1. Biochemistry and Chemical Biology
  2. Chromosomes and Gene Expression
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Highly efficient 5' capping of mitochondrial RNA with NAD+ and NADH by yeast and human mitochondrial RNA polymerase

  1. Jeremy G Bird
  2. Urmimala Basu
  3. David Kuster
  4. Aparna Ramachandran
  5. Ewa Grudzien-Nogalska
  6. Atif Towheed
  7. Douglas C Wallace
  8. Megerditch Kiledjian
  9. Dmitry Temiakov
  10. Smita S Patel  Is a corresponding author
  11. Richard H Ebright  Is a corresponding author
  12. Bryce E Nickels  Is a corresponding author
  1. Rutgers University, United States
  2. Heidelberg University, Germany
  3. The Children’s Hospital of Philadelphia, United States
  4. The Children's Hospital of Philadelphia, Perelman School of Medicine, United States
  5. Thomas Jefferson University, United States
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Cite this article as: eLife 2018;7:e42179 doi: 10.7554/eLife.42179

Abstract

Bacterial and eukaryotic nuclear RNA polymerases (RNAPs) cap RNA with the oxidized and reduced forms of the metabolic effector nicotinamide adenine dinucleotide, NAD+ and NADH, using NAD+ and NADH as non-canonical initiating nucleotides for transcription initiation. Here, we show that mitochondrial RNAPs (mtRNAPs) cap RNA with NAD+ and NADH, and do so more efficiently than nuclear RNAPs. Direct quantitation of NAD+- and NADH-capped RNA demonstrates remarkably high levels of capping in vivo: up to ~60% NAD+ and NADH capping of yeast mitochondrial transcripts, and up to ~15% NAD+ capping of human mitochondrial transcripts. The capping efficiency is determined by promoter sequence at, and upstream of, the transcription start site and, in yeast and human cells, by intracellular NAD+ and NADH levels. Our findings indicate mtRNAPs serve as both sensors and actuators in coupling cellular metabolism to mitochondrial transcriptional outputs, sensing NAD+ and NADH levels and adjusting transcriptional outputs accordingly.

https://doi.org/10.7554/eLife.42179.001

Introduction

Chemical modifications of the RNA 5'-end provide a layer of ‘epitranscriptomic’ regulation, influencing RNA fate, including stability, processing, localization, and translation efficiency (Höfer and Jäschke, 2018; Jäschke et al., 2016; Ramanathan et al., 2016; Shuman, 2015). One well-characterized RNA 5'-end modification is the ‘cap’ comprising 7-methylguanylate (m7G) added to many eukaryotic messenger RNAs (Furuichi and Shatkin, 2000; Shatkin, 1976; Shuman, 1995; Wei et al., 1975). Recently, a new RNA 5'-end cap comprising the metabolic effector nicotinamide adenine dinucleotide (NAD) has been shown to be added to certain RNAs isolated from bacterial, yeast, and human cells (Cahová et al., 2015; Chen et al., 2009; Frindert et al., 2018; Jiao et al., 2017; Walters et al., 2017).

In contrast to a m7G cap, which is added to nascent RNA by a capping complex that associates with eukaryotic RNA polymerase II (RNAP II) (Ghosh and Lima, 2010; Martinez-Rucobo et al., 2015; Shuman, 1995; Shuman, 2001; Shuman, 2015), an NAD cap is added by RNAP itself during transcription initiation, by serving as a non-canonical initiating nucleotide (NCIN) (Bird et al., 2016) (reviewed in Barvík et al., 2017; Julius et al., 2018; Vasilyev et al., 2018). NCIN-mediated NAD capping has been demonstrated for bacterial RNAP (Bird et al., 2016; Frindert et al., 2018; Julius and Yuzenkova, 2017; Vvedenskaya et al., 2018) and eukaryotic RNAP II (Bird et al., 2016). Thus, whereas m7G capping occurs after transcription initiation, on formation of the ~20th phosphodiester bond, and occurs only in organisms harboring specialized capping complexes, NAD capping occurs in transcription initiation, on formation of the first phosphodiester bond, and because it is performed by RNAP itself, is likely to occur in most, if not all, organisms.

NAD exists in oxidized and reduced forms: NAD+ and NADH, respectively (Figure 1A). Capping with NAD+ has been demonstrated both in vitro and in vivo (Bird et al., 2016; Frindert et al., 2018; Julius and Yuzenkova, 2017; Vvedenskaya et al., 2018). Capping with NADH has been demonstrated in vitro (Bird et al., 2016; Julius and Yuzenkova, 2017).

Figure 1 with 1 supplement see all
S. cerevisiae and human mtRNAPs cap RNA with NADand NADH in vitro.

(A) Structures of ATP, NAD+, and NADH. Grey, identical atoms; black, distinct atoms. (B) Processing of RNA 5' ends by RppH and NudC. A, adenosine; N+, NAD+ nicotinamide; N, NADH nicotinamide; p, phosphate. (C and D) NCIN capping with NAD+ and NADH by S. cerevisiae mtRNAP (C) and human mtRNAP (D). Top, promoter derivatives. Middle, initial RNA products of in vitro transcription reactions with ATP, NAD+, or NADH as initiating nucleotide and [α32P]-GTP as extending nucleotide. Bottom, full-length RNA products of in vitro transcription reactions with ATP, NAD+, or NADH as initiating nucleotide and [α32P]-GTP, ATP, UTP, and 3'-deoxy-CTP (C), or [α32P]-GTP, ATP, and UTP (D) as extending nucleotides. Products were treated with RppH or NudC as indicated. Grey box and arrow, transcription start site (TSS);+31 and+17/18, position of last NTP incorporated into full-length RNA products; M, 10-nt marker.

https://doi.org/10.7554/eLife.42179.002

Jäschke and co-workers developed a method that combines click-chemistry-mediated covalent capture and high-throughput sequencing, ‘NAD captureSeq,’ to detect NAD+-capped RNA (Cahová et al., 2015; Winz et al., 2017). Jäschke and co-workers used this method to identify NAD+-capped RNAs in bacterial cells (Escherichia coli and Bacillus subtilis; Cahová et al., 2015; Frindert et al., 2018). Parker, Kiledjian, and co-workers used the same method to identify NAD+-capped RNAs in eukaryotic cells (Saccharomyces cerevisiae and human cell line HEK293T; Jiao et al., 2017; Walters et al., 2017). Notably, the identified Saccharomyces cerevisiae NAD+-capped RNAs included not only RNAs produced by nuclear RNAPs, but also RNAs produced by mitochondrial RNAP (mtRNAP). The eukaryotic nuclear RNAPs--RNAP I, II, and III--are multi-subunit RNAPs closely related in sequence and structure to bacterial RNAP (Cramer, 2002; Darst, 2001; Ebright, 2000; Werner and Grohmann, 2011); in contrast, mtRNAPs are single-subunit RNAPs that are unrelated in sequence and structure to multi-subunit RNAPs and, instead, are related to DNA polymerases, reverse transcriptases, and DNA-dependent RNAPs from T7-like bacteriophages (Cermakian et al., 1996; Cheetham and Steitz, 2000; Hillen et al., 2018; Masters et al., 1987; McAllister and Raskin, 1993; Ringel et al., 2011; Sousa, 1996).

The identification of NAD+-capped mitochondrial RNAs in S. cerevisiae raises the question of whether eukaryotic single-subunit mtRNAPs--like the structurally unrelated bacterial and eukaryotic nuclear multi-subunit RNAPs--can perform NCIN-mediated capping. A recent review discussed evidence supporting the hypothesis that human mtRNAP can perform NCIN capping (Julius et al., 2018). Here, we show that single-subunit S. cerevisiae mtRNAP and human mtRNAP perform NCIN-mediated capping with NAD+ and NADH in vitro, and do so substantially more efficiently than bacterial and eukaryotic multi-subunit RNAPs. Further, we show that capping efficiency is determined by promoter sequence, we demonstrate very high levels of NAD+ and NADH capping--up to ~60% of mitochondrial transcripts in vivo, and we demonstrate that the extents of capping in vivo, and distributions of NAD+ capping vs. NADH capping in vivo are influenced by intracellular levels of NAD+ and NADH.

Results

S. cerevisiae and human mtRNAPs cap RNA with NAD+ and NADH in vitro

To assess whether mtRNAP can cap RNA with NAD+ and NADH, we performed in vitro transcription experiments (Figure 1 and Figure 1—figure supplement 1). We analyzed S. cerevisiae mtRNAP and a DNA template carrying the S. cerevisiae mitochondrial 21S promoter (Deshpande and Patel, 2014), and, in parallel, human mtRNAP and a DNA template containing a derivative of the human mitochondrial light-strand promoter, LSPAGU (Sologub et al., 2009) (Figure 1C–D, top). We performed reactions using either ATP, NAD+, or NADH as the initiating entity and using [α32P]-GTP as the extending nucleotide (Figure 1C–D, middle). We observed efficient formation of an initial RNA product in all cases (Figure 1C–D, middle). The initial RNA products obtained with ATP, but not with NAD+ or NADH, were processed by RppH, which previous work has shown to process 5'-triphosphate RNAs to 5'-monophosphate RNAs (Deana et al., 2008) (Figure 1B), whereas the initial RNA products obtained with NAD+ or NADH, but not with ATP, were processed by NudC, which previous work has shown to process 5'-NAD+- and 5'-NADH-capped RNAs to 5'-monophosphate RNAs (Cahová et al., 2015; Höfer et al., 2016) (Figure 1B). The results establish that S. cerevisiae mtRNAP and human mtRNAP are able to generate initial RNA products using NAD+ and NADH as NCINs.

We next assessed whether the initial RNA products formed using NAD+ and NADH as NCINs can be extended to yield full-length RNA products (Figure 1C–D, bottom, and Figure 1—figure supplement 1). We performed parallel transcription experiments using either ATP, NAD+, or NADH as the initiating entity and using [α32P]-GTP, ATP, UTP, and 3'-deoxy-CTP (Figure 1C, bottom) or [α32P]-GTP, ATP, and UTP (Figure 1D, bottom) as extending nucleotides. We observed efficient formation of full-length RNA products in all cases, and we observed that full-length RNA products obtained with NAD+ or NADH, but not with ATP, were sensitive to NudC treatment (Figure 1C–D, bottom). We also observed efficient formation of full-length RNA products in transcription experiments performed using [α32P]-ATP or [32P]-NAD+ as the initiating entity and using non-radiolabeled extending nucleotides (Figure 1—figure supplement 1). Full-length products obtained with [32P]-NAD+, but not with [α32P]-ATP, were insensitive to treatment with alkaline phosphatase (which processes 5' phosphates). Furthermore, NudC treatment of full-length products obtained with [32P]-NAD+ yielded products that were sensitive to alkaline phosphatase (Figure 1—figure supplement 1). The results establish that S. cerevisiae mtRNAP and human mtRNAP not only generate initial RNA products, but also generate full-length RNA products, using NAD+ and NADH as NCINs.

S. cerevisiae and human mtRNAPs cap RNA with NAD+ and NADH more efficiently than bacterial and nuclear RNAPs

We next determined the relative efficiencies of NCIN-mediated initiation vs. ATP-mediated initiation, (kcat/KM)NCIN / (kcat/KM)ATP, for mtRNAPs (Figure 2 and Figure 2—figure supplement 1; methods as in Bird et al., 2017). We performed reactions with S. cerevisiae mtRNAP and DNA templates carrying the S. cerevisiae mitochondrial 21S promoter or 15S promoter (Figure 2A and Figure 2—figure supplement 1), and, in parallel, with human mtRNAP and DNA templates carrying the human mitochondrial light-strand promoter (LSP) or heavy-strand promoter (HSP1) (Figure 2B and Figure 2—figure supplement 1). We obtained values of (kcat/KM)NCIN / (kcat/KM)ATP of ~0.3 to ~0.4 for NCIN-mediated initiation with NAD+ and NADH by S. cerevisiae mtRNAP and ~0.2 to ~0.6 for NCIN-mediated initiation with NAD+ and NADH by human mtRNAP. These values imply that NCIN-mediated initiation with NAD+ or NADH is up to 40% as efficient as initiation with ATP for S. cerevisiae mtRNAP and up to 60% as efficient as initiation with ATP for human mtRNAP.

Figure 2 with 2 supplements see all
S. cerevisiae and human mtRNAPs cap RNA with NADand NADH more efficiently than bacterial and nuclear RNAPs.

(A and B) Dependence of NCIN-mediated capping with NAD+ and NADH on [NCIN] / [ATP] ratio for S. cerevisiae mtRNAP (A) and human mtRNAP (B) (mean ± SD; n = 3). DNA templates and representative data are shown in Figure 2—figure supplement 1. (C) Dependence of NCIN-mediated capping with NAD+ and NADH on [NCIN] / [ATP] ratio for mtRNAPs vs. E. coli RNAP and S. cerevisiae RNAP II. Top, tailed template. Grey box and arrow indicate TSS. Bottom, dependence of NCIN-mediated capping with NAD+ and NADH on [NCIN] / [ATP] ratio for S. cerevisiae mtRNAP (Sce mtRNAP), human mtRNAP, E. coli RNAP (Eco RNAP) and S. cerevisiae RNAP II (Sce RNAP II) (mean ± SD; n = 3).

https://doi.org/10.7554/eLife.42179.006

The observed efficiencies of NCIN-mediated initiation with NAD+ or NADH by mtRNAPs are substantially higher than the highest previously reported efficiencies for NCIN-mediated initiation with NAD+ or NADH by cellular RNAPs (~15%; Bird et al., 2017; Bird et al., 2016; Vvedenskaya et al., 2018). To enable direct comparison of efficiencies of NCIN capping by mtRNAPs vs. cellular RNAPs on the same templates under identical reaction conditions, we performed transcription assays using a ‘tailed’ template (Figure 2C, top) that bypasses the requirement for sequence-specific RNAP-DNA interactions and transcription-initiation factor-DNA interactions for transcription initiation (Dedrick and Chamberlin, 1985; Kadesch and Chamberlin, 1982). In these experiments, we observe efficiencies of NCIN-mediated initiation with NAD+ and NADH by mtRNAP that are fully ~10 to ~40 fold higher than efficiencies of NCIN-mediated initiation with NAD+ and NADH by E. coli RNAP and S. cerevisiae RNAP II (Figure 2C, bottom). We conclude that S. cerevisiae mtRNAP and human mtRNAP cap RNA with NAD+ and NADH more efficiently than bacterial RNAP and eukaryotic nuclear RNAP II.

We next used the same tailed template and reaction conditions as in assays performed with mtRNAPs to determine the efficiency of NCIN-mediated initiation with NAD+ and NADH for the single-subunit RNAP of bacteriophage T7 (T7 RNAP) (Figure 2—figure supplement 2). The efficiencies of NCIN-mediated initiation with NAD+ and NADH by T7 RNAP were nearly as high as the efficiencies of NCIN-mediated initiation by mtRNAPs. We conclude that there is a quantitative difference in the efficiency of NCIN capping between members of the single-subunit RNAP family (T7 RNAP and mtRNAPs) and members of the multi-subunit RNAP family (bacterial RNAP and eukaryotic nuclear RNAP II).

Promoter sequence determines efficiency of RNA capping by mtRNAP

In previous work, we have shown that NCIN capping with NAD+ and NADH by bacterial RNAP is determined by promoter sequence, particularly at and immediately upstream of, the transcription start site (TSS) (Bird et al., 2016; Vvedenskaya et al., 2018). NCIN capping by bacterial RNAP occurs only at promoters where the base pair (nontemplate-strand base:template-strand base) at the TSS is A:T (+1A promoters), and occurs most efficiently at the subset of +1A promoters where the base pair immediately upstream of the TSS is purine:pyrimidine (−1R promoters). We have further shown that sequence determinants for NCIN capping by bacterial RNAP reside within the template strand of promoter DNA (i.e., the strand that templates incoming nucleotide substrates) (Vvedenskaya et al., 2018).

To determine whether the specificity for A:T at the TSS (position +1), observed with bacterial RNAP, also is observed with mtRNAP, we assessed NAD+ capping by S. cerevisiae mtRNAP using promoter derivatives having A:T or G:C at position +1 (Figure 3A–B). We observed NAD+ capping in reactions performed using the promoter derivative having A:T at position +1, but not in reactions performed using the promoter derivative having G:C at position +1 (Figure 3B), indicating specificity for A:T at position +1. To determine whether specificity resides in the template strand for A:T at position +1, we analyzed NAD+ capping with S. cerevisiae mtRNAP using template derivatives having noncomplementary nontemplate- and template-strand-nucleotides (A/C or G/T) at position +1 (Figure 3B). We observed NAD+ capping only with the promoter derivative having T as the template strand base at position +1, indicating that specificity at position +1 resides in the template strand.

Figure 3 with 1 supplement see all
Promoter sequence determines efficiency of RNA capping with NAD+ by mtRNAP.

(A) S. cerevisiae mitochondrial 21S promoter DNA depicted in the context of the mtRNAP-promoter open complex. DNA nontemplate strand (NT) on top; DNA template strand (T) on bottom; Unwound, non-base-paired DNA region, ‘transcription bubble,’ indicated by raised and lowered nucleotides; +1 and grey boxes, bases at the TSS; −1, bases immediately upstream of the TSS (the 21S promoter is a −1Y promoter). (B) Products of transcription reactions with NAD+ as initiating nucleotide and [α32P]-CTP as extending nucleotide for templates having complementary or non-complementary nucleotides at position +1. (C) Dependence of NAD+ capping on [NAD+] / [ATP] ratio for homoduplex templates having A:T, G:C, T:A, or C:G at position −1 relative to TSS (mean ± SD; n = 3). Red, −1R promoters; black, −1Y promoters. (D) Dependence of NAD+ capping on [ATP] / [NAD+] ratio for heteroduplex templates having an abasic site (*) on the DNA nontemplate strand (mean ± SD; n = 3). Red, promoters with a template-strand Y; black, promoters with a template-strand R. (E) Sequence preferences at position −1 for S. cerevisiae mtRNAP, E. coli RNAP, and T7 RNAP. Graphs show normalized values of (kcat/KM)NAD+ / (kcat/KM)ATP determined for homoduplex templates having A:T, G:C, T:A, or C:G at position −1 (mean ± SD; n = 3). Normalized values were calculated by dividing the value for each individual promoter by the average value measured for −1R promoters. Data for S. cerevisiae mtRNAP are from panel C, data for E. coli RNAP are from (Vvedenskaya et al., 2018), and data for T7 RNAP are from Figure 3—figure supplement 1.

https://doi.org/10.7554/eLife.42179.011

To determine whether specificity for R:Y at position −1, observed with bacterial RNAP, also is observed with mtRNAP, we analyzed NAD+ capping by S. cerevisiae mtRNAP using promoter derivatives having either R:Y (A:T or G:C) or Y:R (C:G or T:A) at position −1 (Figure 3C). We observed higher efficiencies of NAD+ capping with promoter derivatives having R:Y at position −1 than with promoter derivatives having Y:R (Figure 3C). To determine whether specificity at position −1 resides in the DNA template strand, we performed experiments using promoter derivatives having Y (C or T) or R (A or G) at position −1 of the template strand and having an abasic site (*) on the nontemplate strand (Figure 3D). We observed higher efficiencies of NAD+ capping in reactions performed using promoter derivatives having Y at template-strand position −1 than with those having R. Furthermore, within error, the capping efficiencies for promoter derivatives having Y or R at template-strand position −1 matched the capping efficiencies for homoduplex promoter derivatives (Figure 3C–D), indicating that sequence information for NAD+ capping with S. cerevisiae mtRNAP resides exclusively in the template strand.

We conclude that NCIN capping with NAD+ by mtRNAP is determined by the sequence at, and immediately upstream of, the TSS (positions +1 and −1, respectively). We further conclude that the sequence and strand preferences at positions +1 and −1 for NCIN capping with NAD+ by mtRNAP match the sequence and strand preferences observed for bacterial RNAP (Figure 3C–E) (Bird et al., 2016; Vvedenskaya et al., 2018), suggesting that these sequence and strand preferences may be universal determinants of NCIN capping with NAD+ for all RNAPs. Consistent with this hypothesis, we find that sequence preferences for NCIN capping with NAD+ by bacteriophage T7 RNAP, another member of the single-subunit RNAP family, match the sequence preferences observed for S. cerevisiae mtRNAP and bacterial RNAP (Figure 3E and Figure 3—figure supplement 1). Further consistent with this hypothesis, structural modeling suggests the basis for these sequence and strand preferences is universal: specifically, a strict requirement for template-strand +1T for base pairing to the NAD+ adenine moiety, and a preference for template strand −1Y for ‘pseudo’ base pairing to the NAD+ nicotinamide moiety (Bird et al., 2016; Vvedenskaya et al., 2018).

Detection and quantitation of NAD+- and NADH-capped mitochondrial RNA in vivo: boronate affinity electrophoresis with processed RNA and synthetic standards

Kössel, Jäschke and co-workers have demonstrated that boronate affinity electrophoresis allows resolution of uncapped RNAs from capped RNAs--such as m7G, NAD+ and NADH--that contain a vicinal-diol moiety (Igloi and Kössel, 1985; Igloi and Kössel, 1987; Nübel et al., 2017). However, the procedures of Kössel, Jäschke and co-workers have two limitations: (i) boronate affinity electrophoresis does not allow resolution of RNAs longer than ~200 nt, and (ii) boronate affinity electrophoresis, by itself, is unable to distinguish between different vicinal-diol containing cap structures (m7G, NAD+, NADH, or others). Here, to overcome these limitations, we have combined boronate affinity electrophoresis with use of oligodeoxynucleotide-mediated RNA cleavage (‘DNAzyme’ cleavage) (Joyce, 2001) and use of synthetic NCIN-capped RNA standards generated using NCIN-mediated transcription initiation in vitro (Figure 4). Use of DNAzyme cleavage enables processing of long RNAs to yield defined, short, 5'-end-containing subfragments (Figure 4A). Use of synthetic NCIN-capped RNA standards enables distinction between capped species (Figure 4A,D).

Detection and quantitation of NAD+- and NADH-capped mitochondrial RNA in vivo: boronate affinity electrophoresis with DNAzyme-cleaved cellular RNA and DNAzyme-cleaved synthetic NCIN-capped RNA standards.

(A) Use of DNAzyme (DZ) to process RNA to yield a defined, short 5'-end-containing subfragment, in parallel in vivo (red) and in vitro (blue). Uncapped, 5'-triphosphate (ppp) end generated using ATP-mediated initiation; 5'-NAD+, NAD+-capped end generated using NAD+-mediated initiation; 5'-NADH, NADH-capped end generated using NADH-mediated initiation. (B) Use of boronate affinity electrophoresis to resolve 5'-uncapped, 5'-NAD+, and 5'-NADH containing RNAs. Grey, structure of phenylboronic acid (PB) polyacrylamide gel. (C) PB-polyacrylamide gel (left) and polyacrylamide gel (right) analysis of DNAzyme-generated 5'-end-containing subfragments of S. cerevisiae mitochondrial RNA COX2. Red, observed 5'-end-containing RNA subfragments resolved by PB-polyacylamide-gel (left) or not resolved by polyacrylamide gel (right); identities of these subfragments are defined in Panel D. (D) Comparison of electrophoretic mobilities of observed 5'-end-containing subfragments of COX2 RNA generated in vivo to 5'-end-containing subfragments of synthetic RNA standards generated in vitro. a, NAD+-capped RNA; b, NADH-capped RNA; c, uncapped RNA (mean ± SD; n = 3).

https://doi.org/10.7554/eLife.42179.015

To detect and quantify NCIN capping with NAD+ and NADH in mitochondrial RNA isolated from cells, we employed the following steps: (i) DNAzyme cleavage of target RNAs to generate 5'-end-containing subfragments < 80 nt in length (Figure 4A, top); (ii) DNAzyme treatment of synthetic NAD+- and NADH-capped RNA standards having sequences identical to RNAs of interest (Figure 4A, bottom); (iii) boronate affinity electrophoresis of DNAzyme-generated subfragments of mitochondrial RNA and DNAzyme-generated subfragments of synthetic NAD+- and NADH-capped RNA standards having sequences identical to RNAs of interest (Figure 4B); and (iv) detection of DNAzyme-generated 5'-end-containing subfragments of mitochondrial RNAs and synthetic RNA standards by hybridization with a radiolabeled oligodeoxyribonucleotide probe (Figure 4C–D).

We selected for analysis two S. cerevisiae mitochondrial RNAs that previously had been detected as NAD+-capped: COX2 and 21S (Walters et al., 2017). We isolated S. cerevisiae total RNA and analyzed COX2 and 21S RNAs using the procedure described in the preceding paragraph. The results are presented in Figures 4C–D and 5A (top). For both COX2 and 21S RNAs, we detect at least one RNA species with electrophoretic mobility retarded as compared to that of uncapped RNA, indicating the presence of capped RNA. Treatment with the decapping enzyme NudC eliminates these species, confirming the presence of capping. Comparison of the electrophoretic mobilities to those of synthetic NAD+- and NADH-capped RNA standards indicates that one of the capped species is NAD+-capped RNA, and the other capped species, present under these growth conditions only for COX2 RNA, is NADH-capped RNA. The results show that both COX2 and 21S RNAs are present in NAD+-capped forms, and that COX2 RNA also is present in an NADH-capped-form. Because the hybridization probe detects RNA fragments that contain 5' ends generated by transcription initiation (red subfragment depicted in Figure 4A), the detected NAD+ and NADH caps are concluded to be at 5' ends generated by transcription initiation, as opposed to 5' ends generated by RNA processing. The results confirm that S. cerevisiae mitochondrial RNAs undergo NAD+ capping in cells, show that S. cerevisiae mitochondrial RNAs undergo NAD+ capping at 5' ends generated by transcription initiation (as opposed to 5' ends generated by RNA processing), and show that S. cerevisiae mitochondrial RNAs also undergo NADH capping in cells.

Detection and quantitation of NAD+- and NADH-capped mitochondrial RNA in vivo: effects of intracellular NADand NADH levels in S. cerevisiae and human cells.

(A) Changes in intracellular NAD+/NADH ratios result in changes in levels of NAD+- and NADH-capped mitochondrial RNA (mean ± SD; n = 3). Gel images show representative data for S. cerevisiae COX2 RNA (left) and 21S RNA (right). Blue annotations as in Figure 4. (B) Changes in intracellular NAD(H) levels result in changes in levels of NAD+- and NADH-capped mitochondrial RNA (mean ± SD; n = 3). Gel images show representative data for LSP-derived RNAs. Red, NAD(H) biosynthesis inhibitor FK866; NAMPT, Nicotinamide phosphoribosyltransferase; NMNAT, Nicotinamide mononucleotide adenylyltransferase.

https://doi.org/10.7554/eLife.42179.017

The observed levels of NAD+- and NADH-capping of S. cerevisiae mitochondrial RNAs are remarkably high. For COX2 RNA, NAD+-capped RNA comprises ~50% of the total COX2 RNA pool and NADH-capped RNA comprises ~10% of the total COX2 RNA pool (Figures 4C–D and 5A, top). For 21S RNA, NAD+-capped RNA comprises ~30% of the total 21S RNA pool (Figure 5A, top). These levels of NCIN capping are ~2 to ~50 times higher than levels of NCIN-capping in exponentially growing E. coli (less than 1% to ~20% for NAD+-capping; not previously detected for NADH-capping) (Bird et al., 2016; Cahová et al., 2015; Nübel et al., 2017; Vvedenskaya et al., 2018).

We performed analogous experiments analyzing RNAs produced by transcription from the human mitochondrial LSP promoter (Figure 5B, top). We isolated and analyzed total RNA from HEK293T cells. We observed an NAD+-capped species comprising ~15% of the total LSP-derived RNA pool (Figure 5B, top). The results establish that human mitochondrial RNAs undergo NAD+ capping in cells and show that human mitochondrial RNAs undergo NAD+ capping at 5' ends generated by transcription initiation (as opposed to 5' ends generated by RNA processing).

Detection and quantitation of NAD+- and NADH-capped mitochondrial RNA in vivo: mtRNAPs serve as both sensors and actuators in coupling cellular metabolism to mitochondrial gene expression

Mitochondria are the primary locus of metabolism and energy transformation in the eukaryotic cell, serving as the venue for the tricarboxylic acid cycle (TCA) cycle and oxidative phosphorylation. The TCA cycle reduces NAD+ to NADH and oxidative phosphorylation oxidizes NADH to NAD+. Our finding that mtRNAPs perform NCIN capping with NAD+ and NADH at efficiencies that vary in a simple mass-action-dependent fashion with [NAD+] / [ATP] and [NADH] / [ATP] ratios in vitro (Figures 1 and 2), and our finding that mtRNAPs perform efficient NCIN capping to yield NAD+- and NADH-capped mitochondrial RNAs in vivo (Figures 4 and 5), raise the possibility that mtRNAPs may serve as both sensors and actuators in coupling metabolism to mitochondrial gene expression in vivo.

As a first test of this hypothesis, we assessed whether changing intracellular [NAD+] / [NADH] ratios results in changes in NAD+ and NADH capping of mitochondrial RNAs. We isolated total RNA from S. cerevisiae grown under conditions that result in either high or low [NAD+] / [NADH] ratios (Bekers et al., 2015; Canelas et al., 2008): respiration (glycerol/ethanol; aerobic) or fermentation (glucose; anaerobic). We analyzed the same two mitochondrial RNAs as above: COX2 and 21S (Figure 5A). We observed marked changes in levels of NAD+ and NADH capping for both analyzed mitochondrial RNAs. For COX2 RNA, on changing from the growth condition yielding a high [NAD+] / [NADH] ratio to the growth condition yielding a low [NAD+] / [NADH] ratio, we observe a decrease in levels of NAD+ capping (from ~50% to ~20%) and an anti-correlated increase in the level of NADH capping (from ~10% to ~40%). Notably, the total level of NAD+ and NADH capping, NAD(H) capping, remains constant under the two conditions (~60%), indicating that the relative levels of NCIN-mediated initiation and ATP-mediated initiation do not change (Figure 5A). For 21S RNA, the same pattern is observed (Figure 5A): on changing from the growth condition yielding a high [NAD+] / [NADH] ratio to the growth condition yielding a low [NAD+] / [NADH] ratio, the level of NAD+ capping decreases (from ~30% to ~10%), the level of NADH capping increases (from 0% to ~20%), and the total level of NAD+ and NADH capping remains constant (~30%). The results indicate that changing the [NAD+] / [NADH] ratio changes transcription outputs in vivo.

As a second test of this hypothesis, we assessed whether changing intracellular total NAD(H) levels results in changes in NCIN capping of mitochondrial RNAs (Figure 5B). We isolated RNA from human HEK293T cells grown under conditions yielding either high or low intracellular NAD(H) levels: standard growth media or growth media in the presence of the NAD(H)-biosynthesis inhibitor FK866 (Hasmann and Schemainda, 2003; Khan et al., 2006). We analyzed the same LSP-derived mitochondrial RNA as in the preceding section (Figure 5B). On changing from the growth condition yielding high intracellular NAD(H) levels to the growth condition yielding low NAD(H) levels, we observe a marked change in the total level of NCIN capping (from ~15% to 0%). The results indicate that changing NAD(H) levels changes levels of NCIN-capped mitochondrial RNAs in vivo.

Taken together, the results of the two experiments in Figure 5 indicate that mtRNAP serves as both sensor and actuator in coupling [NAD+] / [NADH] ratios to relative levels of NAD+- and NADH-capped mitochondrial RNAs (Figure 5A), and in coupling total NAD(H) levels to total levels of NCIN-capped mitochondrial RNAs (Figure 5B), thereby coupling cellular metabolism to mitochondrial transcription outputs. We suggest that mtRNAPs serve as sensors through their mass-action-dependence in selecting NAD+ vs. NADH vs. ATP as the initiating entity during transcription initiation, and serve as actuators by incorporating NAD+ vs. NADH vs. ATP at the RNA 5' end during transcription initiation.

Discussion

Our results show that S. cerevisiae and human mtRNAPs cap RNA with NAD+ and NADH (Figure 1), show that S. cerevisiae and human mtRNAPs cap RNA with NAD+ and NADH more efficiently than bacterial and eukaryotic nuclear RNAPs (Figure 2), and show that capping efficiency by mtRNAPs is determined by promoter sequence (Figure 3). Our results further show that the proportions of mitochondrial RNAs that are capped with NAD+ and NADH are remarkably high--up to ~60% (Figures 4 and 5)--and that these proportions change in response to cellular NAD+ and NADH levels (Figure 5).

We and others previously have shown that NCIN capping by cellular RNAPs has functional consequences, including modulating RNA stability and modulating RNA translatability (Bird et al., 2016; Cahová et al., 2015; Jiao et al., 2017). Our results here showing that S. cerevisiae and human mitochondrial RNAs are capped at substantially higher levels than non-mitochondrial RNAs--up to ~60% for analyzed S. cerevisiae mitochondrial RNAs and up to ~15% for analyzed human mitochondrial RNAs (Figures 4 and 5)--suggest that NCIN capping in mitochondria occurs at a higher efficiency, and has a higher importance, than NCIN capping in other cellular compartments. Four other considerations support this hypothesis. First, mtRNAPs are substantially more efficient at NAD+ and NADH capping than bacterial and eukaryotic nuclear RNAPs (Figure 2C). Second, levels of NAD+ and NADH relative to ATP in mitochondria are substantially higher than levels of NAD+ and NADH relative to ATP in bacteria and eukaryotic nuclei (Cambronne et al., 2016; Chen et al., 2016; Park et al., 2016). Third, all S. cerevisiae and human mitochondrial promoters are +1A promoters (18 promoters in S. cerevisiae mitochondria; two promoters in human mitochondria) (Biswas, 1999; Chang and Clayton, 1984; Taanman, 1999), in contrast to bacterial and eukaryotic nuclear RNAP promoters, for which approximately half are +1A promoters (Chen et al., 2013; Haberle et al., 2014; Saito et al., 2013; Thomason et al., 2015; Tsuchihara et al., 2009). Fourth, we observe capping with both NAD+ and NADH for mitochondrial RNAs in vivo (Figures 4 and 5), whereas, to date, capping has been observed with only NAD+ for non-mitochondrial RNAs in vivo, raising the possibility that, in mitochondria, but not in other cellular compartments, NAD+ and NADH caps dictate different RNA fates and, correspondingly, different transcription outputs.

Our results showing that levels of NAD+ and NADH capping by mtRNAP correlate with changes in intracellular levels of NAD+ and NADH (Figure 5) indicate that mtRNAP serves as a sensor, sensing [NAD+] / [ATP] and [NADH] / [ATP] ratios, when selecting initiating entities and, simultaneously, serves as an actuator by altering RNA 5' ends when selecting initiating entities. Because all S. cerevisiae and human mitochondrial promoters are +1A promoters, this dual sensor/actuator activity of mtRNAP will occur at, and couple metabolism to gene expression at, all S. cerevisiae and human mitochondrial promoters. This dual sensor/actuator activity of mtRNAP obviates the need for a dedicated signal-processing machinery for coupling metabolism to gene expression, instead employing a pan-eukaryotic housekeeping enzyme for signal processing. As such, this dual sensor/actuator activity of mtRNAP provides a remarkably economical, parsimonious mechanism of coupling metabolism to gene expression.

Materials and methods

Key resources table
Reagent type
(species) or
resource
DesignationSource or
reference
IdentifiersAdditional information
Strain, strain
background
(E. coli)
BL21(DE3) bacteriaNEBC2527H
Strain, strain
background
(E. coli)
NiCo21(DE3) bacteriaNEBC2529H
Strain, strain
background
(E. coli)
Artic Express (DE3)
bacteria
Fisher ScientificNC9444283
Strain, strain
background
(S. cerevisiae)
246.1.1
(MATa ura3 trp1 leu2 his4)
Gift of A. Vershon
Cell line
(human)
HEK293T
(human embryonic
kidney cells)
ATCCCRL-3216
Recombinant
DNA reagent
pIA900Gift of I.
Artsimovitch
Recombinant
DNA reagent
pET NudC-His(Bird et al., 2016)
Recombinant
DNA reagent
pJJ1399gift of J. Jaehning
Recombinant
DNA reagent
pTrcHisC-Mtf1gift of J. Jaehning
Recombinant
DNA reagent
pPROEXHTb-POLRMT
(43–1230)−6xHis
(Ramachandran et al., 2017)
Recombinant
DNA reagent
pPROEXHTb-TFAM
(43-245)−6xHis
(Ramachandran et al., 2017)
Recombinant
DNA reagent
pT7TEV-HMBP4(Yakubovskaya et al., 2014)
Recombinant
DNA reagent
pAR1219(Jia et al., 1996)
Sequence-based
reagent
DK64Integrated DNA
Technologies (IDT)
tailed template with
PEG6 linker
GGCTCGCCTCGGCTCG/iSp18/
CGAGCCGAGGCGAGCGTCACCAA
Sequence-based
reagent
JB459IDThuman LSP DNA template
 + 1 AGU variant nontemplate
strand
GTGTTAGTTGGGGGGTGACTGTT
AAAAGTGCATACCGCCAAAGTATA
AAATTTGTGGGCC
Sequence-based
reagent
JB460IDThuman LSP DNA template
+ 1 AGU variant template
strand
GGCCCACAAATTTTATACTTTGGC
GGTATGCACTTTTAACAGTCACCC
CCCAACTAACAC
Sequence-based
reagent
JB469IDTT7φ2.5–35 n nontemplate
strand (−1T)
CAGTAATACGACTCACTATTAGCGAA
GCGGGCATGCGGCCAGCCATAGC
CGATCA
Sequence-based
reagent
JB470IDTT7φ2.5–35 n template
strand (−1A)
TGATCGGCTATGGCTGGCCGCATGCC
CGCTTCGCTAATAGTGAGTCGTA
TTACTG
Sequence-based
reagent
JB471IDTT7φ2.5–35 n nontemplate
strand (−1A)
CAGTAATACGACTCACTATAAGCGAAGC
GGGCATGCGGCCAGCCATAG
CCGATCA
Sequence-based
reagent
JB472IDTT7φ2.5–35 n template
strand (−1T)
TGATCGGCTATGGCTGGCCGCATGCCC
GCTTCGCTTATAGTGAGTCGTATTACTG
Sequence-based
reagent
JB473IDTT7φ2.5–35 n nontemplate
strand (−1G)
CAGTAATACGACTCACTATGAGCGAAG
CGGGCATGCGGCCAGCCATAG
CCGATCA
Sequence based
reagent
JB474IDTT7φ2.5 35 n template
strand (−1C)
TGATCGGCTATGGCTGGCCGCATGCC
CGCTTCGCTCATAGTGAGTCGTATTACTG
Sequence-based
reagent
JB475IDTT7φ2.5–35 n nontemplate
strand (−1C)
CAGTAATACGACTCACTATCAGCGAA
GCGGGCATGCGGCCAGCCA
TAGCCGATCA
Sequence-based
reagent
JB476IDTT7φ2.5–35 n template
strand (−1G)
TGATCGGCTATGGCTGGCCGCATGC
CCGCTTCGCTGATAGTG
AGTCGTATTACTG
Sequence-based
reagent
JB515IDTprobe for human LSP-generated
RNA (complementary to
positions + 2 to+31)
CACCAGCCTAACCAGATTTCAA
ATTTTATC
Sequence-based
reagent
JB525IDTprobe for S. cerevisiae
21S RNA (complementary
to positions + 9 to+42)
CTATATAATAAATATTTCAAATC
TATTATTCTAC
Sequence-based
reagent
JB526IDTS. cerevisiae 21S RNA
DNAzyme; cleaves transcript
at position + 53
ACTCCATGATTAGGCTAGCTACAA
CGACTCTTTAAATCT
Sequence-based
reagent
JB555IDTprobe for S. cerevisiae
COX2 RNA (complementary
to positions + 8 to+46)
ATCTTAACCTTTAGACTCTTTTGTC
TATTTATAATATGT
Sequence-based
reagent
JB557IDTS. cerevisiae COX2 DNAzyme;
cleaves at position + 57
TCTTAATAAATCTAAGGCTAGCTACA
ACGAATTTTAATAAATCTT
Sequence-based
reagent
JB559IDThuman LSP-generated RNA
DNAzyme; cleaves at
position + 67
GCACTTAAACAGGCTAGCTACAA
CGAATCTCTGCCA
Sequence-based
reagent
JB560IDTS. cerevisiae COX2 −40
to + 125 nontemplate strand
oligo (for generation of
in vitro transcription template)
TATATAATAATAAATTATAAATAAATTTT
AATTAAAAGTAGTATTAACATATTATAAA
TAGACAAAAGAGTCTAAAGGTTAAGATT
TATTAAAATGTTAGATTTATTAAGATTAC
AATTAACAAC
Sequence-based
reagent
JB561IDTS. cerevisiae COX2 −40 to + 3
forward primer (for generation
of in vitro transcription template)
TATATAATAATAAATTATAAATAAATTTT
AATTAAAAGTAGT
Sequence-based
reagent
JB562IDTS. cerevisiae COX2 + 83 to+125
reverse primer (for generation
of in vitro transcription template)
GTTGTTAATTGTAATCTTAATAAATCTAA
CATTTTAATAAATC
Sequence-based
reagent
UB1IDThuman LSP DNA template (−43
to + 19) nontemplate strand
ATGTGTTAGTTGGGGGGTGACTGTTAA
AAGTGCATACCGCCAAAAGATAAAATT
TGAAATCTG
Sequence-based
reagent
UB2IDThuman LSP DNA template
(−43 to + 19) template
strand
CAGATTTCAAATTTTATCTTTTGGCGGT
ATGCACTTTTAACAGTCACCCCCCAAC
TAACACAT
Sequence-based
reagent
UB3IDThuman HSP1 DNA template
(−43 to + 20) nontemplate
strand
ACACACCGCTGCTAACCCCATACCCCGA
ACCAACCAAACCCCAAAGACACCCGCC
ACAGTTTA
Sequence-based
reagent
UB4IDThuman HSP1 DNA template
(−43 to + 20) template
strand
TAAACTGTGGCGGGTGTCTTTGGGGT
TTGGTTGGTTCGGGGTATGGGGTTA
GCAGCGGTGTGT
Sequence-based
reagent
UB5IDTS. cerevisiae 15S DNA
template (−25 to + 1;
C-less cassette) nontemplate
strand
ATAATTTATTTATTATTATATAAGTAAT
AAATAATTGTTTTATATAATAAGAA
TTCTCCTTC
Sequence-based
reagent
UB6IDTS. cerevisiae 15S DNA template
(−25 to + 1; C-less cassette)
template strand
GAAGGAGAATTCTTATTATATAAAACA
ATTATTTATTACTTATATAATAATAA
ATAAATTAT
Sequence-based
reagent
UB7IDTS. cerevisiae 21S DNA
template (−25 to + 1;
C-less cassette) nontemplate
strand
TATTATTATTATTATATATATAAGTAG
TAAAAAGTAGAATAATAGATTT
GAAATACC
Sequence-based
reagent
UB8IDTS. cerevisiae 21S DNA
template (−25 to + 1;
C-less cassette) template
strand
GAAGGAGACCAACCACAAACACACA
ACAACCACCAACTACTTATATAATAA
TAAATAAATTAT
Sequence-based
reagent
UB9IDTS. cerevisiae 21S DNA
template (−25 to + 1; C-less
and A-less cassette) nontemplate
strand
ATAATTTATTTATTATTATATAAGTAG
TTGGTGGTTGTTGTGTGTTTGTG
GTTGGTCTCCTTC
Sequence-based
reagent
UB10IDTS. cerevisiae 21S DNA
template (−25to + 1;
C-less and A-less cassette)
template strand
GAAGGAGACCAACCACAAACACAC
AACAACCACCAACTACTTATATAA
TAATAAATAAATTAT
Sequence-based
reagent
UB11IDTS. cerevisiae 21S
nontemplate strand
(−1A)
ATAATTTATTTATTATTATATAAGAA
GTTGGTGGTTGTTGTGTGTTTGTG
GTTGGTCTCCTTC
Sequence-based
reagent
UB12IDTS. cerevisiae 21S template
strand (−1T)
GAAGGAGACCAACCACAAACACA
CAACAACCACCAACTTCTTATATA
ATAATAAATAAATTAT
Sequence-based
reagent
UB13IDTS. cerevisiae 21S
nontemplate strand
(−1G)
ATAATTTATTTATTATTATATAAGG
AGTTGGTGGTTGTTGTGTGTTTG
TGGTTGGTCTCCTTC
Sequence-based
reagent
UB14IDTS. cerevisiae 21S
template strand
(−1C)
GAAGGAGACCAACCACAAACACA
CAACAACCACCAACTCCTTATAT
AATAATAAATAAATTAT
Sequence-based
reagent
UB15IDTS. cerevisiae 21S
nontemplate strand
(−1C)
ATAATTTATTTATTATTATATAAG
CAGTTGGTGGTTGTTGTGTGT
TTGTGGTTGGTCTCCTTC
Sequence-based
reagent
UB16IDTS. cerevisiae 21S
template strand
(−1G)
GAAGGAGACCAACCACAAACACA
CAACAACCACCAACTGCTTATATA
ATAATAAATAAATTAT
Sequence-based
reagent
UB17IDTS. cerevisiae 21S
nontemplate strand
(+1C)
ATAATTTATTTATTATTATATAAGTC
GTTGGTGGTTGTTGTGTGTTTGT
GGTTGGTCTCCTTC
Sequence-based
reagent
UB18IDTS. cerevisiae 21S
template strand
(+1G)
GAAGGAGACCAACCACAAACACAC
AACAACCACCAACGACTTATATAA
TAATAAATAAATTAT
Sequence-based
reagent
JB527IDTS. cerevisiae 21S
nontemplate strand
(−1 abasic)
ATAATTTATTTATTATTATATAAG/
idSp/AGTTGGTGGTTGTTGTGTGT
TTGTGGTTGGTCTCCTTC
Peptide, recombinant
protein (S. cerevisiae)
Rpo41 (mtRNAP)(Tang et al., 2009)
Peptide, recombinant
protein (S. cerevisiae)
Mtf1(Paratkar and Patel, 2010)
Peptide, recombinant
protein (Human)
POLRMT (mtRNAP)(Ramachandran et al., 2017)
Peptide, recombinant
protein (Human)
TFAM(Ramachandran et al., 2017)
Peptide, recombinant
protein (Human)
TFB2(Yakubovskaya et al., 2014)
Peptide, recombinant
protein (S. cerevisiae)
RNA
polymerase II
Gift of C. Kaplan
Peptide, recombinant
protein (E. coli)
RNA polymerase
core (β'−6xHis)
(Artsimovitch et al., 2003)
Peptide, recombinant
protein
T7 RNA polymerase(Jia et al., 1996)
Peptide, recombinant
protein (E. coli)
NudC(Cahová et al., 2015)
Peptide, recombinant
protein
Phusion Flash HF
master mix
ThermoFisherF-548L
Peptide, recombinant
protein
T4 Polynucleotide
Kinase
NEBM0201L
Peptide, recombinant
protein
RNA 5'
pyrophosphohydrolase
(RppH)
NEBM0356S
Peptide, recombinant
protein
FastAP Alkaline
Phosphatase
Thermo FisherEF0651
Commercial assay
or kit
Monarch PCR and DNA
clean up kit
NEBT1030S
Chemical compound,
drug
Nuclease-free water
(not DEPC-treated)
ThermoFisherAM9932
Chemical compound,
drug
Bacto agarVWR90000–760
Chemical compound,
drug
Bacto tryptoneVWR90000–286
Chemical compound,
drug
Bacto yeast extractVWR90000–726
Chemical compound,
drug
D-Glucose
monhydrate
Amresco0643–1 kg
Chemical compound,
drug
GlycerolEMD Millipore55069521
Chemical compound,
drug
DMEM mediumThermo Fisher11965–092
Chemical compound,
drug
Fetal Bovine SerumAtlanta BiologicalS11150H
Chemical compound,
drug
dNTP solution mix,
10 mM of each NTP
NEBN0447S
Chemical compound,
drug
NTP set (ultra-pure),
100 mM solutions
GE Healthcare27-2025-01
Chemical compound,
drug
NAD+Roche
(Sigma-Aldrich)
10127965001
Chemical compound,
drug
NADHRoche
(Sigma-Aldrich)
10107735001
Chemical compound,
drug
Tris base (Amresco)VWR97061–800
Chemical compound,
drug
Boric Acid (ACS grade)VWR97061–980
Chemical compound,
drug
EDTA disodium salt
dyhydrate
VWR97061–018
Chemical compound,
drug
0.5 M EDTA pH 8ThermoFisherAM9260G
Chemical compound,
drug
Dibasic Sodium
phosphate
EMD MilliporeSX0715-1
Chemical compound,
drug
Sodium ChlorideEMD MilliporeSX0420-3
Chemical compound,
drug
Potassium ChlorideEMD Millipore7300–500 GM
Chemical compound,
drug
Sodium CitrateEMD Millipore7810–1 KG
Chemical compound,
drug
Sodium Acetate,
trihydrate
VWRMK736406
Chemical compound,
drug
Ficoll 400VWRAAB22095-18
Chemical compound,
drug
PolyvinylpyrrolidoneEMD Millipore7220–1 KG
Chemical compound,
drug
Diethyl Pyrocarbonate
(DEPC)
VWRAAB22753-14
Chemical compound,
drug
Formamide,
deionized
VWREM-4610
Chemical compound,
drug
Sodium
dodecylsulfate (SDS)
VWR97064–470
Chemical compound,
drug
Magnesium chloride
hexahydrate
VWREM-5980
Chemical compound,
drug
Magnesium sulfate
heptahydrate
VWREM-MX0070-1
Chemical compound,
drug
Glycerol (ACS grade)VWREMGX0185-5
Chemical compound,
drug
Bovine Serum Albumin
(BSA) fraction V
VWR101174–932
Chemical compound,
drug
Bromophenol BlueVWREM-BX1410-7
Chemical compound,
drug
Xylene CyanolSigma-AldrichX4126-10G
Chemical compound,
drug
Amaranth DyeVWR200030–400
Chemical compound,
drug
Temed (JT Baker)VWRJT4098-1
Chemical compound,
drug
Ammonium PersulfateVWR97064–594
Chemical compound,
drug
Dithiothreitol (DTT)Gold BioDTT50
Chemical compound,
drug
Glycogen from Oyster
(type II)
Sigma-AldrichG8751
Chemical compound,
drug
Hydrochloric Acid
(ACS plus)
Fisher ScientificA144-212
Chemical compound,
drug
Ethyl AlcoholPharmco-AAPER111000200
Chemical compound,
drug
GeneMate LE Quick
Dissolve agarose
BioExpressE-3119–500
Chemical compound,
drug
SequaGel sequencing
system
National
Diagnostics
EC833
Chemical compound,
drug
Nytran SuPerCharge
Nylon Membrane
VWR10416296
Chemical compound,
drug
SigmaSpin G25 cleanup
columns
Sigma-AldrichS5059
Chemical compound,
drug
32P NAD+ 250 uCiPerkin ElmerBLU023X250UC
Chemical compound,
drug
γ-32P ATP Easy
Tide 1 mCi
Perkin ElmerBLU502Z001MC
Chemical compound,
drug
α-32P CTP Easy
Tide 250 uCi
Perkin ElmerBLU508H250UC
Chemical compound,
drug
α-32P GTP Easy
Tide 250 uCi
Perkin ElmerBLU506H250UC
Chemical compound,
drug
α-32P UTP Easy
Tide 250 uCi
Perkin ElmerBLU507H250UC
Chemical compound,
drug
Decade MarkerThermo FisherAM7778
Chemical compound,
drug
TRI ReagentMolecular
Research Center
TR118
Chemical compound,
drug
Acid phenol:chloroform
(CHCl3) pH 4.5
ThermoFisherAM9720
Chemical compound,
drug
FK866 hydrochloride
hyrate
Sigma-AldrichF8557
Software, algorithmExcelMicrosoft365
Software, algorithmImageQuantGE HealthcareTL 5.1, TL v8.2
Software, algorithmSigmaPlotSystat Software Inc.Version 10
Software, algorithmPymolSchrodinger, LLChttp://www.pymol.org
Software, algorithmIllustratorAdobeVersion CS6
OtherTyphoon RBG ImagerGE Healthcare
OtherNanoDrop 2000C
spectrophotometer
Thermo Fisher
OtherUV CrosslinkerFisher ScientificFB-UVXL-1000
OtherHybridization oven 5420VWR97005–252
OtherSequi-Gen GT sequencing
systems (21 × 50) (38 × 30)
Bio-Rad1653871 and 1653873

Proteins

S. cerevisiae mtRNAP (Rpo41) was prepared from E. coli strain BL21(DE3) transformed with pJJ1399 (gift of Judith A. Jaehning) using culture and induction procedures, polyethyleneimine treatment, ammonium sulfate precipitation, Ni-sepharose, DEAE sepharose and Heparin-sepharose chromatography as in (Tang et al., 2009). S. cerevisiae Mtf1 was prepared from E. coli strain BL21(DE3) transformed with pTrcHisC-Mtf1 (Paratkar and Patel, 2010) and purified using culture and induction procedures, polyethyleneimine treatment, ammonium sulfate precipitation, and tandem DEAE sepharose and Ni-sepharose chromatography as in (Tang et al., 2009).

Human mtRNAP (POLRMT) and TFAM were purified from E. coli strain BL21(DE3) transformed with pPROEXHTb-POLRMT(43–1230)−6xHis (Ramachandran et al., 2017) or pPROEXHTb-TFAM(43-245)−6xHis (Ramachandran et al., 2017), respectively, using culture and induction procedures, polyethyleneimine treatment, ammonium sulfate precipitation, and Ni-sepharose and heparin-sepharose chromatography as in (Ramachandran et al., 2017). Human TFB2 was purified from E. coli strain ArcticExpress (DE3) (Stratagene) transformed with pT7TEV-HMBP4 (Yakubovskaya et al., 2014), using culture and induction procedures, polyethyleneimine treatment, ammonium sulfate precipitation, Ni-sepharose and heparin-sepharose chromatography, and size exclusion chromatography as in (Yakubovskaya et al., 2014).

T7 RNAP was prepared from E. coli strain BL21 transformed with pAR1219 using culture and inductions procedures, SP-Sephadex, CM-Sephadex and DEAE-Sephacel chromatography as described in (Jia et al., 1996).

E. coli RNAP core enzyme was prepared from E. coli strain NiCo21(DE3) transformed with plasmid pIA900 (Artsimovitch et al., 2003) using culture and induction procedures, immobilized-metal-ion affinity chromatography on Ni-NTA agarose, and affinity chromatography on Heparin HP as in (Artsimovitch et al., 2003).

S. cerevisiae RNA polymerase II core enzyme (gift of Craig Kaplan) was prepared as described in (Barnes et al., 2015).

E. coli NudC was prepared from E. coli strain NiCo21(DE3) transformed with plasmid pET NudC-His (Bird et al., 2016) using metal-ion chromatography and size-exclusion chromatography as in (Cahová et al., 2015).

RNA 5' pyrophosphohydrolase (RppH) and T4 polynucleotide kinase (PNK) were purchased from New England Biolabs (NEB). FastAP Thermosensitive Alkaline Phosphatase was purchased from Thermo Fisher Scientific. Molar concentrations of purified proteins were determined by light absorbance at 280 nm and the calculated respective molar extinction coefficients.

Oligodeoxyribonucleotides

Sequences of the oligodeoxyribonucleotides used in this work are provided in Key Resources Table. All oligodeoxyribonucleotides were purchased from Integrated DNA Technologies (IDT) with standard desalting purification unless otherwise specified.

Linear in vitro transcription templates used for transcription assays shown in Figures 1, 2 and 3 were generated by mixing complementary equimolar amounts of nontemplate- and template-strand DNA in 10 mM Tris HCl pH 8.0, incubating the mixture at 95°C for 5 min, and cooling the mixture by 0.5°C per minute to 25°C.

Transcription templates used to generate in vitro RNA standards for Northern analysis (Figures 4 and 5) were generated by PCR. Reactions contained a mixture of 5 nM template oligo, 0.5 µM forward primer, 0.5 µM reverse primer, and Phusion HF Master Mix (Thermo Scientific). Reaction products were isolated using a Monarch PCR and DNA cleanup kit (NEB).

The radiolabeled 10-nt marker labeled ‘M’ in gel images shown in Figures 1, 4 and 5 was generated using the Decade Marker System (Thermo Fisher Scientific), PNK (NEB) and [γ32P]-ATP (Perkin Elmer; 6,000 Ci/mmol).

In vitro transcription assays

Assays performed with S. cerevisiae mtRNAP were based on procedures described in (Deshpande and Patel, 2014). Assays performed with human mtRNAP were based on procedures described in (Ramachandran et al., 2017).

For initial RNA product assays in Figure 1C, Figure 1—figure supplement 1, and Figure 3, 1 µM DNA template, 1 µM S. cerevisiae mtRNAP, and 1 µM Mtf1 were incubated at 25°C for 10 min in Sce-mtRNAP reaction buffer (50 mM Tris-acetate pH 7.5, 100 mM potassium glutamate, 10 mM magnesium acetate, 0.01% Tween-20, 1 mM DTT, and 5% glycerol). A mixture containing the initiating nucleotide (200 µM ATP, 1 mM NAD+, or 1 mM NADH) and extending nucleotide (10 µM of non-radiolabeled GTP plus 6 mCi of [α32P]-GTP [Perkin Elmer; 3,000 Ci/mmol]) was added, and assays were incubated at 25°C for 30 min. For assays in Figure 1CFigure 1—figure supplement 1 and radiolabeled initial products were isolated using a Nanosep 3 kDa cutoff spin concentrator (Pall). For assays in Figure 3B, reactions were stopped with 10 µl RNA loading dye (95% deionized formamide, 18 mM EDTA, 0.25% SDS, xylene cyanol, bromophenol blue, amaranth) and were analyzed by electrophoresis on 7.5 M urea, 1x TBE, 20% polyacrylamide gels (UreaGel System; National Diagnostics), followed by storage-phosphor imaging (Typhoon 9400 variable-mode imager; GE Life Science).

For the initial RNA product assays in Figure 1D and Figure 1—figure supplement 1, 1 µM DNA template, 1 µM human mtRNAP, 1 µM TFAM, and 1 µM TFB2M were incubated at 25°C for 10 min in human-mtRNAP reaction buffer (50 mM Tris-acetate pH 7.5, 50 mM sodium glutamate, 10 mM magnesium acetate, 1 mM DTT, and 0.05% Tween-20). A mixture containing the initiating nucleotide (200 µM ATP, 1 mM NAD+, or 1 mM NADH) and extending nucleotide (10 µM of non-radiolabeled GTP plus 6 mCi of [α32P]-GTP at [Perkin Elmer; 3,000 Ci/mmol]) was added, and assays were incubated at 25°C for 60 min. Radiolabeled initial RNA products were isolated using a Nanosep 3 kDa cutoff spin concentrator (Pall).

A portion of the recovered initial RNA products were mixed with either 10 U of RppH or 400 nM NudC and incubated at 37°C for 30 min. Reactions were stopped by addition of 10 µl RNA loading dye. Samples were analyzed by electrophoresis on 7.5 M urea, 1x TBE, 20% polyacrylamide gels (UreaGel System; National Diagnostics), followed by storage-phosphor imaging (Typhoon 9400 variable-mode imager; GE Life Science).

For full-length product assays in Figure 1C and Figure 1—figure supplement 1 (panel B), 1 µM DNA template, 1 µM S. cerevisiae mtRNAP, and 1 µM Mtf1 were incubated at 25°C for 10 min in Sce-mtRNAP reaction buffer. A mixture containing the initiating nucleotide (1 mM ATP, 1 mM NAD+, or 1 mM NADH for Figure 1C; 200 µM non-radiolabeled ATP plus 10 µCi [γ32P]-ATP [Perkin Elmer; 6,000 Ci/mmol] or 1 mM NAD+ plus 20 µCi [α32P]-NAD+ [Perkin Elmer; 800 Ci/mmol] for Figure 1—figure supplement 1) and extending nucleotides (200 µM GTP, 200 µM 3'-deoxy-CTP, 20 µM ATP, 200 µM non-radiolabeled UTP, and 6 mCi of [α32P]-UTP [Perkin Elmer; 3000 Ci/mmol] for Figure 1C; 200 µM GTP, 200 µM 3'-deoxy-CTP, 20 µM ATP, 200 µM UTP for Figure 1—figure supplement 1) was added, and assays were incubated at 25°C for 30 min. Reactions were stopped by addition of stop solution (0.6 M Tris HCl pH 8.0, 18 mM EDTA, 0.1 mg/ml glycogen), samples were extracted with acid phenol:chloroform (5:1, pH 4.5; Thermo Fisher Scientific), and RNA products were recovered by ethanol precipitation and resuspended in NudC reaction buffer (10 mM Tris HCl pH 8.0, 50 mM NaCl, 10 mM MgCl2, 1 mM DTT).

For full-length product assays in Figure 1D and Figure 1—figure supplement 1 (panel C), 1 µM DNA template, 1 µM human mtRNAP, 1 µM TFAM, and 1 µM TFB2M were incubated at 25°C for 10 min in human-mtRNAP reaction buffer. A mixture containing the initiating nucleotide (1 mM ATP, 1 mM NAD+, or 1 mM NADH for Figure 1D; 200 µM non-radiolabeled ATP plus 10 µCi [γ32P]-ATP [Perkin Elmer; 6000 Ci/mmol] or 1 mM NAD+ plus 20 µCi [α32P]-NAD+ [Perkin Elmer; 800 Ci/mmol] for Figure 1—figure supplement 1) and extending nucleotides (200 µM GTP, 20 µM ATP, 200 µM non-radiolabeled UTP, and 6 mCi of [α32P]-UTP [Perkin Elmer; 3000 Ci/mmol] for Figure 1D; 200 µM GTP, 20 µM ATP, 200 µM UTP for Figure 1—figure supplement 1) was added, and assays were incubated at 25°C for 60 min. Reactions were stopped by addition of stop solution, samples were extracted with acid phenol:chloroform (5:1) (pH 4.5; Thermo Fisher Scientific), RNA products were recovered by ethanol precipitation and resuspended in NudC reaction buffer.

Full-length RNA products were incubated at 37°C for 30 min with 400 nM NudC alone (Figure 1C–D and Figure 1—figure supplement 1), 0.25 U FastAP Thermosensitive Alkaline Phosphatase alone (Figure 1—figure supplement 1), or both NudC and FastAP (Figure 1—figure supplement 1). Reactions were stopped by addition of an equal volume of RNA loading dye. Samples were analyzed by electrophoresis on 7.5 M urea, 1x TBE, 20% polyacrylamide gels (UreaGel System; National Diagnostics), followed by storage-phosphor imaging (Typhoon 9400 variable-mode imager; GE Life Science).

Determination of efficiency of NCIN-mediated initiation vs. ATP-mediated initiation, (kcat/KM)NCIN / (kcat/KM)ATP,in vitro: full-length product assays

For experiments in Figure 2A–B and Figure 2—figure supplement 1, 1 µM of template DNA, 1 µM of mtRNAP, and 1 µM transcription factor(s) (Mtf1 for S. cerevisiae mtRNAP; TFAM and TFB2M for human mtRNAP) were incubated at 25°C for 10 min in Sce-mtRNAP or human reaction buffer. A mixture containing 200 µM ATP, 200 µM UTP, 200 µM non-radiolabeled GTP, and 6 mCi [α32P]-GTP at 3000 Ci/mmol and NCIN (0, 50, 100, 200, 400, 800, 1600, 3200, 6400 µM) was added, and assays were incubated at 25°C for 30 min. Reactions were stopped by addition of an equal volume of RNA loading dye. Samples were analyzed by electrophoresis on 7.5 M urea, 1x TBE, 20% polyacrylamide gels (UreaGel System; National Diagnostics) supplemented with 0.2% 3-acrylamidophenylboronic acid (Boron Molecular), followed by storage-phosphor imaging (Typhoon 9400 variable-mode imager; GE Life Science).

Bands corresponding to uncapped (pppRNA) and NCIN-capped (NCIN-RNA) full-length products were quantified using ImageQuant software. The ratio of NCIN-RNA to total RNA [NCIN-RNA / (pppRNA + NCIN RNA)] was plotted vs. the relative concentrations of NCIN vs. ATP ([NCIN] / [ATP]) on a semi-log plot (SigmaPlot) and non-linear regression was used to fit the data to the equation: y = (ax) / (b + x); where y is [NCIN-RNA / (pppRNA + NCIN RNA)], x is ([NCIN] / [ATP]), and a and b are regression parameters. The resulting fit yields the value of x for which y = 0.5. The relative efficiency (kcat/KM)NCIN / (kcat/KM)ATP is equal to 1/x. Data for determination of relative efficiencies are means of three technical replicates.

Determination of efficiency of NCIN-mediated initiation vs. ATP-mediated initiation, (kcat/KM)NCIN / (kcat/KM)ATP,in vitro: initial product assays (Bird et al., 2017)

For experiments in Figure 3C–D, and Figure 3—figure supplement 1, 1 µM of template DNA, 1 µM of RNAP, and 1 µM transcription factor(s) (Mtf1 for S. cerevisiae mtRNAP; TFAM and TFB2M for human mtRNAP; none for T7 RNAP) were incubated at 25°C for 10 min in Sce-mtRNAP, human mtRNAP reaction buffer, or T7 RNAP reaction buffer (40 mM Tris HCl pH 7.9, 6 mM MgCl2, 2 mM DTT, 2 mM Spermidine). A mixture containing 1 mM NCIN, ATP (0, 25, 50, 100, 200, 400, 800, 1600 µM), 20 µM non-radiolabeled GTP, and 6 mCi [α32P]-GTP at 3000 Ci/mmol was added, and assays were incubated at 25°C for 30 min. Reactions were stopped by addition of an equal volume of RNA loading dye. Samples were analyzed by electrophoresis on 7.5 M urea, 1x TBE, 20% polyacrylamide gels (UreaGel System; National Diagnostics) supplemented with 0.2% 3-acrylamidophenylboronic acid (Boron Molecular), followed by storage-phosphor imaging (Typhoon 9400 variable-mode imager; GE Life Science).

For experiments in Figure 2C and Figure 2—figure supplements 2, 200 nM of tailed template and 500 nM RNAP (S. cerevisiae mtRNAP, human mtRNAP, S. cerevisiae RNAP II, T7 RNAP, or E. coli RNAP) were incubated at 25°C for 15 min in reaction buffer containing 10 mM Tris pH 8.0, 50 mM potassium glutamate, 10 mM MgCl2, 2 mM DTT, and 50 ug/ml BSA. A mixture containing NCIN (1 mM NCIN for mtRNAPs, T7 RNAP, and E. coli RNAP; 4 mM for S. cerevisiae RNAP II), ATP (0, 25, 50, 100, 200, 400, 800, 1600 µM for mtRNAPs and S. cerevisiae RNAP II; 0, 6.25, 12.5, 25, 50, 100, 200, 400 µM for E. coli RNAP), 10 µM non-radiolabeled CTP, and 6 mCi [α32P]-CTP (Perkin Elmer; 3000 Ci/mmol) was added, and assays were incubated at 25°C for 1 hr. Reactions were stopped by addition of an equal volume of RNA loading dye. Samples were analyzed by electrophoresis on 7.5 M urea, 1x TBE, 20% polyacrylamide gels (UreaGel System; National Diagnostics), followed by storage-phosphor imaging (Typhoon 9400 variable-mode imager; GE Life Science).

Bands corresponding to uncapped (pppApC) and NCIN-capped (NCINpC) initial RNA products were quantified using ImageQuant software. The ratio of NCINpC to total RNA (NCINpC / [pppApC + NCINpC]) was plotted vs. the relative concentrations of NCIN vs. ATP ([NCIN] / [ATP]) on a semi-log plot (SigmaPlot) and non-linear regression was used to fit the data to the equation: y = (ax) / (b + x); where y is [NCINpC / (pppApC + NCINpC)], x is ([NCIN] / [ATP]), and a and b are regression parameters. The resulting fit yields the value of x for which y = 0.5. The relative efficiency (kcat/KM)NCIN / (kcat/KM)ATP is equal to 1/x. Data for determination of relative efficiencies are means of three technical replicates.

Detection and quantitation of NAD+- and NADH-capped mitochondrial RNA in vivo: isolation of total cellular RNA from S. cerevisiae

For analysis of NAD+ and NADH capping during respiration, S. cerevisiae strain 246.1.1 [(Tatchell et al., 1981); MATα ura3 trp1 leu2 his4; gift of Andrew Vershon, Rutgers University] was grown at 30°C in 25 ml YPEG (24 g Bacto-tryptone, 20 g Bacto-yeast extract, 30 mL ethanol, 3% glycerol per liter) in 125 ml flasks (Bellco) shaken at 220 rpm. When cell density reached an OD600 ~1.8 (approximately 24 hr) the cell suspension was centrifuged to collect cells (5 min, 10,000 g at 4°C), supernatants were removed, and cell pellets were resuspended in 0.8 mL RNA extraction buffer (0.5 mM NaOAc pH 5.5, 10 mM EDTA, 0.5% SDS).

For analysis of NAD+ and NADH capping during fermentation, S. cerevisiae strain 246.1.1 was grown at 30°C in 100 ml YPD [24 g Bacto-tryptone, 20 g Bacto-yeast extract, 2% (w/v) glucose per liter] in 125 ml flasks (Bellco) with airlocks to prevent oxygenation without shaking for 42 hr. The cell suspension was centrifuged to collect cells (5 min, 10,000 g at 4°C), supernatants were removed, and cell pellets were resuspended in 0.8 mL RNA extraction buffer (0.5 mM NaOAc pH 5.5, 10 mM EDTA, 0.5% SDS).

To extract RNA, an equal volume of acid phenol:chloroform (5:1, pH 4.5; Thermo Fisher Scientific) was added to cells in resuspension buffer and mixed by vortexing. Samples were incubated at 65°C for 5 min, −80°C for 5 min, then centrifuged (15 min, 21,000 g, 4°C) to separate the aqueous and organic phases. The aqueous phase was collected and acid phenol:chloroform extraction was performed two more times on this solution. RNA transcripts were recovered by ethanol precipitation and resuspended in RNase free H2O.

Detection and quantitation of NAD+- and NADH-capped mitochondrial RNA in vivo: isolation of total cellular RNA from human cells

Human embryonic kidney HEK293T cells (obtained from ATCC, tested negative for mycoplasma) were maintained under 5% CO2 at 37°C in DMEM medium (Thermo Fisher Scientific) supplemented with 10% fetal bovine serum (Atlanta Biologicals), 100 units/ml penicillin, and 100 µg/ml streptomycin. HEK293T cells were seeded in 100 mm tissue-culture treated plates and grown for 72 hr at 37°C or seeded in 100 mm tissue-culture treated plates, grown for 24 hr at 37°C, treated with 5 nM FK866 (Sigma Aldrich), and grown for an additional 48 hr at 37°C. Total cellular RNA was isolated with TRIzol Reagent according to the manufacture’s protocol (Thermo Fisher Scientific).

Detection and quantitation of NAD+- and NADH-capped mitochondrial RNA in vivo: DNAzyme cleavage

For analysis of NCIN capping of S. cerevisiae mitochondrial RNA, 40 µg of total cellular RNA was mixed with 1 µM DNAzyme (JB557 for S. cerevisiae COX2 RNA; JB526 for S. cerevisiae 21S RNA) in buffer containing 10 mM Tris pH 8.0, 50 mM NaCl, 2 mM DTT, and 10 mM MgCl2 (total volume 50 µl). When present, NudC was added to 400 nM. Reactions were incubated for 60 min at 37°C and 100 µl of stop solution and 500 µl ethanol was added. Samples were centrifuged (30 min, 21,000 g, 4°C), the supernatant removed, and the pellet resuspended in RNA loading dye.

For analysis of NCIN capping of human mitochondrial RNA, 40 µg of total cellular RNA was mixed with 1 µM DNAzyme (JB559 for human LSP-generated RNA) in buffer containing 10 mM Tris pH 8.0, 50 mM NaCl, 2 mM DTT (total volume 50 µl). Samples were heated to 85°C for 5 min, cooled to 37°C. MgCl2 was added to a final concentration of 10 mM and, when present, NudC was added to 400 nM. Reactions were incubated for 60 min at 37°C and 100 µl of stop solution and 500 µl ethanol was added. Samples were centrifuged (30 min, 21,000 g, 4°C), the supernatant removed, and the pellet resuspended in RNA loading dye.

To prepare synthetic RNA standards non-radiolabeled full-length RNA products were generated by in vitro transcription reactions containing 1 µM of template DNA, 1 µM of mtRNAP, 1 µM transcription factor(s), 1 mM initiating nucleotide (ATP, NAD+ or NADH), 200 µM GTP, 200 µM UTP, 200 µM CTP, and 20 µM ATP. Reactions were incubated for 60 min at 25°C, stopped by addition of stop solution, extracted with acid phenol:chloroform (5:1, pH 4.5; Thermo Fisher Scientific) and ethanol precipitated. Full-length RNAs were resuspended in buffer containing 10 mM Tris pH 8.0, 50 mM NaCl, 2 mM DTT, and 10 mM MgCl2 and treated with DNAzyme as described above.

Detection and quantitation of NAD+- and NADH-capped mitochondrial RNA in vivo: hybridization with a radiolabeled oligodeoxyribonucleotide probe

NCIN capping of DNAzyme-generated subfragments of mitochondrial RNA were analyzed by a procedure consisting of: (i) electrophoresis on 7.5 M urea, 1x TBE, 10% polyacrylamide gels supplemented with 0.2% 3-acrylamidophenylboronic acid (Boron Molecular); (ii) transfer of nucleic acids to a Nytran supercharge nylon membrane (GE Healthcare Life Sciences) using a semidry transfer apparatus (Bio-Rad); (iii) immobilization of nucleic acids by UV crosslinking; (iv) incubation with a 32P-labelled oligodeoxyribonucleotide probe complementary to the 5'-end containing subfragments of target RNAs (JB555, COX2 RNA; JB525, 21S RNA; JB515, LSP-derived RNA; 32P-labelled using T4 polynucleotide kinase and [γ32P]-ATP [Perkin Elmer]); (v) high-stringency washing, procedures as in (Goldman et al., 2015); and (vi) storage-phosphor imaging (Typhoon 9400 variable-mode imager; GE Life Science).

Bands corresponding to uncapped and NCIN-capped DNAzyme-generated subfragments were quantified using ImageQuant software. The percentages of uncapped RNA (5'-ppp), NAD+-capped RNA (5'-NAD+), or NADH-capped RNA (5'-NADH) to total RNA were determined from three biological replicates.

Determination of NAD(H) levels in human HEK293T cells

Cells were lysed in 400 μl of NAD/H Extraction Buffer (NAD/H Quantitation Kit, Sigma-Aldrich) and total proteins were extracted with two freeze-thaw cycles (20 min on dry ice, 10 min at RT). Lysates were centrifuged (10 min, 13,000 g, 4°C), the supernatant was isolated, and the protein concentration was determined. NAD(H) was measured using a NAD/H Quantitation Kit (Sigma) from a volume of supernatant containing 10 μg protein.

Molecules of NAD(H) per cell were calculated using a value of 300 pg total protein per cell. The cellular concentration of NAD(H) was then estimated by using a cellular volume of ~1200 µm3 for HEK293T cells and assuming a homogenous distribution of NAD(H) in the cell. The volume of HEK293T cells was calculated using a value of 13 µm for the diameter of HEK293T cells, assuming trypsinized cells adopt a spherical shape. NAD/H concentrations were determined from three biological replicates.

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Decision letter

  1. Alan G Hinnebusch
    Reviewing Editor; Eunice Kennedy Shriver National Institute of Child Health and Human Development, United States
  2. James L Manley
    Senior Editor; Columbia University, United States

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.

Thank you for submitting your article "Highly efficient 5' capping of mitochondrial RNA with NAD+ and NADH by yeast and human mitochondrial RNA polymerase" 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 James Manley as the Senior Editor. The following individual involved in review of your submission has agreed to reveal their identity: Ann Hochschild (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 paper uses a combination of in-depth biochemical analysis and a novel in vivo approach to provide important new insights into NAD/NADH capping of eukaryotic mitochondrial transcripts and of this capping process in general. Using purified RNA polymerases from yeast or human mitochondria (mt), as well yeast nuclear, E. coli and T7 RNA Pol enzymes, they show by careful enzyme kinetics that the mt enzymes are considerably more efficient than the yeast and E. coli enzymes at NAD+ capping in vitro. They further show that the mt enzymes and T7 Pol, all single polypeptide polymerases, exhibit the same promoter sequence specificity for the reaction demonstrated previously for the E. coli multi-subunit enzyme, requiring a T on the template strand for the TSS (+1 position) and a purine:pyrimidine base pair on the non-template:template strands at the -1 position. They introduce a significant advance in the techniques for quantifying the occurrence of NAD+ and NADH, vs uncapped mt transcripts produced in vivo, and show that the proportions of NAD+/NADH vs. uncapped transcripts for two yeast and one human mt transcripts are much higher than observed previously for bacterial transcripts and yeast cytoplasmic mRNAs. Finally, they show that the ratio of NAD+ vs NADH capped mRNAs for the two yeast mt mRNAs vary in response to altered growth conditions expected to change the NAD+/NADH ratio in cells, providing evidence that mt RNAP could be utilized as a sensor of energy metabolism that would generate altered ratios of NAD+ vs NADH-capped mt mRNAs. In view of their findings that a large fraction of the mt transcripts harbor one or the other cap, this has the potential of altering mt gene expression through possible influences of these caps on mRNA translation or turnover.

Although NAD+ capped yeast mt mRNAs had been detected previously, this work is significant in providing direct evidence that mt RNAPs carry out this reaction and utilize the same promoter sequences found for multi-subunit RNAPs to direct capping; and they also function much more efficiently than do the latter. The results also provide evidence that the capped mt transcripts detected in cells represent capping during transcription initiation vs. capping of degradation products. By measuring the precise proportions of NAD+-capped, NADH-capped, and uncapped mt mRNAs in cells-which is much higher than seen previously for bacterial or yeast cytoplasmic mRNAs-they show that the ratio of NAD+ to NADH-capped mRNAs varies with the energy status of yeast cells.

Essential revisions:

It is requested that you discuss how the relative levels of NAD/NADH capping of mitochondrial versus cytoplasmic mRNAs could be affected by different concentrations of NAD/NADH in the mitochondria versus eukaryotic nuclei or E. coli cells in addition to the different capping efficiencies of the respective RNA polymerases, and cite any information available about these concentrations.

It is requested that you provide a much better explanation of how the "fork-junction" template works and how it bypasses the requirement for sequence-specific RNAP-DNA interactions and transcription-initiation factor-DNA interactions for transcription initiation, as neither the text nor cartoon were found to be adequate.

It is requested that you state more explicitly how the conclusion that "mitochondrial RNAs undergo NAD+ capping at 5' ends generated by transcription initiation (as opposed 5' ends generated by RNA processing" follows from the data provided.

It is requested that you discuss what is known about whether the capping ratio affects the amount of transcript, the stability of the transcript, or the efficiency with which it is translated.

https://doi.org/10.7554/eLife.42179.022

Author response

Essential revisions:

It is requested that you discuss how the relative levels of NAD/NADH capping of mitochondrial versus cytoplasmic mRNAs could be affected by different concentrations of NAD/NADH in the mitochondria versus eukaryotic nuclei or E. coli cells in addition to the different capping efficiencies of the respective RNA polymerases, and cite any information available about these concentrations.

We have modified the text in the second paragraph of the Discussion to address this point. The revised text is:

“First, mtRNAPs are substantially more efficient at NAD+ and NADH capping than bacterial and eukaryotic nuclear RNAPs (Figure 2C). Second, levels of NAD+ and NADH relative to ATP in mitochondria are substantially higher than levels of NAD+ and NADH relative to ATP in bacteria and eukaryotic nuclei (Cambronne et al., 2016; W. W. Chenet al., 2016; Parket al., 2016).”

It is requested that you provide a much better explanation of how the "fork-junction" template works and how it bypasses the requirement for sequence-specific RNAP-DNA interactions and transcription-initiation factor-DNA interactions for transcription initiation, as neither the text nor cartoon were found to be adequate.

We have revised the text to indicate that the template used for these assays is a “tailed” template and provided references establishing use of a tailed template to bypass the requirement for sequence-specific RNAP-DNA interactions and transcription-initiation factor-DNA interactions. The revised text is:

“…we performed transcription assays using a “tailed” template (Figure 2C, top) that bypasses the requirement for sequence-specific RNAP-DNA interactions and transcription-initiation factor-DNA interactions for transcription initiation (Dedrick and Chamberlin, 1985; Kadesch and Chamberlin, 1982).”

It is requested that you state more explicitly how the conclusion that "mitochondrial RNAs undergo NAD+ capping at 5' ends generated by transcription initiation (as opposed 5' ends generated by RNA processing" follows from the data provided.

We have revised the text to clarify this point by adding the following sentence to the Results section:

“Because the hybridization probe detects RNA fragments that contain 5' ends generated by transcription initiation (red sub-fragment depicted in Figure 4A), the detected NAD+ and NADH caps are concluded to be at 5' ends generated by transcription initiation, as opposed to 5' ends generated by RNA processing.”

It is requested that you discuss what is known about whether the capping ratio affects the amount of transcript, the stability of the transcript, or the efficiency with which it is translated.

We have modified the text at the beginning of the second paragraph of the Discussion. The revised text is:

“We and others previously have shown that NCIN capping by cellular RNAPs has functional consequences, including modulating RNA stability and modulating RNA translatability.”

https://doi.org/10.7554/eLife.42179.023

Article and author information

Author details

  1. Jeremy G Bird

    1. Department of Genetics and Waksman Institute, Rutgers University, United States
    2. Department of Chemistry and Waksman Institute, Rutgers University, United States
    Contribution
    Formal analysis, Investigation, Visualization, Methodology, Writing—review and editing
    Contributed equally with
    Urmimala Basu
    Competing interests
    No competing interests declared
  2. Urmimala Basu

    1. Department of Biochemistry and Molecular Biology, Robert Wood Johnson Medical School, Rutgers University, United States
    2. Biochemistry PhD Program, School of Graduate Studies, Rutgers University, United States
    Contribution
    Formal analysis, Funding acquisition, Investigation, Visualization, Methodology, Writing—review and editing
    Contributed equally with
    Jeremy G Bird
    Competing interests
    No competing interests declared
  3. David Kuster

    1. Department of Genetics and Waksman Institute, Rutgers University, United States
    2. Department of Chemistry and Waksman Institute, Rutgers University, United States
    3. Biochemistry Center Heidelberg, Heidelberg University, Germany
    Contribution
    Formal analysis, Investigation, Visualization, Methodology, Writing—review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-8157-9223
  4. Aparna Ramachandran

    Department of Biochemistry and Molecular Biology, Robert Wood Johnson Medical School, Rutgers University, United States
    Contribution
    Investigation, Methodology, Writing—review and editing
    Competing interests
    No competing interests declared
  5. Ewa Grudzien-Nogalska

    Department of Cell Biology and Neuroscience, Rutgers University, United States
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  6. Atif Towheed

    Center for Mitochondrial and Epigenomic Medicine, The Children’s Hospital of Philadelphia, United States
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  7. Douglas C Wallace

    1. Center for Mitochondrial and Epigenomic Medicine, The Children’s Hospital of Philadelphia, United States
    2. Department of Pediatrics, Division of Human Genetics, The Children's Hospital of Philadelphia, Perelman School of Medicine, United States
    Contribution
    Supervision, Funding acquisition
    Competing interests
    No competing interests declared
  8. Megerditch Kiledjian

    Department of Cell Biology and Neuroscience, Rutgers University, United States
    Contribution
    Supervision, Funding acquisition, Writing—review and editing
    Competing interests
    No competing interests declared
  9. Dmitry Temiakov

    Department of Biochemistry and Molecular Biology, Sidney Kimmel Cancer Center, Thomas Jefferson University, United States
    Contribution
    Conceptualization, Funding acquisition, Methodology, Writing—review and editing
    Competing interests
    No competing interests declared
  10. Smita S Patel

    Department of Biochemistry and Molecular Biology, Robert Wood Johnson Medical School, Rutgers University, United States
    Contribution
    Conceptualization, Supervision, Funding acquisition, Visualization, Project administration, Writing—review and editing
    For correspondence
    patelss@rwjms.rutgers.edu
    Competing interests
    No competing interests declared
  11. Richard H Ebright

    Department of Chemistry and Waksman Institute, Rutgers University, United States
    Contribution
    Conceptualization, Supervision, Funding acquisition, Visualization, Writing—original draft, Project administration
    For correspondence
    ebright@waksman.rutgers.edu
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-8915-7140
  12. Bryce E Nickels

    Department of Genetics and Waksman Institute, Rutgers University, United States
    Contribution
    Conceptualization, Supervision, Funding acquisition, Visualization, Writing—original draft, Project administration
    For correspondence
    bnickels@waksman.rutgers.edu
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-7449-8831

Funding

American Heart Association (16PRE30400001)

  • Urmimala Basu

National Institutes of Health (NS021328)

  • Douglas C Wallace

National Institutes of Health (MH108592)

  • Douglas C Wallace

National Institutes of Health (OD010944)

  • Douglas C Wallace

U.S. Department of Defense (W81XWH-16-1-0401)

  • Douglas C Wallace

National Institutes of Health (GM126488)

  • Megerditch Kiledjian

National Institutes of Health (GM104231)

  • Dmitry Temiakov

National Institutes of Health (GM118086)

  • Smita S Patel

National Institutes of Health (GM041376)

  • Richard H Ebright

National Institutes of Health (GM118059)

  • Bryce E Nickels

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

Senior Editor

  1. James L Manley, Columbia University, United States

Reviewing Editor

  1. Alan G Hinnebusch, Eunice Kennedy Shriver National Institute of Child Health and Human Development, United States

Publication history

  1. Received: September 19, 2018
  2. Accepted: December 10, 2018
  3. Accepted Manuscript published: December 10, 2018 (version 1)
  4. Accepted Manuscript updated: December 12, 2018 (version 2)
  5. Version of Record published: December 18, 2018 (version 3)

Copyright

© 2018, Bird et al.

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