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. 2019 Mar 26;8:e43204. doi: 10.7554/eLife.43204

The cryo-EM structure of a 12-subunit variant of RNA polymerase I reveals dissociation of the A49-A34.5 heterodimer and rearrangement of subunit A12.2

Lucas Tafur 1,2, Yashar Sadian 1, Jonas Hanske 1, Rene Wetzel 1, Felix Weis 1, Christoph W Müller 1,✉
Editors: John Kuriyan3, Cynthia Wolberger4
PMCID: PMC6435322  PMID: 30913026

Abstract

RNA polymerase (Pol) I is a 14-subunit enzyme that solely transcribes pre-ribosomal RNA. Cryo-electron microscopy (EM) structures of Pol I initiation and elongation complexes have given first insights into the molecular mechanisms of Pol I transcription. Here, we present cryo-EM structures of yeast Pol I elongation complexes (ECs) bound to the nucleotide analog GMPCPP at 3.2 to 3.4 Å resolution that provide additional insight into the functional interplay between the Pol I-specific transcription-like factors A49-A34.5 and A12.2. Strikingly, most of the nucleotide-bound ECs lack the A49-A34.5 heterodimer and adopt a Pol II-like conformation, in which the A12.2 C-terminal domain is bound in a previously unobserved position at the A135 surface. Our structural and biochemical data suggest a mechanism where reversible binding of the A49-A34.5 heterodimer could contribute to the regulation of Pol I transcription initiation and elongation.

Research organism: S. cerevisiae

Introduction

RNA polymerase I (Pol I) is a eukaryotic, 14-subunit enzyme that solely transcribes pre-ribosomal (rRNA) from ribosomal DNA (rDNA) repeats. Although all three eukaryotic RNA polymerases (Pol I, Pol II and Pol III) share a structurally conserved 10-subunit core and a 2-subunit stalk, they have evolved distinct structural features, including accessory subunits, and rely each on a unique specialized set of general transcription factors (Engel et al., 2018; Khatter et al., 2017; Vannini and Cramer, 2012). Before the first structures became available, functional studies already suggested that Pol I had adapted to accommodate the transcriptional needs of ribosome production resulting in differences in its regulation, initiation and elongation compared to the well-studied Pol II system to promote fast initiation and processivity (Albert et al., 2012). Accordingly, Pol I relies on a simpler transcription initiation machinery compared to Pol II (Keener et al., 1998), and similar to Pol III, has incorporated Pol II transcription factor-like subunits during evolution (Engel et al., 2018; Khatter et al., 2017; Vannini and Cramer, 2012).

In recent years, structural information available for Saccharomyces cerevisiae (yeast) Pol I has increased dramatically, revealing the structural basis of the Pol I-specific functional adaptations. In the first crystal structures, Pol I formed a dimer, thereby locking the enzyme in an inactive conformation (Engel et al., 2013; Fernández-Tornero et al., 2013). Although the core structure was overall conserved compared to Pol II, the Pol I DNA-binding cleft was very wide, and it was occupied by the C-terminal domain of Pol I-specific subunit A12.2 and a DNA-mimicking loop/expander element that occupies the position of the DNA-RNA transcription bubble. This wide cleft conformation resulted in the unfolding of the bridge helix, a conserved element that connects the two biggest subunits in multi-subunit RNA polymerases and plays an important role during catalysis (Weinzierl, 2011). Subsequent cryo-electron microscopy (cryo-EM) structures of the Pol I elongation complex (EC) revealed that binding to a DNA-RNA scaffold promoted the closure of the DNA-binding cleft, thus freeing the active site for nucleotide binding and causing the folding of the bridge helix (Neyer et al., 2016; Tafur et al., 2016). The EC structures were very similar to those observed for Pol II (Kettenberger et al., 2004) and Pol III (Hoffmann et al., 2015), highlighting that the actively transcribing Pol I adopts a conserved conformation and suggesting that the enzymatic mechanism of nucleotide addition is functionally conserved. However, mutating specific conserved residues in the Pol I and Pol II active sites appear to have different effects in vitro (Viktorovskaya et al., 2013), suggesting that other elongation intermediates might reveal previously uncharacterized differences between Pol I and Pol II.

Structural data of the basal Pol I initiation complex have revealed a very different, much simpler architecture compared to Pol II (Engel et al., 2017; Han et al., 2017; Sadian et al., 2017). This simplification is further supported by the incorporation of transcription factor-like functions into Pol I subunits: In Pol I, the A49-A34.5 heterodimer (hereafter referred also as ‘heterodimer’) has been proposed to function as both, a TFIIF- and TFIIE-like factor, participating during transcription initiation and elongation (Geiger et al., 2010; Vannini and Cramer, 2012). A49 has two domains connected by a linker, each of which appears to have evolved functionally distinct properties. While the A49 C-terminal tandem winged helix domain (tWH) has structural homology to TFIIE, the N-terminal A49 domain forms a dimer with the A34.5 subunit which adopts a triple β-barrel structure that resembles the Rap74/30 module of TFIIF (Geiger et al., 2010). The heterodimer is anchored to the core enzyme by interactions through the A49-A34.5 dimerization domain and by an extended surface between the long C-terminal tail of A34.5 (A34.5-Ct) and Pol I’s second biggest subunit A135 (Engel et al., 2013; Fernández-Tornero et al., 2013). However, as the dimerization domains contribute most to the binding, deletion of either subunit results in a Pol I enzyme lacking both subunits (Gadal et al., 1997; Pilsl et al., 2016).

Since its discovery, Pol I has been shown to exist in two different conformations that differ by the presence of the heterodimer, which can be reversibly dissociated (Huet et al., 1975). The form lacking A49-A34.5, termed Pol I*, has reduced transcriptional specificity and activity compared to the complete Pol I enzyme (Huet et al., 1976). Although the heterodimer appears to increase the processivity of Pol I, details of its function are still unknown. Neither A49 (Liljelund et al., 1992) nor A34.5 (Gadal et al., 1997) are essential genes, and Pol I* has been proposed to co-exist with Pol I in vivo (Gadal et al., 1997). Deletion of topoisomerase I causes a very strong growth defect in yeast only when combined with a deletion of A34.5 (Gadal et al., 1997) suggesting that A34.5 is important for relieving topological stress during rDNA transcription. In vitro, the A49-A34.5 heterodimer has a stronger effect on promoter-dependent transcription than on non-specific transcription, while addition of the A49 tWH domain is sufficient to restore promoter-dependent and non-specific transcription (Pilsl et al., 2016). Overall, the data suggest that the heterodimer is functionally important for transcription initiation and/or (early) elongation. However, the functional and physiological relevance of Pol I* has not been elucidated to date. Furthermore, it is not clear if the heterodimer participates in all phases of transcription, or only during initiation and early elongation.

The Pol I-specific subunit A12.2 also contains additional built-in functionality. A12.2 shares homology with Pol II subunit Rpb9 in its N-terminal domain and the Pol II cleavage factor TFIIS in its C-terminal domain (Ruan et al., 2011). While the role of TFIIS in RNA cleavage is well established (Cheung and Cramer, 2011), Rpb9 appears to regulate transcription elongation (Hemming et al., 2000), proofreading (Knippa and Peterson, 2013) and transcription-coupled DNA repair (Li et al., 2006). The A12.2 C-terminal Zn ribbon domain (A12.2C) is required for the Pol I intrinsic RNA cleavage activity (Kuhn et al., 2007) and adopts a similar position as TFIIS in the cleft in unbound (apo) Pol I (Engel et al., 2013; Fernández-Tornero et al., 2013; Neyer et al., 2016) as well as in Pol I bound only to DNA (Sadian et al., 2017; Tafur et al., 2016), but is excluded from the active site upon formation of the EC (Neyer et al., 2016; Tafur et al., 2016). The exact position of A12.2C, however, has not been determined in the context of an actively transcribing complex. While deletion of A12.2C does not cause any growth defect, deletion of the A12.2 N-terminal Zn ribbon domain (A12.2N) produces a similar effect as deletion of the complete protein (Van Mullem et al., 2002). Interestingly, deletion of either the complete A12.2 or A12.2N also alters the nucleolar localization of Pol I, suggesting that A12.2 is important for Pol I integrity.

Studies to date suggest a functional interplay between the Pol I A49-A34.5 heterodimer and subunit A12.2. The heterodimer stimulates A12.2-mediated RNA cleavage in vitro (Geiger et al., 2010), the latter which is important for Pol I backtrack recovery (Lisica et al., 2016). A12.2N interacts directly with the dimerization domain of A49, thus stabilizing the anchoring of the heterodimer (Engel et al., 2013; Fernández-Tornero et al., 2013). Recently, A12.2 has also been proposed to be important for transcription initiation in vivo and in vitro, especially in the absence of A49 (Darrière et al., 2018). Combined with the reduced number of general transcription factors required for productive transcription initiation, the Pol I A49-A34.5 heterodimer and subunit A12.2 might promote the high initiation rate observed on rDNA repeats (French et al., 2003).

In this work, we describe the cryo-EM structures of Pol I and spontaneously formed Pol I* bound to a DNA-RNA scaffold and the nucleotide analog GMPCPP. These structures reveal a previously unobserved relationship between A12.2 and A34.5, provide the structural basis for the exclusion of the heterodimer from the core enzyme, and suggest mutually exclusive binding of the A49-A34.5 heterodimer and A12.2C during the Pol I transcription cycle.

Results

Cryo-EM structures of the GMPCPP-bound Pol I elongation complexes (EC)

In order to better understand the catalytic mechanism of Pol I, we incubated the Pol I EC with the non-hydrolysable nucleotide analog GMPCPP as previously used for Pol II (Kettenberger et al., 2004; Wang et al., 2006). The Pol I EC was prepared as previously described (Tafur et al., 2016) except that 1 mM MgCl2 was included in the buffer (Materials and methods). 5768 micrograph movies were collected on a FEI Titan Krios equipped with a K2 direct electron detector, and processed with RELION 2.0 (Kimanius et al., 2016). After sorting particles with 2D and 3D classification, an unexplained extra density next to the A135 surface was observed in most of the particles with a closed cleft and strong DNA-RNA density, concomitant with streaky and weak density for the A49-A34.5 heterodimer. To better resolve this density, particles were classified using a mask in this region (Figure 1—figure supplement 1). This revealed that the extra density corresponded to A12.2C (Figure 1). In total, 63% of all particles selected after the first unmasked 3D classification step did not have the heterodimer bound and showed density for A12.2C in this new position (named Pol I* in analogy to RNA polymerase A* (Huet et al., 1975)), while only 37% represented the 14-subunit Pol I. Extensive 3D classification ultimately yielded two different nucleotide-bound ECs: 12-subunit Pol I* EC lacking the heterodimer, which was refined to 3.18 Å resolution, and 14-subunit Pol I EC, which was refined to 3.42 Å resolution (Figure 1—figure supplement 2). The overall conformation of both Pol I forms is very similar, with the exception of the presence/absence of the heterodimer, the previously unobserved position of A12.2C and a slight difference in the conformation of the clamp, and resemble previously published structures (Figure 1) (Neyer et al., 2016; Tafur et al., 2016). Interestingly, an apo Pol I* reconstruction at 3.21 Å resolution was also obtained with a similar conformation as previously observed for the cryo-EM structures of monomeric Pol I (Neyer et al., 2016; Pilsl et al., 2016), highlighting that the presence of the heterodimer and the novel position of A12.2C do not impose any conformational constraints on the Pol I core (Figure 1—figure supplement 3). Models were built using previous Pol I structures as a starting point and were real-space refined, yielding structures with excellent stereochemistry (Table 1).

Figure 1. Structures of the Pol I* EC and Pol I EC bound to GMPCPP.

(A) In the Pol I* EC, the A49-A34.5 is absent and the A12.2C adopts a position on the A135 surface. The overlap between this position and the C-terminal domain of A34.5 is indicated in the Pol I EC as a dashed yellow surface. (B) The 14-subunit Pol I bound to a DNA-RNA scaffold is shown colored according to the subunits indicated in the legend. In this conformation, only up to residue 67 of A12.2 is observed (A12.2 hinge), while the C-terminal domain (A12.2C) is disordered. (C) Comparison between the apo (left), Pol I EC (middle) and Pol I* EC (right) reveals that the A12.2C can alternate between TFIIS-like (apo) or Rpb9-like (right) positions. Movement of the A12.2C is around a hinge at residue 67, also indicated in the Pol I EC (A). The position of the External domain 1 (ED1) and hybrid binding (HB) interaction surfaces are indicated in the Pol I EC. A12.2 is shown as ribbon diagram and yellow surface (not EM density) for easier visualization. See also Figure 1—figure supplements 1–3.

Figure 1.

Figure 1—figure supplement 1. Cryo-EM data and processing.

Figure 1—figure supplement 1.

(A) Representative micrograph. Scale bar = 100 nm. (B) 2D class averages from the initial auto-picked particles. (C) Processing pipeline. The resolution, number of particles and percentage of particles with respect to the initial number of particles after 2D classification is shown below each class. The stalk subunits (A43–A14), the DNA-RNA, the heterodimer (A49-A34.5) and A12.2 are colored as in Figure 1.
Figure 1—figure supplement 2. Average and local resolution estimates for the reconstructions.

Figure 1—figure supplement 2.

(A) Fourier-shell correlation (FSC) curves for the reconstructions. (B) FSC curves for the models versus experimental maps. (C–F) Local resolution estimates for all the reconstructions and angular distribution plots. (G). Representative densities.
Figure 1—figure supplement 3. Structure of the apo Pol I*.

Figure 1—figure supplement 3.

(A) Top and front views for the apo Pol I* model fitted into the sharpened density. The bridge helix is unfolded and shown below the models. (B) Comparison between the A12.2 in the apo Pol I (PDB: 5m3m) (Neyer et al., 2016) and apo Pol I*. In the apo Pol I, the A12.2C is in the TFIIS-like position (orange), while in the apo Pol I*, the A12.2C is in the Rpb9-like position (yellow). Movement of the A12.2C induces a shift in the N-terminal A12.2 Zn ribbon domain A12.2N, which moves towards the lobe. The flexible jaw helices also move slightly compared to the apo Pol I (orange red).
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Table 1. Data collection and refinement statistics.

Pol I (core) EC + GMPCPP Pol I EC + GMPCPP Pol I* EC + GMPCPP Apo Pol I*
Data collection
Particle number 54,017 30,232 182,488 73,660
Pixel size (Å/pix) 1.04 1.04 1.04 1.04
Average resolution (Å) 3.18 3.42 3.18 3.21
B-factor −44.5 −34.2 −92.9 −99.6
EMDB code EMD-0240 EMD-0238 EMD-0239 EMD-0241
Refinement statistics*
PDB code 6HLR 6HKO 6HLQ 6HLS
CC (atoms)† 0.816 0.804 0.796 0.797
RMSD (bonds) 0.007 0.006 0.006 0.007
RMSD (angles) 1.22 1.18 1.18 1.25
Clashscore 4.74 5.27 5.13 5.17
Rotamer outliers (%) 0.12 0.14 0.09 0.32
C-beta deviations (%) 0 0 0 0
Ramachandran plot
Outliers (%) 0 0 0 0
Allowed (%) 4.9 5.64 4.59 5.48
Favored (%) 95.1 94.36 95.41 94.61
Molprobity score 1.58 1.67 1.59 1.65
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*Calculated with Molprobity.

†From PHENIX real space refinement.

The A12.2 C-terminal domain alternates between a TFIIS-like and an Rpb9-like position

In the 12-subunit Pol I* EC, lacking the heterodimer, the A12.2C occupies a novel position next to the A135 surface (Figure 1A). This new position overlaps with the A34.5-Ct in the complete, 14-subunit Pol I EC (Figure 1B), where A12.2C is disordered and only density up to residue 67 is observed (Neyer et al., 2016; Tafur et al., 2016). The new position of A12.2C resembles that of the C-terminal domain of Pol II subunit Rpb9, outside of the DNA-binding cleft (Figure 1C, right), and is distinct from the previously reported TFIIS-like position near the active site (Figure 1C, left). The A12.2C can move between these positions by rotating around a hinge located at residues 66–67 (Figure 1A, Figure 1C, middle). Whereas binding to the TFIIS-like position is only possible when the DNA-binding cleft is open (Engel et al., 2013; Fernández-Tornero et al., 2013) or partially open (Neyer et al., 2016; Sadian et al., 2017; Tafur et al., 2016), binding to the Rpb9-like position can only occur when the heterodimer is absent.

Detailed analysis of Pol I* and Pol I reveals that different interactions occur in two areas of the A135 surface (Figure 2A, Figure 1C, middle). The first area involves part of the A135 External Domain 1 (ED1), which interacts either with the A34.5-Ct (in Pol I) or A12.2C (in Pol I*). The second area corresponds to part of the A135 Hybrid Binding (HB) domain (residues 989 to 1000), which in Pol I interacts with the A34.5-Ct but in Pol I* interacts with the A135 N-terminal tail (A135-Nt), which folds back towards the HB domain (Figure 2A). The A135-Nt effectively acts as a switch, changing its positioning to allow or to prevent A34.5-Ct binding to the HB domain. Both A12.2C and A34.5-Ct form similar interactions with the Pol I core, as both interact with two neighboring asparagine residues in the A135 ED1 (N683 and N684) (Figure 2B,C) and an aspartate residue (D990) in the A135 HB domain (Figure 2D,E).

Figure 2. Interactions of the A12.2C with the A135 External domain one and Hybrid binding domain.

Figure 2.

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(A) Two interfaces are differently arranged in Pol I* versus Pol I. Both A34.5-Ct and A12.2C can bind to the A135 External Domain 1 (ED1, red), and A34.5-Ct and the N-terminal tail of A135 (A135-Nt) can bind to the A135 Hybrid Binding domain (HB). (B). In the ED1, the A12.2C interacts with both A135 N683 and N684 through Y96 and T98, respectively. (C). In the ED1, the A34.5-Ct interacts with A135 N683 and N684 through R154 and Y150, respectively. (D) In the HB surface, the A135-Nt folds back and positions R12 next to D990. (E) A34.5-Ct interacts with D990 from the HB domain through R157. Densities shown for panels B-E are from the sharpened Pol I* and Pol I EC (+GMPCPP). See also Figure 1—figure supplement 2.

In the monomeric apo Pol I, A12.2C can still occupy the TFIIS-like position (Neyer et al., 2016). However, in the apo Pol I*, despite being sufficient space for accommodating A12.2C in the TFIIS-like position, A12.2C is observed in the Rpb9-like position (Figure 1—figure supplement 3). The presence of the heterodimer in the enzyme could thus promote binding of A12.2C to the TFIIS-like site (when accessible) by blocking the Rpb9-like binding site. In apo Pol I*, the change in the position of A12.2C also shifts the A12.2N by ~3 Å towards the jaw, and part of the latter appears to move towards the A12.2 linker, likely to stabilize its position (Figure 1—figure supplement 3). Interestingly, both domains move relative to a region of A12.2 (residues ~ 43–66), which fixes this subunit to the Pol I core. Therefore, the movement of both, the A12.2N and the jaw, accommodate the change in the position of A12.2C.

A12.2C does not displace A49-A34.5 from the Pol I core

At present, it is unclear why most of the particles lack the heterodimer compared to previous Pol I EC structures (Neyer et al., 2016; Tafur et al., 2016). It is possible that differences in sample preparation conditions such as changes in the buffer conditions during freezing or the use of a thin layer of carbon in the cryo-EM grids account for the difference. While the cryo-EM structures show that A34.5-Ct and A12.2C compete for the same binding sites in A135, they don’t allow to distinguish if A12.2C displaces the heterodimer from the Pol I core or if A12.2C binds only once the heterodimer has dissociated from the enzyme. To test these hypotheses, we performed a series of fluorescence anisotropy experiments, using recombinant heterodimer, where a cysteine has been introduced in the A49 linker region for labelling with Alexa Fluor 594, and endogenously purified Pol I* (Pilsl et al., 2016) incubated with DNA (Pol I * EC) (Figure 3A). Because the fluorescent signal was low, we performed the experiments with a heterodimer concentration of 100 nM. Compared to the heterodimer alone, we observed an increase in anisotropy in a concentration-dependent manner as we added Pol I* EC (Figure 3B). The same experiment using wild type Pol I EC gave a right-shifted curve, indicating an exchange between endogenous heterodimer on wild type Pol I and labelled heterodimer. These data suggest that heterodimer binding to Pol I is reversible, and that A12.2C binding A135 as observed in Pol I* does not irreversibly prevent heterodimer binding. Because a 1:1 binding model did not allow fitting the data, no attempt was made to introduce more complex binding models. Incubation of the Pol I*/A49-A34.5 sample with recombinant A12.2C (residues 79 to 125) for 30 min did not reduce the anisotropy (indicating the release of the heterodimer from Pol I) even at 50-fold molar excess (Figure 3C). Although the affinity of A12.2C for the ED1 might further increase when it is constitutively anchored to Pol I by A12.2N. Similarly, incubation of the complex in the presence of GMPCPP did not change the anisotropy of the bound complex even at 20 mM (Figure 3C). These results suggest that binding of the A12.2C to the Rpb9-like position is only possible after the heterodimer has dissociated from Pol I.

Figure 3. Binding of the A49-A34.5 to the Pol I core in vitro.

Figure 3.

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(A) Experimental set up. Recombinant A49-A34.5, fluorescently labeled with Alexa 594 at residue 140, was mixed with the reconstituted Pol I* EC. The change in fluorescence anisotropy reflects the binding of A49-A34.5 to the Pol I core (an increase in anisotropy with respect to the free A49-A34.5 represents the formation of the 14-subunit Pol I). (B) Experimental data showing the change in fluorescence anisotropy upon binding of fluorescent A49-A34.5 to Pol I* as well as the replacement of endogenous heterodimer in wild type Pol I by fluorescent A49-A34.5. The points shown are an average of three replicates, with the standard deviation. (C) The reconstituted and labeled 14-subunit Pol I EC was incubated with increasing amounts of recombinant A12.2C (residues 70–125) for 30 min. Compared to the Pol I EC, no change in anisotropy is observed at either 1, 5 or 50-fold molar excess of A12.2C or with 20 mM GMPCPP.

ED1 determines binding of the C-terminal domain of the Rpb9-like subunit

Comparison of Pol I* with Pol II and Pol III reveals that while the External Domain 2 (ED2) appears to be structurally more conserved, the Pol I ED1 diverges from its Pol II and Pol III counterparts, as it is smaller and lacks an extension that overlaps with A12.2C in the Rpb9-like position (Figure 4A). In Pol II, the Rpb9 C-terminal domain (Rpb9C) also binds the ED1, although differently than A12.2C due to the presence of an extension in the ED1 (Figure 4B). Therefore, the Pol I and Pol II ED1 are specifically tailored to bind A12.2C and Rpb9C, respectively. Interestingly, a similar situation is observed in Pol III (Figure 4C). The Pol III ED1, as in Pol II, also has an extension in a region that overlaps with the position of A12.2C, but in addition, binding of the C11 C-terminal domain (C11C, equivalent to Pol I A12.2C and Pol II Rbp9C) in an Rbp9C-like position is precluded by the presence of a helix from subunit C53. Accordingly, the C11C adopts a position far from the Pol III ED1 (Hoffmann et al., 2015) that differs from the position of both A12.2C and Rpb9C (Figure 4C).

Figure 4. Comparison of the positions of the C-terminal domains of Pol I A12.2, Pol II Rbp9 and Pol III C11.

Figure 4.

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The positions of A12.2 (A), Rpb9 (B) or C11 (C) are shown in yellow for Pol I*, Pol II (Kettenberger et al., 2004) and Pol III (Hoffmann et al., 2015), respectively. While the ED2 is structurally more conserved (light sea green color), the ED1 in Pol II and Pol III are larger than the Pol I ED1 (red). The structure of the ED1 determines the binding mode of Pol I A12.2C and Pol II Rpb9C, while in Pol III the presence of C53 induces a different binding site for C11C far from the ED. The position of the N-terminal tail of the second largest subunit is also indicated for each polymerase, as well as the extension in the ED1 of Pol II and Pol III.

The active site conformations in Pol I * and Pol I are identical

Because the Pol I* EC reconstruction was obtained in the presence of the non-hydrolysable nucleotide analog GMPCPP and 1 mM MgCl2, we carefully compared the Pol I* active site with the active site in the 14-subunit Pol I reconstruction. As no differences were observed between the active sites, we pooled particles from both EC reconstructions, and classified them by restricting the classification to the core enzyme and the DNA-RNA hybrid using a soft mask and higher weight on the data (Scheres, 2016) (Figure 1—figure supplement 1, Materials and methods). This strategy also allowed us to resolve two main features from the active site: the binding and interactions of the incoming nucleotide (NTP) substrate (GMPCPP), and the interactions between Pol I and the +1 and+2 bases from the single-stranded non-template strand (NT) (Figure 5A).

Figure 5. Interactions in the Pol I active site with GMPCPP, and the +1 and+2 bases from the non-template strand.

(A) Pol I can bind the incoming nucleotide (GMPCPP) in the active site, while nucleotides of the opposite, non-template strand (+1 and+2), interact with the Fork loop two and Loop B. (B) GMPCPP is bound by conserved, identical residues in Pol I and Pol II. These include two arginines that interact with the phosphate (R714 and R957), a leucine from the trigger loop that stacks against the DNA base (L1202), and R591 and N625 which recognize the 2’- and 3’-OH groups, respectively. The ‘gating tyrosine’ (Y717), involved in RNA positioning during backtracking (Cheung and Cramer, 2011), and K916 and K924, which bind the 3’-end of the RNA are also indicated. Residues are shown in grey (A190) or tan (A135) for Pol I, while those in Pol II in dark green, and in Pol III in light green. Density for the DNA-RNA hybrid is from the sharpened, Pol I (core) EC (+GMPCPP) reconstruction, while the GMPCPP is from the same reconstruction but from the unsharpened/unmasked map. Density for L1202 is shown at a lower threshold. Residues are boxed according to their proposed role: black box, triphosphate binding; red box, nucleotide base stabilization; dashed box, NTP/dNTP discrimination. (C) Binding of GMPCPP is virtually identical in Pol I (top) and Pol II (bottom, PDB: 4a3j) (Cheung et al., 2011).( D) In the downstream edge of the transcription bubble, the +2 base of the NT strand is flipped into a pocket formed by Fork loop 2 (FL2) and loop B (‘A135 pocket’). These elements interact with the nucleotide through R219, R225 and the conserved D395. (E) These interactions also position the +1 base next to F508 from FL2 (top), resembling the interaction of the +1 base with βW183 in bacterial Pol (bottom, PDB: 6alh) (Kang et al., 2017). See also Figure 5—figure supplement 1.

Figure 5.

Figure 5—figure supplement 1. Conformational heterogeneity in the Pol I EC.

Figure 5—figure supplement 1.

Classification of the pooled Pol I EC particles reveals intermediates with slight differences in the width of the cleft, flipping of the +2 base, density for GMPCPP and trigger loop (TL). Closing of the cleft is accompanied by +2 base flipping, stronger density for GMPCPP and appearance of density for A190 L1202. Flipping of DNA base +2 occurs before complete closing of the cleft by the movement of the Jaw/clamp domains towards each other (between state 3 and 4). The transition from state 1 to 5 includes the sequential movement of the protrusion and wall domains towards the clamp, the jaw/clamp movement that induces +2 base flipping and closing of the cleft by movement of modules 1 and 2.
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As suggested by the conservation of residues in this region, the NTP is positioned in the ‘A’ site, as previously seen in Pol II (Cheung et al., 2011; Wang et al., 2006; Westover et al., 2004) and bacterial RNA polymerase (bcPol) (Vassylyev et al., 2007) (Figure 5B). Accordingly, the phosphate moiety is bound by two invariant arginine residues (A135 R714 and R957). In addition, the conserved A190 N625 and R591, which are involved in NTP/dNTP discrimination, come close to the 3’- and 2’-OH group, respectively. While the corresponding residue to N625 in Pol II (Rpb1 N479) has been shown to interact with either the 3’-OH (Wang et al., 2006) or the 2’-OH (Cheung et al., 2011), the invariant R591 (Rpb1 R446) interacts with the 2’-OH of the ribose in all structures. The NTP is maintained in the correct position by L1202 from the trigger loop, which interacts with the guanosine base. Only up to this residue, weak density can be observed, while the ‘tip’ loop (A190 residues 1203–1212) is unresolved. Overall, the positioning of the NTP substrate in the Pol I active site is virtually identical to that in Pol II (Figure 5C).

An interesting scenario is also observed opposite to the NTP binding site, where Pol I displays features similar to Pol II and bcPol. In both, the Pol I and Pol I* EC, the downstream edge of the transcription bubble is stabilized by interactions of Pol I with nucleotides + 1 and+2 from the NT strand. The +2 base is flipped into a pocket formed by elements from the A135 subunit, namely, the fork loop 2 (FL2) and loop B (Tafur et al., 2016) (Figure 5D). These two elements form a pocket (‘A135 pocket’) which resembles that formed by the β subunit (‘β-pocket’) in bcPol (Zhang et al., 2012). Whereas loop B exposes several positively charged residues towards the cavity of the A135 pocket that likely stabilize the phosphate backbone, a phenylalanine from the A135 FL2 (F508) appears to stack with the +1 base, in an analogous fashion as W183 from the bcPol β subunit (Zhang et al., 2012) (Figure 5E). Finally, the highly conserved D395 also interacts with the +2 base as in bcPol (β subunit D446) (Vvedenskaya et al., 2014) and Pol II (Rpb2 D399) (Cheung and Cramer, 2011), and probably also in Pol III (C128 D370). However, neither Pol II nor Pol III can form the equivalent interactions as in the A135 pocket because their corresponding loop B is differently positioned and far from the +1 and+2 bases. Interestingly, both sets of interactions are formed only when the DNA-binding cleft is completely closed and the jaw and clamp modules move towards each other (Figure 5—figure supplement 1). Thus, while formation of the EC involves the coarse movement of modules 1 and 2, nucleotide stabilization in the active site requires a more subtle, modular rearrangement.

Discussion

Crystal and cryo-EM structures of Pol I in different functional states have revealed not only an overall conformational conservation compared to Pol II and Pol III, but have also shed light on the role of specific subunits, as well as the structural transitions from an inactive dimer to an actively transcribing enzyme (Engel et al., 2018). One of the main differences between the available Pol I structures is the position of A12.2C. In elongating Pol I, A12.2C is excluded from the active site, while no alternative position could be determined presumably because it is disordered. Here, we show that A12.2C can alternate between TFIIS-like and Rpb9-like positions depending on the presence of the A49-A34.5 heterodimer. In the TFIIS-like position, A12.2C is positioned in the DNA-binding cleft and occludes the active site, which is incompatible with NTP incorporation but in accordance with RNA cleavage. When the cleft closes (and thereby clashes with A12.2C in the TFIIS-like position), A12.2C is excluded from the active site and can bind to the A135 ED1. Binding of A12.2C and heterodimer to the ED1 are mutually exclusive, as A12.2C and A34.5-Ct use overlapping binding sites. Exclusion of the heterodimer in this conformation is supported by the movement of the A135-Nt towards the HB domain, which blocks the interaction of the distal part of the A34.5-Ct with this domain. These results suggest a mechanism by which the surface of A135 (in particular, the ED1) plays a pivotal role in specific factor exchange in Pol I. Recent genetic studies have suggested that A12.2 may be involved in modulation of the movement of the jaw/lobe interface especially in the absence of A49, as the A49 linker and tWH domain appear to stabilize the closed conformation of Pol I when bound to DNA (Darrière et al., 2018). As the A12.2C binds to the A135 ED1, which sits next to the A135 lobe, the A12.2C might restrict the movement of the lobe. Thus, while A12.2N regulates the flexibility of the jaw, A12.2C could additionally regulate the movement of the lobe. Together, both A12.2 domains could therefore regulate cleft opening/closing of Pol I upon DNA binding, as well as binding to the +1 and+2 nucleotides in the non-template strand (see above). Restriction of movement of the A135 lobe by A12.2C might be important to maintain the closed state in the absence of A49, as in Pol I*. In contrast, when the heterodimer is present in the complex, A12.2 might destabilize the EC as it can only occupy the TFIIS-like site, thereby preventing cleft closure (Appling et al., 2018). In this scenario, A49 could play an important role in maintaining a narrow cleft, which would also explain (in addition to the direct interaction of their N-terminal domains with A12.2) the stimulatory role of the heterodimer on A12.2-mediated RNA cleavage (Geiger et al., 2010).

In vivo, heterodimer association to Pol I might offer an additional layer of regulation of rDNA transcription (Figure 6). The proportion of initiation-competent Pol I molecules in the cell has been proposed to represent those Pol I particles bound to initiation factor Rrn3 (Milkereit and Tschochner, 1998). In contrast, the number of Pol I* particles in the cell could represent a population of actively transcribing DNA-bound Pol I, but also a pool of pre-active Pol I that can readily initiate transcription upon heterodimer binding and Rrn3 recruitment (in contrast to Pol I dimers, which appear to be a storage form of the enzyme (Torreira et al., 2017)). The number of initiation-competent Pol I molecules could be thus regulated not only by Pol I homo-dimerization and association with Rrn3, but also by changes in the heterodimer concentration in the nucleolus, thereby controlling the ratio of Pol I to Pol I*. Nutrient-dependent regulation of nucleolar localization of the mammalian A49-A34.5 homolog PAF53-PAF49 has been observed (Penrod et al., 2015). PAF49 (A34.5 counterpart) accumulates in the nucleolus in growing cells but disperses to the nucleoplasm upon serum starvation (Yamamoto et al., 2004). In yeast, A34.5 is maintained in the nucleolus by its association with A49 (but also contains a nucleolar localization signal in its C-terminal region), and A49 is required for the high loading rate of Pol I onto rDNA (Albert et al., 2011). Human PAF53-PAF49 can substitute the A49-A34.5 heterodimer in vivo (Albert et al., 2011) suggesting a conserved function (and possibly regulation). Regulation of heterodimer binding to Pol I might also explain why promoter association of Pol I-Rrn3 complexes is low upon nutrient starvation even when the concentration of such complexes is relatively high (Torreira et al., 2017); the levels of the heterodimer might further regulate Pol I initiation rates.

Figure 6. Schematic representation of the possible physiological role of the A49-A34.5 heterodimer in the regulation Pol I activity.

The pool of initiation-competent Pol I particles is controlled by Pol I homo-dimerization (A) and binding of Rrn3 to monomeric Pol I (B). After transcription initiation and promoter escape, during elongation, Pol I can alternate between Pol I and Pol I* conformations. Release of the A49-A34.5 heterodimer would allow the recruitment of elongation factors (C). After dissociating from DNA, Pol I* could bind to the A49-A34.5 heterodimer to replenish the pool of initiation-competent Pol I monomers. The concentration of A49-A34.5 heterodimer in the nucleolus might be also regulated by the nutrient status of the cell as in the mammalian system. Regulated localization of the A49-A34.5 heterodimer would serve to alter the ratio of Pol I to Pol I* in the nucleolus, thereby controlling the initiation rate on the rDNA. See also Figure 6—figure supplement 1.

Figure 6.

Figure 6—figure supplement 1. A49-A34.5 heterodimer release frees the binding site for Spt4/5 and Paf1C.

Figure 6—figure supplement 1.

Heterodimer binding to Pol I is prevented by the binding of A12.2C to the ED1 (1) and binding of the A135-Nt to the HB further prevents interactions of Pol I with the A34.5-Ct (2). Without anchoring of the heterodimer to the Pol I core, the interaction of the A49 tWH with the upstream DNA is abolished (3). Both A49-A34.5 dimerization module and A49 tWH sites are replaced by Paf1C and Spt4/5, respectively, in a manner similar to the transition from initiation to elongation in Pol II where TFIIF and TFIIE are replaced by Paf1C and Spt4/5.
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In addition, the release of the heterodimer from the enzyme would also allow the binding of elongation factors to Pol I. Pol I has been shown to bind to elongation factor Spt5 directly (Viktorovskaya et al., 2011) and its activity is affected by Spt4/5 in vivo (Anderson et al., 2011). In the Pol I EC, canonical binding of Spt4/5 (as in the Pol II EC) is precluded by the A49 tWH (Tafur et al., 2016), as it occupies a position equivalent to the KOW1-L1 domain of Spt5, and by the A49 linker helix spanning the cleft, which clashes with the N-terminal region of Spt5 (Figure 6—figure supplement 1) (Bernecky et al., 2017; Ehara et al., 2017). Interestingly, Spt5 interacts physically and genetically with A49, suggesting a functional interplay between these proteins (Viktorovskaya et al., 2011). Paf1C, another elongation factor, has also been shown to stimulate Pol I transcription in vivo and in vitro (Zhang et al., 2009; Zhang et al., 2010). Paf1C binds to Pol II on the outer surface of subunit Rpb2 (Pol II counterpart of A135) including the Rpb2 ED2 and lobe (Vos et al., 2018; Xu et al., 2017). In this position, it clashes and competes with TFIIF for Pol II binding (Xu et al., 2017). Heterodimer dissociation from Pol I could potentially free the binding site for both Spt4/5 and Paf1C in a mechanism that could be akin to the transition from initiation to elongation in Pol II: while TFIIE (A49 tWH) blocks the Spt4/5 binding site, TFIIF (A49-A34.5 dimerization domain) occupies the binding site of part of Paf1C (Vos et al., 2018; Xu et al., 2017) (Figure 6—figure supplement 1). Thus, binding of elongation factors is mutually exclusive with the presence of initiation factors. Therefore, in Pol I, factor exchange during the transition from initiation to elongation could be accommodated more readily just by the release of the heterodimer and switching to the Pol I* form. In this scenario, A12.2 might further prevent re-association of the heterodimer. A similar allosteric transition during promoter escape mediated by the heterodimer, Spt5 and the stalk has been previously proposed for Pol I (Beckouët et al., 2011). Because we could not observe any effect of free A12.2C on heterodimer binding to Pol I in vitro, release of the heterodimer in vivo might be directly induced by Spt4/5 and Paf1C.

Materials and methods

Pol I EC-GMPCPP complex formation

Endogenous Pol I was purified from yeast cells as previously described (Moreno-Morcillo et al., 2014). Pol I was incubated with a 38 base pair transcription scaffold containing an 11 nucleotide mismatch bubble and a 20 nucleotide RNA as used previously for formation of the Pol I EC (Tafur et al., 2016). The complex was incubated for 1 hr at 4°C in 15 mM HEPES-NaOH (pH 7.5), 150 mM ammonium sulfate, 1 mM MgCl2, 1 mM GMPCPP (Jena Bioscience) and 10 mM DTT. The sample was diluted to ~0.1 mg/mL in the same buffer immediately before grid freezing.

Cryo-EM sample preparation

2.5 μL of sample was deposited on a freshly glow-discharged cryo grid (R 2/1 + 2 nm carbon, Quantifoil), incubated for 30 s, and blotted for 3 s (with a blotting force of ‘3’), at 100% humidity and 4°C in a Vitrobot Mark IV (FEI). Grids were stored in liquid nitrogen until data collection.

Cryo-EM data collection

5768 micrograph movies were collected on a FEI Titan Krios at 300 keV through a Gatan Quantum 967 LS energy filter using a 20 eV slit width in zero-loss mode. The movies were recorded on a Gatan K2 direct electron detector, at a nominal magnification of 135,000x corresponding to a pixel size of 1.04 Å in super resolution mode, using Serial EM. Movies were collected in 40 frames with defocus values from −0.75 to −2.5 μM, with a dose of 0.9775 e- Å−2 s−1 per frame for 16 s.

Cryo-EM data processing

Movies were aligned, motion-corrected and dose-fractionated using MotionCor2 (Zheng et al., 2017). Contrast transfer function (CTF) estimation was done using CTFFIND4 (Rohou and Grigorieff, 2015). All processing steps were performed in Relion 2.0 (Kimanius et al., 2016) unless otherwise indicated. Resolution estimates reported are those obtained after masking and B-factor sharpening (Relion post-processing). Data were divided in five batches to increase processing speed. For each batch, autopicking was followed by a 2D classification step (with data downsized five times) to remove contamination and damaged particles. Good classes were selected, re-extracted and un-binned, and refined against the Pol I EC (PDB: 5m5x) low pass filtered to 40 Å. Then, a 3D classification step was performed without alignment. For all batches the same procedure was followed, except for batch 5, in which 3D classification was performed with data downsized five times. Classes were selected based on the width of the cleft, the position of the clamp, and the DNA-RNA scaffold density, and grouped by similarity. Refinement of the pooled particles with closed cleft and strong DNA-RNA density revealed an extra density and streaky, weak density for the A49-A34.5 heterodimer. To resolve this region, a masked classification was performed. This yielded a class with high resolution in the extra density, allowing the unambiguous assignment of the A12.2 C-terminal domain (A12.2C). Based on these results, all other pooled classes were classified with a mask on this area. Particles were merged depending on whether they showed density for the A49-A34.5 heterodimer (Pol I) or the A12.2C without A49-A34.5 (Pol I*). During the process, additional bad particles were discarded by global 3D classification without a mask nor alignment. After refinement of all good particles for Pol I and Pol I*, additional classification steps were performed to increase the resolvability of the active site. For Pol I* particles, a 3D classification step with a mask on the core and DNA-RNA hybrid yielded a class (182,488 particles) with a better density for GMPCPP, which could be refined to 3.18 Å resolution. An apo form of Pol I* consisting of 73,660 particles was obtained during a global classification step of the initial subset with a closed cleft and strong DNA-RNA density, and was refined to 3.21 Å resolution. For the pooled Pol I particles, a global 3D classification step yielded a class with a closed clamp (EC) and a class bound to DNA-RNA with a slightly more open clamp. The latter was classified one more round, which gave a class in an EC conformation. These particles were merged with the EC particles from the previous 3D classification step, refined (consensus Pol I EC) and classified with a mask on the core, the full DNA-RNA scaffold and the linker helix of A49, which yielded a class with strong GMPCPP density (30,232 particles) that was refined to 3.42 Å resolution. As both Pol I EC and Pol I* EC reconstructions were very similar in the active site, EC particles were merged and classified using different masks. Masked classification based on the full DNA scaffold and rudder produced one class (34,475 particles) with improved density for the upstream DNA duplex and revealing the path of the single stranded non-template strand (ssNT), which was refined to 4.0 Å resolution (without post-processing). Classification based on the core and DNA-RNA scaffold revealed different states differing in the width of the cleft, base flipping at position +2, presence of the GMPCPP and conformation of the trigger loop (shown in Figure 5—figure supplement 1). One of these classes (Pol I (core) EC +GMPCPP), which showed better density for GMPCPP, the +2 base and A190 L1202 was refined to 3.18 Å resolution (54,017 particles). Local resolution was estimated with Blocres (Cardone et al., 2013).

Model building and refinement

Previous Pol I structures in its apo (PDB: 4c3i and 4c2m) and elongating (PDB: 5m5x) forms were used as starting models. The initial placement of GMPCPP in the active site was based on its position in a Pol II EC with bound GMPCPP (Wang et al., 2006) (PDB: 2e2j and 4a3j). Initially, the model for the Pol I (core) EC (+GMPCPP) was built in COOT (Emsley and Cowtan, 2004) and real-space refined in PHENIX (Adams et al., 2010). This model was then rigid body fitted in the Pol I* or Pol I EC (+GMPCPP) maps in UCSF Chimera, further adjusted in COOT, and real-space refined again in PHENIX. For Pol I*, residues 66–125 from A12.2 were taken from the apo crystal structure (PDB: 4c3i), fitted to the density and manually adjusted. The A12.2 linker region was deleted afterwards. Agreement between maps and models was estimated in PHENIX. Model quality was assessed with Molprobity (Chen et al., 2010).

Expression, purification and labeling of recombinant A49-34.5

The cDNA of S. cerevisiae of rpa49 and rpa34 was codon-optimized for bacterial expression hosts and synthesized by GenScript. The two genes were cloned into separate ORFs in a pRSF Duet expression vector (Novagen) for co-expression. Codons for native cysteine residues were exchanged for alanine by mutagenesis PCR. Another mutation in A49 was introduced resulting in A140C to introduce a fluorescent label at this position. The construct was expressed in E. coli BL21 (DE3) Star in TB media by incubation with shaking at 37°C until an OD600nm of 0.8 was reached. The temperature was shifted to 18°C and expression was induced by addition of 0.05 mM IPTG at an OD600nm of 1 to 1.2. After 16 hr, cells were harvested by centrifugation. Cells were lysed using an enzymatic-chemical approach by resuspending in a buffer containing lysozyme, DNaseI and Triton-X 0.1% in 50 mM Tris (pH 7.5), 300 mM NaCl, 10 mM MgCl2, 10 mM β-mercapto ethanol, and 5 mM imidazole. The mixture was stirred at 4°C for 2–4 hr. The lysate was cleared by centrifugation (45,000 g for 90 min at 4°C) and the supernatant incubated with 5 to 10 mL Ni-NTA beads (QIAGEN) while rotating for 1 hr at 4°C. The beads were collected by gravity flow in a Biorad column and washed with 100 mL of washing buffer (50 mM Tris (pH 7.5), 500 mM NaCl, 10 mM β-mercapto ethanol, and 10 mM imidazole). Bound protein was eluted with 10–20 mL elution buffer (50 mM Tris (pH 7.5), 300 mM NaCl, 10 mM β-mercapto ethanol, and 300 mM imidazole). The elution fraction was dialyzed overnight against SP Buffer A (50 mM Tris (pH 7.5), 100 mM NaCl, 10 mM DTT). The next day, the protein solution was loaded onto a 5 mL HiTrap SP column (GE Healthcare) and eluted into 1 mL fractions with a 10 CV gradient from 100 to 1000 mM NaCl in 50 mM Tris (pH 7.5) with 10 mM DTT. Elution fractions of the major peak were analyzed by SDS-PAGE, combined, concentrated, and loaded onto a Superdex 200 (120 mL, GE Healthcare) equilibrated in 25 mM HEPES (pH 7.4), 150 mM NaCl, and 0.5 mM TCEP. The peak fractions were analyzed by SDS-PAGE, combined and concentrated. Protein identity was confirmed by mass spectrometry. The purified heterodimer was directly labeled with maleimide-functionalized Alexa Fluor 594 that was freshly dissolved at 10 mM in DMSO. The dye was added slowly to the protein solution with a final ratio of in 1:10 (protein:dye). The mixture was incubated overnight in the dark while shaking (800 rpm) at 4°C. The reaction was quenched by addition of 10 mM DTT and unreacted dye molecules were removed by size-exclusion chromatography (Superdex 200, 24 mL, GE Healthcare) equilibrated in reconstitution buffer (50 mM ammonium sulfate, 25 mM HEPES (pH 7.4), 10 mM MgCl2, 10 mM DTT). Labeling efficiency was determined by UV-VIS measurements of protein (280 nm) and dye absorbance.

Expression and purification of A12.2C

The part of cDNA of S. cerevisiae rpa12 coding for the A12.2 C-terminal domain (residues 79 to 125) was cloned into a pET24a expression vector with an N-terminal 6xHis tag followed by a TEV cleavage site. The construct was expressed in E. coli BL21 (DE3) Star in TB media by shaking at 37°C until an OD600nm of 0.8 was reached. Expression was induced by adding 0.5 mM IPTG and continued at 37°C for 4 hr. Cells were harvested, re-suspended in lysis buffer (50 mM Tris (pH 8), 500 mM NaCl, 10 mM β-mercaptoethanol, 5 mM imidazole, pH 8), and lysed by sonication. The cleared lysate was incubated with Ni-NTA beads (QIAGEN) for 1 hr at 4°C. Beads were washed with 50 mM Tris (pH 8), 500 M NaCl, 10 mM β-mercapto ethanol, and 10 mM imidazole and incubated in 30 mL wash buffer with 1.5 mg of TEV overnight. The cleaved protein was concentrated to about 2 mL and loaded onto a Superdex 75 (GE Healthcare) equilibrated in 50 mM HEPES (pH 7.5), 150 mM NaCl, and 0.5 mM TCEP. The major peak was collected, analyzed by SDS-PAGE, combined and concentrated. Protein identity was confirmed by mass spectrometry.

Expression and purification of Pol I*

The yeast strain Y2670 harboring Pol I Δrpa49 was generously provided by Herbert Tschochner (Universität Regensburg) (Pilsl et al., 2016). The mutant strain was expressed and purified analogous to the wild type Pol I yielding pure Pol I*.

Fluorescence polarization measurements

Purified Pol I* was incubated with labeled heterodimer at different concentrations overnight at 4°C in 150 mM ammonium sulfate, 15 mM HEPES (pH 7.5) and 10 mM DTT. For measurements using the Pol I* EC, Pol I* was incubated with an equimolar concentration of the same transcription scaffold used for the cryo-EM data for 1 hr at 4°C, previous to the overnight incubation. For the experiments using the A12.2C, recombinant A12.2C was incubated with the labeled Pol I EC for 30 min at room temperature.

Fluorescence polarization of A49(A140C)−34.5 heterodimer labeled with Alexa Fluor 594 was measured on a Jasco FP-6000 fluorometer equipped with polarization filters in a 150 μL volume with a final concentration of 100 nM of the labeled species. Fluorescence intensities at different polarization angles were measured at 594 nm excitation (2.5 nm bandwidth) and 625 nm emission (10 nm bandwidth) wavelengths. The anisotropy was calculated for the free and bound heterodimer by using an excess of Pol I* bound to DNA.

Accession numbers

Models have been deposited in the PDB with codes: 6HKO (Pol I EC +GMPCPP), 6HLQ (Pol I* EC +GMPCPP), 6HLR (Pol I (core) EC +GMPCPP), and 6HLS (apo Pol I*). Cryo-EM maps have been deposited in the EMDB with codes: EMD-0238 (Pol I EC +GMPCPP), EMD-0239 (Pol I* EC +GMPCPP), EMD-0240 (Pol I (core) EC +GMPCPP), EMD-0241 (apo Pol I*) and EMD-0242 (Pol I EC +GMPCPP (upstream DNA focused)).

Acknowledgements

YS, LT, RW and CWM acknowledge support by the ERC Advanced Grant (ERC-2013-AdG340964-POL1PIC). LT acknowledges support by the EMBL International PhD program. JH acknowledges EMBO for a postdoctoral long-term fellowship (ALTF 372–2017). We thank Matthias Vorländer, Florence Baudin, Herman KH Fung, Kathryn Perez and Vladimir Rybin for advice and discussion.

Funding Statement

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Contributor Information

Christoph W Müller, Email: christoph.mueller@embl.de.

John Kuriyan, University of California, Berkeley, United States.

Cynthia Wolberger, Johns Hopkins University School of Medicine, United States.

Funding Information

This paper was supported by the following grants:

  • European Commission ERC-2013-AdG340964-POL1PIC to Lucas Tafur, Yashar Sadian, Jonas Hanske, Rene Wetzel, Christoph W Müller.

  • European Molecular Biology Organization ALTF 372-2017 to Jonas Hanske.

Additional information

Competing interests

No competing interests declared.

Author contributions

Data curation, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing—original draft.

Data curation, Formal analysis, Investigation, Visualization.

Data curation, Investigation, Writing—review and editing.

Responsible for wild type and mutant yeast fermentation and wild type and mutant RNA polymerase I purification.

Data curation, Investigation.

Supervision, Funding acquisition, Project administration, Writing—review and editing.

Additional files

Transparent reporting form
elife-43204-transrepform.docx (249.9KB, docx)
DOI: 10.7554/eLife.43204.014

Data availability

Coordinates and cryo-EM maps have been deposited with the PDB and EMDB, respectively.

The following datasets were generated:

Tafur L, Sadian Y, Weis F, Muller CW. 2018. Pol I (core) EC + GMPCPP. Electron Microscopy Data Bank. EMD-0240

Tafur L, Sadian Y, Weis F, Muller CW. 2018. Pol I (core) EC + GMPCPP. Protein Data Bank. 6HLR

Tafur L, Sadian Y, Weis F, Muller CW. 2018. Pol I EC + GMPCPP. Electron Microscopy Data Bank. EMD-0238

Tafur L, Sadian Y, Weis F, Muller CW. 2018. Pol I EC + GMPCPP. Protein Data Bank. 6HKO

Tafur L, Sadian Y, Weis F, Muller CW. 2018. Pol I* EC + GMPCPP. Electron Microscopy Data Bank. EMD-0239

Tafur L, Sadian Y, Weis F, Muller CW. 2018. Pol I* EC + GMPCPP. Protein Data Bank. 6HLQ

Tafur L, Sadian Y, Weis F, Muller CW. 2018. Apo Pol I*. Electron Microscopy Data Bank. EMD-0241

Tafur L, Sadian Y, Weis F, Muller CW. 2018. Apo Pol I*. Protein Data Bank. 6HLS

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Decision letter

Editor: Cynthia Wolberger1
Reviewed by: Alessandro Vannini

In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.

[Editors’ note: a previous version of this study was rejected after peer review, but the authors submitted for reconsideration. The first decision letter after peer review is shown below.]

Thank you for submitting your work entitled "Structural rearrangement of TFIIS- and TFIIE/TFIIF-like subunits in RNA polymerase I transcription complexes" for consideration by eLife. Your article has been reviewed by three peer reviewers and the evaluation has been overseen by a Reviewing Editor and a Senior Editor. The following individual involved in review of your submission has agreed to reveal their identity: Alessandro Vannini (Reviewer #3).

Our decision has been reached after consultation between the reviewers. Based on these discussions and the individual reviews below, we regret to inform you that your work in its present form will not be considered further for publication in eLife. However, as described below, we would be open to considering a new manuscript that takes into account the concerns and suggestions articulated below.

This paper describes cryo-EM studies of RNA polymerase I (Pol I) that provide new information on several aspects of the A49-A34.5 heterodimer, which is the functional equivalent of the Pol II TFs, TFIIE and TFFIIF, and A12.2, the functional equivalent of TFIIS. The new structures in this manuscript were obtained from a preparation of Pol I bound to an NTP analogue, GMPCPP, in the absence of MgCl2. By sorting and classifying subsets of particles, the authors were able to identify distinct species representing the Pol I elongation complex (EC) + GMPCPP, Pol I core EC + GMCPP, Apo Pol I* (i.e. lacking the A49-A34.5 heterodimer) and Pol I* EC + GMCPP. By comparing these structures with previous structure of all three eukaryotic RNA polymerases, the authors draw several new insights. One interesting observation is the relocation of the A49-A34.5 heterodimer, which is exchanged for the A12 subunit and thus promotes shift between a Pol I and Pol II-like conformation. The structures also provide additional details on the interaction of Pol I with dNTP and stabilization of the transcription bubble.

All three reviewers appreciated the technical quality of the structural studies and the implications of some of the findings, in particular the role of heterodimer exchange with A12. However, it was agreed that the manuscript in its current form is written in a way that is accessible to experts only and presumes a high degree of familiarity with the Pol I literature in particular, as well as with Pol II and III. A significant number of new structural observations are described, although they are not all well-connected in terms of the questions they are addressing. The dimer exchange model was considered one interesting aspect of this new set of structures; however, this model needs to account more explicitly for genetic data suggesting that the heterodimer is present throughout the gene. Additional experimental data supporting this model could also strengthen the paper. A significant revision and refocusing of the paper would be needed to make the work described here more appropriate for the general readership of eLife.

Reviewer #1:

This paper describes cryo-EM studies of RNA polymerase I (Pol I) that provide new information on several aspects of the A49-A34.5 heterodimer, which is the functional equivalent of the Pol II TFs, TFIIE and TFFIIF, and A12.2, the functional equivalent of TFIIS. There have been multiple structures determined of Pol I by cryo-EM and x-ray crystallography, including several by the senior author. The new structures in this manuscript were obtained from a preparation of Pol I bound to an NTP analogue, GMPCPP, in the absence of MgCl2. By sorting and classifying subsets of particles, the authors were able to identify distinct species representing the Pol I elongation complex (EC) + GMPCPP, Pol I core EC + GMCPP, Apo Pol I* (i.e. lacking the A49-A34.5 heterodimer) and Pol I* EC + GMCPP. By comparing these structures with previous structure of all three eukaryotic RNA polymerases, the authors draw several new insights. Chief among these is that binding of the A49-A34.5 heterodimer and A12.2C are incompatible, and thereby gate Pol I conformation shifts between a Pol I and Pol II-like conformation. There are also additional insights into the detailed interactions that stabilize the transcription bubble.

Overall, the manuscript reads like a collection of very detailed structural information on different Pol I conformations with no clear direction. The questions to be addressed are not framed clearly, so the reader is left largely with an unfocused catalogue of structural details without a clear sense of why they matter. The paper is also written in a way that will be accessible primarily to those who work on Pol I and are already familiar with the structure and prior publications. The complicated and unfortunate naming system of Pol I subunits (no fault of the authors) makes the manuscript an even tougher slog, especially without sufficient graphic overviews to guide the reader. In the end, while the authors speculate about the meaning of various observations, for example about the role of the heterodimer in regulating binding of elongation factors, these ideas are not tested experimentally. While these new structures will be of interest to specialists, there are insufficient insights that will be of interest to the general readership of eLife.

Specific points:

The interpretation of additional density (Figure 1—figure supplement 3) that "connects to the DML toward the A12.2 linker,” etc.) is not at all convincing or even clear as described. It is difficult to tell where in the structure the density is located due to poor labeling. Moreover, the legend states that this density is only visible at low contour levels. Without further clarification of which residues this might correspond to, this should be omitted.

Reviewer #2:

In this manuscript, the authors bring forward some detailed, reasonably high resolution models for positions in and around the active site of RNA polymerase I. They present a catalogue of comparisons to the other eukaryotic polymerases as well as the prokaryotic enzyme. The big picture is really quite compelling: These enzymes work via the same mechanism, but they have apparently evolved unique properties that are reflected in the structures. Big picture: I like the manuscript.

There are some significant challenges, though, that can be addressed and would make the story seem better supported.

1) The authors have a robust understanding of the three polymerases, but their comparison of similarities could use more clarification for the non-expert reader.

2) This is a feature of structural biology, but this manuscript seems to make substantial mechanistic conclusions from the structural models. This was particularly evident in the section describing the motions in the +1 and +2 positions. Throughout, there should be more care taken to indicate that these are hypotheses.

3) Does the heterodimer really leave the enzyme during transcription? The manuscript draws on some literature, but is this a well-supported view? Can there be tests?

4) A commentary on dynamics might be interesting. The discussion of these subunits being exclusive, then swapping with transcription factors (Discussion section) would benefit from a consideration of dynamics in solution (or in living cells).

Reviewer #3:

The manuscript by Tafur et al. reports cryo-EM structures of elongating RNA Polymerase I in presence of a nucleotide analog, revealing that the heterodimer A49-A34.5 has been expelled from the complex and the C-term domain of A12.2 is relocated on a surface that was previously occupied by the heterodimer.

The cryo-EM reconstructions are impeccable and of high quality. The main conclusions are justified by the structural findings. This is potentially very interesting also because a form of the enzyme lacking A49-34.5 has been identified in vivo.

I strongly support publication, upon minor textual revisions and clarification of few minor issues

1) Can the author monitor "ejection" of the heterodimer upon incubation with GMPCC? This would be nice to see. This could probably be done on immobilised DNA beads and the composition of Pol I monitored over time in presence of GMPCC.

2) Results section “Cryo-EM structures of the GMPCPP-bound Pol I elongation complex (EC): this is then the authors elaborate on how this apo PolI* is formed? And why wasn't this observed in previous reconstruction of elongating Polymerase I? Could it be that Pol I* is a less stable elongation complex and partially loses the DNA scaffold after ejection of the heterodimer? I think that an in vitro assay (and possibly re-titrating in recombinant heterodimer) would add a lot to the manuscript and help understand the mechanism.

3) The first paragraph of the Introduction is difficult to follow maybe rewrite more concisely?

4) In the Accession numbers section, are the complexes named properly? Which one is the apo PolI*?

5) Last sentence of the Discussion: the fact the general catalytic mechanism in RNA Polymerase is conserved was already clear. I would suggest that the molecular determinants/regions involved in catalysis performs similar functions and this is now successfully proven.

[Editors’ note: what now follows is the decision letter after the authors submitted for further consideration.]

Thank you for submitting your article "The cryo-EM structure of a 12-subunit variant of RNA polymerase I reveals dissociation of the A49-A34.5 heterodimer" for consideration by eLife. Your article has been reviewed by three peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation has been overseen by John Kuriyan as the Senior Editor. The following individual involved in review of your submission has agreed to reveal his identity: Alessandro Vannini (Reviewer #2).

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

Summary:

This is a revision of a previous submission describing cryo-EM studies of RNA polymerase I (Pol I). The revised manuscript is improved in that it focuses on exchange of the A49-A34.5 heterodimer and A12.2, and the differences between Pol I and Pol I*. By sorting and classifying subsets of particles, the authors were able to identify distinct species of Pol I core and Pol I*. Chief among the conclusions is that binding of the A49-A34.5 heterodimer and A12.2C are incompatible, and thereby gate Pol I conformation shifts between a Pol I and Pol II-like conformation. The addition of binding data on A49-A34.5 heterodimer and A12.2C is, in principle, a positive addition and was requested in the previous review. However, there were issues with the binding data that must first be addressed before the manuscript is ready for publication.

Essential revisions:

1) The binding curves and analysis are problematic. According to the methods section, labeled heterodimer was present at 100 nM concentration in the binding studies. If this was indeed the concentration, the equation for the binding isotherm is not valid, as this approximation assumes that the labeled species is present at much lower concentration than Pol I. Either the binding data should be re-analyzed with a quadratic binding equation or the experiments should be repeated at much lower labelled heterodimer concentration.

2) The proposal that sequential binding of the N- and C-terminal domains of the heterodimer cannot explain apparent cooperativity, as the experiment only monitors binding of single monomers. It is possible that the analysis of the data as suggested above, a more complex model or a repeat of the experiment under different conditions could sort this out.

3) There was not enough information given to evaluate the competition/exchange experiment. If the heterodimer indeed binds with a 14 nM KD and the complex with Pol I was present at 100 nM concentration, almost all of the proteins would be in a complex. Exchange with A12.2C would depend upon the off rate of A49-A34.5 and the on rate for A 12.2C. The complex was incubated with A12.2C for 30 minutes. Were longer time points measured? What is the KD of A12.2C for Pol I? Might there be different conditions under which dimer exchange would be observed? The ultimate conclusion, that the heterodimer must first dissociate in order for A12.2C to bind is surely correct in light of the structure, but it is important that the paper has binding data that show this. A good control for the experiment would be exchange with unlabeled heterodimer, as this would provide a good benchmark. Similarly, the authors should consider repeating the GMCPP experiment at lower concentration of Pol I* – heterodimer complex, as it is possible the complex would dissociate if the concentrations were not so far above the (apparent) KD.

eLife. 2019 Mar 26;8:e43204. doi: 10.7554/eLife.43204.033

Author response

[Editors’ note: the author responses to the first round of peer review follow.]

[…] All three reviewers appreciated the technical quality of the structural studies and the implications of some of the findings, in particular the role of heterodimer exchange with A12. However, it was agreed that the manuscript in its current form is written in a way that is accessible to experts only and presumes a high degree of familiarity with the Pol I literature in particular, as well as with Pol II and III. A significant number of new structural observations are described, although they are not all well-connected in terms of the questions they are addressing. The dimer exchange model was considered one interesting aspect of this new set of structures; however, this model needs to account more explicitly for genetic data suggesting that the heterodimer is present throughout the gene. Additional experimental data supporting this model could also strengthen the paper. A significant revision and refocusing of the paper would be needed to make the work described here more appropriate for the general readership of eLife.

We would like to thank all reviewers for their comments and suggestions. We believe that their input has significantly improved our original manuscript. The revised manuscript now focuses on the exchange of the A49-A34.5 heterodimer with the C-terminal domain of subunit A12.2 (A12.2C) as observed in the Pol I* structure, and we discuss our structural findings in the context of known biochemical and genetic data. In addition, we have developed an assay to track the binding and dissociation of the heterodimer to the Pol I core by fluorescence polarization. We observe that under our experimental conditions, the heterodimer binds with high affinity and cannot be displaced by either free A12.2C or GMPCPP. Thus, binding of A12.2C can only occur once the A49-A34.5 heterodimer has been released from the enzyme. Moreover, the features of the active site (GMPCPP binding, and interactions with the non-template strand) are now discussed in the context of the Pol I* variant. Finally, we have revised our figures and regrouped them to increase the clarity of the paper. A detailed answer to each of the reviewer’s comments is listed below.

Reviewer #1:

[…] Overall, the manuscript reads like a collection of very detailed structural information on different Pol I conformations with no clear direction. The questions to be addressed are not framed clearly, so the reader is left largely with an unfocused catalogue of structural details without a clear sense of why they matter. The paper is also written in a way that will be accessible primarily to those who work on Pol I and are already familiar with the structure and prior publications. The complicated and unfortunate naming system of Pol I subunits (no fault of the authors) makes the manuscript an even tougher slog, especially without sufficient graphic overviews to guide the reader. In the end, while the authors speculate about the meaning of various observations, for example about the role of the heterodimer in regulating binding of elongation factors, these ideas are not tested experimentally. While these new structures will be of interest to specialists, there are insufficient insights that will be of interest to the general readership of eLife.

Following the criticisms of the reviewer, we have re-written the manuscript that now focuses on the A49-A34.5 heterodimer exchange with the C-terminal domain of subunit A12.2 (A12.2C) as observed in the Pol I* structure. We have also added experimental data in vitro that helps understanding the dynamics of the binding of the A49-A34.5 heterodimer and A12.2C. Additionally, we have revised the figures to increase the clarity of the manuscript (for example, we have included the Pol I* and Pol I structures in Figure 1 to facilitate comparison and we included a legend indicating the colored subunits and their relationship with Pol II to guide the reader). We believe that our revised manuscript will be of interest not only to Pol I specialists, but also to the broad readership of eLife interested in the conformational dynamics of protein complexes and transcription. Furthermore, the manuscript highlights the power of cryo-EM to resolve distinct intermediates in biochemically homogenous samples.

Specific points:

The interpretation of additional density (Figure 1—figure supplement 3) that "connects to the DML toward the A12.2 linker,” etc.) is not at all convincing or even clear as described. It is difficult to tell where in the structure the density is located due to poor labeling. Moreover, the legend states that this density is only visible at low contour levels. Without further clarification of which residues this might correspond to, this should be omitted.

As suggested by the reviewer, we have now omitted this figure and the corresponding discussion.

Reviewer #2:

[…] There are some significant challenges, though, that can be addressed and would make the story seem better supported.

1) The authors have a robust understanding of the three polymerases, but their comparison of similarities could use more clarification for the non-expert reader.

We have expanded the Introduction to include more background information about the Pol I structure compared to Pol II and Pol III. We also have modified figures and text to improve readability and clarity for non-expert users.

2) This is a feature of structural biology, but this manuscript seems to make substantial mechanistic conclusions from the structural models. This was particularly evident in the section describing the motions in the +1 and +2 positions. Throughout, there should be more care taken to indicate that these are hypotheses.

In the revised version of the manuscript, we have greatly reduced the section describing the motions of the +1 and +2 nucleotides, and only briefly mention these features of the active site of both Pol I and Pol I*. Throughout the revised manuscript, we now avoid making substantial conclusions from the structural models, but rather put them into context of the known literature.

3) Does the heterodimer really leave the enzyme during transcription? The manuscript draws on some literature, but is this a well-supported view? Can there be tests?

To our knowledge, there is no conclusive in vivo evidence that shows that the heterodimer stays or leaves the complex during transcription. The current view is that the heterodimer functions as a built-in general transcription factor with roles in transcription initiation and/or (early) elongation. For the PAF53/PAF49 there is also solid in vivo evidence that the interaction with Pol I is regulated and that starvation leads to the dissociation of the heterodimer from Pol I (Penrod et al., 2015). However, this does not prove that the heterodimer binds and dissociates during the transcription cycle. in vitro, the A49-A34.5 heterodimer is required for promoter-dependent transcription initiation, although the C-terminal A49 tWH domain is sufficient to restore this activity (Pilsl et al., 2016). We have now included additional experimental data that also show that the heterodimer binds with high affinity to Pol I in vitro, and cannot be displaced by the isolated A12.2C or by GMPCPP.

4) A commentary on dynamics might be interesting. The discussion of these subunits being exclusive, then swapping with transcription factors (Discussion section) would benefit from a consideration of dynamics in solution (or in living cells).

We have now included some in vitroexperimental evidence on the dynamics of heterodimer binding to Pol I by fluorescence polarization assays. Whereas we can specifically track the binding and release of the heterodimer from the Pol I core, it couldn’t be displaced from the complex by either free A12.2C or GMPCPP (Figure 3). Although we could not identify conditions (or factors) that allow dissociation of the heterodimer, the heterodimer has to dissociate from the Pol I core to allow A12.2C binding to the Rpb9-like site.

Reviewer #3:

[…] I strongly support publication, upon minor textual revisions and clarification of few minor issues

1) Can the author monitor "ejection" of the heterodimer upon incubation with GMPCC? This would be nice to see. This could probably be done on immobilised DNA beads and the composition of Pol I monitored over time in presence of GMPCC.

Taking into consideration the suggestions of this reviewer and reviewer #2 (see above), we used a fluorescence polarization assay to monitor binding and dissociation of the heterodimer from the Pol I core. Whereas this set up allowed us to track the specific binding of the heterodimer (Figure 3 and Figure 3—figure supplement 1), we could not observe any significant effect of GMPCPP or A12.2C on heterodimer dissociation from the Pol I core. Thus, based on these results, it is likely that the high proportion of Pol I* observed in our cryo-EM data set is due to other factors occurring during grid preparation such as a change in the salt concentration on the grid or eventually also the use of carbon-coated grids, which shifted the proportion of Pol I/Pol I* normally seen in solution. Whereas in vivo, we hypothesize that binding of elongation factors such as Spt4/Spt5 of Paf1 could expel the heterodimer.

2) Results section “Cryo-EM structures of the GMPCPP-bound Pol I elongation complex (EC): this is then the authors elaborate on how this apo PolI* is formed? And why wasn't this observed in previous reconstruction of elongating Polymerase I? Could it be that Pol I* is a less stable elongation complex and partially loses the DNA scaffold after ejection of the heterodimer? I think that an in vitro assay (and possibly re-titrating in recombinant heterodimer) would add a lot to the manuscript and help understand the mechanism.

The presence of apo Pol I* particles in this data set might parallel the proportion of Pol I not bound to DNA at this scaffold concentration (2-fold molar excess). A 2-fold molar excess of scaffold was also used by Neyer et al., who observed about 50% of the particles in an apo Pol I state (Neyer et al., Nature, 2016). We could also observe apo 14-subunit Pol I in our dataset, but did not analyze it further. In our previous data set (Tafur et al., Mol. Cell, 2016) we used a higher scaffold concentration (5-fold molar excess) presumably resulting in fewer apo 14-subunit Pol I particles. The lack of Pol I* particles observed in the previous data might result from the much lower total number of particles available for classification or from different experimental conditions (different salt concentration during grid preparation etc.). Nevertheless, we also observed Pol I classes lacking the heterodimer in the previous reconstruction of elongating Pol I (Tafur et al., Mol. Cell 2016), but no unassigned additional densities were observed and thus these particles were discarded.

3) The first paragraph of the Introduction is difficult to follow maybe rewrite more concisely?

The Introduction has been rewritten.

4) In the Accession numbers section, are the complexes named properly? Which one is the apo PolI*?

We thank the reviewer for pointing out that apo Pol I* had been not included. We now include all models and corresponding PDB/EMDB codes in the manuscript.

5) Last sentence of the Discussion: the fact the general catalytic mechanism in RNA Polymerase is conserved was already clear. I would suggest that the molecular determinants/regions involved in catalysis performs similar functions and this is now successfully proven.

The reviewer is right and in the revised version of the manuscript this sentence has been deleted.

[Editors' note: the author responses to the re-review follow.]

Essential revisions:

1) The binding curves and analysis are problematic. According to the methods section, labeled heterodimer was present at 100 nM concentration in the binding studies. If this was indeed the concentration, the equation for the binding isotherm is not valid, as this approximation assumes that the labeled species is present at much lower concentration than Pol I. Either the binding data should be re-analyzed with a quadratic binding equation or the experiments should be repeated at much lower labelled heterodimer concentration.

We agree with the reviewer that analyzing our binding data is problematic given that heterodimer and Pol I were present at similar concentrations (100 nM heterodimer, 0 to 100 nM Pol I*). Following the suggestion of the reviewer, we tried to fit the data using a quadratic binding equation, but have not be able to converge to a reliable solution suggesting that the binding mode is indeed more complex than we thought. As we have no further information on the number of binding sites and the sequential binding events, we want to refrain from applying a more complex binding model. Alternatively, we have also tried to reduce the heterodimer concentration in the fluorescence polarization experiments. However, reducing the heterodimer concentration led to a very faint fluorescent signal with poor signal-to-noise.

Because the primary objective of our binding experiments has been to compare binding among the different complexes, we therefore suggest to no longer report KD and Hill coefficient. Instead, we just show in Figure 3B the normalized change of anisotropy upon binding of the heterodimer to Pol I* or to wild type Pol I at a given concentration without trying to deduce any quantitative binding parameters. In addition, we also show the inability to displace the heterodimer by an excess of A12.2C or GMPCPP.

In the revised version of the manuscript, the paragraph describing the results in Figure 3B now states: “To test these hypotheses, we performed a series of fluorescence anisotropy experiments, using recombinant heterodimer, where a cysteine has been introduced in the A49 linker region for labelling with Alexa Fluor 594, and endogenously purified Pol I* (Pilsl et al., 2016) incubated with DNA (Pol I * EC) (Figure 3A). […] Because a 1:1 binding model did not allow fitting the data no attempt was made to introduce more complex binding models.”

2) The proposal that sequential binding of the N- and C-terminal domains of the heterodimer cannot explain apparent cooperativity, as the experiment only monitors binding of single monomers. It is possible that the analysis of the data as suggested above, a more complex model or a repeat of the experiment under different conditions could sort this out.

As outlined in our response to point 1), we have removed the KD, Hill coefficient and no longer discuss the cooperativity of binding in the text.Accordingly, we have deleted in the revised version of the manuscript our statement: “Such behavior can be explained by the sequential binding of the N-terminal and C-terminal domains where anchoring of the A49-A34.5 heterodimer to Pol I by the N-terminal dimerization domains promotes the binding of the C-terminal A49 tWH domain (Figure 3B).”

3) There was not enough information given to evaluate the competition/exchange experiment. If the heterodimer indeed binds with a 14 nM KD and the complex with Pol I was present at 100 nM concentration, almost all of the proteins would be in a complex. Exchange with A12.2C would depend upon the off rate of A49-A34.5 and the on rate for A 12.2C. The complex was incubated with A12.2C for 30 minutes. Were longer time points measured? What is the KD of A12.2C for Pol I? Might there be different conditions under which dimer exchange would be observed? The ultimate conclusion, that the heterodimer must first dissociate in order for A12.2C to bind is surely correct in light of the structure, but it is important that the paper has binding data that show this. A good control for the experiment would be exchange with unlabeled heterodimer, as this would provide a good benchmark. Similarly, the authors should consider repeating the GMCPP experiment at lower concentration of Pol I* – heterodimer complex, as it is possible the complex would dissociate if the concentrations were not so far above the (apparent) KD.

We performed the competition assay to test whether excess of A12.2C could compete for the heterodimer binding site, which we saw not to be true. As proposed by the reviewer we also performed the incubation overnight and did not see a difference. As these experiments were only performed once, we did not include them into Figure 3C, but now mention these results without showing the data. We agree with the reviewer that it would be desirable obtaining binding data for A12.2C. However, A12.2C is only a small domain (Mr = 10 kDa) that presumably will not easily displace the bound heterodimer. Obtaining reliable binding data for A12.2C to the Pol I* core is experimentally challenging and in our opinion beyond the scope of this manuscript. In the revised we now state: “Incubation of the sample Pol I*/A49-A34.5 sample with recombinant A12.2C (residues 79 to 125) for 30 min and overnight (data not shown) did not reduce the anisotropy (indicating the release of the heterodimer from Pol I) even at 50-fold molar excess (Figure 3C).”

Associated Data

    This section collects any data citations, data availability statements, or supplementary materials included in this article.

    Data Citations

    1. Tafur L, Sadian Y, Weis F, Muller CW. 2018. Pol I (core) EC + GMPCPP. Electron Microscopy Data Bank. EMD-0240
    2. Tafur L, Sadian Y, Weis F, Muller CW. 2018. Pol I (core) EC + GMPCPP. Protein Data Bank. 6HLR
    3. Tafur L, Sadian Y, Weis F, Muller CW. 2018. Pol I EC + GMPCPP. Electron Microscopy Data Bank. EMD-0238
    4. Tafur L, Sadian Y, Weis F, Muller CW. 2018. Pol I EC + GMPCPP. Protein Data Bank. 6HKO
    5. Tafur L, Sadian Y, Weis F, Muller CW. 2018. Pol I* EC + GMPCPP. Electron Microscopy Data Bank. EMD-0239
    6. Tafur L, Sadian Y, Weis F, Muller CW. 2018. Pol I* EC + GMPCPP. Protein Data Bank. 6HLQ
    7. Tafur L, Sadian Y, Weis F, Muller CW. 2018. Apo Pol I*. Electron Microscopy Data Bank. EMD-0241
    8. Tafur L, Sadian Y, Weis F, Muller CW. 2018. Apo Pol I*. Protein Data Bank. 6HLS

    Supplementary Materials

    Transparent reporting form
    elife-43204-transrepform.docx (249.9KB, docx)
    DOI: 10.7554/eLife.43204.014

    Data Availability Statement

    Coordinates and cryo-EM maps have been deposited with the PDB and EMDB, respectively.

    The following datasets were generated:

    Tafur L, Sadian Y, Weis F, Muller CW. 2018. Pol I (core) EC + GMPCPP. Electron Microscopy Data Bank. EMD-0240

    Tafur L, Sadian Y, Weis F, Muller CW. 2018. Pol I (core) EC + GMPCPP. Protein Data Bank. 6HLR

    Tafur L, Sadian Y, Weis F, Muller CW. 2018. Pol I EC + GMPCPP. Electron Microscopy Data Bank. EMD-0238

    Tafur L, Sadian Y, Weis F, Muller CW. 2018. Pol I EC + GMPCPP. Protein Data Bank. 6HKO

    Tafur L, Sadian Y, Weis F, Muller CW. 2018. Pol I* EC + GMPCPP. Electron Microscopy Data Bank. EMD-0239

    Tafur L, Sadian Y, Weis F, Muller CW. 2018. Pol I* EC + GMPCPP. Protein Data Bank. 6HLQ

    Tafur L, Sadian Y, Weis F, Muller CW. 2018. Apo Pol I*. Electron Microscopy Data Bank. EMD-0241

    Tafur L, Sadian Y, Weis F, Muller CW. 2018. Apo Pol I*. Protein Data Bank. 6HLS

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