Analytical and Bioanalytical Chemistry

, Volume 410, Issue 6, pp 1667–1677 | Cite as

Poly A tail length analysis of in vitro transcribed mRNA by LC-MS

  • Michael Beverly
  • Caitlin Hagen
  • Olga Slack
Research Paper


The 3′-polyadenosine (poly A) tail of in vitro transcribed (IVT) mRNA was studied using liquid chromatography coupled to mass spectrometry (LC-MS). Poly A tails were cleaved from the mRNA using ribonuclease T1 followed by isolation with dT magnetic beads. Extracted tails were then analyzed by LC-MS which provided tail length information at single-nucleotide resolution. A 2100-nt mRNA with plasmid-encoded poly A tail lengths of either 27, 64, 100, or 117 nucleotides was used for these studies as enzymatically added poly A tails showed significant length heterogeneity. The number of As observed in the tails closely matched Sanger sequencing results of the DNA template, and even minor plasmid populations with sequence variations were detected. When the plasmid sequence contained a discreet number of poly As in the tail, analysis revealed a distribution that included tails longer than the encoded tail lengths. These observations were consistent with transcriptional slippage of T7 RNAP taking place within a poly A sequence. The type of RNAP did not alter the observed tail distribution, and comparison of T3, T7, and SP6 showed all three RNAPs produced equivalent tail length distributions. The addition of a sequence at the 3′ end of the poly A tail did, however, produce narrower tail length distributions which supports a previously described model of slippage where the 3′ end can be locked in place by having a G or C after the poly nucleotide region.

Graphical abstract

Determination of mRNA poly A tail length using magnetic beads and LC-MS.


mRNA Poly A Tail length Mass spectrometry Slippage In vitro transcription 


Funding information

Support for this work was provided by DARPA grant HR-0011-13-3-0003.

Compliance with ethical standards

Conflict of interest

The authors declare they have no conflict of interest.


The views, opinions, and/or finding expressed are those of the authors and should not be interpreted as representing the official views or policies of the Department of Defense or the US Government.

Supplementary material

216_2017_840_MOESM1_ESM.pdf (1 mb)
ESM 1 (PDF 1.04 mb)


  1. 1.
    Harrison PF, Powell DR, Clancy JL, Preiss T, Boag PR, Traven A, et al. PAT-seq: a method to study the integration of 3′-UTR dynamics with gene expression in the eukaryotic transcriptome. RNA. 2015;21(8):1502–10.CrossRefGoogle Scholar
  2. 2.
    Aitken CE, Lorsch JR. A mechanistic overview of translation initiation in eukaryotes. Nat Struct Mol Biol. 2012;19(6):568–76.CrossRefGoogle Scholar
  3. 3.
    Moqtaderi Z, Geisberg JV, Struhl K. Secondary structures involving the poly(A) tail and other 3′ sequences are major determinants of mRNA isoform stability in yeast. Microb Cell. 2014;1(4):137–9.CrossRefGoogle Scholar
  4. 4.
    Subtelny AO, Eichhorn SW, Chen GR, Sive H, Bartel DP. Poly(A)-tail profiling reveals an embryonic switch in translational control. Nature. 2014;508(7494):66–71.CrossRefGoogle Scholar
  5. 5.
    Ratcliffe CD, Sahgal P, Parachoniak CA, Ivaska J, Park M. Regulation of cell migration and beta1 integrin trafficking by the endosomal adaptor GGA3. Traffic. 2016;17(6):670–88.CrossRefGoogle Scholar
  6. 6.
    Beilharz TH, Preiss T. Widespread use of poly(A) tail length control to accentuate expression of the yeast transcriptome. RNA. 2007;13(7):982–97.CrossRefGoogle Scholar
  7. 7.
    Peng J, Murray EL, Schoenberg DR. In vivo and in vitro analysis of poly(A) length effects on mRNA translation. Methods Mol Biol. 2008;419:215–30.CrossRefGoogle Scholar
  8. 8.
    Koski GK, Kariko K, Xu S, Weissman D, Cohen PA, Czerniecki BJ. Cutting edge: innate immune system discriminates between RNA containing bacterial versus eukaryotic structural features that prime for high-level IL-12 secretion by dendritic cells. J Immunol. 2004;172(7):3989–93.CrossRefGoogle Scholar
  9. 9.
    Bernstein P, Peltz SW, Ross J. The poly(A)-poly(A)-binding protein complex is a major determinant of mRNA stability in vitro. Mol Cell Biol. 1989;9(2):659–70.CrossRefGoogle Scholar
  10. 10.
    Bernstein P, Ross J. Poly(A), poly(A) binding protein and the regulation of mRNA stability. Trends Biochem Sci. 1989;14(9):373–7.CrossRefGoogle Scholar
  11. 11.
    Chang H, Lim J, Ha M, Kim VN. TAIL-seq: genome-wide determination of poly(A) tail length and 3′ end modifications. Mol Cell. 2014;53(6):1044–52.CrossRefGoogle Scholar
  12. 12.
    Janicke A, Vancuylenberg J, Boag PR, Traven A, Beilharz TH. ePAT: a simple method to tag adenylated RNA to measure poly(A)-tail length and other 3' RACE applications. RNA. 2012;18(6):1289–95.CrossRefGoogle Scholar
  13. 13.
    Murray EL, Schoenberg DR. Assays for determining poly(A) tail length and the polarity of mRNA decay in mammalian cells. Methods Enzymol. 2008;448:483–504.CrossRefGoogle Scholar
  14. 14.
    Meijer HA, Bushell M, Hill K, Gant TW, Willis AE, Jones P, et al. A novel method for poly(A) fractionation reveals a large population of mRNAs with a short poly(A) tail in mammalian cells. Nucleic Acids Res. 2007;35(19):e132.CrossRefGoogle Scholar
  15. 15.
    Salles FJ, Strickland S. Rapid and sensitive analysis of mRNA polyadenylation states by PCR. PCR Methods Appl. 1995;4(6):317–21.CrossRefGoogle Scholar
  16. 16.
    Minasaki R, Rudel D, Eckmann CR. Increased sensitivity and accuracy of a single-stranded DNA splint-mediated ligation assay (sPAT) reveals poly(A) tail length dynamics of developmentally regulated mRNAs. RNA Biol. 2014;11(2):111–23.CrossRefGoogle Scholar
  17. 17.
    Muddiman DC, Null AP, Hannis JC. Precise mass measurement of a double-stranded 500 base-pair (309 kDa) polymerase chain reaction product by negative ion electrospray ionization Fourier transform ion cyclotron resonance mass spectrometry. Rapid Commun Mass Spectrom. 1999;13(12):1201–4.CrossRefGoogle Scholar
  18. 18.
    Hannis JC, Muddiman DC. Detection of double-stranded PCR amplicons at the attomole level electrosprayed from low nanomolar solutions using FT-ICR mass spectrometry. Fresenius J Anal Chem. 2001;369(3–4):246–51.CrossRefGoogle Scholar
  19. 19.
    Ecker DJ, Sampath R, Massire C, Blyn LB, Hall TA, Eshoo MW, et al. Ibis T5000: a universal biosensor approach for microbiology. Nat Rev Microbiol. 2008;6(7):553–8.CrossRefGoogle Scholar
  20. 20.
    Wolk DM, Kaleta EJ, Wysocki VH. PCR-electrospray ionization mass spectrometry: the potential to change infectious disease diagnostics in clinical and public health laboratories. J Mol Diagn. 2012;14(4):295–304.CrossRefGoogle Scholar
  21. 21.
    Beverly M, Dell A, Parmar P, Houghton L. Label-free analysis of mRNA capping efficiency using RNase H probes and LC-MS. Anal Bioanal Chem. 2016;408(18):5021–30.CrossRefGoogle Scholar
  22. 22.
    Gilar M. Analysis and purification of synthetic oligonucleotides by reversed-phase high-performance liquid chromatography with photodiode array and mass spectrometry detection. Anal Biochem. 2001;298(2):196–206.CrossRefGoogle Scholar
  23. 23.
    Apffel A, Chakel JA, Fischer S, Lichtenwalter K, Hancock WS. Analysis of oligonucleotides by HPLC-electrospray ionization mass spectrometry. Anal Chem. 1997;69(7):1320–5.CrossRefGoogle Scholar
  24. 24.
    Huber CG, Oberacher H. Analysis of nucleic acids by on-line liquid chromatography-mass spectrometry. Mass Spectrom Rev. 2001;20(5):310–43.CrossRefGoogle Scholar
  25. 25.
    Hail ME, Elliott B, Anderson K. High-throughput analysis of oligonucleotides using automated electrospray ionization mass spectrometry. Am Biotechnol Lab. 2004;22:12–3.Google Scholar
  26. 26.
    Haukanes BI, Kvam C. Application of magnetic beads in bioassays. Biotechnology (N Y). 1993;11(1):60–3.Google Scholar
  27. 27.
    Berensmeier S. Magnetic particles for the separation and purification of nucleic acids. Appl Microbiol Biotechnol. 2006;73(3):495–504.CrossRefGoogle Scholar
  28. 28.
    Adams NM, Bordelon H, Wang KK, Albert LE, Wright DW, Haselton FR. Comparison of three magnetic bead surface functionalities for RNA extraction and detection. ACS Appl Mater Interfaces. 2015;7(11):6062–9.CrossRefGoogle Scholar
  29. 29.
    Blower MD, Jambhekar A, Schwarz DS, Toombs JA. Combining different mRNA capture methods to analyze the transcriptome: analysis of the Xenopus laevis transcriptome. PLoS One. 2013;8(10):e77700.CrossRefGoogle Scholar
  30. 30.
    Cabada MO, Darnbrough C, Ford PJ, Turner PC. Differential accumulation of two size classes of poly(A) associated with messenger RNA during oogenesis in Xenopus laevis. Dev Biol. 1977;57(2):427–39.CrossRefGoogle Scholar
  31. 31.
    Park JE, Yi H, Kim Y, Chang H, Kim VN. Regulation of poly(A) tail and translation during the somatic cell cycle. Mol Cell. 2016;62(3):462–71.CrossRefGoogle Scholar
  32. 32.
    Jacobsen N, Nielsen PS, Jeffares DC, Eriksen J, Ohlsson H, Arctander P, et al. Direct isolation of poly(A)+ RNA from 4 M guanidine thiocyanate-lysed cell extracts using locked nucleic acid-oligo(T) capture. Nucleic Acids Res. 2004;32(7):e64.CrossRefGoogle Scholar
  33. 33.
    Phelan D, Hondorp K, Choob M, Efimov V, Fernandez J. Messenger RNA isolation using novel PNA analogues. Nucleosides Nucleotides Nucleic Acids. 2001;20(4–7):1107–11.CrossRefGoogle Scholar
  34. 34.
    Holtkamp S, Kreiter S, Selmi A, Simon P, Koslowski M, Huber C, et al. Modification of antigen-encoding RNA increases stability, translational efficacy, and T-cell stimulatory capacity of dendritic cells. Blood. 2006;108(13):4009–17.CrossRefGoogle Scholar
  35. 35.
    Triana-Alonso FJ, Dabrowski M, Wadzack J, Nierhaus KH. Self-coded 3′-extension of run-off transcripts produces aberrant products during in vitro transcription with T7 RNA polymerase. J Biol Chem. 1995;270(11):6298–307.CrossRefGoogle Scholar
  36. 36.
    Wagner LA, Weiss RB, Driscoll R, Dunn DS, Gesteland RF. Transcriptional slippage occurs during elongation at runs of adenine or thymine in Escherichia coli. Nucleic Acids Res. 1990;18(12):3529–35.CrossRefGoogle Scholar
  37. 37.
    Molodtsov V, Anikin M, McAllister WT. The presence of an RNA:DNA hybrid that is prone to slippage promotes termination by T7 RNA polymerase. J Mol Biol. 2014;426(18):3095–107.CrossRefGoogle Scholar
  38. 38.
    Macdonald LE, Zhou Y, McAllister WT. Termination and slippage by bacteriophage T7 RNA polymerase. J Mol Biol. 1993;232(4):1030–47.CrossRefGoogle Scholar
  39. 39.
    Anikin M, Molodtsov V, Temiakov D, McAllister WT. Transcript slippage and recoding. In: Atkins JF, Gesteland RF, editors. Recoding: expansion of decoding rules enrishes gene expression, Nucleic acid and molecular biology: Springer; 2010. p. 409–32.Google Scholar
  40. 40.
    Elango N, Elango S, Shivshankar P, Katz MS. Optimized transfection of mRNA transcribed from a d(a/T)100 tail-containing vector. Biochem Biophys Res Commun. 2005;330(3):958–66.CrossRefGoogle Scholar
  41. 41.
    Grier AE, Burleigh S, Sahni J, Clough CA, Cardot V, Choe DC, et al. pEVL: a linear plasmid for generating mRNA IVT templates with extended encoded poly(A) sequences. Mol Ther Nucleic Acids. 2016;5:e306.CrossRefGoogle Scholar
  42. 42.
    Mockey M, Goncalves C, Dupuy FP, Lemoine FM, Pichon C, Midoux P. mRNA transfection of dendritic cells: synergistic effect of ARCA mRNA capping with poly(A) chains in cis and in trans for a high protein expression level. Biochem Biophys Res Commun. 2006;340(4):1062–8.CrossRefGoogle Scholar

Copyright information

© Springer-Verlag GmbH Germany, part of Springer Nature 2018

Authors and Affiliations

  1. 1.Novartis Institutes of Biomedical ResearchCambridgeUSA

Personalised recommendations