ABSTRACT
Telomerase maintains telomere length by reverse transcribing short G-rich DNA repeat sequences from its internal RNA template. G-rich telomere DNA repeats readily fold into G-quadruplex (GQ) structures in vitro, and the presence of GQ-prone sequences throughout the genome introduces challenges to replication in vivo. Using a combination of ensemble and single-molecule telomerase assays we discovered that GQ folding of the nascent DNA product during processive addition of multiple telomere repeats modulates the kinetics of telomerase catalysis and dissociation. Telomerase reactions performed with telomere DNA primers of varying sequence or using K+ versus Li+ salts yield changes in DNA product profiles consistent with formation of GQ structure within the telomerase-DNA complex. Single-molecule FRET experiments reveal complex DNA structural dynamics during real-time catalysis, supporting the notion of nascent product folding within the active telomerase complex. To explain the observed distributions of telomere products, we fit telomerase time series data to a global kinetic model that converges to a unique set of rate constants describing each successive telomere repeat addition cycle. Our results highlight the potential influence of the intrinsic folding properties of telomere DNA during telomerase catalysis and provide a detailed characterization of GQ modulation of polymerase function.
SIGNIFICANCE Telomeres protect the ends of linear chromosomes from illicit DNA processing events that can threaten genome stability. Telomere structure is built upon repetitive G-rich DNA repeat sequences that have the ability to fold into stable secondary structures called G-quadruplexes (GQs). In rapidly dividing cells, including the majority of human cancers, telomeres are maintained by the specialized telomerase enzyme. Thus, telomerase and its telomere DNA substrates represent important targets for developing novel cancer drugs. In this work, we provide evidence for GQ folding within the newly synthesized DNA product of an actively extending telomerase enzyme. Our results highlight the delicate interplay between the structural properties of telomere DNA and telomerase function.
INTRODUCTION
Telomeres safeguard the ends of chromosomes from illicit DNA processing events that would otherwise threaten genome stability (1, 2). The foundation of telomere structure consists of short G-rich DNA sequence repeats. The majority of telomere DNA is double-stranded and can be up to several kilobases in length, while the ends are processed to terminate with a short 3’ single-stranded G-rich tail (∼50-150 nucleotides in vertebrates) (3, 4). Repetitive G-rich DNA sequences are not unique to telomeres and are found throughout the human genome (5). These G-rich repeats have the capacity to fold into G-quadruplexes (GQs), structures composed of multiple Hoogsteen bonded G-quartet motifs that stack together to yield stable DNA folds (6, 7). GQ folding has been implicated in a variety of biological processes. For example, replication of GQ-prone sequences is problematic and requires contributions from specific DNA helicase enzymes to avoid replication-coupled DNA damage (8–10). Sequences with GQ-folding potential are enriched within promoter sequences of oncogenes where they are thought to regulate gene expression (11). Finally, recent evidence suggests GQ folds can form in vivo in a spatially and temporally regulated manner (12–14). Thus, small molecules that bind and stabilize GQ-folds hold promise as novel cancer drugs; a fact that motivates better understanding of how GQ structure can modulate enzyme function.
Telomerase is an RNA-dependent DNA polymerase that is uniquely adapted to synthesizing G-rich repetitive DNA sequences (15, 16). Telomerase activity combats gradual telomere shortening that occurs with each round of cellular division (17). Left unchecked, telomere shortening induces senescence or cell death in somatic tissues. In contrast, proliferative cells such as stem cells rely upon telomerase activity to maintain telomeres in order to support continued rounds of cell division (15). Genetically inherited loss of function mutations in telomerase subunits cause human disorders characterized by deterioration of proliferative tissue types (18–21). On the other hand, telomerase overexpression contributes to the immortal phenotype of ∼90% of human cancers, and is therefore an important target for development of novel cancer therapies (22).
Telomerase is a ribonucleoprotein (RNP) complex that includes the long non-coding telomerase RNA (TR) and the catalytic telomerase reverse transcriptase (TERT) protein subunit (23, 24). In vertebrates, the 3’ end of TR includes canonical H/ACA box RNA motifs and associated proteins that promote TR stability and efficient telomerase RNP biogenesis (25). To initiate telomerase catalysis, the 3’ ssDNA telomeric tail base pairs with the TR template, forming a short RNA-DNA hybrid that is extended in the TERT active site (Fig. 1A). TERT utilizes a limited region of TR to direct synthesis of a defined GGTTAG hexameric telomere DNA repeat sequence (kpol) (Fig 1A). A unique property of telomerase is the ability to translocate on the DNA product (ktrans) in order to recycle the integral TR template during processive addition of multiple telomere repeats prior to dissociation from the DNA product (koff) (Fig 1A)(26). This repeat addition processivity (RAP) implicitly requires multiple points of contact between telomerase and its DNA substrate, a notion that is consistent with data from a variety of telomerase systems identifying ‘anchor site’ DNA interactions outside the enzyme active site (27–30).
Model telomere DNA substrates harboring integer multiples of four consecutive telomere repeats are inefficient binding substrates for telomerase in vitro while DNA primers with five, six, or seven consecutive repeats are efficiently bound and extended (31). Thus, while GQ structures can inhibit telomerase association, the presence of a small single-stranded DNA overhang in the substrate is sufficient to recover telomerase loading and function. These previous studies illuminated DNA sequence determinants that mediate the initial binding of telomerase to its substrate; however there remained an untested possibility that GQ structure may regulate the behavior of an actively extending telomerase-DNA complex, as suggested by studies of ciliate telomerase (32, 33). Furthermore, we reasoned that the specialized telomerase system provides a powerful opportunity to investigate the influence of GQ-forming sequences on nucleic acid polymerase function.
To study the relationship between DNA structure and human telomerase catalysis we performed direct primer extension assays using dNTP concentrations similar to those found in the cellular environment (34). Our experiments reveal a complex pattern of telomerase DNA product accumulation that indicates the efficiency of template recycling is dependent upon the number of synthesized repeats. Experiments using telomere DNA primers of varying sequence and varying salt conditions support the notion that a GQ can form within the telomerase-DNA complex. Single-molecule FRET experiments provide further support for DNA structural dynamics within actively extending telomerase enzymes. To estimate individual rate constants for successive repeat addition cycles we performed global kinetic modeling of telomerase time-series data. Interestingly, our model converges to a unique solution of rate constants that provides a direct measure of processivities for each cycle of telomere repeat addition. These results are consistent telomere DNA GQ folding serving to promote template recycling, as well as to accelerate product dissociation. We present a mechanistic model that provides a framework to understand the delicate interplay of telomere DNA structure and function during telomerase catalysis.
RESULTS
Telomerase product distribution is sensitive to dNTP concentrations and stoichiometry
When measuring telomerase activity in vitro, it is common to employ direct primer extension assays in the presence of α32P-dGTP. This approach permits reactions to be performed with a large excess of unlabeled DNA substrate, benefits from very high sensitivity of product detection, and circumvents PCR-induced artifacts inherent to the telomere repeat amplification protocol (TRAP) assay. However, the use of α32P-dGTP incorporation to detect product accumulation limits the amount of total dGTP that can be used in the assay, leading to the widely reported practice of using non-physiological dNTP stoichiometry that has the potential to significantly alter the telomerase product distribution (35, 36). To circumvent this problem, we used 5’ radiolabeled DNA primers and cold dNTPs to monitor telomerase activity (Fig. 1B). We compared this approach to standard assays performed with α32P-dGTP using identical dNTP and primer concentrations (Fig. S1). Although the reaction profiles are qualitatively distinct, we observed quantitatively similar product distributions for the two approaches when normalized for the amount of α32P-dGTP incorporation. Importantly, the majority of the input DNA primers are not extended in our experiments, demonstrating our reaction conditions are sufficient to limit distributive telomerase activity (i.e. an individual primer being extended by multiple telomerase enzymes). Processive telomerase activity is also evident when analyzing pulse-chase experiments in which longer DNA products continue to accumulate after addition of a 400-fold excess of cold DNA primer (Fig. S2).
Having established that our end-labeled DNA primer assay is capable of accurately monitoring processive telomerase action, we next sought to analyze the influence of varying dNTP concentrations on the telomerase product distribution (Fig. 1B). Previously published studies defined telomerase processivity as the number of repeats corresponding to the point where the dissociated DNA represents 50% of the total population (R1/2) (37, 38). This approach is analogous to half-time (t1/2) analysis and can be performed by fitting a linear regression to a plot of ln(1-FLB) versus repeat number, where FLB is the fraction left behind (Fig. 1C inset, see Methods for details). Titrating increasing amounts of dGTP in the presence of a large excess of dATP and dTTP yields a significant boost in RAP as has been reported previously for both human and Tetrahymena telomerase (Fig. 1B and 1C) (35, 39–43). However, the use of a large excess of dATP and dTTP is not a good approximation for the physiological dNTP pool which is generally closer to the ∼10 uM range (34). Note that dCTP is not required for the human telomere sequence and its absence does not alter telomerase function (data not shown). When assayed in the presence of equimolar dGTP, dATP, and dTTP, we observe the highest RAP activity of all conditions tested (Fig. 1B, lane 4 and Fig 1C). Moreover, using this approach, we noticed that the previously observed doublet band at the +1 position after each complete telomere repeat is effectively suppressed (Fig. 1B, compare lanes 1-3 with lane 4). We interpret this +1 band to reflect promiscuous use of the non-template RNA base at hTR position U45, which may be enhanced by the large excess of dATP. Based upon these results, we elected to perform all subsequent telomerase assays in our study under conditions of equimolar dNTPs that better reflect cellular dNTP concentrations.
G-quadruplex folding alters the pattern of telomerase product accumulation
Established methods for approximating RAP using the R1/2 value described above assume an exponential decay in the distribution of accumulated telomerase product lengths with each telomere repeat added (37, 38, 44). However, we noted the appearance of plateaus in the product distribution when using equimolar concentrations of dNTPs (Fig. 1C and 2A). For example, when using a standard telomere DNA primer composed of the sequence (TTAGGG)3, we observe a sudden drop in product accumulation between the bands corresponding to the third and fourth telomere repeats added to the primer (Fig. 2A, red asterisk). Surprisingly, the intensities of the subsequent four added repeats are approximately equal, until a second decrease in accumulation occurs between added repeats seven and eight. This pattern of four equally populated product lengths, followed by a decrease in accumulation, continues throughout the detectable range of telomere DNA products.
Telomere DNA primers with at least four contiguous G-rich repeats can fold into a GQ in vitro (45, 46), suggesting the RAP-associated ‘pattern of four’ we observe in our experiments may be due to GQ folding of the DNA product within an actively extending telomerase complex. To test this hypothesis, we altered the 5’ end of the telomere DNA sequence so that it no longer harbored the requisite run of guanines needed to participate in GQ folding (Fig. 2A, lanes 2 and 3). Altering the primer in this way should change the product length where the ‘pattern of four’ appears if the newly synthesized DNA folds into a GQ. Indeed, a modified DNA primer with a 5’ (TG)3 substitution supports telomerase RAP, but the plateaus in the product profile are delayed by one additional repeat, corresponding to the sequence needed to promote GQ formation in the product DNA (Fig. 2A, compare lanes 1 and 2). Similarly, when the first two repeats in the telomere DNA primer were substituted with the TG dinucleotide sequence ((TG)6), the plateaus are delayed by two additional repeats (Fig. 2A, compare lanes 1 and 3). These results were highly reproducible across three independent experimental trials (Fig. 2B) and agree with the hypothesis that GQ folding within the nascent telomere DNA causes the telomerase product profile to deviate from a strictly decreasing decay.
GQ stabilization alters telomerase kinetics and DNA product structural dynamics
The H-bonding configuration of the G-quartet motifs within a GQ fold can be differentially stabilized by coordination of specific monovalent cations, with a rank order of K+ > Na+ > Li+ in terms of degree of stabilization (47). Therefore, we next set out to further dissect the contribution of GQ folding to telomerase activity by altering the salt conditions of our telomerase assays. We observed robust telomerase activity in all salt conditions tested; however, there was a clear reduction in total product accumulation in Li+ when compared to Na+ and K+ (Fig. 3A). The lower total product accumulation in the presence of Li+ is a consequence of slower DNA synthesis kinetics (Fig. S3) and is not completely unexpected based on reported effects of Li+ on DNA polymerases (48). Interestingly, we do not observe the robust “pattern of four” RAP product distribution in the presence of Li+ (Fig. 3A and 3B), the salt condition expected to least stabilize GQ folding during telomerase catalysis. This result lends additional support to the hypothesis that GQ formation in the presence of K+ or Na+ impacts the product distribution of an actively extending telomerase complex.
The results of our ensemble telomerase assays suggest that folding of the nascent DNA product can influence telomerase catalysis. However, this approach does not directly interrogate DNA conformation within an active RNP complex. Therefore, we turned to a single molecule Förster Resonance Energy Transfer (smFRET) based approach that directly monitors DNA structure and dynamics within individual telomerase enzymes (49, 50). In order to ensure our smFRET assay supports telomerase activity in both K+ and Li+ buffers we employed a recently reported method for in situ detection of extended DNA products at the single-molecule level (51). Telomerase RNP complexes harboring a Cy3 dye incorporated into the telomerase RNA subunit were bound to a biotinylated DNA primer and then surface-immobilized onto a streptavidin coated glass slide (Fig. 4A). The telomerase-DNA complexes were incubated in either K+ or Li+ activity buffer as well as with a Cy5-labeled detection oligonucleotide with a sequence that is complementary to the telomere product. In this way, telomere primers that are being actively extended by telomerase (i.e. enzyme is still bound and some of the DNA product becomes accessible) are detected as a FRET signal between the Cy3-labeled enzyme and the Cy5-labeled DNA probe. The appearance of the FRET signal was strictly dependent upon addition of activity buffer containing dNTPs and was time dependent (Fig. 4B). After 20 minutes of incubation, comparable levels of telomerase activity, measured as the total number of telomerase-DNA complexes producing a positive FRET signal, were detected in both K+ and Li+ activity buffers (Fig. 4B). Taken together, these results demonstrate that telomerase is catalytically active in both K+ and Li+ buffers in the single-molecule assay.
Having detected robust telomerase activity in both K+ and Li+ buffers, we next performed smFRET assays with Cy3-labeled telomerase and Cy5-labeled telomere primer in order to directly monitor DNA dynamics during active telomere elongation. (Fig. 4C). Stalled telomerase-DNA complexes harboring these site-specific dye modifications were surface-immobilized as described above. Data collected on stalled complexes prepared in this manner yielded a unimodal FRET distribution centered at ∼0.75 (Fig. 4D, top left panel). Next, the telomerase complexes were activated for DNA synthesis by introducing dNTPs in telomerase activity buffer (containing 50mM KCl), resulting in a significant shift towards lower FRET values and ultimately yielding a single population centered at near-zero FRET (Fig. 4D, left panels). This FRET change upon activation of DNA synthesis is consistent with the Cy5-label site on the telomere DNA moving further away from the telomerase active site, and perhaps folding into a stable structure, during multiple rounds of DNA repeat synthesis. Analysis of real-time single-molecule FRET trajectories collected on actively extending telomerase complexes in this same condition reveals complex DNA conformational dynamics characterized by a general transition to discrete lower FRET states combined with the occurrence of transient increases in FRET (Fig. 4E and Fig. S4).
In order to dissect the contribution of DNA structure to the observed FRET dynamics we performed the same experiment using LiCl instead of KCl in the telomerase activity buffer. Once again, the initial FRET population of stalled telomerase-DNA complexes was centered at ∼0.75 in the presence of Li+ (Fig. 4D, top right panel). However, in contrast to the K+ activity buffer, we did not observe a substantial drop in FRET upon addition of dNTPs (Fig. 4C right panels). Instead we observed a modest shift in the major FRET population to ∼0.6 after 60 minutes incubation time. This result is consistent with individual smFRET trajectories of telomerase-DNA complexes incubated in Li+ activity buffer, where the FRET level appears to remain constant over the course of several minutes (Fig. 4E). Together, these results argue that the formation of a GQ in the nascent telomere product contributes to efficient extrusion of the telomere DNA away from the telomerase active site during the course of activity.
Global kinetic modeling provides a measure of telomerase microscopic processivity
Telomerase processivity can be modeled as a series of consecutive reactions in which nucleotide addition is in competition with DNA dissociation at each step of the reaction. To simplify our telomerase kinetics analysis, we focus on the intense repeat addition bands, assuming the intervening nucleotide addition steps are rapid and accompanied by little DNA dissociation. The macroscopic processivity of telomerase can be conveniently described by the median product length (R1/2= ln2/decay constant) (see Fig. 1C) (37). However, treating the data in this manner has several limitations. First, in a typical primer extension experiment one cannot distinguish between dissociated product and DNA that remained associated with enzyme at the point when the reaction was arrested. In addition, this analysis approach has the effect of suppressing possible deviations in microscopic processivity, which reflects the individual probability of adding another DNA repeat at a specific step of telomerase catalysis. The experiments described in the present study provide clear evidence that telomerase products do not accumulate uniformly and display patterns dependent upon assay conditions, DNA sequence, and/or product length. We therefore developed a kinetic model that can be utilized to globally fit telomerase time-series data in order to extract microscopic processivity values that underlie the observed distributions of telomerase products (Fig. 5A). Using this scheme, we treat this multistep process as a first-order reaction with an effective forward rate constant (kf) for the transition between each repeat and a dissociation rate constant (kd) (Fig. 5A) (note that kf in this model reflects a combination of kpol and ktrans described in Fig. 1A). Using the kinetic scheme depicted in Fig. 5A, we can then define the microscopic processivity (p) at each step as p = kf / (kf + kd).
DNA dissociation is an effectively irreversible process when a large excess of unreacted primer remains, which outcompetes the re-binding of any product DNA. To ensure our experiments complied with this assumption, we analyzed telomerase kinetics following a chase with 400-fold excess of unlabeled primer DNA, which serves to block re-association of the labeled DNA primer following telomerase dissociation (Fig. 5B). Telomerase time course assays were performed in the presence of either K+ or Li+ activity buffer conditions (Fig. 5B). Activity was initiated at time zero in the presence of end-labeled telomere DNA primer and dNTPs, followed by addition of excess chase primer at 20 minutes. The presence of 400-fold excess cold primer prior to enzyme addition was sufficient to eliminate any observable extension of the 50 nM endlabeled DNA primer used in our assays (Fig. 5B, lanes 1 and 9). Time points were collected at regular intervals out to 90 minutes and the concentration of each repeat species (B + B#, C + C#, etc, Fig 5A) was determined at each time point from the band intensity, knowing that the intensity of the initial primer was 50 nM. Individual rate constants were estimated by fitting the concentration time courses globally, using DynaFit (Fig. 5C, data shown for bands E-J). Global fitting converged to a unique set of rate constants that could be extracted with reasonable precision (see Supplementary Methods for details of kinetic modeling) (Fig. 5D). Comparison of the data obtained in the presence of K+ and Li+ revealed that the rate constants, kf and kd, decrease with increasing repeat number and result in lower microscopic processivity values for short DNA products as has been noted previously (37). However, in the presence of K+, the rate constants and microscopic processivity values show a saw-tooth modulation that gives rise to the ‘pattern of four’ clustering of products noted earlier (Fig. 5E, closed symbols). Although the effect is relatively small it is robust as determined by Monte-Carlo analysis and is reproducible between experiments and telomerase preparations (Fig. S5 and S6). Interestingly, the rate constants for repeat addition (kf) and dissociation (kd) were greater in the presence of K+ than in Li+, but the resultant microscopic processivity was lower (Fig. 5D and 5E). Taken together, these results suggest that in the presence of K+, the ability of the DNA product to form a GQ fold (which first arises at step 3, see Discussion for details) appears to promote translocation (increased kf) but at the increased risk of DNA product dissociation (increased kd).
DISCUSSION
The foundation of telomere structure consists of short G-rich repeat sequences, GGTTAG in humans, that have the propensity to fold into G-quadruplex (GQ) structures in vitro and in vivo (6, 7). Short ssDNA oligonucleotides that form GQs are poor telomerase substrates in vitro, due to the occlusion of an accessible single-stranded 3’ end (31, 46). Here, we present evidence that GQ formation can occur within an actively extending telomerase complex in vitro and that formation of such GQs affect the kinetic properties of telomerase. Telomerase activity assays using modified telomere DNA primer sequence and varying salt conditions together support the notion that GQ folding occurs during telomere DNA synthesis. Consistent with this concept, single-molecule FRET experiments directly demonstrate complex folding dynamics of the nascent DNA product in the presence of K+ but not Li+. We describe a detailed kinetic framework for telomerase catalysis and use this model to globally fit telomerase time-series data in order to extract microscopic processivity values for each cycle of telomere DNA repeat synthesis (see below for details). Our kinetic modeling reveals small but significant GQ-dependent changes in the rate constants describing the DNA repeat synthesis reaction (kf) and product dissociation (kd).
Telomerase repeat addition processivity (RAP) can be quantitatively described as the number of repeats corresponding to the point where the dissociated DNA represents 50% of the total population (R1/2) (37). The median number of repeats is a measure of the macroscopic processivity of telomerase that is implicitly assumed to be independent of product length. This method of analysis has been useful in previous studies that sought to characterize telomerase RAP under varying experimental conditions or with mutant telomerase enzymes (37, 52). A caveat of this approach is that typical telomerase primer extension assays cannot distinguish between products that have already dissociated and DNA that remained bound to enzyme when the reaction was stopped. This caveat can be addressed by the kinetic modeling approach described in the present study, or by introducing additional experimental steps to separate dissociated and bound products prior to gel analysis (43). However, one additional advantage of the global kinetic modeling approach is the ability to extract microscopic processivity values that reflect the individual probability of synthesizing each subsequent telomere repeat. Indeed, the results of our global kinetic modeling reveal a saw-tooth modulation of kf and kd, that together explain the ‘pattern of four’ clustering of telomerase product accumulation.
We present a working model for the mechanism of GQ-dependent effects on telomerase repeat addition processivity (Fig. 6). The complex rearrangements that are necessary for template recycling during multiple rounds of telomere repeat synthesis require multiple points of contact between telomerase and its DNA substrate. Minimally, the DNA must engage the enzyme by interacting with the RNA template in the active site and/or making distal contacts with a separate ‘anchor site’ (27–30). Upon completion of a telomere repeat, the 3’ end of the DNA product must dissociate from the template and dynamically realign with the downstream region of the RNA to prime the next round of repeat synthesis (Fig. 6) (50). This large-scale translocation step represents a vulnerable stage of telomerase catalysis, requiring a stable anchor-site interaction to prevent product dissociation. In principle, GQ folding within the DNA product can compete with the anchor-site contact to promote dissociation (Fig. 6, bottom pathway). Alternately, GQ folding can bias the positioning of the 3’ end of the primer to favor realignment for a subsequent round of repeat synthesis while allowing for anchor site contacts to remain intact (Fig. 6, top pathway). Which pathway the enzyme takes will depend on if anchor site interactions are maintained and the register of the 3’ end of the telomere during the formation of a GQ. In the schematic model depicted in Fig. 6, a GQ formed when the most recently synthesized telomere is bound in the active site will disrupt anchor site interactions and promote product dissociation, while a GQ formed when the 3’ telomere end is annealed to the template priming site, will facilitate another round of repeat addition. The latter outcome is mechanistically similar to DNA hairpin induced translocation models proposed for diverse telomerase systems (53, 54).
The finding of GQ-dependent effects on telomerase catalysis in vitro begs the question of whether a similar phenomenon occurs in the cell. While direct demonstration of GQ folding in vivo is challenging, recent experimental advances using GQ specific antibodies suggest that GQ folding can occur in a temporally and spatially regulated manner in vivo (12–14). Additionally, studies of ciliate telomerase have suggested that GQ folding may contribute to product dissociation during active telomere elongation (32, 33). The recently reported cryo-EM structures of the human and Tetrahymena telomerase enzymes provide a platform for investigating the potential for GQ folding within the actively extending telomerase complex (55, 56). Interestingly, when analyzing the EM density of the human telomerase complex, we noticed a structural pocket immediately proximal to the path of the nascent DNA emerging from the active site (Fig. S7). This pocket is flanked by two evolutionarily conserved features that are essential for telomerase processivity: the TEN domain and the TR p6.1 stem loop. The volume of this pocket is remarkably compatible with the dimensions of a single GQ fold, and raises the possibility that the nascent DNA products has ample space to assume a GQ structure within the confines of telomerase RNP complex. In the Tetrahymena structure, the C-terminal domain of the processivity factor, Teb1, is positioned as a lid to the DNA exit pocket, forming an enclosed cavity that is also sufficient to accommodate a single GQ fold. We propose the hypothesis that the Teb1-related shelterin component POT1 may occupy a similar position in the human telomerase complex. Thus, the anticipated binding of POT1 to the DNA as it is threaded out of this DNA exit channel is not mutually exclusive with the possibility of GQ folding within this protected cavity (46, 56). While speculative, the notion of GQ folding within the DNA exit channel is consistent with the amount of DNA protected by exonuclease cleavage as well as previous smFRET measurements that showed a similar amount of the DNA substrate is in close proximity to the template (50, 57). Future structural and biochemical experiments are required to investigate the influence of the shelterin proteins POT1-TPP1 on the folding properties of the nascent telomere DNA product, as well as the in vivo significance of GQ folding during telomerase catalysis.
MATERIALS AND METHODS
Preparation of RNAs
Synthetic, dye-labeled RNA fragments
Synthetic PK hTR fragment 32-62 was ordered from Dharmacon with an internal aminoallyl uridine (5-N-U) on position U42. A single tube of RNA was resuspended in nuclease-free H2O and ethanol precipitated in the presence of 300mM NaOAc pH 5.2 to prevent introduction of reactive amines. The pellet was resuspended in 100 ul 0.1M NaHCO3 and was used to solubilize a single mono-reactive Cy3-dye pack (Amersham). The solution was incubated for 2 h at 37°C. The solution was then ethanol precipitated, again with NaOAc. The RNA was then deprotected by resuspension in deprotection buffer (100mM acetic acid, pH 3.6) followed by vortexing and centrifugation for 10 seconds each. The solution was then heated at 60°C for 30 min followed by ethanol precipitation. The pellet was resuspended in 60 ul 0.1M TEAA, pH 7.5, and was HPLC purified on a reverse phase C18 column (Agilent technologies).
Ligation of RNA fragments
A splinted ligation reaction containing 800 pmol of Cy3-labeled hTR 32-62 fragment, 1600 pmol of in vitro transcribed unlabeled hTR 63-195, 1600 pmol DNA splint and 0.5x ligase buffer (NEB) was set up in a 200ul volume and incubated for 95C for 5 min followed by 30C for 10 min. 200ul ligation mix (1.5x ligase buffer, 8000 units of T4 DNA ligase (NEB), 2 mM ATP and 1U/ul RNasin Plus) was then added to the reaction mixture and incubated overnight at 30°C. 10 units of Turbo DNase was added and incubated for 15min at 37°C. The RNA was then phenol-chloroform extracted and ethanol precipitated prior to PAGE purification.
In vitro transcription
Unlabeled CR4/5 (hTR 239-328) and PK (32-195) was in vitro transcribed using homemade T7 polymerase in 1x RNA polymerase buffer (40mM Tris-HCl pH 7.9, 6mM MgCl2, 1mM DTT, 2mM spermidine) with 1.5 mM NTPs, 22 mM additional MgCl2, 90mM additional DTT and 40 units RNasin Plus. The reaction was incubated overnight at 37°C followed by the addition of 10 units of Turbo DNase for 15min at 37C. The RNA was phenol-chloroform extracted and ethanol precipitated prior to PAGE purification. PK 63-195 was in vitro transcribed as above but with 1mM of each NTP and an additional 5mM GMP in order to preferentially make a 5’ monophosphate fragment for use in splinted ligation.
Telomerase expression
Human telomerase was reconstituted using the Promega TnT Quick Coupled
Transcription/Translation system. In Lo-bind tubes (Eppendorf), 200ul of TnT quick mix was combined with 5ug of pNFLAG hTERT plasmid as well as 1uM of the in vitro transcribed PK and CR4/5 fragments except for when using Cy3-labeled PK, which was added at 0.1uM. The reaction was incubated for 3 h at 30°C. 5 ul of 0.5 M EDTA, pH 8.0, was added to quench excess Mg2+ present in the lysate.
Telomerase purification
Directly after reconstitution, telomerase was purified using the N-terminal FLAG tag on hTERT. To pull down the enzyme, Sigma Anti-FLAG M2-agarose resin was used. 50 ul bead slurry was first washed three times with wash buffer (50 mM Tris-HCl, pH 8.3, 3 mM MgCl2, 2 mM DTT, 100 mM NaCl) with 30 sec spins at 5000rpm at 4°C after each wash. The beads were then blocked in blocking buffer (50mM Tris-HCl pH 8.3, 3mM MgCl2, 2mM DTT, 500ug/ml BSA, 50ug/ml glycogen, 100ug/ml yeast tRNA) for 15 min under gentle agitation at 4°C followed by a 30 sec spin and removal of the supernatant. This blocking step was performed twice. After blocking, the beads were resuspended in 200ul fresh blocking buffer and added to the telomerase lysate. The beads and lysate were incubated for 2 h at 4°C under gentle agitation. The beads were then spun down for 30 sec at 5000 rpm and at 4°C and the supernatant was discarded. The beads were then washed three times in wash buffer containing 300mM NaCl (KCl or LiCl for salt dependence experiments) followed by three washes in wash buffer with 100 mM NaCl (KCl or LiCl for salt dependence experiments). A 30 sec spin at 5000 rpm at 4°C was performed between each wash. To elute the enzyme, the beads were incubated in 60 ul elution buffer (50 mM Tris-HCl, pH 8.3, 3 mM MgCl2, 2 mM DTT, 750 ug/ml 3x FLAG peptide, 20% glycerol) under gentle agitation at 4°C for 1 h. After elution, the beads were removed by centrifugation at 13000 rpm through Nanosep MF 0.45um filters. 5ul aliquots were then aliquoted into Lo-bind tubes and flash frozen in liquid nitrogen and stored at −70°C until use.
End-labeling of primers
50pmol of primer was labeled with alpha-32P ATP using T4 PNK in 1x PNK buffer (70 mM Tris-HCl, pH 7.6, 10 mM MgCl2, 5 mM DTT) in 50 ul. The reaction was incubated for 1h at 37°C followed by heat inactivation of T4 PNK at 65°C for 20min. Centrispin columns (Princeton separations) were used to obtain labeled primer.
Primer extension assays
Telomerase activity assays were performed by using 5 ul of purified telomerase in 1x activity buffer (50 mM Tris-HCl pH 8.3, 50 mM KCl (NaCl or LiCl when indicated), 1 mM MgCl2, 2 mM DTT) in a final volume of 15ul. For experiments containing end-labeled primer, each reaction contained 10 uM each of dATP, dTTP and dGTP as well as 50 nM of indicated 32P-primer. For reactions containing radiolabeled dGTP, each reaction contained the indicated dNTP concentrations as well as 50nM unlabeled (TTAGGG)3 primer. Reactions were incubated for 90 minutes at 30C. Reactions were stopped using 200ul 1x TES buffer (10 mM Tris-HCl, pH 7.5, 1 mM EDTA, 0.1% SDS), followed by phenol-chloroform extraction and ethanol precipitation of the DNA. The pellet was resuspended in 1x formamide gel loading buffer and resolved on a 12% denaturing PAGE gel. The gel was then dried and exposed on a phoshorimager screen and scanned using a Typhoon scanner. Band intensities were quantified using SAFA (58). R1/2 values were calculated by summing each band and all bands below it divided by the total counts for a given lane (to calculate the fraction left behind (flb). The natural log of (1- flb) was then plotted against repeat number and 0.693 was divided by the value of the slope to give the R1/2 value (37).
Single-molecule experiments
Slide preparation
Quartz slides (Finkenbeiner Inc.) were boiled in water for 20 min to remove parafilm, epoxy and coverslips from previous experiments. The quartz slides were then scrubbed with alconox by hand and rinsed with MilliQ water. The slides were then placed in a solution containing 10% w/v Alconox and sonicated for 20 min. The slides were then rinsed and sonicated in MilliQ water for 5 min. The slides were then sonicated in acetone for 15 min followed by direct transfer to a solution containing 1 M KOH and sonicated for 20 min. The slides were then rinsed with water and thoroughly flame dried using a butane torch (BernzOmatic). After flaming, the slides were left to cool on a drying rack in the fume hood. While the slides were cooling, a slide holder containing methanol was sonicated for 5 min and a silanizing solution containing 100 ml methanol, 5 ml of glacial acetic acid and 1 ml of N-(2-aminoethyl)-3- aminopropyltrimethoxysilane (UCT) was prepared during this time. After the slides had cooled down, the methanol in the slide holder was exchanged for the silanizing solution and the slides were sonicated in this solution for 1 min and then allowed to stand in the solution for an additional 20 min at room temperature. While slides were incubating, 400 mg of mPEGSuccinimidyl Valerate MW 5000 (Laysan Bio, Inc.) was resuspended in 800 ul 0.1M NaHCO3. Also, 2 mg of Biotin-PEG-Succinimidyl Valerate MW 5000 (Laysan Bio, Inc.) was resuspended in 200 ul of 0.1M NaHCO3 and mixed with the other PEG solution. After combining, the PEG solution was briefly sonicated to ensure complete resuspension. The slides were rinsed with MilliQ water and dried with nitrogen gas and then put in a humidor box. 150 ul of the PEG solution was applied to each slide and covered with a coverslip. The slides were incubated in the dark overnight. The next day the coverslip and PEG solution was rinsed off using MilliQ water and dried using nitrogen gas. Channels were assembled using parafilm strips as spacers on the pegylated quartz slide and plasma-cleaned coverslips were added as the upper channel face. For real-time experiments, flow cells were assembled by first drilling two holes at opposite ends of a quartz slide. A cut pipette tip was glued into one hole to act as a reservoir, and intramedic® PE100 polyethylene tubing glued to the other to provide a vacuum for buffer exchange. The channel was cut from a piece of double sided tape which was sandwiched between the pegylated quartz slide and a coverslip.
Enzyme immobilization
Channels were incubated with 30 ul 10 mg/ml BSA (NEB) for at least 30 min. 60 ul of 0.2 mg/ml MPS (streptavidin) was then added and incubated for 5 min. The channel was then washed with 150 ul T50 buffer (10 mM Tris-HCl, pH 8, 50 mM NaCl, or LiCl for Li+ experiments)). 2.5 ul of Cy3-telomerase, 0.5 nM primer-Cy5 and Ix imaging buffer (50 mM Tris-HCl, pH 8.3, 50 mM KCl, or LiCl for Li+ experiments, 1 mM MgCl2, 0.8 % glucose, 0.5 mg/ml BSA) in a total volume of 10 ul was incubated for 1h on the benchtop in the dark. Before addition of primer to the enzyme, a 10x primer stock was diluted in 1x T50 with LiCl and then heated for 5 min at 95°C. The 10x primer was snap cooled on ice and 1 ul was subsequently added to the enzyme. The imaging buffer was first saturated with Trolox (triplet state quencher) and then filtered through a 0.22 um filter and brought to pH 8.3. The enzyme/primer complex was then diluted with 40 ul of Ix imaging buffer and then incubated on the slide for 10 minutes. The slide was put on the TIRF objective, the density of immobilized molecules was assessed and 100 ul of imaging buffer + 1 ul gloxy solution (200 ug/ml catalase and 100 mg/ml glucose oxidase in T50) was flowed through the channel.
Primer extension by surface-immobilized telomerase
After the telomerase complexes were stalled on the slide surface, the indicated dNTPs were added as described in the text at a concentration of 200 uM. Reactions were allowed to proceed for the indicated amount of time. For real-time experiments, dNTPs were added to the slide during active data acquisition.
Data acquisition and analysis
Imaging fields containing 300-700 molecules were imaged at 2 frames/second for real-time smFRET experiments and 10 frames/second for standard smFRET experiments. Green laser intensity was set to 3mW for real-time experiments and to 15mW for standard experiments. 20 two-second movies were collected at each indicated time-point for the standard FRET experiments and 15-minute movies were collected for the real-time experiments. Individual traces were parsed out using custom written IDL software to correct for dye-cross talk and background. Individual traces were then filtered in MATLAB and molecules that did not contain and acceptor dye were discarded. FRET intensities were calculated using the equation IA/(IA+yID) where IA is the acceptor intensity, ID is the donor intensity and y is the gamma correction factor. The first two seconds of individual FRET traces were binned into histograms.
Kinetic analysis
For kinetic analysis, the primer extension assay described above was modified by chasing with 20 μM cold (TTAGGG)3 primer after 20 minutes of initiating the reaction, to prevent hot primer and hot product rebinding within the subsequent 70-minute time course. The intensities of the RAP bands on the gel electrophoretogram were converted to absolute concentrations based on the intensity of the initial hot primer band (50 nM). Multiple exposures were analyzed to ensure intense and weak bands remained within the linear range of the phosphorimager. These data were analyzed globally according to the sequential model (Figure 6B) using DynaFit (59, 60). Further details are given in the supplementary Methods.
AUTHOR CONTRIBUTIONS
L.I.J. and M.D.S. designed research, L.I.J., T.C. and R.B. performed research, L.I.J., J.H., J.W.P., C.R.B. and M.D.S. analyzed data and L.I.J. and M.D.S. wrote the paper.
ACKNOWLEDGEMENTS
We thank Dr. Harry Noller and Dr. Alan Zahler for critical reading of the manuscript. This work was supported by National Institutes of Health Grants F99CA212439 (to L.I.J) and R01GM095850 (to M.D.S).