Review Article | Published:

Diverse light responses of cyanobacteria mediated by phytochrome superfamily photoreceptors

Nature Reviews Microbiologyvolume 17pages3750 (2019) | Download Citation


Cyanobacteria are an evolutionarily and ecologically important group of prokaryotes. They exist in diverse habitats, ranging from hot springs and deserts to glaciers and the open ocean. The range of environments that they inhabit can be attributed in part to their ability to sense and respond to changing environmental conditions. As photosynthetic organisms, one of the most crucial parameters for cyanobacteria to monitor is light. Cyanobacteria can sense various wavelengths of light and many possess a range of bilin-binding photoreceptors belonging to the phytochrome superfamily. Vital cellular processes including growth, phototaxis, cell aggregation and photosynthesis are tuned to environmental light conditions by these photoreceptors. In this Review, we examine the physiological responses that are controlled by members of this diverse family of photoreceptors and discuss the signal transduction pathways through which these photoreceptors operate. We highlight specific examples where the activities of multiple photoreceptors function together to fine-tune light responses. We also discuss the potential application of these photosensing systems in optogenetics and synthetic biology.

Access optionsAccess options

Rent or Buy article

Get time limited or full article access on ReadCube.


All prices are NET prices.

Additional information

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


  1. 1.

    Whitton, B. A. & Potts, M. The Ecology of Cyanobacteria: Their Diversity in Time and Space (Kluwer Academic, 2000).

  2. 2.

    Graham, P. J., Nguyen, B., Burdyny, T. & Sinton, D. A penalty on photosynthetic growth in fluctuating light. Sci. Rep. 7, 12513 (2017).

  3. 3.

    Stomp, M., Huisman, J., Stal, L. J. & Matthijs, H. C. Colorful niches of phototrophic microorganisms shaped by vibrations of the water molecule. ISME J. 1, 271–282 (2007).

  4. 4.

    Allahverdiyeva, Y., Suorsa, M., Tikkanen, M. & Aro, E. M. Photoprotection of photosystems in fluctuating light intensities. J. Exp. Bot. 66, 2427–2436 (2015).

  5. 5.

    Gutu, A. & Kehoe, D. M. Emerging perspectives on the mechanisms, regulation, and distribution of light color acclimation in cyanobacteria. Mol. Plant 5, 1–13 (2011).

  6. 6.

    Ho, M. Y., Soulier, N. T., Canniffe, D. P., Shen, G. Z. & Bryant, D. A. Light regulation of pigment and photosystem biosynthesis in cyanobacteria. Curr. Opin. Plant Biol. 37, 24–33 (2017).

  7. 7.

    Sharma, N. K., Rai, A. K. & Stal, L. J. Cyanobacteria: An Economic Perspective (John Wiley & Sons Inc., 2014).

  8. 8.

    Abed, R. M. M., Dobretsov, S. & Sudesh, K. Applications of cyanobacteria in biotechnology. J. Appl. Microbiol. 106, 1–12 (2009).

  9. 9.

    Lau, N. S., Matsui, M. & Abdullah, A. A. Cyanobacteria: photoautotrophic microbial factories for the sustainable synthesis of industrial products. Biomed Res. Int. 2015, 754934 (2015).

  10. 10.

    Rodionova, M. V. et al. Biofuel production: challenges and opportunities. Intl J. Hydrog. Energy 42, 8450–8461 (2017).

  11. 11.

    Singh, R. et al. Uncovering potential applications of cyanobacteria and algal metabolites in biology, agriculture and medicine: current status and future prospects. Front. Microbiol. 8, 515 (2017).

  12. 12.

    Balasubramanian, R., Shen, G., Bryant, D. A. & Golbeck, J. H. Regulatory roles for IscA and SufA in iron homeostasis and redox stress responses in the cyanobacterium Synechococcus sp. strain PCC 7002. J. Bacteriol. 188, 3182–3191 (2006).

  13. 13.

    Quail, P. H. Phytochromes: photosensory perception and signal transduction. Science 268, 675–680 (1995).

  14. 14.

    Rockwell, N. C. & Lagarias, J. C. A brief history of phytochromes. Chem. Phys. Chem. 11, 1172–1180 (2010).

  15. 15.

    Rockwell, N. C. et al. Eukaryotic algal phytochromes span the visible spectrum. Proc. Natl Acad. Sci. USA 111, 3871–3876 (2014). The authors of this study demonstrate that algal phytochromes can sense light across a wider range of light colours — blue, green, orange, red and far-red — than has been shown for plant phytochromes.

  16. 16.

    Auldridge, M. E. & Forest, K. T. Bacterial phytochromes: more than meets the light. Crit. Rev. Biochem. Mol. Biol. 46, 67–88 (2011).

  17. 17.

    Idnurm, A., Verma, S. & Corrochano, L. M. A glimpse into the basis of vision in the kingdom Mycota. Fungal Genet. Biol. 47, 881–892 (2010).

  18. 18.

    Karniol, B., Wagner, J. R., Walker, J. M. & Vierstra, R. D. Phylogenetic analysis of the phytochrome superfamily reveals distinct microbial subfamilies of photoreceptors. Biochem. J. 392, 103–116 (2005).

  19. 19.

    Ikeuchi, M. & Ishizuka, T. Cyanobacteriochromes: a new superfamily of tetrapyrrole-binding photoreceptors in cyanobacteria. Photochem. Photobiol. Sci. 7, 1159–1167 (2008). This work is the first to define the cyanobacteriochromes as a new group within the phytochrome superfamily of photoreceptors.

  20. 20.

    Rockwell, N. C. & Lagarias, J. C. Phytochrome diversification in cyanobacteria and eukaryotic algae. Curr. Opin. Plant Biol. 37, 87–93 (2017). This Review summarizes some of the current viewpoints on the structural diversity, evolutionary origins and functions of phytochromes in plants, cyanobacteria and algae.

  21. 21.

    Wu, S. H. & Lagarias, J. C. Defining the bilin lyase domain: lessons from the extended phytochrome superfamily. Biochemistry 39, 13487–13495 (2000).

  22. 22.

    Campbell, E. L. et al. Genetic analysis reveals the identity of the photoreceptor for phototaxis in hormogonium filaments of Nostoc punctiforme. J. Bacteriol. 197, 782–791 (2015).

  23. 23.

    Mandalari, C., Losi, A. & Gartner, W. Distance-tree analysis, distribution and co-presence of bilin- and flavin-binding prokaryotic photoreceptors for visible light. Photochem. Photobiol. Sci. 12, 1144–1157 (2013).

  24. 24.

    Meeks, J. et al. An overview of the genome of Nostoc punctiforme, a multicellular, symbiotic cyanobacterium. Photosynth. Res. 70, 85–106 (2001).

  25. 25.

    Rockwell, N. C., Martin, S. S., Gulevich, A. G. & Lagarias, J. C. Phycoviolobilin formation and spectral tuning in the DXCF cyanobacteriochrome subfamily. Biochemistry 51, 1449–1463 (2012).

  26. 26.

    Rockwell, N. C., Martin, S. S. & Lagarias, J. C. Identification of cyanobacteriochromes detecting far-red light. Biochemistry 55, 3907–3919 (2016).

  27. 27.

    Yerrapragada, S. et al. Extreme sensory complexity encoded in the 10-megabase draft genome sequence of the chromatically acclimating cyanobacterium Tolypothrix sp. PCC 7601. Genome Announc. 3, e00355–15 (2015).

  28. 28.

    Butler, W. L., Norris, K. H., Siegelman, H. W. & Hendricks, S. B. Detection, assay, and preliminary purification of the pigment controlling photoresponsive development of plants. Proc. Natl Acad. Sci. USA 45, 1703–1708 (1959).

  29. 29.

    Wagner, J. R., Brunzelle, J. S., Forest, K. T. & Vierstra, R. D. A light-sensing knot revealed by the structure of the chromophore-binding domain of phytochrome. Nature 438, 325–331 (2005). This article provides the first insights into the 3D features of phytochrome photosensory regions.

  30. 30.

    Burgie, E. S., Bussell, A. N., Walker, J. M., Dubiel, K. & Vierstra, R. D. Crystal structure of the photosensing module from a red/far-red light-absorbing plant phytochrome. Proc. Natl Acad. Sci. USA 111, 10179–10184 (2014).

  31. 31.

    Thümmler, F., Algorra, P. & Fobo, G. M. Sequence similarities of phytochrome to protein kinases: Implication for the structure, function and evolution of the phytochrome gene family. FEBS Lett. 357, 149–155 (1995).

  32. 32.

    Yeh, K. C., Wu, S.-H., Murphy, J. T. & Lagarias, J. C. A cyanobacterial phytochrome two-component light sensory system. Science 277, 1505–1508 (1997). This manuscript is the first to demonstrate light colour-driven phosphorylation and phosphotransfer activity of a prokaryotic member of the phytochrome superfamily.

  33. 33.

    Buchberger, T. & Lamparter, T. Streptophyte phytochromes exhibit an N-terminus of cyanobacterial origin and a C-terminus of proteobacterial origin. BMC Res. Notes 8, 1–13 (2015).

  34. 34.

    Stock, A. M., Robinson, V. L. & Goudreau, P. N. Two-component signal transduction. Annu. Rev. Biochem. 69, 183–215 (2000).

  35. 35.

    Rockwell, N. C., Su, Y. S. & Lagarias, J. C. Phytochrome structure and signaling mechanisms. Annu. Rev. Plant Biol. 57, 837–858 (2006).

  36. 36.

    Burgie, E. S. & Vierstra, R. D. Phytochromes: an atomic perspective on photoactivation and signaling. Plant Cell 26, 4568–4583 (2014).

  37. 37.

    Anders, K. & Essen, L. O. The family of phytochrome-like photoreceptors: diverse, complex and multi-colored, but very useful. Curr. Opin. Struct. Biol. 35, 7–16 (2015).

  38. 38.

    Anders, K., Daminelli-Widany, G., Mroginski, M. A., von Stetten, D. & Essen, L. O. Structure of the cyanobacterial phytochrome 2 photosensor implies a tryptophan switch for phytochrome signaling. J. Biol. Chem. 288, 35714–35725 (2013).

  39. 39.

    Rockwell, N. C., Martin, S. S., Feoktistova, K. & Lagarias, J. C. Diverse two-cysteine photocycles in phytochromes and cyanobacteriochromes. Proc. Natl Acad. Sci. USA 108, 11854–11859 (2011).

  40. 40.

    Rockwell, N. C., Martin, S. S. & Lagarias, J. C. Red/green cyanobacteriochromes: sensors of color and power. Biochemistry 51, 9667–9677 (2012).

  41. 41.

    Bussell, A. N. & Kehoe, D. M. Control of a four-color sensing photoreceptor by a two-color sensing photoreceptor reveals complex light regulation in cyanobacteria. Proc. Natl Acad. Sci. USA 110, 12834–12839 (2013).

  42. 42.

    Savakis, P. et al. Light-induced alteration of c-di-GMP level controls motility of Synechocystis sp PCC 6803. Mol. Microbiol. 85, 239–251 (2012). This is the first study to show that the second messenger c-di-GMP is directly involved in light-regulated phototaxis in cyanobacteria.

  43. 43.

    Song, J. Y. et al. Near-UV cyanobacteriochrome signaling system elicits negative phototaxis in the cyanobacterium Synechocystis sp PCC 6803. Proc. Natl Acad. Sci. USA 108, 10780–10785 (2011).

  44. 44.

    Lacey, R. F. & Binder, B. M. Ethylene regulates the physiology of the cyanobacterium Synechocystis sp PCC 6803 via an ethylene receptor. Plant Physiol. 171, 2798–2809 (2016).

  45. 45.

    Golden, S. S., Brusslan, J. & Haselkorn, R. Genetic engineering of the cyanobacterial chromosome. Methods Enzymol. 153, 215–231 (1987).

  46. 46.

    Bryant, D. A., de Lorimier, R., Guglielmi, G. & Stevens, S. E. J. Structural and compositional analyses of the phycobilisomes of Synechococcus sp. PCC 7002. Analyses of the wild-type strain and a phycocyanin-less mutant constructed by interposon mutagenesis. Arch. Microbiol. 153, 550–560 (1990).

  47. 47.

    Cai, Y. P. & Wolk, C. P. Use of a conditionally lethal gene in Anabaena sp. strain PCC 7120 to select for double recombinants and to entrap insertion sequences. J. Bacteriol. 172, 3138–3145 (1990). This paper provides important technical approaches that allowed the subsequent development of molecular genetic systems for many cyanobacteria.

  48. 48.

    Vanderplas, J. et al. Genomic integration system based on pBR322 sequences for the cyanobacterium Synechococcus sp PCC7942 - transfer of genes encoding plastocyanin and ferredoxin. Gene 95, 39–48 (1990).

  49. 49.

    MacColl, R. Cyanobacterial Phycobilisomes. J. Struct. Biol. 124, 311–334 (1998).

  50. 50.

    Adir, N. Elucidation of the molecular structures of components of the phycobilisome: reconstructing a giant. Photosynth. Res. 85, 15–32 (2005).

  51. 51.

    Watanabe, M. & Ikeuchi, M. Phycobilisome: architecture of a light-harvesting supercomplex. Photosynth. Res. 116, 265–276 (2013).

  52. 52.

    Stomp, M. et al. The timescale of phenotypic plasticity and its impact on competition in fluctuating environments. Am. Nat. 172, E169–E185 (2008).

  53. 53.

    Bogorad, L. Phycobiliproteins and complementary chromatic adaptation. Annu. Rev. Plant Physiol. Plant Mol. Biol. 26, 369–401 (1975).

  54. 54.

    Kehoe, D. M. & Gutu, A. Responding to color: the regulation of complementary chromatic adaptation. Annu. Rev. Plant Biol. 57, 127–150 (2006).

  55. 55.

    Tandeau de Marsac, N. Occurrence and nature of chromatic adaptation in cyanobacteria. J. Bacteriol. 130, 82–91 (1977).

  56. 56.

    Palenik, B. Chromatic adaptation in marine Synechococcus strains. Appl. Environ. Microbiol. 67, 991–994 (2001).

  57. 57.

    Gan, F. et al. Extensive remodeling of a cyanobacterial photosynthetic apparatus in far-red light. Science 345, 1312–1317 (2014). This study is the first description of FaRLiP, a type of red and/or far-red chromatic acclimation in cyanobacteria that drastically affects the photosynthetic machinery.

  58. 58.

    Gaidukov, N. Die farbervonderung bei den prozessen der komplementoren chromatischen adaptation. Ber. Dtsch. Bot. Ges. 21, 517–522 (1903).

  59. 59.

    Grossman, A. R. A molecular understanding of complementary chromatic adaptation. Photosynth. Res. 76, 207–215 (2003).

  60. 60.

    Oelmuller, R., Grossman, A. R. & Briggs, W. R. Photoreversibility of the effect of red and green light-pulses on the accumulation in darkness of messenger-RNAs coding for phycocyanin and phycoerythrin in Fremyella diplosiphon. Plant Physiol. 88, 1084–1091 (1988).

  61. 61.

    Kehoe, D. M. & Grossman, A. R. Similarity of a chromatic adaptation sensor to phytochrome and ethylene receptors. Science 273, 1409–1412 (1996). This paper provides the first molecular evidence for the existence of phytochrome superfamily members in prokaryotes.

  62. 62.

    Terauchi, K., Montgomery, B. L., Grossman, A. R., Lagarias, J. C. & Kehoe, D. M. RcaE is a complementary chromatic adaptation photoreceptor required for green and red light responsiveness. Mol. Microbiol. 51, 567–577 (2004).

  63. 63.

    Wiltbank, L. B. & Kehoe, D. M. Two cyanobacterial photoreceptors regulate photosynthetic light harvesting by sensing teal, green, yellow and red light. mBio 7, e02130–15 (2016).

  64. 64.

    Hirose, Y. et al. Green/red cyanobacteriochromes regulate complementary chromatic acclimation via a protochromic photocycle. Proc. Natl Acad. Sci. USA 110, 4974–4979 (2013).

  65. 65.

    Kehoe, D. M. & Grossman, A. R. New classes of mutants in complementary chromatic adaptation provide evidence for a novel four-step phosphorelay system. J. Bacteriol. 179, 3914–3921 (1997).

  66. 66.

    Chiang, G. G., Schaefer, M. R. & Grossman, A. R. Complementation of a red-light-indifferent cyanobacterial mutant. Proc. Natl Acad. Sci. USA 89, 9415–9419 (1992).

  67. 67.

    Li, L. & Kehoe, D. M. In vivo analysis of the roles of conserved aspartate and histidine residues within a complex response regulator. Mol. Microbiol. 55, 1538–1552 (2005).

  68. 68.

    Li, L., Alvey, R. M., Bezy, R. P. & Kehoe, D. M. Inverse transcriptional activities during complementary chromatic adaptation are controlled by the response regulator RcaC binding to red and green light-responsive promoters. Mol. Microbiol. 68, 286–297 (2008).

  69. 69.

    Bezy, R. P., Wiltbank, L. & Kehoe, D. M. Light-dependent attenuation of phycoerythrin gene expression reveals convergent evolution of green light sensing in cyanobacteria. Proc. Natl Acad. Sci. USA 108, 18542–18547 (2011).

  70. 70.

    Gutu, A., Nesbit, A. D., Alverson, A. J., Palmer, J. D. & Kehoe, D. M. Unique role for translation initiation factor 3 in the light color regulation of photosynthetic gene expression. Proc. Natl Acad. Sci. USA 110, 16253–16258 (2013).

  71. 71.

    Bennett, A. & Bogorad, L. Complementary chromatic adaptation in a filamentous blue-green alga. J. Cell Biol. 58, 419–435 (1973).

  72. 72.

    Bordowitz, J. R. & Montgomery, B. L. Photoregulation of cellular morphology during complementary chromatic adaptation requires sensor-kinase-class protein RcaE in Fremyella diplosiphon. J. Bacteriol. 190, 4069–4074 (2008).

  73. 73.

    Singh, S. P. & Montgomery, B. L. Morphogenes bolA and mreB mediate the photoregulation of cellular morphology during complementary chromatic acclimation in Fremyella diplosiphon. Mol. Microbiol. 93, 167–182 (2014).

  74. 74.

    Singh, S. P. & Montgomery, B. L. Regulation of BolA abundance mediates morphogenesis in Fremyella diplosiphon. Front. Microbiol. 6, 1215 (2015).

  75. 75.

    Pattanaik, B., Busch, A. W. U., Flu, P. S., Chen, J. & Montgomery, B. L. Responses to iron limitation are impacted by light quality and regulated by RcaE in the chromatically acclimating cyanobacterium Fremyella diplosiphon. Microbiology 160, 992–1005 (2014).

  76. 76.

    Singh, S. P. & Montgomery, B. L. Reactive oxygen species are involved in the morphology-determining mechanism of Fremyella diplosiphon cells during complementary chromatic adaptation. Microbiology 158, 2235–2245 (2012).

  77. 77.

    Montgomery, B. L., Lechno-Yossef, S. & Kerfeld, C. A. Interrelated modules in cyanobacterial photosynthesis: the carbon-concentrating mechanism, photorespiration, and light perception. J. Exp. Bot. 67, 2931–2940 (2016).

  78. 78.

    Rohnke, B. A., Singh, S. P., Pattanaik, B. & Montgomery, B. L. RcaE-dependent regulation of carboxysome structural proteins has a central role in environmental determination of carboxysome morphology and abundance in Fremyella diplosiphon. mSphere 3, e00617–17 (2018).

  79. 79.

    Hirose, Y., Shimada, T., Narikawa, R., Katayama, M. & Ikeuchi, M. Cyanobacteriochrome CcaS is the green light receptor that induces the expression of phycobilisome linker protein. Proc. Nat. Acad. Sci. USA 105, 9528–9533 (2008).

  80. 80.

    Tandeau de Marsac, N. & Cohen-Bazire, G. Molecular composition of cyanobacterial phycobilisomes. Proc. Natl Acad. Sci. USA 74, 1635–1639 (1977).

  81. 81.

    Kondo, K., Geng, X. X., Katayama, M. & Ikeuchi, M. Distinct roles of CpcG1 and CpcG2 in phycobilisome assembly in the cyanobacterium Synechocystis sp PCC 6803. Photosynth. Res. 84, 269–273 (2005).

  82. 82.

    Watanabe, M. et al. Attachment of phycobilisomes in an antenna-photosystem I supercomplex of cyanobacteria. Proc. Natl Acad. Sci. USA 111, 2512–2517 (2014).

  83. 83.

    Abe, K. et al. Engineering of a green-light inducible gene expression system in Synechocystis sp PCC6803. Microb. Biotechnol. 7, 177–183 (2014).

  84. 84.

    Hirose, Y., Narikawa, R., Katyama, M. & Ikeuchi, M. Cyanobacteriochrome CcaS regulates phycoerythrin accumulation in Nostoc punctiforme, a group II chromatic adaptor. Proc. Natl Acad. Sci. USA 107, 8854–8859 (2010).

  85. 85.

    Brown, I. I. et al. Polyphasic characterization of a thermotolerant siderophilic filamentous cyanobacterium that produces intracellular iron deposits. Appl. Environ. Microbiol. 76, 6664–6672 (2010).

  86. 86.

    Zhao, C., Gan, F., Shen, G. Z. & Bryant, D. A. RfpA, RfpB, and RfpC are the master control elements of far-red light photoacclimation (FaRLiP). Front. Microbiol. 6, 1303 (2015).

  87. 87.

    Gan, F., Shen, G. & Bryant, D. A. Occurrence of far-red light photoacclimation (FaRLiP) in diverse cyanobacteria. Life 5, 4–24 (2015).

  88. 88.

    Ho, M. Y., Gan, F., Shen, G. Z. & Bryant, D. A. Far-red light photoacclimation (FaRLiP) in Synechococcus sp PCC 7335. II. Characterization of phycobiliproteins produced during acclimation to far-red light. Photosynth. Res. 131, 187–202 (2017).

  89. 89.

    Ho, M. Y., Gan, F., Shen, G. Z., Zhao, C. & Bryant, D. A. Far-red light photoacclimation (FaRLiP) in Synechococcus sp PCC 7335: I. Regulation of FaRLiP gene expression. Photosynth. Res. 131, 173–186 (2017).

  90. 90.

    Wilde, A. & Mullineaux, C. W. Light-controlled motility in prokaryotes and the problem of directional light perception. FEMS Microbiol. Rev. 41, 900–922 (2017).

  91. 91.

    Bhaya, D., Bianco, N. R., Bryant, D. & Grossman, A. Type IV pilus biogenesis and motility in the cyanobacterium Synechocystis sp. PCC6803. Mol. Microbiol. 37, 941–951 (2000). This work gives important mechanistic insights into the role of type IV pili in the motility of the cyanobacterium Synechocystis sp. PCC 6803.

  92. 92.

    Choi, J. S. et al. Photomovement of the gliding cyanobacterium Synechocystis sp. PCC 6803. Photochem. Photobiol. 70, 95–102 (1999).

  93. 93.

    Ng, W. O., Grossman, A. R. & Bhaya, D. Multiple light inputs control phototaxis in Synechocystis sp. strain PCC6803. J. Bacteriol. 185, 1599–1607 (2003).

  94. 94.

    Yoshihara, S., Suzuki, F., Fujita, H., Geng, X. X. & Ikeuchi, M. Novel putative photoreceptor and regulatory genes required for the positive phototactic movement of the unicellular motile cyanobacterium Synechocystis sp. PCC 6803. Plant Cell Physiol. 41, 1299–1304 (2000).

  95. 95.

    Fiedler, B., Borner, T. & Wilde, A. Phototaxis in the cyanobacterium Synechocystis sp. PCC 6803: role of different photoreceptors. Photochem. Photobiol. 81, 1481–1488 (2005).

  96. 96.

    Chau, R. M. W., Bhaya, D. & Huang, K. C. Emergent phototactic responses of cyanobacteria under complex light regimes. mBio 8, e02330–16 (2017).

  97. 97.

    Moon, Y. J. et al. The role of cyanopterin in UV/blue light signal transduction of cyanobacterium Synechocystis sp PCC 6803 phototaxis. Plant Cell Physiol. 51, 969–980 (2010).

  98. 98.

    Masuda, S. & Ono, T. Biochemical characterization of the major adenylyl cyclase, Cya1, in the cyanobacterium Synechocystis sp PCC 6803. FEBS Lett. 577, 255–258 (2004).

  99. 99.

    Okajima, K. et al. Biochemical and functional characterization of BLUF-type flavin-binding proteins of two species of cyanobacteria. J. Biochem. 137, 741–750 (2005).

  100. 100.

    Wilde, A., Fiedler, B. & Borner, T. The cyanobacterial phytochrome Cph2 inhibits phototaxis towards blue light. Mol. Microbiol. 44, 981–988 (2002).

  101. 101.

    Moon, Y. J. et al. Cyanobacterial phytochrome Cph2 is a negative regulator in phototaxis toward UV-A. FEBS Lett. 585, 335–340 (2011).

  102. 102.

    Park, C. M. et al. A second photochromic bacteriophytochrome from Synechocystis sp PCC 6803: spectral analysis and down-regulation by light. Biochemistry 39, 10840–10847 (2000).

  103. 103.

    Schwarzkopf, M., Yoo, Y. C., Hueckelhoven, R., Park, Y. M. & Proels, R. K. Cyanobacterial phytochrome2 regulates the heterotrophic metabolism and has a function in the heat and high-light stress response. Plant Physiol. 164, 2157–2166 (2014).

  104. 104.

    Romling, U., Galperin, M. Y. & Gomelsky, M. Cyclic di-GMP: the first 25 years of a universal bacterial second messenger. Microbiol. Mol. Biol. Rev. 77, 1–52 (2013).

  105. 105.

    Sato, S. et al. A large-scale protein-protein interaction analysis in Synechocystis sp PCC6803. DNA Res. 14, 207–216 (2007).

  106. 106.

    Angerer, V. et al. The protein Slr1143 is an active diguanylate cyclase in Synechocystis sp PCC 6803 and interacts with the photoreceptor Cph2. Microbiology 163, 920–930 (2017).

  107. 107.

    Dahlstrom, K. M. & O’Toole, G. A. Symphony of cyclases: specificity in diguanylate cyclase signaling. Annu. Rev Microbiol. 71, 179–195 (2017).

  108. 108.

    Bhaya, D., Takahashi, A. & Grossman, A. R. Light regulation of type IV pilus-dependent motility by chemosensor-like elements in Synechocystis PCC6803. Proc. Natl Acad. Sci. USA 98, 7540–7545 (2001).

  109. 109.

    Yoshihara, S., Katayama, M., Geng, X. & Ikeuchi, M. Cyanobacterial phytochrome-like PixJ1 holoprotein shows novel reversible photoconversion between blue- and green-absorbing forms. Plant Cell Physiol. 45, 1729–1737 (2004).

  110. 110.

    Yoshihara, S. & Ikeuchi, M. Phototactic motility in the unicellular cyanobacterium Synechocystis sp. PCC 6803. Photochem. Photobiol. Sci. 3, 512–518 (2004).

  111. 111.

    Chau, R. M. W., Ursell, T., Wang, S., Huang, K. C. & Bhaya, D. Maintenance of motility bias during cyanobacterial phototaxis. Biophys. J. 108, 1623–1632 (2015).

  112. 112.

    Narikawa, R. et al. Novel photosensory two-component system (PixA-NixB-NixC) involved in the regulation of positive and negative phototaxis of cyanobacterium Synechocystis sp PCC 6803. Plant Cell Physiol. 52, 2214–2224 (2011).

  113. 113.

    Ulijasz, A. T. et al. Cyanochromes are blue/green light photoreversible photoreceptors defined by a stable double cysteine linkage to a phycoviolobilin-type chromophore. J. Biol. Chem. 284, 29757–29772 (2009).

  114. 114.

    Makarova, K. S., Koonin, E. V., Haselkorn, R. & Galperin, M. Y. Cyanobacterial response regulator PatA contains a conserved N-terminal domain (PATAN) with an alpha-helical insertion. Bioinformatics 22, 1297–1301 (2006).

  115. 115.

    Schuergers, N. et al. Cyanobacteria use micro-optics to sense light direction. eLife 5, e12620 (2016). This is a fascinating study that presents the first evidence that cyanobacteria can act as microlenses, allowing them to sense the direction of a light source.

  116. 116.

    Hughes, J. et al. A prokaryotic phytochrome. Nature 386, 663 (1997).

  117. 117.

    Fiedler, B. et al. Involvement of cyanobacterial phytochromes in growth under different light qualities and quantities. Photochem. Photobiol. 79, 551–555 (2004).

  118. 118.

    Hubschmann, T., Yamamoto, H., Gieler, T., Murata, N. & Borner, T. Red and far-red light alter the transcript profile in the cyanobacterium Synechocystis sp PCC 6803: impact of cyanobacterial phytochromes. FEBS Lett. 579, 1613–1618 (2005).

  119. 119.

    Wilde, A., Churin, Y., Schubert, H. & Borner, T. Disruption of a Synechocystis sp. PCC 6803 gene with partial similarity to phytochrome genes alters growth under changing light qualities. FEBS Lett. 406, 89–92 (1997).

  120. 120.

    Anderson, S. L. & McIntosh, L. Light-activated heterotrophic growth of the cyanobacterium Synechocystis sp. strain PCC 6803: a blue-light-requiring process. J. Bacteriol. 173, 2761–2767 (1991).

  121. 121.

    Zilliges, Y. et al. An extracellular glycoprotein is implicated in cell-cell contacts in the toxic cyanobacterium Microcystis aeruginosa PCC 7806. J. Bacteriol. 190, 2871–2879 (2008).

  122. 122.

    Kawano, Y. et al. Cellulose accumulation and a cellulose synthase gene are responsible for cell aggregation in the cyanobacterium Thermosynechococcus vulcanus RKN. Plant Cell Physiol. 52, 957–966 (2011).

  123. 123.

    Enomoto, G. et al. Cyanobacteriochrome SesA Is a diguanylate cyclase that induces cell aggregation in Thermosynechococcus. J. Biol. Chem. 289, 24801–24809 (2014).

  124. 124.

    Enomoto, G., Win, N. N., Narikawa, R. R. & Ikeuchi, M. Three cyanobacteriochromes work together to form a light color-sensitive input system for c-di-GMP signaling of cell aggregation. Proc. Natl Acad. Sci. USA 112, 8082–8087 (2015). This study presents a model describing how three photoreceptors integrate multiple light inputs to regulate the process of cell aggregation.

  125. 125.

    Maeda, K. et al. Genetic identification of factors for extracellular cellulose accumulation in the thermophilic cyanobacterium Thermosynechococcus vulcanus: proposal of a novel tripartite secretion system. Mol. Microbiol. (2018).

  126. 126.

    Enomoto, G., Okuda, Y. & Ikeuchi, M. Tlr1612 is the major repressor of cell aggregation in the light-color-dependent c-di-GMP signaling network of Thermosynechococcus vulcanus. Sci. Rep. 28, 5338 (2018).

  127. 127.

    Yu, J. J. et al. Synechococcus elongatus UTEX 2973, a fast growing cyanobacterial chassis for biosynthesis using light and CO2. Sci. Rep. 5, 8132 (2015).

  128. 128.

    Ungerer, J. & Pakrasi, H. B. Cpf1is a versatile tool for CRISPR genome editing across diverse species of cyanobacteria. Sci. Rep. 6, 39681 (2016).

  129. 129.

    Shcherbakova, D. M., Shemetov, A. A., Kaberniuk, A. A. & Verkhusha, V. V. Natural photoreceptors as a source of fluorescent proteins, biosensors, and optogenetic tools. Annu. Rev. Biochem. 84, 519–550 (2015).

  130. 130.

    Chernov, K. G., Redchuk, T. A., Omelina, E. S. & Verkhushaa, V. V. Near-infrared fluorescent proteins, biosensors, and optogenetic tools engineered from phytochromes. Chem. Rev. 117, 6423–6446 (2017).

  131. 131.

    Boyden, E. S., Zhang, F., Bamberg, E., Nagel, G. & Deisseroth, K. Millisecond-timescale, genetically targeted optical control of neural activity. Nat. Neurosci. 8, 1263–1268 (2005).

  132. 132.

    Levskaya, A. et al. Engineering Escherichia coli to see light — these smart bacteria ‘photograph’ a light pattern as a high-definition chemical image. Nature 438, 441–442 (2005).

  133. 133.

    Gambetta, G. A. & Lagarias, J. C. Genetic engineering of phytochrome biosynthesis in bacteria. Proc. Natl Acad. Sci. USA 98, 10566–10571 (2001).

  134. 134.

    Mukougawa, K., Kanamoto, H., Kobayashi, T., Yokota, A. & Kohchi, T. Metabolic engineering to produce phytochromes with phytochromobilin, phycocyanobilin, or phycoerythrobilin chromophore in Escherichia coli. FEBS Lett. 580, 1333–1338 (2006).

  135. 135.

    Huang, H. H., Camsund, D., Lindblad, P. & Heidorn, T. Design and characterization of molecular tools for a synthetic biology approach towards developing cyanobacterial biotechnology. Nucleic Acids Res. 38, 2577–2593 (2010).

  136. 136.

    Taton, A. et al. Broad-host-range vector system for synthetic biology and biotechnology in cyanobacteria. Nucleic Acids Res. 42, 16 (2014).

  137. 137.

    Zess, E. K., Begemann, M. B. & Pfleger, B. F. Construction of new synthetic biology tools for the control of gene expression in the cyanobacterium Synechococcus sp strain PCC 7002. Biotechnol. Bioeng. 113, 424–432 (2016).

  138. 138.

    Gordon, G. C. et al. CRISPR interference as a titratable. trans-acting regulatory tool for metabolic engineering in the cyanobacterium Synechococcus sp. strain PCC 7002. Metab. Eng. 38, 170–179 (2016).

  139. 139.

    Ruffing, A. M., Jensen, T. J. & Strickland, L. M. Genetic tools for advancement of Synechococcus sp PCC 7002 as a cyanobacterial chassis. Microb. Cell Fact. 15, 190 (2016).

  140. 140.

    Kim, W. J., Lee, S. M., Um, Y., Sim, S. J. & Woo, H. M. Development of SyneBrick vectors as a synthetic biology platform for gene expression in Synechococcus elongatus PCC 7942. Front. Plant Sci. 8, 293 (2017).

  141. 141.

    Wang, B., Eckert, C., Maness, P. C. & Yu, J. P. A genetic toolbox for modulating the expression of heterologous genes in the cyanobacterium Synechocystis sp PCC 6803. ACS Synth. Biol. 7, 276–286 (2018).

  142. 142.

    Immethun, C. M. et al. Physical, chemical, and metabolic state sensors expand the synthetic biology toolbox for Synechocystis sp PCC 6803. Biotechnol. Bioeng. 114, 1561–1569 (2017).

  143. 143.

    Hirose, Y. et al. Complete genome sequence of cyanobacterium Fischerella sp NIES-3754, providing thermoresistant optogenetic tools. J. Biotechnol. 220, 45–46 (2016).

  144. 144.

    Wang, H. & Yang, Y. T. Mini photobioreactors for in vivo real-time characterization and evolutionary tuning of bacterial optogenetic circuit. ACS Synth. Biol. 6, 1793–1796 (2017).

  145. 145.

    Milias-Argeitis, A., Rullan, M., Aoki, S. K., Buchmann, P. & Khammash, M. Automated optogenetic feedback control for precise and robust regulation of gene expression and cell growth. Nat. Commun. 7, 12546 (2016).

  146. 146.

    Tabor, J. J. et al. A synthetic genetic edge detection program. Cell 137, 1272–1281 (2009). This study provides the first example of engineered bacterial cells capable of integrating light and chemical information to detect and respond to edges of light–dark boundaries.

  147. 147.

    Tabor, J. J., Levskaya, A. & Voigt, C. A. Multichromatic control of gene expression in Escherichia coli. J. Mol. Biol. 405, 315–324 (2011).

  148. 148.

    Schmidl, S. R., Sheth, R. U., Wu, A. & Tabor, J. J. Refactoring and optimization of light-switchable Escherichia coli two-component systems. ACS Synth. Biol. 3, 820–831 (2014).

  149. 149.

    Olson, E. J., Tzouanas, C. N. & Tabor, J. J. A photoconversion model for full spectral programming and multiplexing of optogenetic systems. Mol. Syst. Biol. 13, 926 (2017).

  150. 150.

    Fernandez-Rodriguez, J., Moser, F., Song, M. & Voigt, C. A. Engineering RGB color vision into Escherichia coli. Nat. Chem. Biol. 13, 706 (2017).

  151. 151.

    Wu, L., Luo, S. W., Ma, S. Y., Liang, Z. & Wu, J. R. Construction of light-sensing two-component systems in Escherichia coli. Sci. Bull. 62, 813–815 (2017).

  152. 152.

    Ma, S. Y., Luo, S. W., Wu, L., Liang, Z. & Wu, J. R. Re-engineering the two-component systems as light-regulated in Escherichia coli. J. Biosci. 42, 565–573 (2017).

  153. 153.

    Badary, A., Abe, K., Ferri, S., Kojima, K. & Sode, K. The development and characterization of an exogenous green-light-regulated gene expression system in marine cyanobacteria. Mar. Biotechnol. 17, 245–251 (2015).

  154. 154.

    Sugie, Y., Hori, M., Oka, S., Ohtsuka, H. & Aiba, H. Reconstruction of a chromatic response system in Escherichia coli. J. Gen. Appl. Microbiol. 62, 140–143 (2016).

  155. 155.

    Miyake, K. et al. A green-light inducible lytic system for cyanobacterial cells. Biotechnol. Biofuels 7, 56 (2014).

  156. 156.

    Nakajima, M., Abe, K., Ferri, S. & Sode, K. Development of a light-regulated cell-recovery system for non-photosynthetic bacteria. Microb. Cell Fact. 15, 31 (2016).

  157. 157.

    Hori, M., Oka, S., Sugie, Y., Ohtsuka, H. & Aiba, H. Construction of a photo-responsive chimeric histidine kinase in Escherichia coli. J. Gen. Appl. Microbiol. 63, 44–50 (2017).

  158. 158.

    Nakajima, M., Ferri, S., Rogner, M. & Sode, K. Construction of a miniaturized chromatic acclimation sensor from cyanobacteria with reversed response to a light signal. Sci. Rep. 6, 37595 (2016).

  159. 159.

    Ramakrishnan, P. & Tabor, J. J. Repurposing Synechocystis PCC6803 UirS-UirR as a UV-violet/green photoreversible transcriptional regulatory tool in E. coli. ACS Synth. Biol. 5, 733–740 (2016).

  160. 160.

    Ziegler, T. & Möglich, A. Photoreceptor engineering. Front. Mol. Biosci. 2, 30 (2015).

  161. 161.

    Narikawa, R. et al. A biliverdin-binding cyanobacteriochrome from the chlorophyll d-bearing cyanobacterium Acaryochloris marina. Sci. Rep. 5, 10 (2015).

  162. 162.

    Fushimi, K. et al. Photoconversion and fluorescence properties of a red/green-type cyanobacteriochrome AM1_C0023g2 that binds not only phycocyanobilin but also biliverdin. Front. Microbiol. 7, 588 (2016).

  163. 163.

    Muller, K. et al. Synthesis of phycocyanobilin in mammalian cells. Chem. Commun. 49, 8970–8972 (2013).

  164. 164.

    Kyriakakis, P. et al. Biosynthesis of orthogonal molecules using ferredoxin and ferredoxin-NADP+ reductase systems enables genetically encoded PhyB optogenetics. ACS Synth. Biol. 7, 706–717 (2018).

  165. 165.

    Reichhart, E., Ingles-Prieto, A., Tichy, A. M., McKenzie, C. & Janovjak, H. A phytochrome sensory domain permits receptor activation by red light. Angew. Chem. Int. Ed. 55, 6339–6342 (2016).

Download references


The authors thank the referees of this Review for their substantial efforts to help improve the manuscript. They also thank J. Sanfilippo and other members of the Kehoe laboratory, as well as M. Wiltbank, for their valuable thoughts and insights. This work was supported by the Office of the Vice Provost for Research at Indiana University, Bloomington, through its Bridge Funding Program to D.M.K. and by a National Science Foundation Grant to D.M.K. (MCB-1818187).

Reviewer information

Nature Reviews Microbiology thanks A. Wilde and the other anonymous reviewer(s) for their contribution to the peer review of this work.

Author information


  1. Department of Biology, Indiana University, Bloomington, IN, USA

    • Lisa B. Wiltbank
    •  & David M. Kehoe


  1. Search for Lisa B. Wiltbank in:

  2. Search for David M. Kehoe in:


D.M.K. and L.B.W. researched the data for the article, substantially contributed to discussion of content, wrote the article and reviewed and edited the manuscript before submission.

Competing interests

The authors declare no competing interests.

Corresponding author

Correspondence to David M. Kehoe.



The amount of radiant energy received by a surface per unit time and unit area.


The ability to use light to produce food from inorganic molecules.


A small molecule or region that imparts light sensing to a protein.


Pigments produced from porphyrins that can function as chromophores for photoreceptors and photosynthetic light-harvesting proteins.


The series of structural changes that a photoreceptor undergoes upon light absorption before returning to its initial structure.


The biological engineering of specific cellular processes in living cells to be controlled by light.

Photosensory module

A collection of domains within a photoreceptor that binds a bilin chromophore and endows the ability to sense and respond to light.

PAS domain

A widespread protein structural domain that is commonly involved in protein–protein interactions or functions as a sensor in signal transduction pathways. PAS is an abbreviation of Per-Arnt-Sim, the proteins in which this domain was first discovered.

GAF domain

A domain that is widely found in eukaryotes and prokaryotes and is structurally similar to a PAS domain. Among its many roles, it binds chromophores and functions as a photoreceptor. GAF is an abbreviation of cGMP phosphodiesterases, adenylyl cyclases and FhiA, the names of three proteins that contain this domain.

PHY domain

A domain that is structurally related to a GAF domain and is found specifically in the photosensory module of phytochromes and stabilizes the light-activated state. PHY is an abbreviation for phytochrome-specific protein.

Output domain

A domain of a signal transduction protein with the capacity to elicit a molecular or cellular response. In photoreceptors, its function may be modulated by the photosensory module.

MCP domain

A membrane-spanning sensory molecule in bacteria first identified as a chemical sensor that transmits its signal through a two-component system histidine–aspartate phosphotransfer reaction. MCP is an acronym for methyl-accepting chemotaxis protein.

GGDEF domain

A domain that is present in many bacterial proteins and synthesizes the second messenger cyclic diguanosine monophosphate (c-di-GMP), typically under the regulation of another domain. GGDEF is an abbreviation of a five amino acid motif (Gly–Gly–Asp–Glu–Phe) in the domain.

EAL domain

A domain that is responsible for hydrolysis of the second messenger cyclic diguanosine monophosphate (c-di-GMP). EAL is based on the highly conserved sequence motif (Glu–Ala–Leu) near the amino-terminal end of the domain.


Light-harvesting antennae of some photosynthetic organisms. These are water soluble, located on the surface of photosynthetic membranes and composed of pigmented and non-pigmented proteins. In cyanobacteria, they are typically hemidiscoidal structures with rods radiating from an inner core.

Absorbance maximum

max). The wavelength that is maximally absorbed by the chromophore or light-absorbing protein being examined.

Cgi system

A post-transcriptional regulatory pathway that modulates light-responsive gene expression during chromatic acclimation in the cyanobacterium Fremyella diplosiphon Fd33 Cgi is an acronym for control of green light induction.


Genes whose protein product causes a change in cell morphology.


A metabolic process in photosynthetic organisms that causes the oxygenation of ribulose-1,5-bisphosphate, reducing the amount of 3-phosphoglycerate produced during the Calvin–Benson cycle and wasting energy obtained by the light-dependent reactions of photosynthesis.


A bacterial compartment, made of proteins and containing the enzymes ribulose-1,5-bisphosphate carboxylase/oxygenase and carbonic anhydrase, where carbon dioxide is concentrated to increase the efficiency of carbon fixation during photosynthesis.


One of the bilin-containing proteins, or phycobiliproteins, typically found in the central or core region of photosynthetic light-harvesting antennae called phycobilisomes.

Cyclic diguanosine monophosphate

(c-di-GMP). A widespread second messenger in bacterial cells that controls cell movement and many developmental processes.

Twitching motility

A type of motility carried out by bacteria on solid surfaces using hair-like filaments called type IV pili.

PATAN domain

A domain within a response regulator involved in cyanobacterial heterocyst patterning during nitrogen starvation. The PATAN domain is found in many prokaryotes and regulates motility and development through protein–protein interactions. PATAN is an abbreviation of PatA amino terminus.

PilZ domain

Named after the type IV pilus control protein PilZ, these domains are capable of binding cyclic diguanosine monophosphate (c-di-GMP), which causes a conformation change and leads to physiological responses including a shift from a motile to sessile state.

About this article

Publication history