Control of nitrogen fixation in bacteria that associate with cereals


Legumes obtain nitrogen from air through rhizobia residing in root nodules. Some species of rhizobia can colonize cereals but do not fix nitrogen on them. Disabling native regulation can turn on nitrogenase expression, even in the presence of nitrogenous fertilizer and low oxygen, but continuous nitrogenase production confers an energy burden. Here, we engineer inducible nitrogenase activity in two cereal endophytes (Azorhizobium caulinodans ORS571 and Rhizobium sp. IRBG74) and the well-characterized plant epiphyte Pseudomonas protegens Pf-5, a maize seed inoculant. For each organism, different strategies were taken to eliminate ammonium repression and place nitrogenase expression under the control of agriculturally relevant signals, including root exudates, biocontrol agents and phytohormones. We demonstrate that R. sp. IRBG74 can be engineered to result in nitrogenase activity under free-living conditions by transferring a nif cluster from either Rhodobacter sphaeroides or Klebsiella oxytoca. For P. protegens Pf-5, the transfer of an inducible cluster from Pseudomonas stutzeri and Azotobacter vinelandii yields ammonium tolerance and higher oxygen tolerance of nitrogenase activity than that from K. oxytoca. Collectively, the data from the transfer of 12 nif gene clusters between 15 diverse species (including Escherichia coli and 12 rhizobia) help identify the barriers that must be overcome to engineer a bacterium to deliver a high nitrogen flux to a cereal crop.

Access options

Rent or Buy article

Get time limited or full article access on ReadCube.


All prices are NET prices.

Fig. 1: Transfer of nif clusters across species.
Fig. 2: Transfer of the native K. oxytoca nif cluster across species.
Fig. 3: Transfer of the refactored K. oxytoca nif clusters to R. sp. IRBG74.
Fig. 4: Control of nitrogen fixation in A. caulinodans ORS571.
Fig. 5: Nitrogenase activity of the inducible nif clusters in P. protegens Pf-5.
Fig. 6: Control of nitrogenase activity with sensors that respond to diverse chemicals in the rhizosphere.

Data availability

Additional data supporting this study are available from the corresponding author on reasonable request. The RNA-seq and ribosome-profiling data are available in the Sequence Read Archive with the accession code PRJNA579767: K. oxytoca native nif cluster, RNA-seq (SRX7032059, SRX7032060 and SRX7032061) and ribosome-profiling (SRX7034729, SRX7034730, SRX7034731 and SRX7034732); K. oxytoca refactored nif cluster v2.1, RNA-seq (SRX7036110) and ribosome-profiling (SRX7036099); K. oxytoca refactored nif cluster v3.2, RNA-seq (SRX7035703, SRX7035704 and SRX7035705) and ribosome-profiling (SRX7036113, SRX7036114 and SRX7036115).


  1. 1.

    Harlan, J. R. Crops and Man 2nd edn (American Society of Agronomy, 1992).

  2. 2.

    Simmonds, J. Community Matters: a History of Biological Nitrogen Fixation and Nodulation Research, 1965 to 1995. PhD thesis, Rensselaer Polytechnic Institute (2007).

  3. 3.

    Erisman, J., Bleeker, A., Galloway, J. & Sutton, M. Reduced nitrogen in ecology and the environment. Environ. Pollut. 150, 140–149 (2007).

  4. 4.

    Geddes, B. A. et al. Use of plant colonizing bacteria as chassis for transfer of N2-fixation to cereals. Curr. Opin. Biotechnol. 32, 216–222 (2015).

  5. 5.

    Zaim, S., Bekkar, A. A. & Belabid, L. in Rhizobium Biology and Biotechnology (eds Hansen, A. P. et al.) 25–37 (Springer, 2017).

  6. 6.

    Nadarajah, K. K. in Rhizobium Biology and Biotechnology (eds Hansen, A. P. et al.) 83–103 (Springer, 2017).

  7. 7.

    Gutierrez-Zamora, M. & Martınez-Romero, E. Natural endophytic association between Rhizobium etli and maize (Zea mays L.). J. Biotechnol. 91, 117–126 (2001).

  8. 8.

    McInroy, J. A. & Kloepper, J. W. Survey of indigenous bacterial endophytes from cotton and sweet corn. Plant Soil 173, 337–342 (1995).

  9. 9.

    Ramachandran, V. K., East, A. K., Karunakaran, R., Downie, J. A. & Poole, P. S. Adaptation of Rhizobium leguminosarum to pea, alfalfa and sugar beet rhizospheres investigated by comparative transcriptomics. Genome Biol. 12, R106 (2011).

  10. 10.

    Frans, J. et al. in Nitrogen Fixation (Ed. Gresshoff, P. M.) 33–44 (Springer, 1990).

  11. 11.

    Delmotte, N. et al. An integrated proteomics and transcriptomics reference data set provides new insights into the Bradyrhizobium japonicum bacteroid metabolism in soybean root nodules. Proteomics 10, 1391–1400 (2010).

  12. 12.

    James, E. Nitrogen fixation in endophytic and associative symbiosis. Field Crops Res. 65, 197–209 (2000).

  13. 13.

    Ars’ene, F., Katupitiya, S., Kennedy, I. R. & Elmerich, C. Use of IacZ fusions to study the expression of nif genes of Azospirillum brasilense. Mol. Plant Microbe Interact. 7, 748–757 (1994).

  14. 14.

    Boddey, R. M. et al. in Management of Biological Nitrogen Fixation for the Development of More Productive and Sustainable Agricultural Systems (eds Ladha, J. K. & Peoples, M. B.) 195–209 (Springer, 1995).

  15. 15.

    Bremer, E., Janzen, H. & Gilbertson, C. Evidence against associative N2 fixation as a significant N source in long-term wheat plots. Plant Soil 175, 13–19 (1995).

  16. 16.

    Chalk, P. The contribution of associative and symbiotic nitrogen fixation to the nitrogen nutrition of non-legumes. Plant Soil 132, 29–39 (1991).

  17. 17.

    Hurek, T., Reinhold-Hurek, B., Van Montagu, M. & Kellenberger, E. Root colonization and systemic spreading of Azoarcus sp. strain BH72 in grasses. J. Bacteriol. 176, 1913–1923 (1994).

  18. 18.

    ames, E., Olivares, F., Baldani, J. & Döbereiner, J. Herbaspirillum, an endophytic diazotroph colonizing vascular tissue 3 Sorghum bicolor L. Moench. J. Exp. Bot. 48, 785–798 (1997).

  19. 19.

    Kapulnik, Y., Feldman, M., Okon, Y. & Henis, Y. Contribution of nitrogen fixed by Azospirillum to the N nutrition of spring wheat in Israel. Soil Biology Biochem. 17, 509–515 (1985).

  20. 20.

    Katupitiya, S. et al. A mutant of Azospirillum brasilense Sp7 impaired in flocculation with a modified colonization pattern and superior nitrogen fixation in association with wheat. Appl. Environ. Microbiol. 61, 1987–1995 (1995).

  21. 21.

    Malik, K. et al. Association of nitrogen-fixing, plant-growth-promoting rhizobacteria (PGPR) with kallar grass and rice. Plant Soil 194, 37–44 (1997).

  22. 22.

    Reinhold-Hurek, B. & Hurek, T. Interactions of gramineous plants with Azoarcus spp. and other diazotrophs: identification, localization, and perspectives to study their function. Crit. Rev. Plant Sci. 17, 29–54 (1998).

  23. 23.

    Stoltzfus, J., So, R., Malarvithi, P., Ladha, J. & De Bruijn, F. Isolation of endophytic bacteria from rice and assessment of their potential for supplying rice with biologically fixed nitrogen. Plant Soil 194, 25–36 (1997).

  24. 24.

    Triplett, E. W. Diazotrophic endophytes: progress and prospects for nitrogen fixation in monocots. Plant Soil 186, 29–38 (1996).

  25. 25.

    Broek, A. V., Michiels, J., Van Gool, A. & Vanderleyden, J. Spatial-temporal colonization patterns of Azospirillum brasilense on the wheat root surface and expression of the bacterial nifH gene during association. Mol. Plant Microbe Interact. 6, 592–600 (1993).

  26. 26.

    Yanni, Y. G. et al. in Opportunities for Biological Nitrogen Fixation in Rice and Other Non-Legumes (eds Ladha, J. K. et al.) 99–114 (Springer, 1997).

  27. 27.

    Okon, Y. Azospirillum as a potential inoculant for agriculture. Trends Biotechnol. 3, 223–228 (1985).

  28. 28.

    Cocking, E. C. Endophytic colonization of plant roots by nitrogen-fixing bacteria. Plant Soil 252, 169–175 (2003).

  29. 29.

    Woodard, H. & Bly, A. Maize growth and yield responses to seed-inoculated N2-fixing bacteria under Dryland production conditions. J. Plant Nutr. 23, 55–65 (2000).

  30. 30.

    Igiehon, N. O. & Babalola, O. O. Rhizosphere microbiome modulators: contributions of nitrogen fixing bacteria towards sustainable agriculture. Int. J. Environ. Res. Public Health 15, 574 (2018).

  31. 31.

    Biswas, J., Ladha, J. & Dazzo, F. Rhizobia inoculation improves nutrient uptake and growth of lowland rice. Soil Sci. Soc. Am. J. 64, 1644–1650 (2000).

  32. 32.

    Biswas, J. C., Ladha, J. K., Dazzo, F. B., Yanni, Y. G. & Rolfe, B. G. Rhizobial inoculation influences seedling vigor and yield of rice. Agron. J. 92, 880–886 (2000).

  33. 33.

    Cummings, S. P. et al. Nodulation of Sesbania species by Rhizobium (Agrobacterium) strain IRBG74 and other rhizobia. Environ. Microbiol. 11, 2510–2525 (2009).

  34. 34.

    Masson-Boivin, C., Giraud, E., Perret, X. & Batut, J. Establishing nitrogen-fixing symbiosis with legumes: how many rhizobium recipes? Trends Microbiol. 17, 458–466 (2009).

  35. 35.

    Hoover, T. R., Imperial, J., Ludden, P. W. & Shah, V. K. Homocitrate is a component of the iron-molybdenum cofactor of nitrogenase. Biochemistry 28, 2768–2771 (1989).

  36. 36.

    Mandon, K. et al. Role of the fixGHI region of Azorhizobium caulinodans in free-living and symbiotic nitrogen fixation. FEMS Microbiol. Lett. 114, 185–189 (1993).

  37. 37.

    Tsukada, S. et al. Comparative genome-wide transcriptional profiling of Azorhizobium caulinodans ORS571 grown under free-living and symbiotic conditions. Appl. Environ. Microbiol. 75, 5037–5046 (2009).

  38. 38.

    Gopalaswamy, G., Kannaiyan, S., O’Callaghan, K. J., Davey, M. R. & Cocking, E. C. The xylem of rice (Oryza sativa) is colonized by Azorhizobium caulinodans. Proc. Biol. Sci. 267, 103–107 (2000).

  39. 39.

    Webster, G. et al. The flavonoid naringenin stimulates the intercellular colonization of wheat roots by Azorhizobium caulinodans. Plant Cell Environ. 21, 373–383 (1998).

  40. 40.

    Webster, G. et al. in Opportunities for Biological Nitrogen Fixation in Rice and Other Non-Legumes (eds Ladha, J. K. et al.) 115–122 (Springer, 1997).

  41. 41.

    Triplett, E. W., Kaeppler, S. M. & Chelius, M. K. Klebsiella pneumoniae inoculants for enhancing plant growth. US patent 7,393,678 (2008).

  42. 42.

    Fujii, T. et al. Effect of inoculation with Klebsiella oxytoca and Enterobacter cloacae on dinitrogen fixation by rice–bacteria associations. Plant Soil 103, 221–226 (1987).

  43. 43.

    El-Khawas, H. & Adachi, K. Identification and quantification of auxins in culture media of Azospirillum and Klebsiella and their effect on rice roots. Biol. Fertil. Soils 28, 377–381 (1999).

  44. 44.

    Palus, J. A., Borneman, J., Ludden, P. W. & Triplett, E. W. A diazotrophic bacterial endophyte isolated from stems of Zea mays L. and Zea luxurians Iltis and Doebley. Plant Soil 186, 135–142 (1996).

  45. 45.

    Rediers, H. et al. Development and application of a dapB-based in vivo expression technology system to study colonization of rice by the endophytic nitrogen-fixing bacterium Pseudomonas stutzeri A15. Appl. Environ. Microbiol. 69, 6864–6874 (2003).

  46. 46.

    Haahtela, K. & Korhonen, T. K. In vitro adhesion of N2-fixing enteric bacteria to roots of grasses and cereals. Appl. Environ. Microbiol. 49, 1186–1190 (1985).

  47. 47.

    Desnoues, N. et al. Nitrogen fixation genetics and regulation in a Pseudomonas stutzeri strain associated with rice. Microbiology 149, 2251–2262 (2003).

  48. 48.

    Paulsen, I. T. et al. Complete genome sequence of the plant commensal Pseudomonas fluorescens Pf-5. Nat. Biotechnol. 23, 873 (2005).

  49. 49.

    Walsh, U. F., Morrissey, J. P. & O’Gara, F. Pseudomonas for biocontrol of phytopathogens: from functional genomics to commercial exploitation. Curr. Opin. Biotechnol. 12, 289–295 (2001).

  50. 50.

    Dos Santos, P. C., Fang, Z., Mason, S. W., Setubal, J. C. & Dixon, R. Distribution of nitrogen fixation and nitrogenase-like sequences amongst microbial genomes. BMC Genomics 13, 162 (2012).

  51. 51.

    Boyd, E. S., Costas, A. M. G., Hamilton, T. L., Mus, F. & Peters, J. W. Evolution of molybdenum nitrogenase during the transition from anaerobic to aerobic metabolism. J. Bacteriol. 197, 1690–1699 (2015).

  52. 52.

    Raymond, J., Siefert, J. L., Staples, C. R. & Blankenship, R. E. The natural history of nitrogen fixation. Molec. Biol. Evol. 21, 541–554 (2004).

  53. 53.

    Rubio, L. M. & Ludden, P. W. Biosynthesis of the iron-molybdenum cofactor of nitrogenase. Annu. Rev. Microbiol. 62, 93–111 (2008).

  54. 54.

    Iki, T., Aono, T. & Oyaizu, H. Evidence for functional differentiation of duplicated nifH genes in Azorhizobium caulinodans. FEMS Microbiol. Lett. 274, 173–179 (2007).

  55. 55.

    Ratet, P., Pawlowski, K., Schell, J. & Bruijn, F. The Azorhizobium caulinodans nitrogen-fixation regulatory gene, nifA, is controlled by the cellular nitrogen and oxygen status. Mol. Microbiol. 3, 825–838 (1989).

  56. 56.

    Poudel, S. et al. Electron transfer to nitrogenase in different genomic and metabolic backgrounds. J. Bacteriol. 200, 00757–00717 (2018).

  57. 57.

    Boyd, E. & Peters, J. W. New insights into the evolutionary history of biological nitrogen fixation. Front. Microbiol. 4, 201 (2013).

  58. 58.

    Shah, V. K., Stacey, G. & Brill, W. J. Electron transport to nitrogenase. Purification and characterization of pyruvate: flavodoxin oxidoreductase. The nifJ gene product. J. Biol. Chem. 258, 12064–12068 (1983).

  59. 59.

    Schmehl, M. et al. Identification of a new class of nitrogen fixation genes in Rhodobacter capsalatus: a putative membrane complex involved in electron transport to nitrogenase. Mol. Gen. Genet. 241, 602–615 (1993).

  60. 60.

    Edgren, T. & Nordlund, S. The fixABCX genes in Rhodospirillum rubrum encode a putative membrane complex participating in electron transfer to nitrogenase. J. Bacteriol. 186, 2052–2060 (2004).

  61. 61.

    Pascuan, C., Fox, A. R., Soto, G. & Ayub, N. D. Exploring the ancestral mechanisms of regulation of horizontally acquired nitrogenases. J. Mol. Evol. 81, 84–89 (2015).

  62. 62.

    Yan, Y. et al. Nitrogen fixation island and rhizosphere competence traits in the genome of root-associated Pseudomonas stutzeri A1501. Proc. Natl Acad. Sci. USA 105, 7564–7569 (2008).

  63. 63.

    Kechris, K. J., Lin, J. C., Bickel, P. J. & Glazer, A. N. Quantitative exploration of the occurrence of lateral gene transfer by using nitrogen fixation genes as a case study. Proc. Natl Acad. Sci. USA 103, 9584–9589 (2006).

  64. 64.

    Thöny, B., Anthamatten, D. & Hennecke, H. Dual control of the Bradyrhizobium japonicum symbiotic nitrogen fixation regulatory operon fixR nifA: analysis of cis-and trans-acting elements. J. Bacteriol. 171, 4162–4169 (1989).

  65. 65.

    Li, X.-X., Liu, Q., Liu, X.-M., Shi, H.-W. & Chen, S.-F. Using synthetic biology to increase nitrogenase activity. Microb. Cell Fact. 15, 43 (2016).

  66. 66.

    Han, Y. et al. Interspecies transfer and regulation of Pseudomonas stutzeri A1501 nitrogen fixation Island in Escherichia coli. J. Microbiol. Biotechnol. 25, 1339–1348 (2015).

  67. 67.

    Dixon, R. A. & Postgate, J. R. Genetic transfer of nitrogen fixation from Klebsiella pneumoniae to Escherichia coli. Nature 237, 102 (1972).

  68. 68.

    Cannon, F., Dixon, R., Postgate, J. & Primrose, S. Chromosomal integration of Klebsiella nitrogen fixation genes in Escherichia coli. Microbiology 80, 227–239 (1974).

  69. 69.

    Cannon, F., Dixon, R., Postgate, J. & Primrose, S. Plasmids formed in nitrogen-fixing Escherichia coli–Klebsiella pneumoniae hybrids. Microbiology 80, 241–251 (1974).

  70. 70.

    Wang, L. et al. A minimal nitrogen fixation gene cluster from Paenibacillus sp. WLY78 enables expression of active nitrogenase in Escherichia coli. PLoS Genet. 9, e1003865 (2013).

  71. 71.

    Temme, K., Zhao, D. & Voigt, C. A. Refactoring the nitrogen fixation gene cluster from Klebsiella oxytoca. Proc. Natl Acad. Sci. USA 109, 7085–7090 (2012).

  72. 72.

    Smanski, M. J. et al. Functional optimization of gene clusters by combinatorial design and assembly. Nat. Biotechnol. 32, 1241 (2014).

  73. 73.

    Zhang, L., Liu, X., Li, X. & Chen, S. Expression of the N2 fixation gene operon of Paenibacillus sp. WLY78 under the control of the T7 promoter in Escherichia coli BL21. Biotechnol. Lett. 37, 1999–2004 (2015).

  74. 74.

    Postgate, J. R. & Kent, H. M. Qualitative evidence for expression of Klebsiella pneumoniae nif in Pseudomonas putida. Microbiology 133, 2563–2566 (1987).

  75. 75.

    Setten, L. et al. Engineering Pseudomonas protegens Pf-5 for nitrogen fixation and its application to improve plant growth under nitrogen-deficient conditions. PLoS ONE 8, e63666 (2013).

  76. 76.

    Dixon, R. & Kahn, D. Genetic regulation of biological nitrogen fixation. Nat. Rev. Microbiol. 2, 621–631 (2004).

  77. 77.

    Fischer, H.-M. Environmental regulation of rhizobial symbiotic nitrogen fixation genes. Trends Microbiol. 4, 317–320 (1996).

  78. 78.

    Tsoy, O. V., Ravcheev, D. A., Čuklina, J. & Gelfand, M. S. Nitrogen fixation and molecular oxygen: comparative genomic reconstruction of transcription regulation in Alphaproteobacteria. Front. Microbiol. 7, 1343 (2016).

  79. 79.

    Hill, S., Kennedy, C., Kavanagh, E., Goldberg, R. B. & Hanau, R. Nitrogen fixation gene (nifL) involved in oxygen regulation of nitrogenase synthesis in K. peumoniae. Nature 290, 424–426 (1981).

  80. 80.

    Mandon, K. et al. Poly-β-hydroxybutyrate turnover in Azorhizobium caulinodans is required for growth and affects nifA expression. J. Bacteriol. 180, 5070–5076 (1998).

  81. 81.

    Little, R., Reyes‐Ramirez, F., Zhang, Y., van Heeswijk, W. C. & Dixon, R. Signal transduction to the Azotobacter vinelandii NIFL–NIFA regulatory system is influenced directly by interaction with 2-oxoglutarate and the PII regulatory protein. EMBO J. 19, 6041–6050 (2000).

  82. 82.

    Poole, P., Ramachandran, V. & Terpolilli, J. Rhizobia: from saprophytes to endosymbionts. Nat. Rev. Microbiol. 16, 291–303 (2018).

  83. 83.

    Kong, Q., Wu, Q., Ma, Z. & Shen, S. Oxygen sensitivity of the nifLA promoter of Klebsiella pneumoniae. J. Bacteriol. 166, 353–356 (1986).

  84. 84.

    Brooks, S. J., Collins, J. J. & Brill, W. J. Repression of nitrogen fixation in Klebsiella pneumoniae at high temperature. J. Bacteriol. 157, 460–464 (1984).

  85. 85.

    Peoples, M., Ladha, J. & Herridge, D. Biological nitrogen fixation: an efficient source of nitrogen for sustainable agricultural production? Plant Soil 174, 3–28 (1995).

  86. 86.

    Barakat, M., Cheviron, B. & Angulo-Jaramillo, R. Influence of the irrigation technique and strategies on the nitrogen cycle and budget: a review. Agric. Water Manag. 178, 225–238 (2016).

  87. 87.

    Burger, M. & Jackson, L. E. Microbial immobilization of ammonium and nitrate in relation to ammonification and nitrification rates in organic and conventional cropping systems. Soil Biol. Biochem. 35, 29–36 (2003).

  88. 88.

    MacNeil, D. & Brill, W. Mutations in nif genes that cause Klebsiella pneumoniae to be derepressed for nitrogenase synthesis in the presence of ammonium. J. Bacteriol. 144, 744–751 (1980).

  89. 89.

    Bali, A., Blanco, G., Hill, S. & Kennedy, C. Excretion of ammonium by a nifL mutant of Azotobacter vinelandii fixing nitrogen. Appl. Environ. Microbiol. 58, 1711–1718 (1992).

  90. 90.

    Brewin, B., Woodley, P. & Drummond, M. The basis of ammonium release in nifL mutants of Azotobacter vinelandii. J. Bacteriol. 181, 7356–7362 (1999).

  91. 91.

    Arsène, F., Kaminski, P. A. & Elmerich, C. Control of Azospirillum brasilense NifA activity by PII: effect of replacing Tyr residues of the NifA N-terminal domain on NifA activity. FEMS Microbiol. Lett. 179, 339–343 (1999).

  92. 92.

    Souza, E., Pedrosa, F., Drummond, M., Rigo, L. & Yates, M. Control of Herbaspirillum seropedicae NifA activity by ammonium ions and oxygen. J. Bacteriol. 181, 681–684 (1999).

  93. 93.

    Paschen, A., Drepper, T., Masepohl, B. & Klipp, W. Rhodobacter capsulatus nifA mutants mediating nif gene expression in the presence of ammonium. FEMS Microbiol. Lett. 200, 207–213 (2001).

  94. 94.

    Rey, F. E., Heiniger, E. K. & Harwood, C. S. Redirection of metabolism for biological hydrogen production. Appl. Environ. Microbiol. 73, 1665–1671 (2007).

  95. 95.

    Smanski, M. J. et al. Synthetic biology to access and expand nature’s chemical diversity. Nat. Rev. Microbiol. 14, 135–149 (2016).

  96. 96.

    Scupham, A. J. et al. Inoculation with Sinorhizobium meliloti RMBPC-2 increases alfalfa yield compared with inoculation with a nonengineered wild-type strain. Appl. Environ. Microbiol. 62, 4260–4262 (1996).

  97. 97.

    Fox, A. R. et al. Major cereal crops benefit from biological nitrogen fixation when inoculated with the nitrogen‐fixing bacterium Pseudomonas protegens Pf-5 X940. Environ. Microbiol. 18, 3522–3534 (2016).

  98. 98.

    Suthar, H., Hingurao, K., Vaghashiya, J. & Parmar, J. Fermentation: a process for biofertilizer production. Microorg. Green Revol. 6, 229–252 (2017).

  99. 99.

    Lucy, M., Reed, E. & Glick, B. R. Applications of free living plant growth-promoting rhizobacteria. Antonie Van Leeuwenhoek 86, 1–25 (2004).

  100. 100.

    Mahmood, A., Turgay, O. C., Farooq, M. & Hayat, R. Seed biopriming with plant growth promoting rhizobacteria: a review. FEMS Microbiol. Ecol. 92, fiw112 (2016).

  101. 101.

    Velusamy, P., Immanuel, J. E., Gnanamanickam, S. S. & Thomashow, L. Biological control of rice bacterial blight by plant-associated bacteria producing 2, 4-diacetylphloroglucinol. Can. J. Microbiol. 52, 56–65 (2006).

  102. 102.

    Mishra, J. Development and Evaluation of Pseudomonas Based Bioformulation for Disease Control and Growth Enhancement of Zea Mays L. PhD thesis, Babasaheb Bhimrao Ambedkar University (2016).

  103. 103.

    Weller, D. M. et al. Role of 2,4-diacetylphloroglucinol-producing fluorescent Pseudomonas spp. in the defense of plant roots. Plant Biol. 9, 4–20 (2007).

  104. 104.

    Kaiser, C. et al. Exploring the transfer of recent plant photosynthates to soil microbes: mycorrhizal pathway vs direct root exudation. New Phytol. 205, 1537–1551 (2015).

  105. 105.

    Mus, F. et al. Symbiotic nitrogen fixation and the challenges to its extension to nonlegumes. Appl. Environ. Microbiol. 82, 3698–3710 (2016).

  106. 106.

    Perrine-Walker, F. M., Prayitno, J., Rolfe, B. G., Weinman, J. J. & Hocart, C. H. Infection process and the interaction of rice roots with rhizobia. J. Exp. Bot. 58, 3343–3350 (2007).

  107. 107.

    Gough, C. et al. Specific flavonoids promote intercellular root colonization of Arabidopsis thaliana by Azorhizobium caulinodans ORS571. Mol. Plant Microbe Interact. 10, 560–570 (1997).

  108. 108.

    O’Callaghan, K. J. et al. Effects of glucosinolates and flavonoids on colonization of the roots of Brassica napus by Azorhizobium caulinodans ORS571. Appl. Environ. Microbiol. 66, 2185–2191 (2000).

  109. 109.

    Mongiardini, E. J. et al. The rhizobial adhesion protein RapA1 is involved in adsorption of rhizobia to plant roots but not in nodulation. FEMS Microbiol. Ecol. 65, 279–288 (2008).

  110. 110.

    Matilla, M. A., Espinosa-Urgel, M., Rodríguez-Herva, J. J., Ramos, J. L. & Ramos-González, M. I. Genomic analysis reveals the major driving forces of bacterial life in the rhizosphere. Genome Biol. 8, R179 (2007).

  111. 111.

    Casas, M. I., Duarte, S., Doseff, A. I. & Grotewold, E. Flavone-rich maize: an opportunity to improve the nutritional value of an important commodity crop. Front. Plant Sci. 5, 440 (2014).

  112. 112.

    Ogo, Y., Ozawa, K., Ishimaru, T., Murayama, T. & Takaiwa, F. Transgenic rice seed synthesizing diverse flavonoids at high levels: a new platform for flavonoid production with associated health benefits. Plant Biotechnol. J. 11, 734–746 (2013).

  113. 113.

    Bacilio-Jiménez, M. et al. Chemical characterization of root exudates from rice (Oryza sativa) and their effects on the chemotactic response of endophytic bacteria. Plant Soil 249, 271–277 (2003).

  114. 114.

    Pini, F. et al. Lux bacterial biosensors for in vivo spatiotemporal mapping of root secretion. Plant Physiol. 174, 1289–1306 (2017).

  115. 115.

    Ryan, P. R., Dessaux, Y., Thomashow, L. S. & Weller, D. M. Rhizosphere engineering and management for sustainable agriculture. Plant Soil 321, 363–383 (2009).

  116. 116.

    Dessaux, Y., Grandclément, C. & Faure, D. Engineering the rhizosphere. Trends Plant Sci. 21, 266–278 (2016).

  117. 117.

    Geddes, B. A. et al. Engineering transkingdom signalling in plants to control gene expression in rhizosphere bacteria. Nat. Comm. 10, 3430 (2019).

  118. 118.

    Arnold, W., Rump, A., Klipp, W., Priefer, U. B. & Pühler, A. Nucleotide sequence of a 24,206-base-pair DNA fragment carrying the entire nitrogen fixation gene cluster of Klebsiella pneumoniae. J. Mol. Biol. 203, 715–738 (1988).

  119. 119.

    de Salamone, I. G., Döbereiner, J., Urquiaga, S. & Boddey, R. M. Biological nitrogen fixation in Azospirillum strain-maize genotype associations as evaluated by the 15N isotope dilution technique. Biol. Fert. Soils 23, 249–256 (1996).

  120. 120.

    Pedrosa, F. & Elmerich, C. in Associative and Endophytic Nitrogen-fixing Bacteria and Cyanobacterial Associations (eds Elmerich, C. & Newton, W. E.) 41–71 (Springer, 2007).

  121. 121.

    Welsh, E. A. et al. The genome of Cyanothece 51142, a unicellular diazotrophic cyanobacterium important in the marine nitrogen cycle. Proc. Natl Acad. Sci. USA 105, 15094–15099 (2008).

  122. 122.

    Hamilton, T. L. et al. Transcriptional profiling of nitrogen fixation in Azotobacter vinelandii. J. Bacteriol. 193, 4477–4486 (2011).

  123. 123.

    Oda, Y. et al. Functional genomic analysis of three nitrogenase isozymes in the photosynthetic bacterium Rhodopseudomonas palustris. J. Bacteriol. 187, 7784–7794 (2005).

  124. 124.

    Haselkorn, R. & Kapatral, V. in Genomes and Genomics of Nitrogen-fixing Organisms (eds Palacios, R. & Newton, W. E.) 71–82 (Springer, 2005).

  125. 125.

    Jeong, H.-S. & Jouanneau, Y. Enhanced nitrogenase activity in strains of Rhodobacter capsulatus that overexpress the rnf genes. J. Bacteriol. 182, 1208–1214 (2000).

  126. 126.

    Curatti, L., Brown, C. S., Ludden, P. W. & Rubio, L. M. Genes required for rapid expression of nitrogenase activity in Azotobacter vinelandii. Proc. Natl Acad. Sci. USA 102, 6291–6296 (2005).

  127. 127.

    Kaminski, P. A. et al. Characterization of the fixABC region of Azorhizobium caulinodans ORS571 and identification of a new nitrogen fixation gene. Mol. Gen. Genet. 214, 496–502 (1988).

  128. 128.

    Wientjens, R. The Involvement of the fixABCX Genes and the Respiratory Chain in the Electron Transport to Nitrogenase in Azotobacter vinelandii. PhD thesis, Agricultural University (1993).

  129. 129.

    McDermott, J. E. et al. A model of cyclic transcriptomic behavior in the cyanobacterium Cyanothece sp. ATCC 51142. Mol. Biosyst. 7, 2407–2418 (2011).

  130. 130.

    Sevilla, M., Burris, R. H., Gunapala, N. & Kennedy, C. Comparison of benefit to sugarcane plant growth and 15N2 incorporation following inoculation of sterile plants with Acetobacter diazotrophicus wild-type and nif mutant strains. Mol. Plant Microbe Interact. 14, 358–366 (2001).

  131. 131.

    Eskin, N., Vessey, K. & Tian, L. Research progress and perspectives of nitrogen fixing bacterium, Gluconacetobacter diazotrophicus, in monocot plants. Int. J. Agron. (2014).

  132. 132.

    Lee, S., Reth, A., Meletzus, D., Sevilla, M. & Kennedy, C. Characterization of a major cluster of nif, fix, and associated genes in a sugarcane endophyte, Acetobacter diazotrophicus. J. Bacteriol. 182, 7088–7091 (2000).

  133. 133.

    Shanks, R. M. et al. Saccharomyces cerevisiae-based molecular tool kit for manipulation of genes from gram-negative bacteria. Appl. Environ. Microbiol. 72, 5027–5036 (2006).

  134. 134.

    Hakoyama, T. et al. Host plant genome overcomes the lack of a bacterial gene for symbiotic nitrogen fixation. Nature 462, 514–517 (2009).

  135. 135.

    Li, G.-W., Oh, E. & Weissman, J. S. The anti-Shine–Dalgarno sequence drives translational pausing and codon choice in bacteria. Nature 484, 538–541 (2012).

  136. 136.

    Lalanne, J.-B. et al. Evolutionary convergence of pathway-specific enzyme expression stoichiometry. Cell 173, 749–761 (2018).

  137. 137.

    Li, G.-W., Burkhardt, D., Gross, C. & Weissman, J. S. Quantifying absolute protein synthesis rates reveals principles underlying allocation of cellular resources. Cell 157, 624–635 (2014).

  138. 138.

    Poza-Carrión, C., Jiménez-Vicente, E., Navarro-Rodríguez, M., Echavarri-Erasun, C. & Rubio, L. M. Kinetics of nif gene expression in a nitrogen-fixing bacterium. J. Bacteriol. 196, 595–603 (2014).

  139. 139.

    Tezcan, F. A., Kaiser, J. T., Howard, J. B. & Rees, D. C. Structural evidence for asymmetrical nucleotide interactions in nitrogenase. J. Am. Chem. Soc. 137, 146–149 (2014).

  140. 140.

    Klugkist, J. & Haaker, H. Inhibition of nitrogenase activity by ammonium chloride in Azotobacter vinelandii. J. Bacteriol. 157, 148–151 (1984).

  141. 141.

    Song, M. et al. Control of type III protein secretion using a minimal genetic system. Nat. Comm. 8, 14737 (2017).

  142. 142.

    Guo, C.-J. et al. Discovery of reactive microbiota-derived metabolites that inhibit host proteases. Cell 168, 517–526 (2017).

  143. 143.

    Ren, H., Hu, P. & Zhao, H. A plug-and‐play pathway refactoring workflow for natural product research in Escherichia coli and Saccharomyces cerevisiae. Biotechnol. Bioeng. 114, 1847–1854 (2017).

  144. 144.

    MacLellan, S. R., MacLean, A. M. & Finan, T. M. Promoter prediction in the rhizobia. Microbiology 152, 1751–1763 (2006).

  145. 145.

    Ramírez-Romero, M. A., Masulis, I., Cevallos, M. A., González, V. & Davila, G. The Rhizobium etli σ70 (SigA) factor recognizes a lax consensus promoter. Nucleic Acids Res. 34, 1470–1480 (2006).

  146. 146.

    Becker, A. et al. Riboregulation in plant-associated α-proteobacteria. RNA Biol. 11, 550–562 (2014).

  147. 147.

    Robledo, M., Frage, B., Wright, P. R. & Becker, A. A stress-induced small RNA modulates alpha-rhizobial cell cycle progression. PLoS Genet. 11, e1005153 (2015).

  148. 148.

    Giacomini, A., Ollero, F. J., Squartini, A. & Nuti, M. P. Construction of multipurpose gene cartridges based on a novel synthetic promoter for high-level gene expression in Gram-negative bacteria. Gene 144, 17–24 (1994).

  149. 149.

    Khan, S. R., Gaines, J., Roop, R. M. & Farrand, S. K. Broad-host-range expression vectors with tightly regulated promoters and their use to examine the influence of TraR and TraM expression on Ti plasmid quorum sensing. Appl. Environ. Microbiol. 74, 5053–5062 (2008).

  150. 150.

    Tett, A. J. et al. Regulatable vectors for environmental gene expression in Alphaproteobacteria. Appl. Environ. Microbiol. 78, 7137–7140 (2012).

  151. 151.

    Mostafavi, M. et al. Analysis of a taurine-dependent promoter in Sinorhizobium meliloti that offers tight modulation of gene expression. BMC Microbiol. 14, 295 (2014).

  152. 152.

    Anderson, J. et al. BglBricks: a flexible standard for biological part assembly. J. Biol. Eng. 4, 1 (2010).

  153. 153.

    Farasat, I. et al. Efficient search, mapping, and optimization of multi-protein genetic systems in diverse bacteria. Mol. Sys. Biol. 10, 731 (2014).

  154. 154.

    Cambray, G. et al. Measurement and modeling of intrinsic transcription terminators. Nucleic Acids Res. 41, 5139–5148 (2013).

  155. 155.

    Chen, Y.-J. et al. Characterization of 582 natural and synthetic terminators and quantification of their design constraints. Nat. Methods 10, 659–664 (2013).

  156. 156.

    Temme, K., Hill, R., Segall-Shapiro, T. H., Moser, F. & Voigt, C. A. Modular control of multiple pathways using engineered orthogonal T7 polymerases. Nucleic Acids Res. 40, 8773–8781 (2012).

  157. 157.

    Price, M. N., Arkin, A. P. & Alm, E. The life-cycle of operons. PLoS Genet. 2, e96 (2006).

  158. 158.

    Touchon, M. & Rocha, E. P. Coevolution of the organization and structure of prokaryotic genomes. Cold Spring Harb. Persp. Biol. 8, a018168 (2016).

  159. 159.

    Price, M. N., Huang, K. H., Arkin, A. P. & Alm, E. Operon formation is driven by co-regulation and not by horizontal gene transfer. Genome Res. 15, 809–819 (2005).

  160. 160.

    Kaminski, P. A. & Elmerich, C. The control of Azorhizobium caulinodans nifA expression by oxygen, ammonia and by the HF‐I‐like protein, NrfA. Mol. Microbiol. 28, 603–613 (1998).

  161. 161.

    Pawlowski, K., Klosse, U. & De Bruijn, F. J. Characterization of a novel Azorhizobium caulinodans ORS571 two-component regulatory system, NtrY/NtrX, involved in nitrogen fixation and metabolism. Mol. Gen. Genet. 231, 124–138 (1991).

  162. 162.

    Kaminski, P. & Elmerich, C. Involvement of fixLJ in the regulation of nitrogen fixation in Azorhizobium caulinodans. Mol. Microbiol. 5, 665–673 (1991).

  163. 163.

    Kaminski, P., Mandon, K., Arigoni, F., Desnoues, N. & Elmerich, C. Regulation of nitrogen fixation in Azorhizobium caulinodans: identification of a fixK‐like gene, a positive regulator of nifA. Mol. Microbiol. 5, 1983–1991 (1991).

  164. 164.

    Michel-Reydellet, N. & Kaminski, P. A. J. Azorhizobium caulinodans PII and GlnK proteins control nitrogen fixation and ammonia assimilation. J. Bacteriol. 181, 2655–2658 (1999).

  165. 165.

    Drepper, T. et al. Role of GlnB and GlnK in ammonium control of both nitrogenase systems in the phototrophic bacterium Rhodobacter capsulatus. Microbiology 149, 2203–2212 (2003).

  166. 166.

    Martínez-García, E. & de Lorenzo, V. Molecular tools and emerging strategies for deep genetic/genomic refactoring of Pseudomonas. Curr. Opin. Biotechnol. 47, 120–132 (2017).

  167. 167.

    Calero, P., Jensen, S. I. & Nielsen, A. T. Broad-host-range ProUSER vectors enable fast characterization of inducible promoters and optimization of p-coumaric acid production in Pseudomonas putida KT2440. ACS Synth. Biol. 5, 741–753 (2016).

  168. 168.

    Choi, K.-H. & Schweizer, H. P. mini-Tn7 insertion in bacteria with single attTn7 sites: example Pseudomonas aeruginosa. Nat. Protoc. 1, 153–161 (2006).

  169. 169.

    Poole, R. K. & Hill, S. Respiratory protection of nitrogenase activity in Azotobacter vinelandii—roles of the terminal oxidases. Biosci. Rep. 17, 303–317 (1997).

  170. 170.

    Sabra, W., Zeng, A.-P., Lünsdorf, H. & Deckwer, W.-D. Effect of oxygen on formation and structure of Azotobacter vinelandii alginate and its role in protecting nitrogenase. Appl. Environ. Microbiol. 66, 4037–4044 (2000).

  171. 171.

    Schlesier, J., Rohde, M., Gerhardt, S. & Einsle, O. A conformational switch triggers nitrogenase protection from oxygen damage by Shethna protein II (FeSII). J. Am. Chem. Soc. 138, 239–247 (2015).

  172. 172.

    Ledbetter, R. N. et al. The electron bifurcating FixABCX protein complex from Azotobacter vinelandii: generation of low-potential reducing equivalents for nitrogenase catalysis. Biochemistry 56, 4177–4190 (2017).

  173. 173.

    Li, B. et al. Root exudates drive interspecific facilitation by enhancing nodulation and N2 fixation. Proc. Natl Acad. Sci. USA 113, 6496–6501 (2016).

  174. 174.

    Wang, Y., Zhang, J., Sun, Y., Feng, J. & Zhang, X. Evaluating the potential value of vatural product cuminic acid against plant pathogenic fungi in cucumber. Molecules 22, 1914 (2017).

  175. 175.

    Zhou, X. & Wu, F. Vanillic acid changed cucumber (Cucumis sativus L.) seedling rhizosphere total bacterial, Pseudomonas and Bacillus spp. communities. Sci. Rep. 8, 4929 (2018).

  176. 176.

    Sun, Y., Wang, Y., Han, L., Zhang, X. & Feng, J. Antifungal activity and action mode of cuminic acid from the seeds of Cuminum cyminum L. against Fusarium oxysporum f. sp. Niveum (FON) causing fusarium wilt on watermelon. Molecules 22, 2053 (2017).

  177. 177.

    Wang, Y., Sun, Y., Zhang, Y., Zhang, X. & Feng, J. Antifungal activity and biochemical response of cuminic acid against Phytophthora capsici Leonian. Molecules 21, 756 (2016).

  178. 178.

    Bais, H. P., Weir, T. L., Perry, L. G., Gilroy, S. & Vivanco, J. M. The role of root exudates in rhizosphere interactions with plants and other organisms. Annu. Rev. Plant Biol. 57, 233–266 (2006).

  179. 179.

    Rasmann, S. & Turlings, T. C. Root signals that mediate mutualistic interactions in the rhizosphere. Curr. Opin. Plant Biol. 32, 62–68 (2016).

  180. 180.

    Soto, M. J., Sanjuan, J. & Olivares, J. Rhizobia and plant-pathogenic bacteria: common infection weapons. Microbiology 152, 3167–3174 (2006).

  181. 181.

    Poonguzhali, S., Madhaiyan, M. & Sa, T. Quorum-sensing signals produced by plant-growth promoting Burkholderia strains under in vitro and in planta conditions. Res. Microbiol. 158, 287–294 (2007).

  182. 182.

    Sanchez-Contreras, M., Bauer, W. D., Gao, M., Robinson, J. B. & Allan Downie, J. Quorum-sensing regulation in rhizobia and its role in symbiotic interactions with legumes. Philos. Trans. R. Soc. B 362, 1149–1163 (2007).

  183. 183.

    Chagas, F. O., de Cassia Pessotti, R., Caraballo-Rodríguez, A. M. & Pupo, M. T. Chemical signaling involved in plant–microbe interactions. Chem. Soc. Rev. 47, 1652–1704 (2018).

  184. 184.

    Schikora, A., Schenk, S. T. & Hartmann, A. Beneficial effects of bacteria–plant communication based on quorum sensing molecules of the N-acyl homoserine lactone group. Plant Mol. Biol. 90, 605–612 (2016).

  185. 185.

    Westervelt, P., Bloom, M. L., Mabbott, G. A. & Fekete, F. A. The isolation and identification of 3, 4-dihydroxybenzoic acid formed by nitrogen-fixing Azomonas macrocytogenes. FEMS Microbiol. Lett. 30, 331–335 (1985).

  186. 186.

    Collinson, S. K., Doran, J. L. & Page, W. J. Production of 3, 4-dihydroxybenzoic acid by Azomonas macrocytogenes and Azotobacter paspali. Can. J. Microbiol. 33, 169–175 (1987).

  187. 187.

    Bhattacharya, A., Sood, P. & Citovsky, V. The roles of plant phenolics in defence and communication during Agrobacterium and Rhizobium infection. Mol. Plant Path. 11, 705–719 (2010).

  188. 188.

    Mathesius, U. et al. Extensive and specific responses of a eukaryote to bacterial quorum-sensing signals. Proc. Natl Acad. Sci. USA 100, 1444–1449 (2003).

  189. 189.

    Hernández-Reyes, C., Schenk, S. T., Neumann, C., Kogel, K. H. & Schikora, A. N-acyl-homoserine lactones-producing bacteria protect plants against plant and human pathogens. Microbial Biotechnol. 7, 580–588 (2014).

  190. 190.

    Pérez-Montaño, F. et al. Rice and bean AHL-mimic quorum-sensing signals specifically interfere with the capacity to form biofilms by plant-associated bacteria. Res. Microbiol. 164, 749–760 (2013).

  191. 191.

    Bressan, M. et al. Exogenous glucosinolate produced by Arabidopsis thaliana has an impact on microbes in the rhizosphere and plant roots. ISME J. 3, 1243–1257 (2009).

  192. 192.

    Badri, D. V. et al. An ABC transporter mutation alters root exudation of phytochemicals that provoke an overhaul of natural soil microbiota. Plant Physiol. 151, 2006–2017 (2009).

  193. 193.

    Abdel-Ghany, S. E., Day, I., Heuberger, A. L., Broeckling, C. D. & Reddy, A. S. Production of phloroglucinol, a platform chemical, in Arabidopsis using a bacterial gene. Sci. Rep. 6, 38483 (2016).

  194. 194.

    Oger, P., Petit, A. & Dessaux, Y. Genetically engineered plants producing opines alter their biological environment. Nat. Biotechnol. 15, 369 (1997).

  195. 195.

    Mondy, S. et al. An increasing opine carbon bias in artificial exudation systems and genetically modified plant rhizospheres leads to an increasing reshaping of bacterial populations. Mol. Ecol. 23, 4846–4861 (2014).

  196. 196.

    Meyer, A. J., Segall-Shapiro, T. H., Glassey, E., Zhang, J. & Voigt, C. A. Escherichia coli “Marionette” strains with 12 highly optimized small-molecule sensors. Nat. Chem. Biol. 15, 196–204 (2018).

  197. 197.

    Poole, P. S., Schofiel, N. A., Reid, C. J., Drew, E. M. & Walshaw, D. L. Identification of chromosomal genes located downstream of dctD that affect the requirement for calcium and the lipopolysaccharide layer of Rhizobium leguminosarum. Microbiology 140, 2797–2809 (1994).

  198. 198.

    Wilson, P. W. & Knight, S. G. Experiments in Bacterial Physiology (Burgess, 1952).

  199. 199.

    Udvardi, M. & Poole, P. S. Transport and metabolism in legume-rhizobia symbioses. Ann. Rev. Plant Biol. 64, 781–805 (2013).

  200. 200.

    Ferri, L., Gori, A., Biondi, E. G., Mengoni, A. & Bazzicalupo, M. Plasmid electroporation of Sinorhizobium strains: the role of the restriction gene hsdR in type strain Rm1021. Plasmid 63, 128–135 (2010).

  201. 201.

    Selbitschka, W. et al. Characterization of recA genes and recA mutants of Rhizobium meliloti and Rhizobium leguminosarum biovar viciae. Mol. Gen. Genet. 229, 86–95 (1991).

  202. 202.

    Prell, J., Boesten, B., Poole, P. & Priefer, U. B. The Rhizobium leguminosarum bv. viciae VF39 γ-aminobutyrate (GABA) aminotransferase gene (gabT) is induced by GABA and highly expressed in bacteroids. Microbiology 148, 615–623 (2002).

  203. 203.

    Silva-Rocha, R. et al. The Standard European Vector Architecture (SEVA): a coherent platform for the analysis and deployment of complex prokaryotic phenotypes. Nucleic Acids Res. 41, D666–D675 (2012).

  204. 204.

    Edgar, R. C. MUSCLE: multiple sequence alignment with high accuracy and high throughput. Nucleic Acids Res. 32, 1792–1797 (2004).

  205. 205.

    Yan, Y. et al. Global transcriptional analysis of nitrogen fixation and ammonium repression in root-associated Pseudomonas stutzeri A1501. BMC Genomics 11, 11 (2010).

  206. 206.

    Jacob, G., Schaefer, J., Garbow, J. & Stejskal, E. Solid-state NMR studies of Klebsiella pneumoniae grown under nitrogen-fixing conditions. J. Biol. Chem. 262, 254–259 (1987).

  207. 207.

    Blomberg, P., Wagner, E. & Nordström, K. Control of replication of plasmid R1: the duplex between the antisense RNA, CopA, and its target, CopT, is processed specifically in vivo and in vitro by RNase III. EMBO J. 9, 2331–2340 (1990).

  208. 208.

    Gorochowski, T. E. et al. Genetic circuit characterization and debugging using RNA‐seq. Mol. Syst. Biol. 13, 952 (2017).

Download references


This work was supported by the National Science Foundation (grant no. NSF-1331098), the Abdul Latif Jameel Water and Food Security Lab (J-WAFS) at the Massachusetts Institute of Technology, the US National Science Foundation Synthetic Biology Engineering Research Center (grant no. SynBERC EEC0540879) and the Office of Naval Research Multidisciplinary University Research Initiative (MURI grant no. N00014-13-1-0074). We thank G. O’Toole of Dartmouth College for the yeast shuttle vectors.

Author information

M.-H.R. and C.A.V. conceived the study and designed the experiments. J.Z. and M.-H.R. performed the RNA-seq and ribosome-profiling experiments and analysed the data. T.T. and M.-H.R. performed the opine experiments and analysed the data. D.K., J.-M.A., F.M. and J.W.P. performed the 15N-incorporation experiments and analysed the data. B.A.G. and P.S.P. performed the diazotrophic growth experiments and analysed the data. M.-H.R. performed all other experiments and analysed the data. M.-H.R. and C.A.V. wrote the manuscript with input from all of the authors.

Correspondence to Christopher A. Voigt.

Ethics declarations

Competing interests

M.-H.R. and C.A.V. have filed a patent application (US provisional application no. 62/820,765) on this work.

Additional information

Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Extended data

Extended Data Fig. 1 Nitrogenase activity in wild-type R. sp. IRBG74 and a Δnif mutant strain, in which the native nif clusters are deleted.

(a) The nif clusters in R. sp. IRBG74. The deleted regions generated by the suicide plasmids pMR45–46 are marked (Methods). (b) Transfer of native nif constructs into R. sp. IRBG74. Nitrogenase activity was detected only from the transfer of the R. sphaeroides nif cluster into R. sp. IRBG74 MR17 (ΔhsdR, recA) but not into R. sp. IRBG74 MR19 (ΔhsdR, recA Δnif). Expression of A. caulinodans nifV on the plasmid pMR49 in R. sp. IRBG74 MR17 was induced by 0.5 mM IPTG. The co-transfer of the complete A. caulinodans nif cluster on the two plasmids pMR19 and pMR20 did not yield activity in R. sp. IRBG74 MR17. Error bars represent standard deviation from three independent experiments on different days. Asterisk indicate ethylene production below the detection limit. Rsp, R. sphaeroides; Aca, A. caulinodans. (c) Plasmid maps used in (b).

Extended Data Fig. 2 Ribosome profiling data for the K. oxytoca nif cluster.

Ribosome profiling data for the K. oxytoca nif cluster in its native host (top) and when transferred into different strains are shown.

Extended Data Fig. 3 The effect of NifA overexpression on the nifH promoter activity in R. sp. IRBG74.

(a) The design of the reporter constructs used to measure nifH promoter activity shown. The nifH promoter activity was analysed in R. sp. IRBG74 using flow cytometry. Overexpression of R. sp. IRBG74 NifA increased the activity of the R. sp. IRBG74 nifH promoter but failed to complement or enhance the activities of the other nifH promoters including K. oxytoca, P. stutzeri and A. caulinodans. Error bars represent standard deviation from three independent experiments on different days. WT, wild-type; Rsp, R. sp. IRBG74; Kox, K. oxytoca M5al; Pst, P. stutzeri A1501; Aca, A. caulinodans ORS571 (b) Plasmid maps used to assess the effect of nifA overexpression in R. sp. IRBG74.

Extended Data Fig. 4 Nitrogenase activity when different inducible nif clusters are transferred to E. coli MG1655.

(a) The same universal controller system based on K. oxytoca nifA was optimized and used for all three clusters (Supplementary Figure 15, 17b). The controller plasmid pMR104 and genetic parts are provided in Supplementary Table 3 and 4. (b) The nif clusters from K. oxytoca, P. stutzeri, and A. vinelandii are shown. The deleted regions corresponding the NifLA regulators are marked, and their corresponding genomic locations are provided in Supplementary Table 3. The dotted lines indicate that multiple regions from the genome were cloned and combined to form the nif cluster. The clusters were carried on the plasmids pMR23–25 (Supplementary Table 3). (c) The induction of the nifH promoters from each species by the controller are shown (+, 50 μM IPTG). (d) The nitrogenase activities of the native cluster (intact nifLA) are compared to the inducible clusters in the presence and absence of 50 μM IPTG. The dashed lines indicate the activity of the native clusters in the wild-type context (top to bottom, K. oxytoca M5al, P. stutzeri A1501 and A. vinelandii DJ). (e) Regulation of nitrogenase activity by ammonium. Ammonium tolerance of nitrogenase from the native (black bar) and inducible (grey bar) systems was tested in the presence of 17.1 mM ammonium acetate and 50 μM IPTG (inducible). Asterisks indicate ethylene production below the detection limit. (f) Regulation of nitrogenase activity by oxygen. The native nif cluster is compared to the inducible version including the controller plasmid and 50 μM IPTG. Nitrogenase activities were measured after 3 h of incubation at constant oxygen concentrations (0 to 3%) in the headspace (Methods). Error bars represent standard deviation from three independent experiments on different days.

Extended Data Fig. 5 Transfer of the refactored nif cluster v3.2 in P. protegens Pf-5.

(a) Controllers whose output is T7 RNAP integrated on the genome of P. protegens Pf-5 are described. Substituted genetic parts for the controller optimization compared to the controller module pKT249 in E. coli MG1655 are highlighted in red. The response functions for the controllers with the reporter plasmid pMR81 was measured in the P. protegens Pf- 5 controller strain MR7. The controller driving the expression of GFP by the T7 promoter led to 96-fold induction by IPTG. (b) The genetic parts used to build the refactored v3.2 nif gene cluster are shown (provided in Supplementary Table 4). (c) The activity of the refactored nif cluster v3.2. Nitrogenase expression was induced by 1 mM IPTG. (d) The function of the transcriptional parts of the cluster v3.2 was analysed by RNA- seq (Supplementary Figure 18). The performance of the promoters (left) and terminators (right) was calculated (Methods). (e) The translation efficiency of the nif genes v3.2 as calculated using ribosome profiling and RNA-seq. Lines connect points that occur in the same operon. (f) The ribosome density (RD) is compared for the refactored v3.2 nif genes in P. protegens Pf-5 versus that measured for the nif genes from the native K. oxytoca cluster in K. oxytoca (→Klebsiella: R2 = 0.68). Error bars represent standard deviation from three independent experiments on different days.

Extended Data Fig. 6 Control of nitrogenase fixation in A. caulinodans ORS571 under changing environmental conditions.

(a) The effect of the absence or presence of 10 mM ammonium chloride is shown. The WT NifA from A. caulinodans ORS571 is compared to different combinations of amino acid substitutions. NifA/RpoN expression is induced by 1 mM IPTG (+) for A. caulinodans ΔnifA containing the controller plasmid pMR124–127 (+). An asterisk indicates ethylene production below the detection limit. (b) The nitrogenase activity is shown as a function of the oxygen concentration in the headspace (Methods). The native nif cluster (wild-type A. caulinodans ORS571, black) is compared to the inducible version (grey) including the controller plasmid and 1 mM IPTG. Error bars represent standard deviation from three independent experiments on different days.

Extended Data Fig. 7 Ammonium repression of the transferred nif clusters in E. coli MG1655 and P. protegens Pf-5.

Nitrogenase sensitivity to ammonium was measured by acetylene reduction assay in the absence (-) or presence (+) of 17.1 mM ammonium acetate. The sensitivity of the native and inducible nif clusters in E. coli MG1655 (a) and P. protegens Pf- 5 (b). Note that the data are from Fig. 4 and Supplementary Figure 8. (c) The specific nitrogenase activities of the native A. vinelandii nif cluster are compared to the inducible A. vinelandii cluster in the presence (+) and absence (-) of 17.1 mM ammonium acetate in P. protegens Pf-5. The nif clusters from the inducible version were induced by 50 μM and 0.5 mM IPTG in E. coli MG1655 and P. protegens Pf-5, respectively. Asterisks indicate ethylene production below the detection limit. Error bars represent standard deviation from three independent experiments on different days.

Extended Data Fig. 8 The effect of oxygen on the activity of the nifH promoters.

Expression from the nifH promoters was analysed in E. coli MG1655 containing the controller plasmid pMR104, P. protegens Pf-5 MR10 (for K. oxytoca) and MR9 (for P. stutzeri and A. vinelandii) at varying initial oxygen levels in the headspace. The three nifH promoters were induced with 0.05 mM IPTG and 0.5 mM IPTG in E. coli MG1655 and P. protegens Pf-5, respectively, and incubated at varying initial oxygen concentrations. Error bars represent standard deviation from three independent experiments on different days.

Extended Data Fig. 9 The effect of the rnf and fix complex on nitrogenase activity.

The modified nif clusters of A. vinelandii on the plasmids pMR25–28 were analysed in the controller strain P. protegens Pf-5 MR9. The deleted regions from the clusters were provided in Supplementary Table 3. Nitrogenase was induced with 0.5 mM IPTG. Dots in the DNA line indicate where multiple regions were cloned from genomic DNA and combined to form one large plasmid-borne nif cluster. An asterisk indicates ethylene production below the detection limit. Error bars represent standard deviation from three independent experiments on different days.

Extended Data Fig. 10 Regulation of nitrogenase activity in the E. coli MG1655 ‘Marionette’ strain.

(a) Controller plasmids used to drive expression of T7 promoters. (b) Inducibility of the T7 promoter by the controller plasmids encoding T7 RNAP under the regulation of the 12 sensors was tested with a reporter plasmid pMR123 (right). (c) Inducible control of nitrogenase activity in response to 12 inducers was tested with each of 12 controller plasmid and the plasmid pMR138 (right) carrying the refactored nif cluster v2.1 on pBBR1 origin. The choline-Cl inducible system was omitted for activity assay as the system was not inducible. For the DAPG-, DHBA-, and vanillic acid-inducible system, the refactored cluster v2.1 was carried on a lower copy number plasmid pMR31 (right) as there was no colony formation from the transformation of the plasmid pMR138. The inducer concentrations are: 400 μM arabinose, 1 mM choline-Cl, 500 nM 3OC14HSL, 50 μM cuminic acid, 25 nM 3OC6HSL, 25 μM DAPG, 500 μM DHBA, 1 mM IPTG, 100 nM aTc, 250 μM naringenin, 50 μM vanillic acid, and 250 μM salicylic acid. Plasmid and genetic parts are provided in Supplementary Table 3 and 4. Error bars represent standard deviation from three independent experiments on different days.

Supplementary information

Supplementary Information

Supplementary Figs. 1–22 and Supplementary Tables 1–6.

Reporting Summary

Supplementary Data 1

Plasmid pMR3_Native nif cluster of Klebsiella oxytoca M5al.

Supplementary Data 2

Plasmid pMR5_Native nif cluster of Pseudomonas stutzeri A1501.

Supplementary Data 3

Plasmid pMR7_Native nif cluster of Azotobacter vinelandii DJ.

Supplementary Data 4

Plasmid pMR9_Native nif cluster of Cyanothece ATCC51142.

Supplementary Data 5

Plasmid pMR11_Native nif cluster of Paenibacillus polymyxa WLY78.

Supplementary Data 6

Plasmid pMR13_Native nif cluster of Azospirillium brasilense Sp7.

Supplementary Data 7

Plasmid pMR15_Native nif cluster of Rhodobacter sphaeroides 2.4.1.

Supplementary Data 8

Plasmid pMR17_Native nif cluster of Rhodopseudomonas palustris CGA009.

Supplementary Data 9

Plasmid pMR19_Native nif cluster of Azorhizobium caulinodans ORS571 (Part 1 of 2).

Supplementary Data 10

Plasmid pMR20_Native nif cluster of Azorhizobium caulinodans ORS571 (Part 2 of 2).

Supplementary Data 11

Plasmid pMR21_Native nif cluster of Gluconacetobacter diazotrophicus PA1 5.

Supplementary Data 12

Plasmid pMR29_Refactored nif cluster v2.1.

Supplementary Data 13

Plasmid pMR38_Refactored nif cluster v3.2.

Rights and permissions

Reprints and Permissions

About this article

Verify currency and authenticity via CrossMark

Cite this article

Ryu, M., Zhang, J., Toth, T. et al. Control of nitrogen fixation in bacteria that associate with cereals. Nat Microbiol 5, 314–330 (2020).

Download citation

Further reading