Structural study of elements of Tetrahymena telomerase RNA stem–loop IV domain important for function

  1. Rebecca J. Richards1,
  2. Haihong Wu1,
  3. Lukas Trantirek1,3,
  4. Catherine M. O'Connor2,
  5. Kathleen Collins2, and
  6. Juli Feigon1
  1. 1Department of Chemistry and Biochemistry, and Molecular Biology Institute, University of California, Los Angeles, California 90095-1569, USA
  2. 2Department of Molecular and Cell Biology, University of California, Berkeley, California 94720-3204, USA
 
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Abstract

Tetrahymena telomerase RNA (TER) contains several regions in addition to the template that are important for function. Central among these is the stem–loop IV domain, which is involved in both catalysis and RNP assembly, and includes binding sites for both the holoenzyme assembly protein p65 and telomerase reverse transcriptase (TERT). Stem–loop IV contains two regions with high evolutionary sequence conservation: a central GA bulge between helices, and a terminal loop. We solved the solution structure of loop IV and modeled the structure of the helical region containing the GA bulge, using NMR and residual dipolar couplings. The central GA bulge with flanking C–G base pairs induces a ∼50° semi-rigid bend in the helix. Loop IV is highly structured, and contains a conserved C–U base pair at the top of the helical stem. Analysis of new and previous biochemical data in light of the structure provides a rationale for some of the sequence conservation in this region of TER. The results suggest that during holoenzyme assembly the protein p65 recognizes a bend in stem IV, and this binding to central stem IV helps to position the structured loop IV for interaction with TERT and other region(s) of TER.

Keywords

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INTRODUCTION

Telomerase is a ribonucleoprotein (RNP) complex that maintains the 3′ ends of chromosomes (Greider and Blackburn 1985; Chan and Blackburn 2004). Telomerases identified from diverse eukaryotes all contain both an RNA component (TER), which includes a template sequence required for telomeric DNA synthesis, and a catalytic protein component telomerase reverse transcriptase (TERT). Together, TER and TERT provide the minimal telomerase subunits required to reconstitute enzymatic activity in vitro (Kelleher et al. 2002). Telomerase synthesizes telomeric repeats (TTGGGG in Tetrahymena) by processive nucleotide addition across the template to generate a single repeat (nucleotide addition processivity, NAP) and processive repeat addition by reuse of the same template (repeat addition processivity, RAP) (Greider and Blackburn 1989; Greider 1991; Lue 2004). The endogenously assembled telomerase holoenzymes contain additional protein subunits essential for activity in vivo, whose roles in RNP assembly, stabilization, and localization have been less well defined (Harrington 2003; Chen and Greider 2004).

The Tetrahymena thermophila telomerase RNP holoenzyme has been characterized by affinity purification, and consists of TERT, TER, and several other associated proteins (Witkin and Collins 2004). The TERT protein contains a region of homology with other reverse transcriptases as well as telomerase-specific regions that are also required for activity, including the essential TEN (TERT essential N-terminal) and RBD (RNA binding domain) domains implicated in TER binding (Lingner et al. 1997; Nakamura and Cech 1998; Lai et al. 2001; O'Connor et al. 2005; Jacobs et al. 2006). Of the accessory proteins, p65 has been shown to form a complex with TER and enhance the assembly of TERT with TER in vitro (Prathapam et al. 2005; O'Connor and Collins 2006). This role for p65 in RNP assembly is likely to account for its genetic requirement for TER and TERT accumulation in vivo.

T. thermophila TER is 159 nucleotides in length and contains four conserved helical regions, stems I–IV, including a potential pseudoknot of stems IIIa and IIIb (Fig. 1A; Romero and Blackburn 1991; Lingner et al. 1994; McCormick-Graham and Romero 1995). The template sequence is bordered on its 5′ side by a template boundary element (TBE) and on its 3′ side by a template recognition element (TRE), both of which are required for correct template definition and usage (Autexier and Greider 1995; Lai et al. 2002; Miller and Collins 2002; Richards et al. 2006). In addition to these template-adjacent elements, the pseudoknot and stem–loop IV have also been shown to have roles in telomerase function (Autexier and Greider 1998; Gilley and Blackburn 1999; Licht and Collins 1999; Sperger and Cech 2001; Lai et al. 2002, 2003; Mason et al. 2003).

FIGURE 1.
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FIGURE 1.

(A) Sequence and secondary structure of the T. thermophila telomerase RNA. The four helical regions as well as the template, TRE, and TBE are labeled. (B) Sequence conservation of stem–loop IV mapped on the proposed secondary structure, modified from the analysis of 17 Tetrahymenine species (Ye and Romero 2002). Absolutely conserved nucleotides are identified while variable nucleotides are shown as pyrimidines (open circles), purines (closed circles), and any nucleotide and/or gapped position (gray circles). Bold dashes between nucleotides indicate positions where at least 15 of the 17 sequences are conserved to base pair (includes G–U base pairs). The length of the proximal stem below the bulged U is variable. (C,D) Sequence and secondary structure representation of stem IV NMR constructs for (C) loop IV and (D) the GA bulge. Residues are numbered to match the full-length T. thermophila TER. Gray nucleotides indicate non-native sequences added during construct design.


Comparison of 17 TER sequences from the Tetrahymenine group ciliated protozoa and predicted secondary structures (Ye and Romero 2002) reveals that stem IV has the highest sequence conservation of TER helical elements (Fig. 1B). Stem–loop IV in T. thermophila consists of two predicted helical regions with a conserved GA bulge in the middle, and is capped by a highly conserved heptaloop (Fig. 1A,B). Both loop IV (last four base pairs and heptaloop; nucleotides 128–142) and the GA bulge region have been shown to have separate but interdependent roles in the function of stem–loop IV. Stem–loop IV influences the activity of telomerase in multiple ways, contributing to both the catalytic cycle and RNP assembly. The contribution of stem–loop IV to NAP and RAP has been shown to be dependent on loop IV (Sperger and Cech 2001; Lai et al. 2003; Mason et al. 2003). Crosslinking and in vitro binding experiments have also identified this loop as a potential interaction site for TERT (Lai et al. 2003; O'Connor et al. 2005). Stem IV, and in particular, the conserved bulged GA nucleotides in the center of the stem, has been implicated in the loop IV-dependent assembly of TERT with TER (Prathapam et al. 2005). Deletion of the GA bulge influences both the contribution of loop IV to catalytic activity and the correct folding of TERT with TER in vitro as assayed by nuclease footprinting (Sperger and Cech 2001).

In an early study of ciliate TER, a structural kink at the GA bulge was proposed as a site for protein binding (Bhattacharyya and Blackburn 1994). Recently, the template-proximal stem IV and GA bulge, along with stem I, have been identified as the core binding site of p65 (O'Connor and Collins 2006). The p65 protein has been shown to initiate the hierarchical assembly of the Tetrahymena holoenzyme, enhancing assembly of TERT with TER (Prathapam et al. 2005; O'Connor and Collins 2006). Substitution of distal stem–loop IV with a tetraloop does not significantly reduce p65 binding but does eliminate the p65 stimulation of TERT binding to TER. Loop IV deletion reduces activity in the context of the p65–TERT–TER RNP, as it does for TERT–TER alone.

To provide structural insight into the role of the conserved elements of stem–loop IV in telomerase function, we solved the solution structure of loop IV and modeled the structure of the helical region containing the GA bulge using NMR and residual dipolar couplings (RDCs). We find that the GA bulge between proximal and distal stem IV induces an ∼50° semi-rigid bend in the stem that flexes in a defined direction. Deletion analysis indicates that this sequence specific bend is the only deviation of stem IV from the helical structure that strongly influences activity. The loop IV is highly structured, and contains a conserved C–U base pair at the top of the helical stem. Mutational analysis supports the importance of the C–U pair stacked on the closing base pair of the helix.

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RESULTS

The GA bulge introduces a large bend in stem IV

The GA bulge and the C–G base pairs on either side are completely conserved among TERs from Tetrahymenine ciliates, but there is considerable variability in the sequence of the rest of the stem except for the four base pairs adjacent to the loop. We investigated whether the GA bulge in the context of helices closed by C–G pairs adjacent to the bulge caused this region to be conformationally flexible or if it induced a defined bend in the RNA. Enzymatic probing indicates that both the proximal and distal stem IV are double stranded in the free RNA (Bhattacharyya and Blackburn 1994; Sperger and Cech 2001). We designed an RNA construct that contained the GA bulge with four base pairs on either side, including the conserved C–G base pairs (GABIV) (Fig. 1D). The helix is stabilized for our NMR studies by a c(UUCG)g tetraloop at the distal end. The first two nucleotides in the proximal stem are Gs for optimal in vitro transcription yields, and the nonconserved bulge U nucleotide was deleted. We calculated the relative orientation of the stem regions (domains P and D) on either side of the GA bulge, using order tensor analysis of residual dipolar couplings (Fig. 2A,B). The order tensor solutions were calculated separately for domains P and D using a total of 10 and 12 RDCs, respectively. Domains P and D were independently rotated into the principal axis system of the order tensor, yielding four interhelical orientations that were consistent with the RDC data. Two of these orientations could be discarded because they lead to antiparallel helical orientations, while the third one could be discarded due to violation of the linkage geometry. The conformation of GABIV that is consistent with both RDCs and nucleic acid stereochemistry is shown in Figure 2. The results indicate that the GA bulge in stem IV introduces a defined kink with an interdomain angle between the two helical regions of 50 ± 15°. Sequential H1′–H8/H6 NOE cross-peaks were observed from the 5′ end along the entire sequence except for the expected break in the UUCG tetraloop. The NOE patterns and chemical shifts of G121 and A122 are consistent with insertion of these bases in the helix, stacking on each other and the flanking C–G base pairs to form a structured bulge. We, therefore, calculated a model structure with bases of G121 and A122 inserted into the helix. The model was initially calculated using sparse sequential NOEs over the GA bulge and artificial constraints imposed on domains P and D and the UUCG tetraloop, and subsequently refined against the RDCs (Fig. 2A,C). The GA bulge creates a large wedge that bends the helix into the major groove at the bulge toward the strand without the bulge.

FIGURE 2.
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FIGURE 2.

(A) Model structure of the GA bulge RNA construct (GABIV) calculated as described. (B) Cylinder representation of the proximal and distal helices showing the bend introduced by the GA bulge and the interdomain motion. (C) Stereoview of the model structure of the GA bulge RNA with the UUCG tetraloop deleted for clarity. Nucleotides are colored A (orange), U (blue), C (green), and G (red).


Bending motions of the GA bulge have a directional preference

The results described above indicate that the GA bulge induces a defined bend. We anticipated that the GA bulge would result in conformational flexibility as is commonly seen in RNA internal loops (Al-Hashimi et al. 2002; Dayie et al. 2002). To address the question of whether domain D and P are conformationally flexible or held rigid with respect to one another, the domain-specific order parameters (υ, η) were compared (Tolman et al. 2001). As shown in Figure 3A, calculated υ values for domain P are quenched by ∼15% relative to values computed for domain D. This attenuation is preserved even when all the RDC data belonging to the ribose sugar rings are excluded. This negates static or dynamic deviation in local stem geometries as a source for the observed discrepancy. Instead, the observed differences in υ values indicate that the two domains experience slightly different molecular alignments and that the two domains are not held rigid with respect to one another, but rather undergo small interdomain fluctuations. To obtain information on the directional character of interdomain motions, domain-specific asymmetry parameters were compared (Al-Hashimi et al. 2002). As shown in Figure 3B, computed η values for domain D are quenched by ∼40% relative to values calculated for domain P. This suggests that the interdomain motions have a very strong directional preference. Assuming a two-state model for interdomain motion, the amplitude of the motion can be estimated to be <20°. Taken together, these results indicate that, even in the absence of protein, the GA bulge induces a large semi-rigid bend that flexes in a defined direction.

FIGURE 3.
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FIGURE 3.

Histogram plots of generalized degrees of (A) order parameter υ and (B) asymmetry parameter η, calculated independently for domain P (empty boxes) and D (black boxes).


Loop IV is highly structured

The role of stem–loop IV in NAP and RAP has been shown to be mediated by loop IV alone. Loop IV added in trans to nucleotides 1–107 of T. thermophila TER stimulates processivity as effectively as the entire stem IV domain (Lai et al. 2003; Mason et al. 2003). In order to study loop IV, we first synthesized an RNA stem–loop containing the wild-type sequence with an additional G–C pair for optimal transcription. This RNA formed primarily dimer (duplex) under all NMR conditions tested (data not shown), and therefore, we extended the helix by an additional two G–C base pairs (Fig. 1C) (SLIV) to increase stability, reduce dimer formation, and increase transcription efficiency.

The solution structure of SLIV was solved using multidimensional NMR spectroscopy at 20°C on unlabeled and uniformly 13C,15N-labeled RNA samples (see Materials and Methods). The proton spectra of loop IV were well dispersed, and NOESY spectra gave rise to a large number of NOEs for all of the nucleotides. The structure determination of loop IV included 452 NOE distance restraints for an average of 22 NOE restraints per nucleotide, 115 dihedral angle restraints, and 17 residual dipolar couplings (Table 1). SLIV has a well-defined structure with an RMSD to the mean of 0.71 ± 0.15 Å for all heavy atoms for the 20 lowest energy structures (Fig. 4A).

FIGURE 4.
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FIGURE 4.

Solution structure of the RNA construct of loop IV (SLIV). (A) Twenty lowest energy structures of the entire SLIV. (B) Twenty lowest energy structures of the C–U base pair closing loop IV. (C,D) Stereoviews of the loop and stem A–U base pair showing view into (C) the major groove and (D) the minor groove of the loop. Nucleotides are colored as in Figure 2.


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TABLE 1

Structural and refinement statistics for loop IV


The SLIV structure has a seven–base-pair stem that forms a standard A-form helix closed by the top A–U pair (Fig. 4A). The seven nucleotide loop IV is surprisingly well structured (Fig. 4B,C). Stacked on top of the A–U base pair is a noncanonical C–U base pair formed by the first and last nucleotides in the loop, with a single potential hydrogen bond between one of the C132 amino protons and the U138 carbonyl oxygen at the 4 position (Fig. 4D). Although we were unable to directly detect this hydrogen bond, the donor and acceptor groups are within hydrogen bond distance in all of the calculated structures, and its existence is supported by the U128 carbonyl chemical shift in the hydrogen bonded range and the observation of a H4(1) amino proton resonance. On the 5′ side of the loop, A131, C132, A133, and C134 all stack on each other with A-form stacking interactions and dihedral angles. The base of C134, the only nonconserved nucleotide in the loop, is rotated slightly so that its Watson-Crick face is pointing into the major groove. U135 is positioned directly at the top of the loop, with its Watson-Crick face exposed to solvent. The turn in the phosphodiester backbone occurs between U135 and A136. The A136 nucleotide has a glycosidic torsion angle in the high anti range, positioning the base below the ribose of U135 and over its own ribose. The base of U137 lies outside the loop on the major groove side almost perpendicular to the helical axis, and is the least well defined of the loop nucleotides. The sugar moieties of nucleotides U135, A136, and U137 all have South-type (C2′-endo) sugar puckering. The last nucleotide in the loop, U138, stacks on the A-form helix in a noncanonical C–U base pair, as discussed above. Interestingly, the Watson-Crick faces of all of the nucleotides in the loop above the C132–U138 base pair are exposed on the major groove side of the loop.

The sequence of the loop is completely conserved on the 3′ side (U135–U138) among 17 Tetrahymenine ciliate TERs. On the 5′ side, C134 is frequently an A, and in one of 17 cases a U, and five of the 17 sequences have an extra nucleotide between C132 and A133 (Fig. 1B; Ye and Romero 2002). Since C134 is stacked out and over U133 with no other interactions with the loop, a U or A could easily be accommodated in this position without changing the loop fold. It is less clear what effect an extra nucleotide between C132 and A133 would have on the loop structure. Either a C or an A could lie in the minor groove with minimal effect on the rest of the loop, but we cannot predict whether they would be out or stacked in. The presence of this extra nucleotide in the loop is correlated with a change in the identity of the nucleotides at the top of the distal stem (see below).

Consistent with the lower sequence conservation on the 5′ side of the loop, T. thermophila TER mutations A133U and C134G have little effect on TERT–TER activity in vitro (Sperger and Cech 2001; O'Connor et al. 2005). TER mutations C132G and U138A, which would disrupt the noncanonical C–U base pair, as well as U137A, have a dramatic effect on the overall level of TERT–TER activity. TER mutations U135A and A136U, the two remaining nucleotides that are located at the top of the loop, have only a small effect on overall activity, although they are completely conserved.

Activity and RNP assembly do not require a bulged nucleotide in proximal or distal stem IV

We investigated the importance of the stem IV unpaired nucleotides and predicted base pairs not supported by phylogenetic covariation, including single bulged U nucleotides in the distal and proximal stems, the GA bulge, and the base pair at the end of distal stem IV below loop IV. We tested deletion and substitution variants of TER for activity with TERT in the presence or absence of p65 (Fig. 5).

FIGURE 5.
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FIGURE 5.

Primer extension assays of TERT assembled with wild-type and mutant TER in the (A) absence and (B) presence of p65. TER substitutions are indicated above the lanes. Successive strong products from the bottom of the gel up correspond to synthesis of the completed first, second, and third repeats added to primer.


In T. thermophila, the stem IV GA bulge is flanked by two helical regions, each eight base pairs long and containing a single bulged U. Among Tetrahymenine species, template-proximal stem IV varies in length from six to nine base pairs, and neither the position nor presence of a bulged nucleotide is conserved. We checked the importance of the bulged U117, which might induce additional conformational flexibility in proximal stem IV, for telomerase activity. The enzymatic activity of TERT with TER(ΔU117) was comparable to wild type (Fig. 5A). This result is consistent with the lack of conservation of a bulged nucleotide in proximal stem IV and with the base paring in this region inferred from RNase VI cleavage (Sperger and Cech 2001).

The distal stem IV is A–U rich, usually eight base pairs long with one bulged nucleotide or mismatch. The distal stem IV 5′ strand (nucleotides 123–131 in T. thermophila) is always nine nucleotides and the 3′ strand (nucleotides 139–146 in T. thermophila) is eight or nine nucleotides. T. thermophila distal stem IV has a nonconserved bulge, U127. In almost 50% of the Tetrahymenine ciliates, this bulge is either replaced by an additional base pair in distal stem IV or it would be possible to form an eight–base-pair stem with a bulged nucleotide in a different location. We find that deletion of U127 does not alter NAP or RAP, but it does decrease the overall activity generated by TERT and TER alone (Fig. 5A). However, in the presence of p65, a wild-type level of activity is restored (Fig. 5B). These results suggest that the bulged U127 affects TERT–TER interaction in a manner that is not essential for activity in the physiological context of the p65-assembled RNP (see Discussion).

The GA bulge is required for activity and RNP assembly

Previous mutational analysis has indicated that deletion of the GA bulge can have a large negative impact on overall activity in the context of TERT–TER reconstituted in vitro (Sperger and Cech 2001; O'Connor et al. 2005). Substitution of GA with UU or UC gives near wild-type activity when TER is added to TERT in excess and activity is assayed without purification from assembly in rabbit reticulocyte lysate (Autexier and Greider 1998; Sperger and Cech 2001; O'Connor et al. 2005). In reconstitution extracts with lower levels of TER (Fig. 5), GA bulge deletion has a large negative impact on activity as well, and activity is not rescued by the presence of p65. Combined, all of the results described above for stem IV bulged nucleotides suggest that the features of stem IV important for activity in the context of the holoenzyme are two helices with a central conserved bend formed sequence specifically by a GA bulge.

The helical stem of distal loop IV is important for activity

Of the four base pairs adjacent to loop IV, only the stem-capping A–U pair is not conserved nor conserved to Watson-Crick base pair among Tetrahymenine species (Fig. 1B). Since A131–U139 forms a stable base pair in the T. thermophila loop IV structure (Fig. 4), we asked whether disruption of this base pair affected TER function. We made the mutations A131C and U139G as well as the compensatory double mutation and tested the activity of each with TERT in the presence or absence of p65. In reactions with TERT alone, both A131C and U139G greatly decreased the overall level of activity, but activity was rescued to wild-type levels with the compensatory pair of mutations (Fig. 5A). This result is consistent with a requirement for a helix below the loop, presumably to provide a stable stacking platform for formation of the C132–U138 base pair in the structured loop. An indirect role for this base pair is consistent with activity assay results in reactions containing p65, which rescued the decrease in activity seen for the single mutations (Fig. 5B). p65 could stabilize the mismatches in the context of overall stabilization of the distal helix or by increasing TERT binding affinity. Of the five Tetrahymenine species that do not have a closing A–U pair, two have a C–G and three have an A–G, and all five of them have an extra loop nucleotide, C or A respectively. In these species, binding of p65 or TERT to TER may compensate for a decrease in distal helix stability. Analysis of the sequences in the distal stem of all 17 Tetrahymenine species shows that if one constrains the two nucleotides below the loop to pair (as Watson-Crick A–U, C–G, or mismatch A–G), all could form a helix of eight base pairs plus one bulge or nine base pairs including a maximum of one mismatch (U–U, G–U, or the terminal A–G). Thus, in the presence of p65 and TERT, loop IV would be at fixed distance and orientation relative to the GA bulge.

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DISCUSSION

Stem–loop IV plays a critical role in Tetrahymena TER folding, telomerase RNP assembly, and catalytic activity. These roles are mediated in part by direct interactions of stem–loop IV with p65 and TERT. Here we have shown that in the free RNA, the GA bulge forms a defined bend between the distal and proximal stems, and that wild-type enzymatic activity in the context of p65, TER, and TERT does not require any other bulged nucleotides in these helices. We have further shown that the highly conserved loop IV forms a structured loop at the end of a distal helix of defined length. Thus, binding of p65 at the GA bulge would place the terminal loop in a specific orientation for assembly with TERT and possible tertiary contacts with TER.

Does telomerase RNP assembly and activity rely on a preformed central bend in an otherwise helical stem IV?

To address the structural role of the conserved GA bulge in the center of stem IV, we determined the relative orientation of helices flanking the GA bulge with flanking C–G base pairs. It has previously been proposed based on chemical and enzymatic probing that the GA bulge in free TER induces a kink that might be a site for protein binding (Bhattacharyya and Blackburn 1994). We find that the GA bulge induces an interhelical angle of ∼50° with stacking of the two bases into the helix. Both the position of the two bases (stacked into the helix or out) and their stacking interactions would depend on the flanking base pairs, which provides a likely explanation for the conservation of the C–G base pairs on either side of the GA bulge.

The A–U rich proximal and distal stem IV helices in T. thermophila TER both contain a single bulged U nucleotide, and would not be expected to be very stable in the free RNA. Both the TERΔU117 and TERΔU127 mutants had activity comparable to wild type in the presence of p65, suggesting that helix irregularity or flexibility other than the GA bulge is not important for catalytic function in the physiological context of the p65–TER–TERT RNP. These activity assay results support our structural model for central stem IV. Chemical and enzymatic probing of stem IV in the absence of TERT are consistent with the presence of a continuous helix in this domain including the bulged U nucleotides (Bhattacharyya and Blackburn 1994; Sperger and Cech 2001). In contrast to results obtained with deletion of either bulged U nucleotide, deletion of the GA bulge nucleotides decreases activity for both the TERT–TER and p65–TER–TERT complexes, particularly when RNP assembly is limiting (Fig. 5; Autexier and Greider 1998; Sperger and Cech 2001; O'Connor et al. 2005; Prathapam et al. 2005).

Studies with T. thermophila TER and purified p65 have demonstrated that stem IV is a key determinant of p65 binding and p65 mediated stimulation of TERT–TER interaction. The bulged U117 is within the core of the p65 binding site (illustrated in Fig. 6; O'Connor and Collins 2006). Deletion of this nucleotide has no impact on p65 binding or assembly of the p65–TER–TERT RNP (C.M. O'Connor and K. Collins, unpubl.). In contrast, deletion of the GA bulge does affect binding of p65, and specifically the p65 C-terminal domain, to TER (O'Connor and Collins 2006). The GA bulge interaction with p65 is also crucial for establishing the hierarchical telomerase RNP assembly pathway (Prathapam et al. 2005; O'Connor and Collins 2006). These findings, the activity assay data described above, and our structural studies mutually reinforce the significance of a bulge in otherwise helical elements of stem IV for telomerase function. However, if stem–loop IV is provided in trans from the template, neither the central stem IV helical elements nor the conserved GA bulge is required for wild-type catalytic activity with TER and TERT in vitro (Lai et al. 2003; Mason et al. 2003). Combined, all of the data indicate a structural role for central stem IV in hierarchical telomerase RNP assembly via p65 binding and in positioning loop IV.

FIGURE 6.
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FIGURE 6.

Secondary structure model of T. thermophila TER with locations of p65 (O'Connor and Collins 2006) and TERT (Lai et al. 2001; O'Connor et al. 2005; Jacobs et al. 2006) domain binding sites. This model is meant as a visualization tool, and no attempt has been made to illustrate an interaction between loop IV and another region of the TER or to define its relative location.


Bulge sequence as well as length should be important for the semi-rigid well-defined bend we report here for TER central stem IV. Replacement of GA with UU would be expected to result in a different degree and/or direction of bend. Consistent with this, replacement of GA with UU decreases binding of p65 (O'Connor and Collins 2006). The decrease in p65 binding could be due to disruption of sequence-specific as well as structure-specific recognition by p65. Why then can TER GA → UU and GA → UC mutations result in wild-type activity for in vitro reconstituted TERT–TER (Autexier and Greider 1998; Sperger and Cech 2001; O'Connor et al. 2005)? We propose that in the absence of p65, the positioning of distal stem–loop IV needs to be flexible in order for loop IV to be able to transiently adopt a conformation relative to the rest of TER that is suitable for assembly and activity with TERT (discussed below). A requirement for flexibility in stem IV in assembly reactions with TERT and TER alone may account for the lower overall activity of the distal stem IV U127 deletion in the absence of p65. Deletion of U127 would decrease the flexibility of the stem and/or change its overall orientation. Binding of p65 would be expected to favor the correct orientation of loop IV for assembly of TERT with TER, thereby rescuing the activity defect.

Does the structured loop form a binding site for TERT and/or another motif of TER?

The helical region of stem IV and p65 are important for correct positioning of loop IV, focusing additional interest on the role of loop IV itself in telomerase activity. In the presence of TERT, loop IV mutations prevent a conformational change in TER detected by enzymatic footprinting in the region of the pseudoknot (Sperger and Cech 2001). We found that loop IV is highly structured in free TER and contains a conserved noncanonical C132–U138 base pair above the helical stem. Furthermore, we show that formation of this stem via a closing base pair below the C–U pair is stimulatory for catalytic activity. The significance of the remainder of the loop structure, when analyzed in light of the extensive biochemical data, is not immediately apparent. Single and double nucleotide substitutions of loop residues have revealed only three loop nucleotides that are crucial for TERT–TER activity, which are C132, U137, and U138 (Sperger and Cech 2001; O'Connor et al. 2005). U137 is the most conformationally flexible nucleotide in the loop in the free RNA, while C132 and U138 form the base pair that sets up the overall loop fold. RNase VI footprinting of TER in the presence of TERT showed that the single nucleotide changes that affected TERT–TER activity in vitro also imposed an RNA folding defect, monitored as a change in the cleavage pattern in the pseudoknot region, while the nucleotide changes that did not have a significant effect on activity (A133, C134, U135, and A136) also showed wild-type cleavage pattern (Sperger and Cech 2001). Thus, mutations at only C132, U137, and U138 appear to disrupt essential interactions of loop IV and other TER motif(s) and/or loop IV and TERT. It is important to note that the structure of loop IV could change in the context of its long-range tertiary interactions or TERT binding.

TERT contains two identified RNA interacting domains, the TEN domain, which has low affinity interactions with TER dependent on loop IV and the 3′ flanking TRE region, and a high-affinity RNA binding domain (RBD) that interacts with the TBE and possibly stem I (Fig. 6; O'Connor et al. 2005). The p65 enhancement of TERT assembly with TER can be recapitulated using only TER, the C-terminal domain of p65, and TERT RBD. However, this effect is abolished by loop IV deletion or loop mutations that disrupt activity (O'Connor and Collins 2006; unpublished data). These results combined with the structure determined here suggest that loop IV is involved in a conformational change in TER via a long-range RNA interaction that is stabilized by p65 binding to central stem IV including the region kinked by the GA bulge. The most likely nucleotides supporting this long-range interaction are C132, U137, and U138, which are also essential for activity. This loop IV–TER interaction may also form part of the TER interaction site for TERT, accounting for both the loss of activity and disruption of TERT binding with TER (U137A and U138A) mutants (Sperger and Cech 2001; Mason et al. 2003; O'Connor et al. 2005). Other loop nucleotides may be involved only in TERT binding rather than local loop folding or long-range TER interactions, and therefore be less crucial to in vitro activity in the context of single substitutions. Analysis of the crystal structure of the TEN domain along with cross-linking and mutagenesis revealed potential binding sites for TER and single-stranded telomeric DNA (Jacobs et al. 2006). A speculative extension of our model consistent with the accumulated data is that loop IV contributes to positioning the template and/or other elements of the active site. Future studies of the interaction of loop IV with TER and TERT will be necessary to test this model.

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MATERIALS AND METHODS

RNA synthesis and purification

Unlabeled and 13C,15N fully labeled RNA oligonucleotides GABIV and SLIV were synthesized using in vitro transcription with T7 RNA polymerase, synthetic DNA oligonucleotides, and purified as described previously (Dieckmann and Feigon 1997). Unlabeled nucleoside triphosphates (NTPs) were purchased from Pharmacia, and individual 13C, 15N-labeled NTPs were purchased from Silantes. Purified RNA was annealed at dilute concentration (1–10 μM) in water and adjusted to desired salt conditions by the addition of the appropriate buffer (10 mM potassium phosphate pH 6.5, 100 mM KCl, 50 μM EDTA, and 0.02% NaN3 for GABIV and 10 mM sodium phosphate, pH 5.8, 20 mM KCl, 50 μM EDTA and 0.2% NaN3 for SLIV). GABIV was subsequently concentrated by ultrafiltration to ∼1 mM, while the SLIV concentration (∼0.6 mM) and salt concentration were kept lower to prevent dimer formation.

NMR spectroscopy

NMR spectra were collected on Bruker DRX 500, 600, and 800 MHz spectrometers. For SLIV, exchangeable proton spectra were measured in 95% H2O/5% D2O at 283 K, and nonexchangeable proton spectra were measured in D2O at 293 K. Essentially complete assignments were obtained for SLIV. Exchangeable protons and nitrogens in the Watson-Crick base pairs were assigned using 2D NOESY, 1H-15N HMQC, and 15N-correlated CPMG-NOESY spectra (Cromsigt et al. 2001). Hydrogen bonding patterns were confirmed for the Watson-Crick base pairs using JNN-HNN COSY (Luy and Marino 2000). Nonexchangeable protons and carbons were assigned using 2D NOESY, homonuclear TOCSY, 1H-13C CT-HSQC, 2D HCCH-COSY, 3D HCCH-TOCSY, and 3D HMQC-NOESY (Cromsigt et al. 2001). 3JH3′P and 3JCP were measured using 31P spin echo difference CT-HSQCs to determine the β and ε torsion angles (Legault et al. 1995; Kolk et al. 1998). 1H-13C heteronuclear NOE experiments were collected to qualitatively investigate the flexibility of the loop. Most of the nucleotides in the loop have comparable heteronuclear NOE difference values of 1.0–1.2, and A136 and U137 have only slightly higher values of ∼1.3. RDCs were measured as the difference in 1JCH in the presence and absence of 3% C12E5/hexanol (Rückert and Otting 2000) using modified 2D 1H-13C HSQC without 13C decoupling in the direct 1H-dimension on an unlabeled sample.

For GABIV, assignment of H6/8, H5, H2, H1′ protons and attached carbons was achieved using a series of two-dimensional experiments including 2D NOESY, homonuclear TOCSY, and 2D 1H-13C HSQC acquired at 298 K in D2O on unlabeled RNA. RDCs were calculated as the difference between 1JCH measured in the absence and presence C12E6/hexanol. The random uncertainty (σ) in 1DCH, estimated from multiple measurements was, on average, <1.5 Hz. All spectra for both constructs were processed using Bruker XWINNMR 2.6 and analyzed using SPARKY software (University of California–San Francisco).

Modeling of the GA bulge structure using order matrix analysis of RDCs

The rigid-body modeling procedure using RDCs (Losonczi et al. 1999; Dosset et al. 2001; Al-Hashimi et al. 2002) as implemented in the MODULE program (Dosset et al. 2001) was used for determination of relative orientation of the domain D and domain P in GABIV (Fig. 3). Interdomain motion was assessed using fragment specific order parameters (υ, η) (Al-Hashimi et al. 2002) computed using REDCAT (Valafar and Prestegard 2003). An input model of the stem IV was built from idealized A-form RNA geometries generated using Insight II (MSI) and the geometry of the UUCG tetraloop (PDB ID: 1HLX). To account for a possibility of small local fluctuations and to accommodate small departures from the assumed local geometry, the input uncertainties of the RDC data were increased by 1.5 Hz (Al-Hashimi et al. 2002). The interhelical angle was calculated using 3DNA (Lu and Olson 2003).

The three-dimensional model for the GA bulge was constructed using the standard simulated annealing protocol (Varani et al. 1996) within NIH-XPLOR (Schwieters et al. 2003). A starting fold was generated from a randomized extended template. Artificial constraints corresponding to idealized A-form of RNA were imposed on domain D and P and constraints for the UUCG tetraloop were extracted from the solution structure of the P1 helix from group I self-splicing introns (PDB ID: 1HLX) (Allain and Varani 1995). The single-stranded nucleotides corresponding to the GA bulge were modeled into the helix based on interresidual H6/8(n)–H6/8(n + 1) and H1'(n)–H6/8(n +1) NOEs. In the next step, the starting fold was refined against RDCs.

Structure calculation for loop IV

Interproton distances from 2D NOESY and 3D NOESY-HMQC spectra were generated as described, except for the classification of semi-quantitative NOEs, which were as follows: strong (1.8–3.5 Å), medium (2.5–4.5 Å), weak (3.5–5.5 Å), and very weak (4.5–6.5 Å). There were a total of 245 intranucleotide and 207 internucleotide nonredundant distance restraints measured. We used 115 dihedral angle constraints for α, β, γ, δ, ε, and ζ backbone angles. The restraints for β, δ, and ε were determined experimentally as described above. The α, γ, δ, and ζ dihedral angles for the nucleotides involved in base pairs in the helix were constrained to the A-form value. Nucleotides with observable H1′–H2′ and H1′–H3′ correlations in a 50-msec mixing time TOCSY were constrained as C2′-endo (South; 145 ± 30°), nucleotides with observable H1′–H2′ correlations only were constrained as intermediate (120 ± 30°), while all other nucleotides which had no detectable H1′–H2′ correlation were constrained as C3′-endo (North; 82 ± 30°). Intranucleotide H1′ to aromatic NOEs from a 50-msec 2D NOESY indicated that all nucleotides were anti and could thus be constrained to a χ value of −160 ± 30°, except for A136. The intranucleotide H1′ to aromatic NOE for A136 was much more intense than any other residues. Its χ angle was left unconstrained, but structure calculations produced an angle in the high anti range. Initial structure calculations included hydrogen bonding distance restraints and weak planarity restraints for the Watson-Crick base pairs only (force constant of 1 kcal/Å2). An additional hydrogen bond was included in later structure calculations for the noncanonical C–U base pair.

Starting from an extended, unfolded RNA conformation the structures were calculated using XPLOR-NIH 2.9.8 (Brunger 1992; Schwieters et al. 2003) using NOE-derived distances and dihedral angle restraints. The folding and refinement stages followed standard XPLOR protocols. Structures with no experimental restraint violations from the initial 200 calculated structures were then subjected to refinement against 17 RDCs. The protocol involved slow cooling from 1000 K to 100 K in 18 cycles of molecular dynamics corresponding to a total of 18 psec. During this stage, the force constant for RDCs was slowly increased from 0.001 to 0.2 kcal mol−1Hz−2. The experimentally derived RDCs together with the lowest energy structure from the structure calculation employing NOE and dihedral restraints were used as input information. The grid search for the values of the magnitude and asymmetry of the alignment tensors produced optimal values of Da = −36.0 Hz and R = 0.6, respectively. The force constants used in the final stage of structure calculations were 50 kcal mol−1Å−2, 200 kcal mol−1deg−2, and 0.2 kcal mol−1Hz−2 for NOEs, dihedral angles, and RDCs, respectively. Structural statistics for the 20 lowest energy structures for SLIV are presented in Table 1. Structures were viewed and analyzed using MOLMOL (Koradi et al. 1996) and Insight II (MSI).

Telomerase activity assays

TERT and p65 were expressed in separate RRL synthesis reactions. TERT with or without p65 was assembled with TER with an ∼10-fold excess of TER (25 ng) for 20 min at 30°C. Telomerase RNA was then diluted 10-fold with buffer containing 20 mM Tris-HCl, pH 8.0, 1 mM MgCl2, 10% glycerol, and 5 mM DTT to constitute half of the activity assay reaction volume. Reactions contained 4 μM cold dGTP, 0.6 μM radiolabeled dGTP, 0.4 mM dTTP, and 1 μM (TG)8T2G3. Reactions were incubated 1 h at 30°C, phenol extracted, ethanol precipitated, and resuspended in TE and formamide. Products were resolved on a denaturing 12% acrylamide gel.

Coordinate deposition

Coordinates for the 20 lowest energy structures of loop IV have been deposited in the Protein Data Bank (PDB ID 2H2X). Coordinates for the model structure for the GA bulge are available upon request.

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ACKNOWLEDGMENTS

This work was supported by NSF Grant MCB051770 to J.F. and NIH GM54198 to K.C. We thank Mr. Evan Feinstein for help with figure preparation.

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Footnotes

  • ↵3 Present address: Department of Molecular Biology and Biochemistry, University of South Bohemia, Branisovska 31, Ceske Budejovice, 370 05, Czech Republic.

  • Reprint requests to: Juli Feigon, Department of Chemistry and Biochemistry, and Molecular Biology Institute, University of California, Los Angeles, CA 90095-1569, USA; e-mail: feigon{at}mbi.ucla.edu; fax: (310) 825-0982.

  • Article published online ahead of print. Article and publication date are at http://www.rnajournal.org/cgi/doi/10.1261/rna.112306.

    • Received April 10, 2006.
    • Accepted May 18, 2006.
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