Enzymatic Degradation of Linear Dinucleotide Intermediates of Cyclic Dinucleotides

  • Mona W. Orr
  • Vincent T. LeeEmail author


Bacterial cyclic dinucleotides (cyclic di-GMP, cyclic di-AMP, and cyclic GMP-AMP) are signaling molecules that bind to intracellular receptors to regulate a wide range of processes. In response to environmental changes, bacteria alter the rate of both synthesis and degradation to control the concentration of cyclic dinucleotides. Degradation occurs in a two-step process. The first step is carried out by enzymes specific to each cyclic dinucleotide and results in the formation of a linear dinucleotide. The second step is the hydrolysis of the linear dinucleotide into mononucleotides. Some phosphodiesterases that degrade the cyclic dinucleotide to the linear form are also capable of further hydrolysis to mononucleotides in vitro. However, not all species that utilize cyclic-dinucleotide signaling have these enzymes. Recently, it was shown that exoribonucleases specific for very short RNA substrates also degrade the linear dinucleotide intermediates of cyclic dinucleotide turnover. These results indicate that there is a potential overlap between RNA degradation and cyclic dinucleotide signaling suggesting the possibility of cross talk between signaling and RNA turnover.


pGpG pApA pApG Oligoribonuclease NrnA NrnB NrnC 


  1. 1.
    Ross P et al (1987) Regulation of cellulose synthesis in Acetobacter xylinum by cyclic diguanylic acid. Nature 325:279–281CrossRefGoogle Scholar
  2. 2.
    Jenal U, Reinders A, Lori C (2017) Cyclic di-GMP: second messenger extraordinaire. Nat Rev Microbiol 15:271–284. CrossRefPubMedGoogle Scholar
  3. 3.
    Römling U, Galperin MY, Gomelsky M (2013) Cyclic di-GMP: the first 25 years of a universal bacterial second messenger. Microbiol Mol Biol Rev 77:1–52. CrossRefPubMedPubMedCentralGoogle Scholar
  4. 4.
    Tal R et al (1998) Three cdg operons control cellular turnover of cyclic di-GMP in Acetobacter xylinum: genetic organization and occurrence of conserved domains in isoenzymes. J Bacteriol 180:4416–4425CrossRefGoogle Scholar
  5. 5.
    Ryjenkov DA, Simm R, Römling U, Gomelsky M (2006) The PilZ domain is a receptor for the second messenger c-di-GMP: the PilZ domain protein YcgR controls motility in enterobacteria. J Biol Chem 281:30310–30314CrossRefGoogle Scholar
  6. 6.
    Schmidt AJ, Ryjenkov DA, Gomelsky M (2005) The ubiquitous protein domain EAL is a cyclic diguanylate-specific phosphodiesterase: enzymatically active and inactive EAL domains. J Bacteriol 187:4774–4781CrossRefGoogle Scholar
  7. 7.
    Simm R, Morr M, Kader A, Nimtz M, Römling U (2004) GGDEF and EAL domains inversely regulate cyclic di-GMP levels and transition from sessility to motility. Mol Microbiol 53:1123–1134CrossRefGoogle Scholar
  8. 8.
    Tamayo R, Patimalla B, Camilli A (2010) Growth in a biofilm induces a hyperinfectious phenotype in Vibrio cholerae. Infect Immun 78:3560–3569. IAI.00048-10 [pii]CrossRefPubMedPubMedCentralGoogle Scholar
  9. 9.
    McKee RW, Kariisa A, Mudrak B, Whitaker C, Tamayo R (2014) A systematic analysis of the in vitro and in vivo functions of the HD-GYP domain proteins of Vibrio cholerae. BMC Microbiol 14:272. CrossRefPubMedPubMedCentralGoogle Scholar
  10. 10.
    Stelitano V, Giardina G, Paiardini A, Castiglione N, Cutruzzola F, Rinaldo S (2013) C-di-GMP hydrolysis by Pseudomonas aeruginosa HD-GYP phosphodiesterases: analysis of the reaction mechanism and novel roles for pGpG. PLoS One 8:e74920. CrossRefPubMedPubMedCentralGoogle Scholar
  11. 11.
    Galperin MY, Nikolskaya AN, Koonin EV (2001) Novel domains of the prokaryotic two-component signal transduction systems. FEMS Microbiol Lett 203:11–21CrossRefGoogle Scholar
  12. 12.
    Witte G, Hartung S, Buttner K, Hopfner KP (2008) Structural biochemistry of a bacterial checkpoint protein reveals diadenylate cyclase activity regulated by DNA recombination intermediates. Mol Cell 30:167–178. S1097-2765(08)00166-4 [pii]CrossRefPubMedGoogle Scholar
  13. 13.
    Commichau FM, Dickmanns A, Gundlach J, Ficner R, Stulke J (2015) A jack of all trades: the multiple roles of the unique essential second messenger cyclic di-AMP. Mol Microbiol 97:189–204. CrossRefPubMedGoogle Scholar
  14. 14.
    Corrigan RM, Campeotto I, Jeganathan T, Roelofs KG, Lee VT, Gründling A (2013) Systematic identification of conserved bacterial c-di-AMP receptor proteins. Proc Natl Acad Sci USA 110:9084–9089. 1300595110 [pii]CrossRefPubMedGoogle Scholar
  15. 15.
    Huynh TN, Woodward JJ (2016) Too much of a good thing: regulated depletion of c-di-AMP in the bacterial cytoplasm. Curr Opin Microbiol 30:22–29. CrossRefPubMedPubMedCentralGoogle Scholar
  16. 16.
    Davies BW, Bogard RW, Young TS, Mekalanos JJ (2012) Coordinated regulation of accessory genetic elements produces cyclic di-nucleotides for V. cholerae virulence. Cell 149:358–370. S0092-8674(12)00290-5 [pii]CrossRefPubMedPubMedCentralGoogle Scholar
  17. 17.
    Kellenberger CA et al (2015) GEMM-I riboswitches from Geobacter sense the bacterial second messenger cyclic AMP-GMP. Proc Natl Acad Sci USA 112:5383–5388. CrossRefPubMedGoogle Scholar
  18. 18.
    Nelson JW, Sudarsan N, Phillips GE, Stav S, Lunse CE, McCown PJ, Breaker RR (2015) Control of bacterial exoelectrogenesis by c-AMP-GMP. Proc Natl Acad Sci USA 112:5389–5394. CrossRefPubMedGoogle Scholar
  19. 19.
    Lacey MM, Partridge JD, Green J (2010) Escherichia coli K-12 YfgF is an anaerobic cyclic di-GMP phosphodiesterase with roles in cell surface remodelling and the oxidative stress response. Microbiology 156:2873–2886. CrossRefPubMedGoogle Scholar
  20. 20.
    Cohen D et al (2015) Oligoribonuclease is a central feature of cyclic diguanylate signaling in Pseudomonas aeruginosa. Proc Natl Acad Sci USA 112:11359–11364. CrossRefPubMedGoogle Scholar
  21. 21.
    Orr MW, Donaldson GP, Severin GB, Wang J, Sintim HO, Waters CM, Lee VT (2015) Oligoribonuclease is the primary degradative enzyme for pGpG in Pseudomonas aeruginosa that is required for cyclic-di-GMP turnover. Proc Natl Acad Sci USA 112:E5048–E5057. CrossRefPubMedGoogle Scholar
  22. 22.
    Goldman SR, Sharp JS, Vvedenskaya IO, Livny J, Dove SL, Nickels BE (2011) NanoRNAs prime transcription initiation in vivo. Mol Cell 42:817–825. S1097-2765(11)00419-9 [pii]CrossRefPubMedPubMedCentralGoogle Scholar
  23. 23.
    Vvedenskaya IO, Sharp JS, Goldman SR, Kanabar PN, Livny J, Dove SL, Nickels BE (2012) Growth phase-dependent control of transcription start site selection and gene expression by nanoRNAs. Genes Dev 26:1498–1507. CrossRefPubMedPubMedCentralGoogle Scholar
  24. 24.
    Druzhinin SY, Tran NT, Skalenko KS, Goldman SR, Knoblauch JG, Dove SL, Nickels BE (2015) A conserved pattern of primer-dependent transcription initiation in Escherichia coli and Vibrio cholerae revealed by 5′ RNA-seq. PLoS Genet 11:e1005348. CrossRefPubMedPubMedCentralGoogle Scholar
  25. 25.
    Povolotsky TL, Hengge R (2012) ‘Life-style’ control networks in Escherichia coli: signaling by the second messenger c-di-GMP. J Biotechnol 160:10–16. CrossRefPubMedGoogle Scholar
  26. 26.
    Orr MW et al (2018) A subset of exoribonucleases serve as degradative enzymes for pGpG in c-di-GMP signaling. J Bacteriol 200:e00300-18. CrossRefPubMedPubMedCentralGoogle Scholar
  27. 27.
    Datta AK, Niyogi K (1975) A novel oligoribonuclease of Escherichia coli. II. Mechanism of action. J Biol Chem 250:7313–7319PubMedGoogle Scholar
  28. 28.
    Niyogi SK, Datta AK (1975) A novel oligoribonuclease of Escherichia coli. I. Isolation and properties. J Biol Chem 250:7307–7312PubMedGoogle Scholar
  29. 29.
    Deutscher MP (2015) Twenty years of bacterial RNases and RNA processing: how we’ve matured. RNA (New York, NY) 21:597–600. CrossRefGoogle Scholar
  30. 30.
    Stevens A, Niyogi SK (1967) Hydrolysis of oligoribonucleotides by an enzyme fraction from Escherichia coli. Biochem Biophys Res Commun 29:550–555CrossRefGoogle Scholar
  31. 31.
    Nirenberg M, Leder P (1964) RNA codewords and protein synthesis. The effect of trinucleotides upon the binding of sRNA to ribosomes. Science 145:1399–1407CrossRefGoogle Scholar
  32. 32.
    Zhang X, Zhu L, Deutscher MP (1998) Oligoribonuclease is encoded by a highly conserved gene in the 3′-5′ exonuclease superfamily. J Bacteriol 180:2779–2781CrossRefGoogle Scholar
  33. 33.
    Ghosh S, Deutscher MP (1999) Oligoribonuclease is an essential component of the mRNA decay pathway. Proc Natl Acad Sci USA 96:4372–4377CrossRefGoogle Scholar
  34. 34.
    Mechold U, Ogryzko V, Ngo S, Danchin A (2006) Oligoribonuclease is a common downstream target of lithium-induced pAp accumulation in Escherichia coli and human cells. Nucleic Acids Res 34:2364–2373. CrossRefPubMedPubMedCentralGoogle Scholar
  35. 35.
    Mechold U, Fang G, Ngo S, Ogryzko V, Danchin A (2007) YtqI from Bacillus subtilis has both oligoribonuclease and pAp-phosphatase activity. Nucleic Acids Res 35:4552–4561. CrossRefPubMedPubMedCentralGoogle Scholar
  36. 36.
    Fang M, Zeisberg WM, Condon C, Ogryzko V, Danchin A, Mechold U (2009) Degradation of nanoRNA is performed by multiple redundant RNases in Bacillus subtilis. Nucleic Acids Res 37:5114–5125. CrossRefPubMedPubMedCentralGoogle Scholar
  37. 37.
    Liu MF et al (2012) Identification of a novel nanoRNase in Bartonella. Microbiology 158:886–895. CrossRefPubMedGoogle Scholar
  38. 38.
    Zuo Y, Deutscher MP (2001) Exoribonuclease superfamilies: structural analysis and phylogenetic distribution. Nucleic Acids Res 29:1017–1026CrossRefGoogle Scholar
  39. 39.
    Aravind L, Koonin EV (1998) A novel family of predicted phosphoesterases includes Drosophila prune protein and bacterial RecJ exonuclease. Trends Biochem Sci 23:17–19CrossRefGoogle Scholar
  40. 40.
    Yuan Z, Gao F, Yin K, Gu L (2018) NrnC, an RNase D-like protein from Agrobacterium, is a novel octameric nuclease that specifically degrades dsDNA but leaves dsRNA intact. Front Microbiol 9:3230. CrossRefPubMedGoogle Scholar
  41. 41.
    Zuo Y et al (2007) Crystal structure of RNase T, an exoribonuclease involved in tRNA maturation and end turnover. Structure 15:417–428. CrossRefPubMedPubMedCentralGoogle Scholar
  42. 42.
    Zuo Y, Deutscher MP (2002) The physiological role of RNase T can be explained by its unusual substrate specificity. J Biol Chem 277:29654–29661. CrossRefPubMedGoogle Scholar
  43. 43.
    Li Z, Pandit S, Deutscher MP (1998) 3′ exoribonucleolytic trimming is a common feature of the maturation of small, stable RNAs in Escherichia coli. Proc Natl Acad Sci USA 95:2856–2861. CrossRefPubMedGoogle Scholar
  44. 44.
    Hsiao YY, Duh Y, Chen YP, Wang YT, Yuan HS (2012) How an exonuclease decides where to stop in trimming of nucleic acids: crystal structures of RNase T-product complexes. Nucleic Acids Res 40:8144–8154. CrossRefPubMedPubMedCentralGoogle Scholar
  45. 45.
    Schmier BJ, Nelersa CM, Malhotra A (2017) Structural basis for the bidirectional activity of Bacillus nanoRNase NrnA. Sci Rep 7:11085. CrossRefPubMedPubMedCentralGoogle Scholar
  46. 46.
    Wakamatsu T, Kim K, Uemura Y, Nakagawa N, Kuramitsu S, Masui R (2011) Role of RecJ-like protein with 5′-3′ exonuclease activity in oligo(deoxy)nucleotide degradation. J Biol Chem 286:2807–2816. CrossRefPubMedGoogle Scholar
  47. 47.
    Zuo Y, Wang Y, Malhotra A (2005) Crystal structure of Escherichia coli RNase D, an exoribonuclease involved in structured RNA processing. Structure 13:973–984. CrossRefPubMedGoogle Scholar
  48. 48.
    Huynh TN, Luo S, Pensinger D, Sauer JD, Tong L, Woodward JJ (2015) An HD-domain phosphodiesterase mediates cooperative hydrolysis of c-di-AMP to affect bacterial growth and virulence. Proc Natl Acad Sci USA 112:E747–E756. CrossRefPubMedGoogle Scholar
  49. 49.
    Bai Y et al (2013) Two DHH subfamily 1 proteins in Streptococcus pneumoniae possess cyclic di-AMP phosphodiesterase activity and affect bacterial growth and virulence. J Bacteriol 195:5123–5132. CrossRefPubMedPubMedCentralGoogle Scholar
  50. 50.
    Corrigan RM, Abbott JC, Burhenne H, Kaever V, Gründling A (2011) c-di-AMP is a new second messenger in Staphylococcus aureus with a role in controlling cell size and envelope stress. PLoS Pathog 7:e1002217. CrossRefPubMedPubMedCentralGoogle Scholar
  51. 51.
    Manikandan K, Sabareesh V, Singh N, Saigal K, Mechold U, Sinha KM (2014) Two-step synthesis and hydrolysis of cyclic di-AMP in Mycobacterium tuberculosis. PLoS One 9:e86096. CrossRefPubMedPubMedCentralGoogle Scholar
  52. 52.
    Tang Q, Luo Y, Zheng C, Yin K, Ali MK, Li X, He J (2015) Functional analysis of a c-di-AMP-specific phosphodiesterase MsPDE from Mycobacterium smegmatis. Int J Biol Sci 11:813–824. CrossRefPubMedPubMedCentralGoogle Scholar
  53. 53.
    Yang J, Bai Y, Zhang Y, Gabrielle VD, Jin L, Bai G (2014) Deletion of the cyclic di-AMP phosphodiesterase gene (cnpB) in Mycobacterium tuberculosis leads to reduced virulence in a mouse model of infection. Mol Microbiol 93:65–79. CrossRefPubMedPubMedCentralGoogle Scholar
  54. 54.
    Ye M et al (2014) DhhP, a cyclic di-AMP phosphodiesterase of Borrelia burgdorferi, is essential for cell growth and virulence. Infect Immun 82:1840–1849. CrossRefPubMedPubMedCentralGoogle Scholar
  55. 55.
    He Q et al (2016) Structural and biochemical insight into the mechanism of Rv2837c from Mycobacterium tuberculosis as a c-di-NMP phosphodiesterase. J Biol Chem 291:14386–14387. CrossRefPubMedPubMedCentralGoogle Scholar
  56. 56.
    Wang F, He Q, Su K, Wei T, Xu S, Gu L (2018) Structural and biochemical characterization of the catalytic domains of GdpP reveals a unified hydrolysis mechanism for the DHH/DHHA1 phosphodiesterase. Biochem J 475:191–205. CrossRefPubMedGoogle Scholar
  57. 57.
    Bowman L, Zeden MS, Schuster CF, Kaever V, Grundling A (2016) New insights into the cyclic di-adenosine monophosphate (c-di-AMP) degradation pathway and the requirement of the cyclic dinucleotide for acid stress resistance in Staphylococcus aureus. J Biol Chem 291:26970–26986. CrossRefPubMedPubMedCentralGoogle Scholar
  58. 58.
    Gao J et al (2015) Identification and characterization of phosphodiesterases that specifically degrade 3′3′-cyclic GMP-AMP. Cell Res 25:539–550. CrossRefPubMedPubMedCentralGoogle Scholar
  59. 59.
    Whiteley AT et al (2019) Bacterial cGAS-like enzymes synthesize diverse nucleotide signals. Nature 567(7747):194–199. CrossRefPubMedPubMedCentralGoogle Scholar

Copyright information

© Springer Nature Switzerland AG 2020

Authors and Affiliations

  1. 1.Eunice Kennedy Shriver National Institute of Child Health and Human Development, National Institutes of HealthBethesdaUSA
  2. 2.Department of Cell Biology and Molecular GeneticsUniversity of MarylandCollege ParkUSA

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