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Distinct effects of DNA lesions on RNA synthesis by Escherichia coli RNA polymerase
Biochemical and Biophysical Research Communications 510 (2019) 122e127
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Biochemical and Biophysical Research Communications journal homepage: www.elsevier.com/locate/ybbrc
Distinct effects of DNA lesions on RNA synthesis by Escherichia coli RNA polymerase Danil Pupov 1, Artem Ignatov 1, Aleksei Agapov, Andrey Kulbachinskiy* Institute of Molecular Genetics, Russian Academy of Sciences, Moscow, 123182, Russia
a r t i c l e i n f o
a b s t r a c t
Article history: Received 3 January 2019 Accepted 12 January 2019 Available online 18 January 2019
DNA lesions can severely compromise genome stability and lead to cell death if unrepaired. RNA polymerase (RNAP) is known to serve as a sensor of DNA damage and to attract DNA repair factors to the damaged template sites. Here, we systematically investigated the ability of Escherichia coli RNAP to transcribe DNA templates containing various types of DNA lesions, and analyzed their effects on transcription fidelity. We showed that transcription is strongly inhibited on templates containing cyclobutane thymine dimers, 1,N6-ethenoadenine and abasic sites, while 8-oxoguanine and thymine glycol have mild effects on transcription efficiency. Similarly to many polymerases, E. coli RNAP follows the “A” rule during nucleotide insertion opposite abasic sites and bulky lesions, and can also incorporate and efficiently extend an adenine nucleotide opposite 8-oxoguanine. Mutations in RNAP regions around the templating nucleotide decrease the efficiency of translesion synthesis, likely by altering the RNAPtemplate contacts in the active site. Thus, DNA lesions can lead to distinct outcomes in transcription, depending on the severity of the damage and contacts of the damaged template with the active site of RNAP. © 2019 Elsevier Inc. All rights reserved.
Keywords: RNA polymerase Translesion RNA synthesis Transcription fidelity Transcription-coupled repair
1. Introduction Genomic DNA in all organisms is continuously modified by multiple endogenous and exogenous factors. Some of these modifications are introduced by specialized enzymes and serve for regulation of gene expression, DNA repair, or defense against foreign DNA. Others appear spontaneously or are introduced by genotoxic agents and may impair various genetic processes and lead to genome instability. In particular, many DNA lesions are known to affect replication dramatically, leading to the replication fork stalling and mutagenesis. Most organisms possess specialized DNA polymerases that are able to bypass certain types of lesions but usually have a much lower replication fidelity [1,2]. Transcription can also be impaired by lesions in the template
Abbreviations: RNAP, RNA polymerase; CPD, cyclobutane pyrimidine (thymine) dimer; εA, 1,N6-ethenoadenine; AP, apurinic/apyrimidinic site; 8oxoG, 8oxoguanine; TG, thymine glycol; TEC, transcription elongation complex; BH, bridge helix. * Corresponding author. Institute of Molecular Genetics, 2 Kurchatov sq., Moscow, 123182, Russia. E-mail address: [email protected] (A. Kulbachinskiy). 1 These authors contributed equally to this work. https://doi.org/10.1016/j.bbrc.2019.01.062 0006-291X/© 2019 Elsevier Inc. All rights reserved.
DNA strand, which may lead to stalling of RNA polymerase (RNAP) or to transcriptional mutagenesis, thus producing mutant RNAs and proteins [3e7]. At the same time, RNAP can act as a sensor for DNA lesions, by attracting the DNA repair machinery to the damaged template sites during transcription coupled repair (TCR) (reviewed in Ref. [8]). Furthermore, stalled transcription complexes can severely compromise genome stability by colliding with the replication machinery [9,10]. Bacterial cells contain a single RNAP, and the process of DNA damage recognition and repair should be highly coordinated with transcription to allow efficient gene expression and DNA replication. However, the ability of bacterial RNAP to transcribe damaged DNA templates has not been systematically studied. Only a handful of lesions have been analyzed in the bacterial transcription system in vitro. In particular, it was shown that similarly to DNA polymerases, the abasic site and 8-oxoguanine promote mutagenic insertion of ATP in the RNA transcript, with transient RNAP pausing [4e6]. At the same time, the molecular mechanisms of translesion RNA synthesis remain poorly understood for most types of damaged nucleotides. In this work, we directly compared the effects on transcription by E. coli RNAP of several types of lesions commonly found in the genomic DNA: thymine dimer (CPD, cyclobutane pyrimidine dimer); 1,N6-ethenoadenine (εA); abasic site (AP); 8-oxoguanine
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(8oxoG), and thymine glycol (TG). We for the first time demonstrated that CPD and εA severely inhibit the activity of bacterial RNAP in vitro, allowing only very slow incorporation of ATP (the same specificity as observed for the AP-site), while TG has only a weak effect on transcription. Furthermore, we showed that 8oxoG is highly mutagenic because it enables not only insertion of a noncomplementary ATP but also its efficient extension by RNAP. Finally, we studied the effects of amino acid substitutions in the active site of E. coli RNAP on transcription of damaged DNA and identified mutations with altered transcription efficiency.
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annealed oligonucleotides (10 nM final RNA concentration) and incubated for 10 min at 20 C or 37 C, followed by NTP addition (100 mM each, unless otherwise indicated). The reactions were stopped after various time intervals, as indicated in the figures, and the RNA products were resolved by 18% denaturing PAGE, followed by phosphorimaging.
3. Results 3.1. Diverse effects of various types of DNA lesions on transcription
2. Materials and methods 2.1. Proteins and nucleic acids Wild-type and mutant core E. coli RNAPs were purified from Eco BL21(DE3) cells expressing all four core RNAP subunits from the plasmids pVS10 (for the K334A, T790A, Y795A and R798A mutants) or pIA679 (for the R542A mutant) as described [11]. The core RNAP mutations were obtained by site-directed mutagenesis. RNAPs were purified by Polymin P precipitation, Heparin-Sepharose, Ni2þaffinity and MonoQ anionic-exchange chromatography steps as described [11,12]. The s subunit was purified using the Ni2þ-affinity and DEAE ion-exchange chromatography [12]. DNA and RNA oligonucleotides used in the assays (Fig. 1) were purchased from DNA Synthesis, Syntol (Moscow, Russia), and TriLink BioTechnologies (San Diego, CA). 2.2. Analysis of RNA synthesis in vitro Transcription assays were performed in minimal transcription elongation complexes (TECs) assembled from synthetic DNA and RNA oligonucleotides and the E. coli core RNAP as described [13]. The 50 -labeled RNA transcript (0.5 mM final concentration) was mixed with the template (1 mM) and nontemplate DNA oligonucleotides (5 mM) in transcription buffer containing 40 mM Tris-HCl, pH 7.9, 40 mM NaCl and 10 mM MgCl2, heated to 65 C, and cooled down to 20 C at 1 /min. For the AP site, the template was assembled for 10 min at 37 C. Core RNAP (25 nM) was mixed with the
To analyze the effects of DNA lesions on transcription, we assembled minimal TECs from synthetic RNA and DNA oligonucleotides and the core enzyme of E. coli RNAP (Fig. 1). Such minimal templates were previously shown to form stable complexes with RNAP and mimic the principal RNAP-nucleic acids interactions in the natural TEC [13]. In most complexes, the RNA 30 -end was located immediately upstream of the damaged template base, so that the incoming nucleotide should be incorporated opposite the lesion; in the case of CPD, the RNA 30 -end was located 2 nt upstream of the lesion (Fig. 1B). The lengths of 50 -labeled starting RNAs in all complexes were 12 or 11 nt, as indicated in the figures. We also assembled control complexes containing undamaged DNA oligonucleotides of the same sequence (for the AP-site, we used the same control template as for the TG lesion). We first tested the ability of wild-type E. coli RNAP to extend RNA in these complexes in the presence of all four NTP substrates. In the case of control undamaged templates, RNA was readily extended by at least 7e10 nt, several nucleotides past the site where we intended to place the lesions (Fig. 2, left column). Although not all transcripts reached the end of the template (the expected RNA extension by 15e19 nucleotides, depending on the template) e because efficient RNA synthesis requires the presence of 14 nt downstream DNA duplex [13] e this allowed us to analyze the effects of the lesions on RNA extension opposite and immediately past the lesion. The presence of damaged nucleotides had significant and diverse effects on transcription, depending on the nature of the lesion. For the bulky CPD modification, we observed almost
Fig. 1. The structure of bacterial RNAP and composition of analyzed DNA templates. (A) The structure of the active site of T. thermophilus RNAP in the transcription elongation complex (2O5J [15]). DNA is orange, the templating nucleotide (þ1) is carmine, the incoming NTP is black, RNA is yellow, magnesium ions bound in the active site are magenta, the bridge helix (BH) is turquoise. Amino acid residues mutated in this study are shown. (B) Minimal templates containing analyzed DNA lesions. The damaged nucleotides are shown in bold italics and underlined. (For interpretation of the references to colour in this figure legend, the reader is referred to the Web version of this article.)
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before and after nucleotide insertion opposite the lesion, as evident from inefficient extension of the starting 12 nt and extended 13 nt RNA products in comparison with the control template (Fig. 2, WT RNAP in the third row). To detect possible defects in RNA synthesis caused by the nonbulky 8oxoG and TG lesions, which were expected to have milder effects on transcription, we used a lower reaction temperature (20 C vs. 37 C) and shorter reaction times in comparison with the first set of lesions. In the case of 8oxoG, RNAP stalling was observed after RNA extension by 2 nucleotides, one nucleotide past the lesion, suggesting that 8oxoG does not prevent nucleotide incorporation but may hamper further RNAP translocation, when located inside the RNA-DNA hybrid (Fig. 2, fourth row). In the case of TG, we observed weak but detectable RNAP stalling at the site of lesion (at RNA length of 13 nt) that was absent in the case of undamaged template (Fig. 2, bottom row). At the same time, most RNAs were efficiently extended past the lesion.
3.2. Transcription of damaged templates by RNAP variants with mutations in the active center
Fig. 2. Activities of wild-type (WT) and mutant RNAP variants on damaged DNA templates. Control reactions performed with the wild-type RNAP on corresponding undamaged templates are shown in the left column. The type of DNA lesion analyzed in each row is shown on the right. For the CPD, εA and AP templates, the reactions were performed at 37 C for total 2 h (the time points are 10, 30 , 100 , 300 , 600 , 1200 ). For the 8oxoG and TG templates, the reactions were performed at 20 C for 10’ (10”, 30”, 10, 20 , 40 and 100 time points). Positions of the starting 11 nt or 12 nt RNAs are shown on the left; positions of DNA lesions are indicated with arrowheads and asterisks on the right.
complete block of transcription when the RNA 30 -end reached the first damaged nucleotide (the 30 -T residue of the dimer), after rapid initial extension by one nucleotide (Fig. 2, wild-type RNAP in the first row). This block was slowly overcome after prolonged incubation, resulting in the insertion of a single nucleotide opposite the 30 -T of CPD, as evident from the accumulation of the 13 nt RNA product. Although its further extension was very inefficient, there was no accumulation of the next extension product, after incorporation of the second nucleotide opposite the 50 -T of CPD, and longer RNA products were observed as a result of read-through synthesis. Thus, there appears to be no strong barrier to elongation after incorporation of the two nucleotides opposite CPD. The 1,N6-ethenoadenine modification, which prevents WatsonCrick pairing with thymine, blocked RNA synthesis even stronger than the CPD lesion (Fig. 2, WT RNAP in the second row), with very little RNA extension observed even at the longest incubation time. Markedly, these read-through RNA products were inefficiently extended past the lesion, suggesting that it blocks further RNAP translocation. In comparison with CPD and εA, the AP site was relatively readily transcribed by RNAP, although pauses were observed both
To get insight into the recognition of damaged nucleotides in the active site of RNAP, we obtained mutant variants of E. coli RNAP with alanine substitutions of several amino acid residues involved in interactions with the transcribed DNA strand (Fig. lA): (i) substitution K334A in the switch2 region in the b0 subunit; K334 contacts the phosphate group of the þ1 template nucleotide and stabilizes the template DNA strand in the active site [14,15]; (ii) substitution R542A in the fork-loop 2 region in the b subunit; R542 contacts the þ1 phosphate and the þ2 base of the downstream template DNA [15]; (iii) substitution T790A in the b0 subunit bridge helix (BH); T790 is the closest to the þ1 templating base in the TEC structure [15]; (iv) substitution Y795A in the BH; this residue contacts the þ2 and þ 3 phosphates in the template strand [15]; (v) substitution R798A in the BH; this residue contacts the 1 and þ2 template phosphates [15]. All mutant RNAPs were generated by site-directed mutagenesis, expressed and purified from E. coli cells. Analysis of the activity of these RNAPs on a linear DNA fragment containing the T7A1cons promoter showed that they all were capable of promoter-dependent transcription initiation (Fig. S1). We further tested the activities of these mutants in the control minimal TECs containing undamaged template DNAs. It was shown that at 37 C (conditions used for analysis of the CPD, εA and AP lesions) the activities of all five mutant RNAPs on undamaged DNA were comparable to the wild-type RNAP, with most RNA primer extended at the first time point (10”; Fig. S2). At the same time, at 20 C (conditions used for analysis of the 8oxoG and TG lesions), the K334A, T790A, Y795A and R798A mutants revealed lower levels of activity (Fig. S2). In particular, the K334A and T790A mutants extended RNA primers with slower kinetics and synthesized shorter RNA products, likely as a result of altered RNAP-DNA contacts and impaired catalysis. We then analyzed RNA synthesis by the mutant RNAPs on damaged DNA templates. The activities of the R542A, Y795A and R798A mutants were similar to the wild-type RNAP on all tested templates (Fig. S3). At the same time, the K334A and especially T790A substitutions additionally impaired RNA extension on most damaged templates in comparison with the wild-type RNAP, even at 37 C (Fig. 2, compare two right columns with the WT column). For example, no RNA extension past the CPD lesion and much lower activity on the AP template were observed for the T790A RNAP. Interestingly, the K334A substitution increased RNA extension past CPD (Fig. 2, first row), possibly as a result of more efficient translocation of this bulky lesion into the active site of RNAP.
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3.3. Effects of DNA lesions on transcription fidelity Many DNA lesions are known to affect the fidelity of nucleotide incorporation by DNA polymerases, thus resulting in mutagenesis [2]. Similarly, nucleotide misincorporation by RNAP may produce mutated RNA variants. We therefore tested the effects of various DNA lesions on the fidelity of nucleotide incorporation by E. coli RNAP, by analyzing RNA extension in the presence of individual NTPs. We observed efficient incorporation of both CTP and ATP on the 8oxoG template, while only CTP (and with low efficiency UTP) was incorporated in the control reaction (Fig. 3A). This agrees with published data showing that 8oxoG allows incorporation of adenine nucleotides in addition to the complementary cytosine by various RNA and DNA polymerases [2,5e7]. The TG lesion allowed efficient incorporation of the complementary adenine residue, in agreement with its weak effects on transcription (Fig. 3B). Similarly, only ATP was incorporated in the case of the AP template, but with a much lower efficiency than in the case of the control TEC containing a thymine residue in the template strand (Fig. 3B). Only very weak ATP incorporation was observed in the case of the CPD and εA lesions, with no incorporation of other three nucleotides (Fig. S4). In the previous experiment, we observed efficient incorporation of both C and A opposite template 8oxoG. To establish which of these nucleotides can be preferably incorporated into the fulllength RNA, we compared the extension efficiencies for 12 nt RNA transcripts containing either C or A at their 30 -ends opposite undamaged G or 8oxoG. We revealed that the correct 30 C RNA was inefficiently extended past the lesion, resulting in a major pause one nucleotide downstream of it (Fig. 4, compare 30 C-8oxoG and 30 C-G reactions). In contrast, the misincorporated 30 A RNA could be efficiently extended on the 8oxoG template but not on the control undamaged template (compare the 30 A-8oxoG and 30 A-G reactions).
4. Discussion Here, we investigated the effects of several DNA lesions, which are well known to affect DNA replication, on the efficiency and fidelity of RNA synthesis by bacterial RNAP. We revealed significant variations in the ability of RNAP to transcribe damaged DNA templates depending on the type of the lesion, as described below, and identified amino acid residues possibly involved in translesion synthesis.
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CPD is the most common UV light-induced DNA lesion that severely inhibits DNA replication [2]. We demonstrated that CPD also strongly impairs transcription by bacterial RNAP, with very slow nucleotide insertion opposite both thymine dimer residues. Similarly to our observations, it was shown that nucleotide insertion opposite CPD by eukaryotic RNAP II is dramatically slowed down in comparison with undamaged templates [3,16e18]. This is likely explained by impaired RNAP translocation and may be accompanied by RNAP backtracking [18,19]. Furthermore, certain substitutions in the trigger loop of RNAP II were shown to stimulate translesion synthesis and cell survival after UV irradiation, possibly by promoting nucleotide insertion [19]. In contrast, our analyzed substitutions in the BH and switch2 of E. coli RNAP inhibited transcription. We revealed that another common lesion εA can completely block transcription, likely by disrupting complementary pairing with the incoming NTP. Although εA has never been studied with any RNAP, a similar guanine modification, 1,N2-ethenoguanine, could block RNA synthesis by eukaryotic RNAP II [20], suggesting that the effects of this type of modification are similar for bacterial and eukaryotic RNAPs. The AP-site was previously shown to induce transient RNAP pausing and ATP insertion by both E. coli and eukaryotic RNAPs, followed by further RNA extension [4,7]. For eukaryotic RNAP, the AP-site was recently demonstrated to slow down nucleotide incorporation both opposite and after the lesion [21]. We also observed pausing by bacterial RNAP before and after ATP insertion demonstrating that the mechanism of AP-site bypass is similar for these RNAPs. Interestingly, all three lesions that significantly inhibited transcription e CPD, εA and the AP site, e promoted insertion of adenine residues opposite the damaged nucleotide. Thus, bacterial RNAP seems to follow the so-called ‘A-rule’ for incorporation of nucleotides opposite AP-sites and bulky lesions, similarly to many DNA polymerases [22]. ATP is a preferred nucleotide for the active site of both bacterial [23] and eukaryotic [21] RNAPs, which likely promotes nontemplated ATP insertion. Indeed, RNAP II was shown to insert A opposite CPD in a nontemplated manner [19]. Other bulky lesions may be transcribed by RNAP in a similar way. 8oxoG is the most common oxidative DNA lesion that has been studied in both eukaryotic and bacterial transcription systems. When located in the template DNA strand, it promotes misincorporation of adenine and causes weak transcriptional pausing by both bacterial and eukaryotic RNAPs [5e7,24]. Misincorporation
Fig. 3. Fidelity of NTP incorporation opposite various DNA lesions by wild-type E. coli RNAP. (A) Single-nucleotide incorporation opposite 8-oxoguanine (right) or guanine in the corresponding control template (left). (B) Nucleotide incorporation opposite the AP-site (middle), thymine glycol (right), and corresponding control template (left). The reactions were performed for 3000 at 20 C with each of the four NTPs taken at indicated concentrations.
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Fig. 4. The efficiency of RNA extension after 8-oxoguanine depending on the RNA 3′-end structure. The TECs contained 12 nt RNA with the 30 -end (either C or A) located opposite either undamaged guanine (G) or 8-oxoguanine (8oxoG). Transcription was performed at 20 C for 10’ (10”, 30”, 10, 20 , 40 and 100 time points). Position of the starting 12 nt RNA is indicated on the left; position of the 8-oxoguanine lesion is shown with an arrowhead and asterisk on the right.
of ATP likely occurs due to the formation of a Hoogsteen base-pair with 8oxoG bound in the syn conformation [25]. We confirmed that bacterial RNAP can insert both CTP and ATP opposite the lesion and also demonstrated that adenine-containing transcripts are preferably elongated after misincorporation, likely because of a more favorable conformation of the noncanonical 8oxoG-A pair during translocation of the RNA-DNA hybrid within RNAP. In agreement with this, the major pause on the 8oxoG template was observed downstream of the lesion (Fig. 2). Thus, the misincorporated adenine can be preferably extended on the 8oxoG templates both in vitro and in vivo resulting in transcriptional mutagenesis [5,25]. Thymine glycol severely inhibits DNA synthesis by replicative DNA polymerases and requires the action of specialized DNA polymerases for translesion synthesis (e.g. Ref. [26]). Eukaryotic and archaeal RNAPs were shown to pause after nucleotide insertion opposite TG [27,28]. Surprisingly, we found that transcription by bacterial RNAP is only slightly affected by this modification, probably because the RNAP active site is more tolerant to the duplex distortion caused by the nonplanar TG configuration, suggesting that this lesion may not be a subject for TCR. Finally, we analyzed several substitutions of amino acid residues in the RNAP active site involved in direct interactions with the transcribed DNA strand. Unexpectedly, some of these substitutions (R542A, Y795A and R798A) lacked any prominent effects on transcription of both normal and damaged DNA templates, despite affecting conserved RNAP residues involved in template interactions. At the same time, two substitutions, K334A in switch2 and T790 in the BH, inhibited transcription, and this effect was exacerbated on most damaged templates. The T790A substitution was previously shown to decrease RNAP activity in a number of assays, possibly by changing the BH conformation and/or its contacts with the template-NTP pair [29], likely explaining its effects on translesion synthesis. The effect of the K334A substitution may result from changes in RNAP contacts with the template DNA strand downstream of the active site and in the conformation of the clamp domain that holds the DNA-RNA duplex [14]. At the same time, the K334A substitution stimulated transcription past CPD, in contrast to other lesions, suggesting that it may help to overcome transcription stalling caused by this bulky lesion, possibly by loosening RNAP-DNA contacts near the active site. Overall, our results shed new light into the mechanisms of translesion RNA synthesis and reveal the ability of bacterial RNAP to pause, stall, or transcribe past various DNA lesions. This opens a way for further analysis of the molecular details of translesion synthesis by bacterial RNAPs, and its modulation by transcription and DNA repair factors.
Acknowledgements We thank I. Artsimovitch for plasmids and for the CPD template. This work was supported by the Grant of the Ministry of Higher Education and Science of the Russian Federation 14.W03.31.0007. Appendix A. Supplementary data Supplementary data to this article can be found online at https://doi.org/10.1016/j.bbrc.2019.01.062. Transparency document Transparency document related to this article can be found online at https://doi.org/10.1016/j.bbrc.2019.01.062. References [1] N. Chatterjee, G.C. Walker, Mechanisms of DNA damage, repair, and mutagenesis, Environ. Mol. Mutagen. 58 (2017) 235e263. [2] R.P. Fuchs, S. Fujii, Translesion DNA synthesis and mutagenesis in prokaryotes, Cold Spring Harb. Perspect. Biol. 5 (2013) a012682. [3] B.A. Donahue, S. Yin, J.S. Taylor, D. Reines, P.C. Hanawalt, Transcript cleavage by RNA polymerase II arrested by a cyclobutane pyrimidine dimer in the DNA template, Proc. Natl. Acad. Sci. U. S. A. 91 (1994) 8502e8506. [4] W. Zhou, P.W. Doetsch, Effects of abasic sites and DNA single-strand breaks on prokaryotic RNA polymerases, Proc. Natl. Acad. Sci. U. S. A. 90 (1993) 6601e6605. [5] D. Bregeon, Z.A. Doddridge, H.J. You, B. Weiss, P.W. Doetsch, Transcriptional mutagenesis induced by uracil and 8-oxoguanine in Escherichia coli, Mol. Cell 12 (2003) 959e970. [6] A. Viswanathan, P.W. Doetsch, Effects of nonbulky DNA base damages on Escherichia coli RNA polymerase-mediated elongation and promoter clearance, J. Biol. Chem. 273 (1998) 21276e21281. [7] I. Kuraoka, M. Endou, Y. Yamaguchi, T. Wada, H. Handa, K. Tanaka, Effects of endogenous DNA base lesions on transcription elongation by mammalian RNA polymerase II. Implications for transcription-coupled DNA repair and transcriptional mutagenesis, J. Biol. Chem. 278 (2003) 7294e7299. [8] G. Spivak, Transcription-coupled repair: an update, Arch. Toxicol. 90 (2016) 2583e2594. [9] D. Dutta, K. Shatalin, V. Epshtein, M.E. Gottesman, E. Nudler, Linking RNA polymerase backtracking to genome instability in E. coli, Cell 146 (2011) 533e543. [10] K.S. Lang, A.N. Hall, C.N. Merrikh, M. Ragheb, H. Tabakh, A.J. Pollock, J.J. Woodward, J.E. Dreifus, H. Merrikh, Replication-transcription conflicts generate R-loops that orchestrate bacterial stress survival and pathogenesis, Cell 170 (2017) 787e799 e718. [11] V. Svetlov, I. Artsimovitch, Purification of bacterial RNA polymerase: tools and protocols, Methods Mol. Biol. 1276 (2015) 13e29. [12] D. Esyunina, M. Turtola, D. Pupov, I. Bass, S. Klimasauskas, G. Belogurov, A. Kulbachinskiy, Lineage-specific variations in the trigger loop modulate RNA proofreading by bacterial RNA polymerases, Nucleic Acids Res. 44 (2016) 1298e1308. [13] N. Korzheva, A. Mustaev, E. Nudler, V. Nikiforov, A. Goldfarb, Mechanistic model of the elongation complex of Escherichia coli RNA polymerase, Cold
D. Pupov et al. / Biochemical and Biophysical Research Communications 510 (2019) 122e127 Spring Harbor Symp. Quant. Biol. 63 (1998) 337e345. [14] G.A. Belogurov, M.N. Vassylyeva, A. Sevostyanova, J.R. Appleman, A.X. Xiang, R. Lira, S.E. Webber, S. Klyuyev, E. Nudler, I. Artsimovitch, D.G. Vassylyev, Transcription inactivation through local refolding of the RNA polymerase structure, Nature 457 (2009) 332e335. [15] D.G. Vassylyev, M.N. Vassylyeva, J. Zhang, M. Palangat, I. Artsimovitch, R. Landick, Structural basis for substrate loading in bacterial RNA polymerase, Nature 448 (2007) 163e168. [16] V.S. Kalogeraki, S. Tornaletti, P.K. Cooper, P.C. Hanawalt, Comparative TFIISmediated transcript cleavage by mammalian RNA polymerase II arrested at a lesion in different transcription systems, DNA Repair (Amst) 4 (2005) 1075e1087. [17] J.S. Mei Kwei, I. Kuraoka, K. Horibata, M. Ubukata, E. Kobatake, S. Iwai, H. Handa, K. Tanaka, Blockage of RNA polymerase II at a cyclobutane pyrimidine dimer and 6-4 photoproduct, Biochem. Biophys. Res. Commun. 320 (2004) 1133e1138. [18] F. Brueckner, U. Hennecke, T. Carell, P. Cramer, CPD damage recognition by transcribing RNA polymerase II, Science 315 (2007) 859e862. [19] C. Walmacq, A.C. Cheung, M.L. Kireeva, L. Lubkowska, C. Ye, D. Gotte, J.N. Strathern, T. Carell, P. Cramer, M. Kashlev, Mechanism of translesion transcription by RNA polymerase II and its role in cellular resistance to DNA damage, Mol. Cell 46 (2012) 18e29. [20] A. Dimitri, A.K. Goodenough, F.P. Guengerich, S. Broyde, D.A. Scicchitano, Transcription processing at 1,N2-ethenoguanine by human RNA polymerase II and bacteriophage T7 RNA polymerase, J. Mol. Biol. 375 (2008) 353e366. [21] W. Wang, C. Walmacq, J. Chong, M. Kashlev, D. Wang, Structural basis of
[22]
[23]
[24]
[25] [26]
[27]
[28] [29]
127
transcriptional stalling and bypass of abasic DNA lesion by RNA polymerase II, Proc. Natl. Acad. Sci. U. S. A. 115 (2018) E2538eE2545. S. Weerasooriya, V.P. Jasti, A.K. Basu, Replicative bypass of abasic site in Escherichia coli and human cells: similarities and differences, PLoS One 9 (2014), e107915. R.S. Basu, B.A. Warner, V. Molodtsov, D. Pupov, D. Esyunina, C. FernandezTornero, A. Kulbachinskiy, K.S. Murakami, Structural basis of transcription initiation by bacterial RNA polymerase holoenzyme, J. Biol. Chem. 289 (2014) 24549e24559. S. Tornaletti, L.S. Maeda, R.D. Kolodner, P.C. Hanawalt, Effect of 8-oxoguanine on transcription elongation by T7 RNA polymerase and mammalian RNA polymerase II, DNA Repair (Amst) 3 (2004) 483e494. G.E. Damsma, P. Cramer, Molecular basis of transcriptional mutagenesis at 8oxoguanine, J. Biol. Chem. 284 (2009) 31658e31663. J.H. Yoon, G. Bhatia, S. Prakash, L. Prakash, Error-free replicative bypass of thymine glycol by the combined action of DNA polymerases kappa and zeta in human cells, Proc. Natl. Acad. Sci. U. S. A. 107 (2010) 14116e14121. I. Kuraoka, K. Suzuki, S. Ito, M. Hayashida, J.S. Kwei, T. Ikegami, H. Handa, Y. Nakabeppu, K. Tanaka, RNA polymerase II bypasses 8-oxoguanine in the presence of transcription elongation factor TFIIS, DNA Repair (Amst) 6 (2007) 841e851. A.M. Gehring, T.J. Santangelo, Archaeal RNA polymerase arrests transcription at DNA lesions, Transcription 8 (2017) 288e296. M. Jovanovic, P.C. Burrows, D. Bose, B. Camara, S. Wiesler, X. Zhang, S. Wigneshweraraj, R.O. Weinzierl, M. Buck, Activity map of the Escherichia coli RNA polymerase bridge helix, J. Biol. Chem. 286 (2011) 14469e14479.
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Distinct effects of DNA lesions on RNA synthesis by Escherichia coli RNA polymerase Danil Pupov 1, Artem Ignatov 1, Aleksei Agapov, Andrey Kulbachinskiy* Institute of Molecular Genetics, Russian Academy of Sciences, Moscow, 123182, Russia
a r t i c l e i n f o
a b s t r a c t
Article history: Received 3 January 2019 Accepted 12 January 2019 Available online 18 January 2019
DNA lesions can severely compromise genome stability and lead to cell death if unrepaired. RNA polymerase (RNAP) is known to serve as a sensor of DNA damage and to attract DNA repair factors to the damaged template sites. Here, we systematically investigated the ability of Escherichia coli RNAP to transcribe DNA templates containing various types of DNA lesions, and analyzed their effects on transcription fidelity. We showed that transcription is strongly inhibited on templates containing cyclobutane thymine dimers, 1,N6-ethenoadenine and abasic sites, while 8-oxoguanine and thymine glycol have mild effects on transcription efficiency. Similarly to many polymerases, E. coli RNAP follows the “A” rule during nucleotide insertion opposite abasic sites and bulky lesions, and can also incorporate and efficiently extend an adenine nucleotide opposite 8-oxoguanine. Mutations in RNAP regions around the templating nucleotide decrease the efficiency of translesion synthesis, likely by altering the RNAPtemplate contacts in the active site. Thus, DNA lesions can lead to distinct outcomes in transcription, depending on the severity of the damage and contacts of the damaged template with the active site of RNAP. © 2019 Elsevier Inc. All rights reserved.
Keywords: RNA polymerase Translesion RNA synthesis Transcription fidelity Transcription-coupled repair
1. Introduction Genomic DNA in all organisms is continuously modified by multiple endogenous and exogenous factors. Some of these modifications are introduced by specialized enzymes and serve for regulation of gene expression, DNA repair, or defense against foreign DNA. Others appear spontaneously or are introduced by genotoxic agents and may impair various genetic processes and lead to genome instability. In particular, many DNA lesions are known to affect replication dramatically, leading to the replication fork stalling and mutagenesis. Most organisms possess specialized DNA polymerases that are able to bypass certain types of lesions but usually have a much lower replication fidelity [1,2]. Transcription can also be impaired by lesions in the template
Abbreviations: RNAP, RNA polymerase; CPD, cyclobutane pyrimidine (thymine) dimer; εA, 1,N6-ethenoadenine; AP, apurinic/apyrimidinic site; 8oxoG, 8oxoguanine; TG, thymine glycol; TEC, transcription elongation complex; BH, bridge helix. * Corresponding author. Institute of Molecular Genetics, 2 Kurchatov sq., Moscow, 123182, Russia. E-mail address: [email protected] (A. Kulbachinskiy). 1 These authors contributed equally to this work. https://doi.org/10.1016/j.bbrc.2019.01.062 0006-291X/© 2019 Elsevier Inc. All rights reserved.
DNA strand, which may lead to stalling of RNA polymerase (RNAP) or to transcriptional mutagenesis, thus producing mutant RNAs and proteins [3e7]. At the same time, RNAP can act as a sensor for DNA lesions, by attracting the DNA repair machinery to the damaged template sites during transcription coupled repair (TCR) (reviewed in Ref. [8]). Furthermore, stalled transcription complexes can severely compromise genome stability by colliding with the replication machinery [9,10]. Bacterial cells contain a single RNAP, and the process of DNA damage recognition and repair should be highly coordinated with transcription to allow efficient gene expression and DNA replication. However, the ability of bacterial RNAP to transcribe damaged DNA templates has not been systematically studied. Only a handful of lesions have been analyzed in the bacterial transcription system in vitro. In particular, it was shown that similarly to DNA polymerases, the abasic site and 8-oxoguanine promote mutagenic insertion of ATP in the RNA transcript, with transient RNAP pausing [4e6]. At the same time, the molecular mechanisms of translesion RNA synthesis remain poorly understood for most types of damaged nucleotides. In this work, we directly compared the effects on transcription by E. coli RNAP of several types of lesions commonly found in the genomic DNA: thymine dimer (CPD, cyclobutane pyrimidine dimer); 1,N6-ethenoadenine (εA); abasic site (AP); 8-oxoguanine
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(8oxoG), and thymine glycol (TG). We for the first time demonstrated that CPD and εA severely inhibit the activity of bacterial RNAP in vitro, allowing only very slow incorporation of ATP (the same specificity as observed for the AP-site), while TG has only a weak effect on transcription. Furthermore, we showed that 8oxoG is highly mutagenic because it enables not only insertion of a noncomplementary ATP but also its efficient extension by RNAP. Finally, we studied the effects of amino acid substitutions in the active site of E. coli RNAP on transcription of damaged DNA and identified mutations with altered transcription efficiency.
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annealed oligonucleotides (10 nM final RNA concentration) and incubated for 10 min at 20 C or 37 C, followed by NTP addition (100 mM each, unless otherwise indicated). The reactions were stopped after various time intervals, as indicated in the figures, and the RNA products were resolved by 18% denaturing PAGE, followed by phosphorimaging.
3. Results 3.1. Diverse effects of various types of DNA lesions on transcription
2. Materials and methods 2.1. Proteins and nucleic acids Wild-type and mutant core E. coli RNAPs were purified from Eco BL21(DE3) cells expressing all four core RNAP subunits from the plasmids pVS10 (for the K334A, T790A, Y795A and R798A mutants) or pIA679 (for the R542A mutant) as described [11]. The core RNAP mutations were obtained by site-directed mutagenesis. RNAPs were purified by Polymin P precipitation, Heparin-Sepharose, Ni2þaffinity and MonoQ anionic-exchange chromatography steps as described [11,12]. The s subunit was purified using the Ni2þ-affinity and DEAE ion-exchange chromatography [12]. DNA and RNA oligonucleotides used in the assays (Fig. 1) were purchased from DNA Synthesis, Syntol (Moscow, Russia), and TriLink BioTechnologies (San Diego, CA). 2.2. Analysis of RNA synthesis in vitro Transcription assays were performed in minimal transcription elongation complexes (TECs) assembled from synthetic DNA and RNA oligonucleotides and the E. coli core RNAP as described [13]. The 50 -labeled RNA transcript (0.5 mM final concentration) was mixed with the template (1 mM) and nontemplate DNA oligonucleotides (5 mM) in transcription buffer containing 40 mM Tris-HCl, pH 7.9, 40 mM NaCl and 10 mM MgCl2, heated to 65 C, and cooled down to 20 C at 1 /min. For the AP site, the template was assembled for 10 min at 37 C. Core RNAP (25 nM) was mixed with the
To analyze the effects of DNA lesions on transcription, we assembled minimal TECs from synthetic RNA and DNA oligonucleotides and the core enzyme of E. coli RNAP (Fig. 1). Such minimal templates were previously shown to form stable complexes with RNAP and mimic the principal RNAP-nucleic acids interactions in the natural TEC [13]. In most complexes, the RNA 30 -end was located immediately upstream of the damaged template base, so that the incoming nucleotide should be incorporated opposite the lesion; in the case of CPD, the RNA 30 -end was located 2 nt upstream of the lesion (Fig. 1B). The lengths of 50 -labeled starting RNAs in all complexes were 12 or 11 nt, as indicated in the figures. We also assembled control complexes containing undamaged DNA oligonucleotides of the same sequence (for the AP-site, we used the same control template as for the TG lesion). We first tested the ability of wild-type E. coli RNAP to extend RNA in these complexes in the presence of all four NTP substrates. In the case of control undamaged templates, RNA was readily extended by at least 7e10 nt, several nucleotides past the site where we intended to place the lesions (Fig. 2, left column). Although not all transcripts reached the end of the template (the expected RNA extension by 15e19 nucleotides, depending on the template) e because efficient RNA synthesis requires the presence of 14 nt downstream DNA duplex [13] e this allowed us to analyze the effects of the lesions on RNA extension opposite and immediately past the lesion. The presence of damaged nucleotides had significant and diverse effects on transcription, depending on the nature of the lesion. For the bulky CPD modification, we observed almost
Fig. 1. The structure of bacterial RNAP and composition of analyzed DNA templates. (A) The structure of the active site of T. thermophilus RNAP in the transcription elongation complex (2O5J [15]). DNA is orange, the templating nucleotide (þ1) is carmine, the incoming NTP is black, RNA is yellow, magnesium ions bound in the active site are magenta, the bridge helix (BH) is turquoise. Amino acid residues mutated in this study are shown. (B) Minimal templates containing analyzed DNA lesions. The damaged nucleotides are shown in bold italics and underlined. (For interpretation of the references to colour in this figure legend, the reader is referred to the Web version of this article.)
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before and after nucleotide insertion opposite the lesion, as evident from inefficient extension of the starting 12 nt and extended 13 nt RNA products in comparison with the control template (Fig. 2, WT RNAP in the third row). To detect possible defects in RNA synthesis caused by the nonbulky 8oxoG and TG lesions, which were expected to have milder effects on transcription, we used a lower reaction temperature (20 C vs. 37 C) and shorter reaction times in comparison with the first set of lesions. In the case of 8oxoG, RNAP stalling was observed after RNA extension by 2 nucleotides, one nucleotide past the lesion, suggesting that 8oxoG does not prevent nucleotide incorporation but may hamper further RNAP translocation, when located inside the RNA-DNA hybrid (Fig. 2, fourth row). In the case of TG, we observed weak but detectable RNAP stalling at the site of lesion (at RNA length of 13 nt) that was absent in the case of undamaged template (Fig. 2, bottom row). At the same time, most RNAs were efficiently extended past the lesion.
3.2. Transcription of damaged templates by RNAP variants with mutations in the active center
Fig. 2. Activities of wild-type (WT) and mutant RNAP variants on damaged DNA templates. Control reactions performed with the wild-type RNAP on corresponding undamaged templates are shown in the left column. The type of DNA lesion analyzed in each row is shown on the right. For the CPD, εA and AP templates, the reactions were performed at 37 C for total 2 h (the time points are 10, 30 , 100 , 300 , 600 , 1200 ). For the 8oxoG and TG templates, the reactions were performed at 20 C for 10’ (10”, 30”, 10, 20 , 40 and 100 time points). Positions of the starting 11 nt or 12 nt RNAs are shown on the left; positions of DNA lesions are indicated with arrowheads and asterisks on the right.
complete block of transcription when the RNA 30 -end reached the first damaged nucleotide (the 30 -T residue of the dimer), after rapid initial extension by one nucleotide (Fig. 2, wild-type RNAP in the first row). This block was slowly overcome after prolonged incubation, resulting in the insertion of a single nucleotide opposite the 30 -T of CPD, as evident from the accumulation of the 13 nt RNA product. Although its further extension was very inefficient, there was no accumulation of the next extension product, after incorporation of the second nucleotide opposite the 50 -T of CPD, and longer RNA products were observed as a result of read-through synthesis. Thus, there appears to be no strong barrier to elongation after incorporation of the two nucleotides opposite CPD. The 1,N6-ethenoadenine modification, which prevents WatsonCrick pairing with thymine, blocked RNA synthesis even stronger than the CPD lesion (Fig. 2, WT RNAP in the second row), with very little RNA extension observed even at the longest incubation time. Markedly, these read-through RNA products were inefficiently extended past the lesion, suggesting that it blocks further RNAP translocation. In comparison with CPD and εA, the AP site was relatively readily transcribed by RNAP, although pauses were observed both
To get insight into the recognition of damaged nucleotides in the active site of RNAP, we obtained mutant variants of E. coli RNAP with alanine substitutions of several amino acid residues involved in interactions with the transcribed DNA strand (Fig. lA): (i) substitution K334A in the switch2 region in the b0 subunit; K334 contacts the phosphate group of the þ1 template nucleotide and stabilizes the template DNA strand in the active site [14,15]; (ii) substitution R542A in the fork-loop 2 region in the b subunit; R542 contacts the þ1 phosphate and the þ2 base of the downstream template DNA [15]; (iii) substitution T790A in the b0 subunit bridge helix (BH); T790 is the closest to the þ1 templating base in the TEC structure [15]; (iv) substitution Y795A in the BH; this residue contacts the þ2 and þ 3 phosphates in the template strand [15]; (v) substitution R798A in the BH; this residue contacts the 1 and þ2 template phosphates [15]. All mutant RNAPs were generated by site-directed mutagenesis, expressed and purified from E. coli cells. Analysis of the activity of these RNAPs on a linear DNA fragment containing the T7A1cons promoter showed that they all were capable of promoter-dependent transcription initiation (Fig. S1). We further tested the activities of these mutants in the control minimal TECs containing undamaged template DNAs. It was shown that at 37 C (conditions used for analysis of the CPD, εA and AP lesions) the activities of all five mutant RNAPs on undamaged DNA were comparable to the wild-type RNAP, with most RNA primer extended at the first time point (10”; Fig. S2). At the same time, at 20 C (conditions used for analysis of the 8oxoG and TG lesions), the K334A, T790A, Y795A and R798A mutants revealed lower levels of activity (Fig. S2). In particular, the K334A and T790A mutants extended RNA primers with slower kinetics and synthesized shorter RNA products, likely as a result of altered RNAP-DNA contacts and impaired catalysis. We then analyzed RNA synthesis by the mutant RNAPs on damaged DNA templates. The activities of the R542A, Y795A and R798A mutants were similar to the wild-type RNAP on all tested templates (Fig. S3). At the same time, the K334A and especially T790A substitutions additionally impaired RNA extension on most damaged templates in comparison with the wild-type RNAP, even at 37 C (Fig. 2, compare two right columns with the WT column). For example, no RNA extension past the CPD lesion and much lower activity on the AP template were observed for the T790A RNAP. Interestingly, the K334A substitution increased RNA extension past CPD (Fig. 2, first row), possibly as a result of more efficient translocation of this bulky lesion into the active site of RNAP.
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3.3. Effects of DNA lesions on transcription fidelity Many DNA lesions are known to affect the fidelity of nucleotide incorporation by DNA polymerases, thus resulting in mutagenesis [2]. Similarly, nucleotide misincorporation by RNAP may produce mutated RNA variants. We therefore tested the effects of various DNA lesions on the fidelity of nucleotide incorporation by E. coli RNAP, by analyzing RNA extension in the presence of individual NTPs. We observed efficient incorporation of both CTP and ATP on the 8oxoG template, while only CTP (and with low efficiency UTP) was incorporated in the control reaction (Fig. 3A). This agrees with published data showing that 8oxoG allows incorporation of adenine nucleotides in addition to the complementary cytosine by various RNA and DNA polymerases [2,5e7]. The TG lesion allowed efficient incorporation of the complementary adenine residue, in agreement with its weak effects on transcription (Fig. 3B). Similarly, only ATP was incorporated in the case of the AP template, but with a much lower efficiency than in the case of the control TEC containing a thymine residue in the template strand (Fig. 3B). Only very weak ATP incorporation was observed in the case of the CPD and εA lesions, with no incorporation of other three nucleotides (Fig. S4). In the previous experiment, we observed efficient incorporation of both C and A opposite template 8oxoG. To establish which of these nucleotides can be preferably incorporated into the fulllength RNA, we compared the extension efficiencies for 12 nt RNA transcripts containing either C or A at their 30 -ends opposite undamaged G or 8oxoG. We revealed that the correct 30 C RNA was inefficiently extended past the lesion, resulting in a major pause one nucleotide downstream of it (Fig. 4, compare 30 C-8oxoG and 30 C-G reactions). In contrast, the misincorporated 30 A RNA could be efficiently extended on the 8oxoG template but not on the control undamaged template (compare the 30 A-8oxoG and 30 A-G reactions).
4. Discussion Here, we investigated the effects of several DNA lesions, which are well known to affect DNA replication, on the efficiency and fidelity of RNA synthesis by bacterial RNAP. We revealed significant variations in the ability of RNAP to transcribe damaged DNA templates depending on the type of the lesion, as described below, and identified amino acid residues possibly involved in translesion synthesis.
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CPD is the most common UV light-induced DNA lesion that severely inhibits DNA replication [2]. We demonstrated that CPD also strongly impairs transcription by bacterial RNAP, with very slow nucleotide insertion opposite both thymine dimer residues. Similarly to our observations, it was shown that nucleotide insertion opposite CPD by eukaryotic RNAP II is dramatically slowed down in comparison with undamaged templates [3,16e18]. This is likely explained by impaired RNAP translocation and may be accompanied by RNAP backtracking [18,19]. Furthermore, certain substitutions in the trigger loop of RNAP II were shown to stimulate translesion synthesis and cell survival after UV irradiation, possibly by promoting nucleotide insertion [19]. In contrast, our analyzed substitutions in the BH and switch2 of E. coli RNAP inhibited transcription. We revealed that another common lesion εA can completely block transcription, likely by disrupting complementary pairing with the incoming NTP. Although εA has never been studied with any RNAP, a similar guanine modification, 1,N2-ethenoguanine, could block RNA synthesis by eukaryotic RNAP II [20], suggesting that the effects of this type of modification are similar for bacterial and eukaryotic RNAPs. The AP-site was previously shown to induce transient RNAP pausing and ATP insertion by both E. coli and eukaryotic RNAPs, followed by further RNA extension [4,7]. For eukaryotic RNAP, the AP-site was recently demonstrated to slow down nucleotide incorporation both opposite and after the lesion [21]. We also observed pausing by bacterial RNAP before and after ATP insertion demonstrating that the mechanism of AP-site bypass is similar for these RNAPs. Interestingly, all three lesions that significantly inhibited transcription e CPD, εA and the AP site, e promoted insertion of adenine residues opposite the damaged nucleotide. Thus, bacterial RNAP seems to follow the so-called ‘A-rule’ for incorporation of nucleotides opposite AP-sites and bulky lesions, similarly to many DNA polymerases [22]. ATP is a preferred nucleotide for the active site of both bacterial [23] and eukaryotic [21] RNAPs, which likely promotes nontemplated ATP insertion. Indeed, RNAP II was shown to insert A opposite CPD in a nontemplated manner [19]. Other bulky lesions may be transcribed by RNAP in a similar way. 8oxoG is the most common oxidative DNA lesion that has been studied in both eukaryotic and bacterial transcription systems. When located in the template DNA strand, it promotes misincorporation of adenine and causes weak transcriptional pausing by both bacterial and eukaryotic RNAPs [5e7,24]. Misincorporation
Fig. 3. Fidelity of NTP incorporation opposite various DNA lesions by wild-type E. coli RNAP. (A) Single-nucleotide incorporation opposite 8-oxoguanine (right) or guanine in the corresponding control template (left). (B) Nucleotide incorporation opposite the AP-site (middle), thymine glycol (right), and corresponding control template (left). The reactions were performed for 3000 at 20 C with each of the four NTPs taken at indicated concentrations.
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Fig. 4. The efficiency of RNA extension after 8-oxoguanine depending on the RNA 3′-end structure. The TECs contained 12 nt RNA with the 30 -end (either C or A) located opposite either undamaged guanine (G) or 8-oxoguanine (8oxoG). Transcription was performed at 20 C for 10’ (10”, 30”, 10, 20 , 40 and 100 time points). Position of the starting 12 nt RNA is indicated on the left; position of the 8-oxoguanine lesion is shown with an arrowhead and asterisk on the right.
of ATP likely occurs due to the formation of a Hoogsteen base-pair with 8oxoG bound in the syn conformation [25]. We confirmed that bacterial RNAP can insert both CTP and ATP opposite the lesion and also demonstrated that adenine-containing transcripts are preferably elongated after misincorporation, likely because of a more favorable conformation of the noncanonical 8oxoG-A pair during translocation of the RNA-DNA hybrid within RNAP. In agreement with this, the major pause on the 8oxoG template was observed downstream of the lesion (Fig. 2). Thus, the misincorporated adenine can be preferably extended on the 8oxoG templates both in vitro and in vivo resulting in transcriptional mutagenesis [5,25]. Thymine glycol severely inhibits DNA synthesis by replicative DNA polymerases and requires the action of specialized DNA polymerases for translesion synthesis (e.g. Ref. [26]). Eukaryotic and archaeal RNAPs were shown to pause after nucleotide insertion opposite TG [27,28]. Surprisingly, we found that transcription by bacterial RNAP is only slightly affected by this modification, probably because the RNAP active site is more tolerant to the duplex distortion caused by the nonplanar TG configuration, suggesting that this lesion may not be a subject for TCR. Finally, we analyzed several substitutions of amino acid residues in the RNAP active site involved in direct interactions with the transcribed DNA strand. Unexpectedly, some of these substitutions (R542A, Y795A and R798A) lacked any prominent effects on transcription of both normal and damaged DNA templates, despite affecting conserved RNAP residues involved in template interactions. At the same time, two substitutions, K334A in switch2 and T790 in the BH, inhibited transcription, and this effect was exacerbated on most damaged templates. The T790A substitution was previously shown to decrease RNAP activity in a number of assays, possibly by changing the BH conformation and/or its contacts with the template-NTP pair [29], likely explaining its effects on translesion synthesis. The effect of the K334A substitution may result from changes in RNAP contacts with the template DNA strand downstream of the active site and in the conformation of the clamp domain that holds the DNA-RNA duplex [14]. At the same time, the K334A substitution stimulated transcription past CPD, in contrast to other lesions, suggesting that it may help to overcome transcription stalling caused by this bulky lesion, possibly by loosening RNAP-DNA contacts near the active site. Overall, our results shed new light into the mechanisms of translesion RNA synthesis and reveal the ability of bacterial RNAP to pause, stall, or transcribe past various DNA lesions. This opens a way for further analysis of the molecular details of translesion synthesis by bacterial RNAPs, and its modulation by transcription and DNA repair factors.
Acknowledgements We thank I. Artsimovitch for plasmids and for the CPD template. This work was supported by the Grant of the Ministry of Higher Education and Science of the Russian Federation 14.W03.31.0007. Appendix A. Supplementary data Supplementary data to this article can be found online at https://doi.org/10.1016/j.bbrc.2019.01.062. Transparency document Transparency document related to this article can be found online at https://doi.org/10.1016/j.bbrc.2019.01.062. References [1] N. Chatterjee, G.C. Walker, Mechanisms of DNA damage, repair, and mutagenesis, Environ. Mol. Mutagen. 58 (2017) 235e263. [2] R.P. Fuchs, S. Fujii, Translesion DNA synthesis and mutagenesis in prokaryotes, Cold Spring Harb. Perspect. Biol. 5 (2013) a012682. [3] B.A. Donahue, S. Yin, J.S. Taylor, D. Reines, P.C. Hanawalt, Transcript cleavage by RNA polymerase II arrested by a cyclobutane pyrimidine dimer in the DNA template, Proc. Natl. Acad. Sci. U. S. A. 91 (1994) 8502e8506. [4] W. Zhou, P.W. Doetsch, Effects of abasic sites and DNA single-strand breaks on prokaryotic RNA polymerases, Proc. Natl. Acad. Sci. U. S. A. 90 (1993) 6601e6605. [5] D. Bregeon, Z.A. Doddridge, H.J. You, B. Weiss, P.W. Doetsch, Transcriptional mutagenesis induced by uracil and 8-oxoguanine in Escherichia coli, Mol. Cell 12 (2003) 959e970. [6] A. Viswanathan, P.W. Doetsch, Effects of nonbulky DNA base damages on Escherichia coli RNA polymerase-mediated elongation and promoter clearance, J. Biol. Chem. 273 (1998) 21276e21281. [7] I. Kuraoka, M. Endou, Y. Yamaguchi, T. Wada, H. Handa, K. Tanaka, Effects of endogenous DNA base lesions on transcription elongation by mammalian RNA polymerase II. Implications for transcription-coupled DNA repair and transcriptional mutagenesis, J. Biol. Chem. 278 (2003) 7294e7299. [8] G. Spivak, Transcription-coupled repair: an update, Arch. Toxicol. 90 (2016) 2583e2594. [9] D. Dutta, K. Shatalin, V. Epshtein, M.E. Gottesman, E. Nudler, Linking RNA polymerase backtracking to genome instability in E. coli, Cell 146 (2011) 533e543. [10] K.S. Lang, A.N. Hall, C.N. Merrikh, M. Ragheb, H. Tabakh, A.J. Pollock, J.J. Woodward, J.E. Dreifus, H. Merrikh, Replication-transcription conflicts generate R-loops that orchestrate bacterial stress survival and pathogenesis, Cell 170 (2017) 787e799 e718. [11] V. Svetlov, I. Artsimovitch, Purification of bacterial RNA polymerase: tools and protocols, Methods Mol. Biol. 1276 (2015) 13e29. [12] D. Esyunina, M. Turtola, D. Pupov, I. Bass, S. Klimasauskas, G. Belogurov, A. Kulbachinskiy, Lineage-specific variations in the trigger loop modulate RNA proofreading by bacterial RNA polymerases, Nucleic Acids Res. 44 (2016) 1298e1308. [13] N. Korzheva, A. Mustaev, E. Nudler, V. Nikiforov, A. Goldfarb, Mechanistic model of the elongation complex of Escherichia coli RNA polymerase, Cold
D. Pupov et al. / Biochemical and Biophysical Research Communications 510 (2019) 122e127 Spring Harbor Symp. Quant. Biol. 63 (1998) 337e345. [14] G.A. Belogurov, M.N. Vassylyeva, A. Sevostyanova, J.R. Appleman, A.X. Xiang, R. Lira, S.E. Webber, S. Klyuyev, E. Nudler, I. Artsimovitch, D.G. Vassylyev, Transcription inactivation through local refolding of the RNA polymerase structure, Nature 457 (2009) 332e335. [15] D.G. Vassylyev, M.N. Vassylyeva, J. Zhang, M. Palangat, I. Artsimovitch, R. Landick, Structural basis for substrate loading in bacterial RNA polymerase, Nature 448 (2007) 163e168. [16] V.S. Kalogeraki, S. Tornaletti, P.K. Cooper, P.C. Hanawalt, Comparative TFIISmediated transcript cleavage by mammalian RNA polymerase II arrested at a lesion in different transcription systems, DNA Repair (Amst) 4 (2005) 1075e1087. [17] J.S. Mei Kwei, I. Kuraoka, K. Horibata, M. Ubukata, E. Kobatake, S. Iwai, H. Handa, K. Tanaka, Blockage of RNA polymerase II at a cyclobutane pyrimidine dimer and 6-4 photoproduct, Biochem. Biophys. Res. Commun. 320 (2004) 1133e1138. [18] F. Brueckner, U. Hennecke, T. Carell, P. Cramer, CPD damage recognition by transcribing RNA polymerase II, Science 315 (2007) 859e862. [19] C. Walmacq, A.C. Cheung, M.L. Kireeva, L. Lubkowska, C. Ye, D. Gotte, J.N. Strathern, T. Carell, P. Cramer, M. Kashlev, Mechanism of translesion transcription by RNA polymerase II and its role in cellular resistance to DNA damage, Mol. Cell 46 (2012) 18e29. [20] A. Dimitri, A.K. Goodenough, F.P. Guengerich, S. Broyde, D.A. Scicchitano, Transcription processing at 1,N2-ethenoguanine by human RNA polymerase II and bacteriophage T7 RNA polymerase, J. Mol. Biol. 375 (2008) 353e366. [21] W. Wang, C. Walmacq, J. Chong, M. Kashlev, D. Wang, Structural basis of
[22]
[23]
[24]
[25] [26]
[27]
[28] [29]
127
transcriptional stalling and bypass of abasic DNA lesion by RNA polymerase II, Proc. Natl. Acad. Sci. U. S. A. 115 (2018) E2538eE2545. S. Weerasooriya, V.P. Jasti, A.K. Basu, Replicative bypass of abasic site in Escherichia coli and human cells: similarities and differences, PLoS One 9 (2014), e107915. R.S. Basu, B.A. Warner, V. Molodtsov, D. Pupov, D. Esyunina, C. FernandezTornero, A. Kulbachinskiy, K.S. Murakami, Structural basis of transcription initiation by bacterial RNA polymerase holoenzyme, J. Biol. Chem. 289 (2014) 24549e24559. S. Tornaletti, L.S. Maeda, R.D. Kolodner, P.C. Hanawalt, Effect of 8-oxoguanine on transcription elongation by T7 RNA polymerase and mammalian RNA polymerase II, DNA Repair (Amst) 3 (2004) 483e494. G.E. Damsma, P. Cramer, Molecular basis of transcriptional mutagenesis at 8oxoguanine, J. Biol. Chem. 284 (2009) 31658e31663. J.H. Yoon, G. Bhatia, S. Prakash, L. Prakash, Error-free replicative bypass of thymine glycol by the combined action of DNA polymerases kappa and zeta in human cells, Proc. Natl. Acad. Sci. U. S. A. 107 (2010) 14116e14121. I. Kuraoka, K. Suzuki, S. Ito, M. Hayashida, J.S. Kwei, T. Ikegami, H. Handa, Y. Nakabeppu, K. Tanaka, RNA polymerase II bypasses 8-oxoguanine in the presence of transcription elongation factor TFIIS, DNA Repair (Amst) 6 (2007) 841e851. A.M. Gehring, T.J. Santangelo, Archaeal RNA polymerase arrests transcription at DNA lesions, Transcription 8 (2017) 288e296. M. Jovanovic, P.C. Burrows, D. Bose, B. Camara, S. Wiesler, X. Zhang, S. Wigneshweraraj, R.O. Weinzierl, M. Buck, Activity map of the Escherichia coli RNA polymerase bridge helix, J. Biol. Chem. 286 (2011) 14469e14479.