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Epitranscriptomics: RNA Modifications in Bacteria and Archaea

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  • Authors: Katharina Höfer1, Andres Jäschke2
  • Editors: Gisela Storz3, Kai Papenfort4
  • VIEW AFFILIATIONS HIDE AFFILIATIONS
    Affiliations: 1: Institute of Pharmacy and Molecular Biotechnology, Im Neuenheimer Feld 364, Heidelberg University, 69120 Heidelberg, Germany; 2: Institute of Pharmacy and Molecular Biotechnology, Im Neuenheimer Feld 364, Heidelberg University, 69120 Heidelberg, Germany; 3: Division of Molecular and Cellular Biology, Eunice Kennedy Shriver National Institute of Child Health and Human Development, Bethesda, MD; 4: Department of Biology I, Microbiology, LMU Munich, Martinsried, Germany
  • Source: microbiolspec June 2018 vol. 6 no. 3 doi:10.1128/microbiolspec.RWR-0015-2017
  • Received 27 November 2017 Accepted 26 January 2018 Published 01 June 2018
  • Andres Jäschke, [email protected]
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  • Abstract:

    The increasingly complex functionality of RNA is contrasted by its simple chemical composition. RNA is generally built from only four different nucleotides (adenine, guanine, cytosine, and uracil). To date, >160 chemical modifications are known to decorate RNA molecules and thereby alter their function or stability. Many RNA modifications are conserved throughout bacteria, archaea, and eukaryotes, while some are unique to each branch of life. Most known modifications occur at internal positions, while there is limited diversity at the termini. The dynamic nature of RNA modifications and newly discovered regulatory functions of some of these RNA modifications gave birth to a new field, now often referred to as “epitranscriptomics.” This review highlights the major developments in this field and summarizes detection principles for internal as well as 5′-terminal mRNA modifications in prokaryotes and archaea to investigate their biological significance.

  • Citation: Höfer K, Jäschke A. 2018. Epitranscriptomics: RNA Modifications in Bacteria and Archaea. Microbiol Spectrum 6(3):RWR-0015-2017. doi:10.1128/microbiolspec.RWR-0015-2017.

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/content/journal/microbiolspec/10.1128/microbiolspec.RWR-0015-2017
2018-06-01
2019-08-18

Abstract:

The increasingly complex functionality of RNA is contrasted by its simple chemical composition. RNA is generally built from only four different nucleotides (adenine, guanine, cytosine, and uracil). To date, >160 chemical modifications are known to decorate RNA molecules and thereby alter their function or stability. Many RNA modifications are conserved throughout bacteria, archaea, and eukaryotes, while some are unique to each branch of life. Most known modifications occur at internal positions, while there is limited diversity at the termini. The dynamic nature of RNA modifications and newly discovered regulatory functions of some of these RNA modifications gave birth to a new field, now often referred to as “epitranscriptomics.” This review highlights the major developments in this field and summarizes detection principles for internal as well as 5′-terminal mRNA modifications in prokaryotes and archaea to investigate their biological significance.

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Figures

Image of FIGURE 1
FIGURE 1

Identification of RNA modifications. (A) Identification of RNA modifications by a combination of digest to single nucleotides and chromatography. Total RNA is digested by nucleases (nuclease P1) and single building blocks separated by TLC or MS coupled with LC. (B) Identification of RNA modifications and their associated transcripts. Modified RNA is specifically enriched by a protocol that is based on antibody treatment (immunoprecipitation), an enzymatic reaction, or a chemical treatment. Afterwards, enriched, modified RNA is converted into cDNA to produce a library for NGS. The reads are mapped to the genome to identify the sequence of the transcripts that bear a specific RNA modification. Known or possible 5′-terminal (C) or internal (D) mRNA modifications in bacteria and archaea.

Source: microbiolspec June 2018 vol. 6 no. 3 doi:10.1128/microbiolspec.RWR-0015-2017
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Image of FIGURE 2
FIGURE 2

Methods to detect internal RNA modifications. (A) Identification of m5C-modified mRNA by bisulfite sequencing. Selective conversion of cytosine to uracil by bisulfite ions, whereas m5C remains as cytosine. After reverse transcription, a cDNA library is prepared, analyzed by high-throughput sequencing, and mapped to the genome. Cytosine is mapped as thymine, whereas m5C residues read as cytosine. (B) Ψ-Seq: identification of pseudouridine-modified RNA. mRNA is treated with CMC, which selectively reacts with Ψ and causes a stop during reverse transcription. cDNA libraries are amplified and sequenced. The reads from a CMC-treated and nontreated control are compared to map pseudouridine-modified RNA. (C) Identification of methylated adenosine by m6A-Seq. Total RNA is fragmented to 100-nucleotide-long RNAs. m6A-specific antibodies are used to immunoprecipitate RNA. RNA is reverse-transcribed to cDNA and analyzed by high-throughput sequencing. These reads produce a peak whose summit reflects an underlying m6A residue. (D) Identification of adenosine-to-inosine editing: ICE-Seq. ICE is based on cyanoethylation of inosine by acrylonitrile combined with reverse transcription and high-throughput sequencing. Inosine pairs with cytosine (control), whereas the chemical modification of inosine results in a stop during reverse transcription. Erased G signals originate from inosines in the sequence map of cDNAs and are finally used to identify A-to-I editing sites. (E) Mapping of 2′-O-methylated nucleotides (Nm) by Nm-Seq. Fragmented RNA is subjected to iterative oxidation-elimination-dephosphorylation cycles that remove 2′-hydroxylated nucleotides in the 3′-to-5′ direction. Internal 2′-O-methylation sites stay intact, and fragments ending with 2′-hydroxyl are finally blocked by an incomplete oxidation-elimination cycle. 2′-O-methylated RNA fragments are ligated to an adapter. After library construction and high-throughput sequencing, reads are mapped to the genome. At 2′-OMe sites, an asymmetric coverage profile is observed whose uniform 3′ end corresponds to the methylation position.

Source: microbiolspec June 2018 vol. 6 no. 3 doi:10.1128/microbiolspec.RWR-0015-2017
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Image of FIGURE 3
FIGURE 3

Identification of 5′-terminal RNA modifications. (A) Differential RNA-Seq to identify primary transcripts. Total RNA is treated with a 5′-P-dependent exonuclease that degrades specifically 5′-P-RNA. 5′-PPP-RNA is converted into 5′-P-RNA enzymatically by TAP or similar enzymes. The 5′ end is ligated to an adapter sequence and the RNA reverse-transcribed into cDNA, which is analyzed by high-throughput sequencing. The reads are mapped to the genome to identify TSSs. (B) Schematic representation of the NAD captureSeq protocol that allows the identification of NAD-capped RNAs. Total RNA is treated with ADPRC from , which specifically catalyzes the transglycosylation reaction of NAD with 4-pentyn-1-ol. The product of the reaction is biotinylated by CuAAC (click reaction). The NAD-capped RNA is captured as well as enriched on streptavidin beads and ligated to an adapter. After on-bead reverse transcription and a second adapter ligation, the obtained cDNA is amplified by PCR and submitted to high-throughput sequencing.

Source: microbiolspec June 2018 vol. 6 no. 3 doi:10.1128/microbiolspec.RWR-0015-2017
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FIGURE 4

Biosynthesis and removal of 5′-terminal RNA modification. (A) Synthesis of m7G-capped RNA in eukaryotes. 5′-PPP-RNA is synthesized by the RNA polymerase. The 5′-γ-phosphate of the nascent pre-mRNA is hydrolyzed by an RNA triphosphatase to 5′-PP-RNA. In the next step, a guanine monophosphate nucleoside is transferred to the 5′-diphosphate mRNA end by RNA guanylyltransferase to generate G-PPP-RNA. Finally, the guanine N7 position (blue) is methylated by SAM to form m7G-PPP-RNA by SAM. (B) Schematic representation of the synthesis of primary transcripts and cofactor-capped RNA in . Bacterial RNA polymerase is able to initiate transcription with a nucleotide triphosphate or an adenosine-containing cofactor, such as NAD, to generate 5′-PPP-RNA or NAD/cofactor-capped RNA. (C) Decapping of m7G-capped RNA in eukaryotic organisms. A decapping enzyme complex including Dcp2 removes the cap and converts the RNA in 5′-P-RNA that is targeted for degradation by 5′-dependent exonucleases like Xrn1. (D) 5′-End processing in . NAD-RNA is decapped by NudC into 5′-P-RNA. RppH processes primary transcripts and 5′-PP-RNA into 5′-P-RNA, which triggers RNase E-mediated degradation.

Source: microbiolspec June 2018 vol. 6 no. 3 doi:10.1128/microbiolspec.RWR-0015-2017
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FIGURE 5

Methods to relatively quantify and characterize 5′-terminally modified RNAs and their decapping enzymes and . (A) Schematic representation of the identification and relative quantification of 5′-PP-RNA (PACO assay) . The 5′ end of 5′-PP-RNA is capped with a guanosine. Subsequent treatment with alkaline phosphatase removes the exposed 5′-terminal phosphates of 5′-PPP-RNA and 5′-P-RNA but not the protected phosphates of the guanylylated G-PPP-RNA, which is then converted to 5′-P-RNA by treatment with a pyrophosphohydrolase. After adapter ligation and Northern blot analysis, the amount of 5′-PP-RNA can be quantified by a shift. (B) APB PAGE to quantify 5′-NAD-capped RNA . APB interacts with -diols of the ribose, which results in retardation of NAD-capped RNA during gel electrophoresis. (C) Identification of decapping enzymes by specific radioactive labeling of RNA . NAD-capped RNA is transcribed by the bacteriophage T7 RNA polymerase in the presence of radioactively labeled NAD. This technology enables specific radioactive labeling of each RNA with a single radioactive mark specifically located at the 5′ termini. After a decapping reaction, e.g., by NudC, 5′-P-RNA is generated. The accessible radioactive phosphate is removed by alkaline phosphatase treatment. The RNA is analyzed by PAGE.

Source: microbiolspec June 2018 vol. 6 no. 3 doi:10.1128/microbiolspec.RWR-0015-2017
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Tables

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TABLE 1

Internal RNA modifications

Source: microbiolspec June 2018 vol. 6 no. 3 doi:10.1128/microbiolspec.RWR-0015-2017
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TABLE 2

Cofactor-capped RNAs

Source: microbiolspec June 2018 vol. 6 no. 3 doi:10.1128/microbiolspec.RWR-0015-2017

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