1887

Abstract

The cyanobacterium can form lipid droplets (LDs), internal inclusions containing triacylglycerols, carotenoids and alkanes. LDs are enriched for a 17 carbon-long alkane in , and it has been shown that the overexpression of the and genes results in increased LD and alkane production. To identify transcriptional adaptations associated with increased alkane production, we performed comparative transcriptomic analysis of an alkane overproduction strain. RNA-seq data identified a large number of highly upregulated genes in the overproduction strain, including genes potentially involved in rRNA processing, mycosporine-glycine production and synthesis of non-ribosomal peptides, including nostopeptolide A. Other genes encoding helical carotenoid proteins, stress-induced proteins and those for microviridin synthesis were also upregulated. Construction of strains with several upregulated genes or operons on multi-copy plasmids resulted in reduced alkane accumulation, indicating possible negative regulators of alkane production. A strain containing four genes for microviridin biosynthesis completely lost the ability to synthesize LDs. This strain exhibited wild-type growth and lag phase recovery under standard conditions, and slightly faster growth under high light. The transcriptional changes associated with increased alkane production identified in this work will provide the basis for future experiments designed to use cyanobacteria as a production platform for biofuel or high-value hydrophobic products.

Keyword(s): alkane , lipid droplets and microviridin
Funding
This study was supported by the:
  • National Institutes of Health (Award TL4GM118977)
    • Principle Award Recipient: Kevin A. Gomez Pinto
  • National Science Foundation (Award MCB-1413583)
    • Principle Award Recipient: Michael L. Summers
Loading

Article metrics loading...

/content/journal/mgen/10.1099/mgen.0.000432
2020-09-17
2021-07-31
Loading full text...

Full text loading...

/deliver/fulltext/mgen/6/10/mgen000432.html?itemId=/content/journal/mgen/10.1099/mgen.0.000432&mimeType=html&fmt=ahah

References

  1. Peramuna A, Summers ML. Composition and occurrence of lipid droplets in the cyanobacterium Nostoc punctiforme . Arch Microbiol 2014; 196:881–890 [View Article]
    [Google Scholar]
  2. Winters K, Parker PL, Van Baalen C. Hydrocarbons of blue-green algae: geochemical significance. Science 1969; 163:467–468 [View Article]
    [Google Scholar]
  3. Lea-Smith DJ, Biller SJ, Davey MP, Cotton CAR, Perez Sepulveda BM et al. Contribution of cyanobacterial alkane production to the ocean hydrocarbon cycle. Proc Natl Acad Sci U S A 2015; 112:13591–13596 [View Article][PubMed]
    [Google Scholar]
  4. Coates RC, Podell S, Korobeynikov A, Lapidus A, Pevzner P et al. Characterization of cyanobacterial hydrocarbon composition and distribution of biosynthetic pathways. PLoS One 2014; 9:e85140 [View Article]
    [Google Scholar]
  5. Mendez-Perez D, Begemann MB, Pfleger BF. Modular synthase-encoding gene involved in α-olefin biosynthesis in Synechococcus sp. strain PCC 7002. Appl Environ Microbiol 2011; 77:4264–4267 [View Article]
    [Google Scholar]
  6. Schirmer A, Rude MA, Li X, Popova E, del Cardayre SB. Microbial biosynthesis of alkanes. Science 2010; 329:559–562 [View Article]
    [Google Scholar]
  7. Klähn S, Baumgartner D, Pfreundt U, Voigt K, Schön V et al. Alkane biosynthesis genes in cyanobacteria and their transcriptional organization. Front Bioeng Biotechnol 2014; 2:24 [View Article][PubMed]
    [Google Scholar]
  8. Lea-Smith DJ, Ortiz-Suarez ML, Lenn T, Nürnberg DJ, Baers LL et al. Hydrocarbons are essential for optimal cell size, division, and growth of cyanobacteria. Plant Physiology 1928; 2016:172
    [Google Scholar]
  9. Berla BM, Saha R, Maranas CD, Pakrasi HB. Cyanobacterial alkanes modulate photosynthetic cyclic electron flow to assist growth under cold stress. Sci Rep 2015; 5:14894 [View Article]
    [Google Scholar]
  10. Peramuna A, Morton R, Summers M. Enhancing alkane production in cyanobacterial lipid droplets: a model platform for industrially relevant compound production. Life 2015; 5:1111–1126 [View Article]
    [Google Scholar]
  11. Cao Y-X, Xiao W-H, Zhang J-L, Xie Z-X, Ding M-Z et al. Heterologous biosynthesis and manipulation of alkanes in Escherichia coli . Metab Eng 2016; 38:19–28 [View Article]
    [Google Scholar]
  12. Song X, Yu H, Zhu K. Improving alkane synthesis in Escherichia coli via metabolic engineering. Appl Microbiol Biotechnol 2016; 100:757–767 [View Article]
    [Google Scholar]
  13. Jiménez-Díaz L, Caballero A, Pérez-Hernández N, Segura A. Microbial alkane production for jet fuel industry: motivation, state of the art and perspectives. Microb Biotechnol 2017; 10:103–124 [View Article]
    [Google Scholar]
  14. Zargar A, Bailey CB, Haushalter RW, Eiben CB, Katz L et al. Leveraging microbial biosynthetic pathways for the generation of ‘drop-in’ biofuels. Curr Opin Biotechnol 2017; 45:156–163 [View Article]
    [Google Scholar]
  15. Wang J, Zhu K. Microbial production of alka(e)ne biofuels. Curr Opin Biotechnol 2018; 50:11–18 [View Article]
    [Google Scholar]
  16. Sorigué D, Légeret B, Cuiné S, Blangy S, Moulin S et al. An algal photoenzyme converts fatty acids to hydrocarbons. Science 2017; 357:903–907 [View Article][PubMed]
    [Google Scholar]
  17. Choi YJ, Lee SY. Microbial production of short-chain alkanes. Nature 2013; 502:571–574 [View Article]
    [Google Scholar]
  18. Sheppard MJ, Kunjapur AM, Prather KLJ. Modular and selective biosynthesis of gasoline-range alkanes. Metab Eng 2016; 33:28–40 [View Article]
    [Google Scholar]
  19. Allen MB, Arnon DI. Studies on nitrogen-fixing blue-green algae. I. growth and nitrogen fixation by Anabaena cylindrica Lemm. Plant Physiol 1955; 30:366–372 [View Article]
    [Google Scholar]
  20. Summers ML, Wallis JG, Campbell EL, Meeks JC. Genetic evidence of a major role for glucose-6-phosphate dehydrogenase in nitrogen fixation and dark growth of the cyanobacterium Nostoc sp. strain ATCC 29133. J Bacteriol 1995; 177:6184–6194 [View Article]
    [Google Scholar]
  21. Schmidt-Goff CM, Federspiel NA. In vivo and in vitro footprinting of a light-regulated promoter in the cyanobacterium Fremyella diplosiphon . J Bacteriol 1993; 175:1806–1813 [View Article]
    [Google Scholar]
  22. Bolger AM, Lohse M, Usadel B. Trimmomatic: a flexible trimmer for Illumina sequence data. Bioinformatics 2014; 30:2114–2120 [View Article]
    [Google Scholar]
  23. Gordon A. FASTQ/A short-reads pre-processing tools. http://hannonlabcshledu/fastx_toolkit/ ; 2019
  24. Trapnell C, Pachter L, Salzberg SL. TopHat: discovering splice junctions with RNA-seq. Bioinformatics 2009; 25:1105–1111 [View Article]
    [Google Scholar]
  25. Trapnell C, Roberts A, Goff L, Pertea G, Kim D et al. Differential gene and transcript expression analysis of RNA-Seq experiments with TopHat and Cufflinks. Nat Protoc 2012; 7:562–578 [View Article]
    [Google Scholar]
  26. Trapnell C, Hendrickson DG, Sauvageau M, Goff L, Rinn JL et al. Differential analysis of gene regulation at transcript resolution with RNA-seq. Nat Biotechnol 2013; 31:46–53 [View Article]
    [Google Scholar]
  27. Ichihara Ken'ichi, Fukubayashi Y, Ichihara K YF. Preparation of fatty acid methyl esters for gas-liquid chromatography. J Lipid Res 2010; 51:635–640 [View Article]
    [Google Scholar]
  28. Anantharaman V, Aravind L. The NYN domains: novel predicted RNAses with a PIN domain-like fold. RNA Biol 2006; 3:18–27 [View Article]
    [Google Scholar]
  29. Szklarczyk D, Gable A, Lyon D, Junge A, Wyder S et al. STRING v11: protein-protein association networks with increased coverage, supporting functional discovery in genome-wide experimental datasets. Nuc Acids Res 2019; 43:D447–452
    [Google Scholar]
  30. Suh H-J, Lee H-W, Jung J. Mycosporine glycine protects biological systems against photodynamic damage by quenching singlet oxygen with a high efficiency. Photochem Photobiol 2007; 78:109–113 [View Article]
    [Google Scholar]
  31. Oren A, Gunde-Cimerman N. Mycosporines and mycosporine-like amino acids: UV protectants or multipurpose secondary metabolites?. FEMS Microbiol Lett 2007; 269:1–10 [View Article]
    [Google Scholar]
  32. Balskus EP, Walsh CT. The genetic and molecular basis for sunscreen biosynthesis in cyanobacteria. Science 2010; 329:1653–1656 [View Article]
    [Google Scholar]
  33. Gao Q, Garcia-Pichel F. An ATP-Grasp ligase involved in the last biosynthetic step of the iminomycosporine shinorine in Nostoc punctiforme ATCC 29133. J Bacteriol 2011; 193:5923–5928 [View Article]
    [Google Scholar]
  34. Völker U, Engelmann S, Maul B, Riethdorf S, Völker A et al. Analysis of the induction of general stress proteins of Bacillus subtilis . Microbiol 1994; 140:741–752 [View Article]
    [Google Scholar]
  35. Prágai Z, Harwood CR. Regulatory interactions between the PHO and σB-dependent general stress regulons of Bacillus subtilis . Microbiol 2002; 148:1593–1602 [View Article]
    [Google Scholar]
  36. Bao H, Melnicki MR, Kerfeld CA. Structure and functions of orange carotenoid protein homologs in cyanobacteria. Curr Opin Plant Biol 2017; 37:1–9 [View Article]
    [Google Scholar]
  37. López-Igual R, Wilson A, Leverenz RL, Melnicki MR, Bourcier de Carbon C et al. Different functions of the paralogs to the N-terminal domain of the orange carotenoid protein in the cyanobacterium Anabaena sp. PCC 7120. Plant Physiol 2016; 171:1852–1866 [View Article]
    [Google Scholar]
  38. Liaimer A, Jenke-Kodama H, Ishida K, Hinrichs K, Stangeland J et al. A polyketide interferes with cellular differentiation in the symbiotic cyanobacterium Nostoc punctiforme . Environ Microbiol Rep 2011; 3:550–558 [View Article]
    [Google Scholar]
  39. Khater S, Gupta M, Agrawal P, Sain N, Prava J et al. SBSPKSv2: structure-based sequence analysis of polyketide synthases and non-ribosomal peptide synthetases. Nucleic Acids Res 2017; 45:W72–W79 [View Article]
    [Google Scholar]
  40. Dehm D, Krumbholz J, Baunach M, Wiebach V, Hinrichs K et al. Unlocking the spatial control of secondary metabolism uncovers hidden natural product diversity in Nostoc punctiforme . ACS Chem Biol 2019; 14:1271–1279 [View Article]
    [Google Scholar]
  41. Fidor A, Konkel R, Mazur-Marzec H. Bioactive peptides produced by cyanobacteria of the genus Nostoc: a review. Mar Drugs 2019; 17:561–577 [View Article]
    [Google Scholar]
  42. Hoffmann D, Hevel JM, Moore RE, Moore BS. Sequence analysis and biochemical characterization of the nostopeptolide A biosynthetic gene cluster from Nostoc sp. GSV224. Gene 2003; 311:171–180 [View Article]
    [Google Scholar]
  43. Liaimer A, Helfrich EJN, Hinrichs K, Guljamow A, Ishida K et al. Nostopeptolide plays a governing role during cellular differentiation of the symbiotic cyanobacterium Nostoc punctiforme . Proc Natl Acad Sci USA 1862; 2015:112
    [Google Scholar]
  44. Viklund H, Bernsel A, Skwark M, Elofsson A. SPOCTOPUS: a combined predictor of signal peptides and membrane protein topology. Bioinformatics 2008; 24:2928–2929 [View Article]
    [Google Scholar]
  45. Yeats C, Bateman A. The BON domain: a putative membrane-binding domain. Trends Biochem Sci 2003; 28:352–355 [View Article]
    [Google Scholar]
  46. Yim HH, Villarejo M. osmY, a new hyperosmotically inducible gene, encodes a periplasmic protein in Escherichia coli . J Bacteriol 1992; 174:3637–3644 [View Article][PubMed]
    [Google Scholar]
  47. Ahmed MN, Reyna-González E, Schmid B, Wiebach V, Süssmuth RD et al. Phylogenomic analysis of the microviridin biosynthetic pathway coupled with targeted chemo-enzymatic synthesis yields potent protease inhibitors. ACS Chem Biol 2017; 12:1538–1546 [View Article]
    [Google Scholar]
  48. Agrawal C, Sen S, Singh S, Rai S, Singh PK et al. Comparative proteomics reveals association of early accumulated proteins in conferring butachlor tolerance in three N2-fixing Anabaena spp. J Proteomics 2014; 96:271–290 [View Article]
    [Google Scholar]
  49. Takahashi E, Wraight CA. Potentiation of proton transfer function by electrostatic interactions in photosynthetic reaction centers from Rhodobacter sphaeroides: First results from site-directed mutation of the H subunit. Proc Natl Acad Sci U S A 1996; 93:2640–2645 [View Article]
    [Google Scholar]
  50. Panda B, Basu B, Rajaram H, Apte SK. Comparative proteomics of oxidative stress response in three cyanobacterial strains native to Indian paddy fields. J Proteomics 2015; 127:152–160 [View Article]
    [Google Scholar]
  51. Martinis J, Glauser G, Valimareanu S, Stettler M, Zeeman SC et al. ABC1K1/PGR6 kinase: a regulatory link between photosynthetic activity and chloroplast metabolism. Plant J 2014; 77:269–283 [View Article]
    [Google Scholar]
  52. Asayama M, Imamura S, Yoshihara S, Miyazaki A, Yoshida N et al. SigC, the Group 2 sigma factor of RNA polymerase, contributes to the late-stage gene expression and nitrogen promoter recognition in the cyanobacterium Synechocystis sp. strain PCC 6803. Biosci Biotechnol Biochem 2004; 68:477–487 [View Article]
    [Google Scholar]
  53. Hauf S, Möller L, Fuchs S, Halbedel S. PadR-type repressors controlling production of a non-canonical FtsW/RodA homologue and other trans-membrane proteins. Sci Rep 2019; 9:10023 [View Article]
    [Google Scholar]
http://instance.metastore.ingenta.com/content/journal/mgen/10.1099/mgen.0.000432
Loading
/content/journal/mgen/10.1099/mgen.0.000432
Loading

Data & Media loading...

Supplements

Supplementary material 1

EXCEL

Most cited this month Most Cited RSS feed

This is a required field
Please enter a valid email address
Approval was a Success
Invalid data
An Error Occurred
Approval was partially successful, following selected items could not be processed due to error