Tertiary interactions identified from the x-ray structure of the Azoarcus indigens ribozyme (Adams et al., 2004b; Adams et al., 2004a).
g
tert
G is as defined in b. The energetic effects were calculated in the Pneumocystis carinii ribozyme (Testa et al., 1997).
tert
h
G is as defined in b. The energetic effects were calculated in the Candida albicans ribozyme (Disney et al., 2000).
i
This energetic effect arises solely from duplex stability of 2'
-OH and 2'
-OMe relative to 2'
-H, and not from docking interactions.
118
h
j
tert
G of GU/GC indicates the stabilization energy by G•U wobble pair relative to GC pair at catalytic site (Knitt et al., 1994; Pyle et al., 1994;
Disney et al., 2000; Disney et al., 2001).
“X” represents no interaction found; “N/D” and “N/A” mean “not-determined” and “not -applicable”, respectively.
119