BMB Reports

Korean Society for Biochemistry and Molecular Biology (생화학분자생물학회)
권호 표지
DOI QR코드

DOI QR Code

Periplasmic glucans isolated from Proteobacteria

  • Lee, Sang-Hoo (Department of Bioanalysis and Mass Spectrometry, Seoul Medical Science Institute) ;
  • Cho, Eun-Ae (Department of Bioscience and Biotechnology, Bio/Molecular Informatics Center, Konkuk University) ;
  • Jung, Seun-Ho (Department of Bioscience and Biotechnology, Bio/Molecular Informatics Center, Konkuk University)
  • Published : 2009.12.31
Download PDF
⟨ Previous Next ⟩

Abstract

Periplasmic glucans (PGs) are general constituents in the periplasmic space of Proteobacteria. PGs from bacterial strains are found in larger amounts during growth on medium with low osmolarity and thus are often been specified as osmoregulated periplasmic glucans (OPGs). Furthermore, they appear to play crucial roles in pathogenesis and symbiosis. PGs have been classified into four families based on the structural features of their backbones, and they can be modified by a variety of non-sugar substituents. It has also recently been confirmed that novel PGs with various degrees of polymerization (DPs) and/or different substituents are produced under different growth conditions among Proteobacteria. In addition to their biological functions as regulators of low osmolarity, PGs have a variety of physico-chemical properties due to their inherent three-dimensional structures, hydrogen-bonding and complex-forming abilities. Thus, much attention has recently been focused on their physico-chemical applications. In this review, we provide an updated classification of PGs, as well as a description of the occurrences of novel PGs with substituents under various bacterial growth environments, the genes involved in PG biosynthesis and the various physico-chemical properties of PGs.

Keywords

References

  1. Bohin, J. P. and Lacroix, J. M. (2006). Osmoregulation in the periplasm; in The Periplasm, Ehrmann. M. (ed.) pp. 325-341. American Society for Microbiology. Washington, DC, USA
  2. McIntire, F. C., Peterson, W. H. and Riker, A. J. (1942) A polysaccharide produced by the crown-gall organism. J. Biol. Chem. 143, 491-496
  3. Van Golde, L. M. G., Schulman, H. and Kennedy, E. P. (1973) Metabolism of membrane phospholipids and its relation to a novel class of oligosaccharides in Escherichia coli. Proc. Natl. Acad. Sci. U.S.A. 70, 1368-1372 https://doi.org/10.1073/pnas.70.5.1368
  4. Kennedy, E. P. Membrane-derived oligosaccharides (periplasmic $\beta$-D-glucans) of Escherichia coli and Salmonella. Cellular and Molecular Biology, Neidhardt, E. C., Curtiss III, R., Ingraham, J. L., Lin, E. C. C., Low, K. B., Maganasik, B., Reznikoff, W. S., Riley, M., Schaechter, M. and Umbarger,H. E. (eds.), 2nd ed., pp. 1064-1074. American Society for Microbiology, Washington DC, USA
  5. Miller, K. J., Kennedy, E. P. and Reinhold, V. N. (1986) Osmotic adaptation in Gram-negative bacteria: possible role for periplasmic oligosaccharides. Science 231, 48-51 https://doi.org/10.1126/science.3941890
  6. Bohin, J. P. (2000) Osmoregulated periplasmic glucans in Proteobacteria. FEMS Microbiol. Lett. 186, 11-19 https://doi.org/10.1111/j.1574-6968.2000.tb09075.x
  7. Cho, E., Jeon, Y. and Jung, S. (2009) Novel succinylated and large-sized osmoregulated periplasmic glucans of Pseudomonas syringae pv. syringae. Carbohydr. Res. 344, 996-1000 https://doi.org/10.1016/j.carres.2009.03.015
  8. Lequette, Y., Rollet, E., Delangle, A., Greenberg, E. P. and Bohin, J. P. (2007) Linear osmoregulated periplasmic glucans are encoded by the opgGH locus of Pseudomonas aeruginosa. Microbiology 153, 3255-3263 https://doi.org/10.1099/mic.0.2007/008953-0
  9. Breedveld, M. W. and Miller, K. J. (1994) Cyclic $\beta$-glucans of the family Rhizobiaceaes. Microbiol. Rev. 58, 145-161
  10. Amemura, A., Hisamatsu, M., Mitani, H. and Harada, T. (1983) Cyclic (1,2)-β-D-glucan and the octasaccharide repeating units of extracellular acidic polysaccharides produced by Rhizobium. Carbohydr. Res. 114, 277-285 https://doi.org/10.1016/0008-6215(83)88194-4
  11. Choma, A. and Komaniecka, I. (2003) Characterisation of Mesorhizobium huakuii cyclic $\beta$-glucan. Acta Biochim. Pol. 50, 1273-1281
  12. Koizumi, K., Okada, Y., Horiyama, S. and Utamura, T. (1983) Separation of cyclic (1$\rightarrow$2)-$\beta$-D-glucans (cyclosophoraoses) produced by Agrobacterium and Rhizobium, and determination of their degree of polymerization by high-performance liquid chromatography. J. Chromatogr. 265, 89-96 https://doi.org/10.1016/S0021-9673(01)96701-9
  13. Komaniecka, I. and Choma, A. (2003) Isolation and characterization of periplasmic cyclic β-glucans of Azorhizobium caulinodans. FEMS Microbiol. Lett. 227, 263-269 https://doi.org/10.1016/S0378-1097(03)00690-6
  14. Altabe, S. G., De Iannino, N. I., De Mendoza, D. and Ugalde, R. A. (1994) New osmoregulated $\beta$(1-3),(1-6) glucosyltransferase(s) in Azospirillum brasilense. J. Bacteriol. 176, 4890-4898 https://doi.org/10.1128/jb.176.16.4890-4898.1994
  15. Altabe, S. G., Talaga, P., Wieruszeski, J. M., Lippens, G., Ugalde, R. and Bohin, J. P. (1998) Periplasmic glucans of Azospirillum brasilense; in Biological nitrogen fixation for the 21st century, Elmerich, C., Kondorosi, A., Newton, W. E. (eds.), p. 390. Kluwer Academic Publishers, Dordrecht
  16. Jung, Y., Park, H., Cho, E. and Jung, S. (2005) Structural analyses of novel glycerophosphorylated α-cyclosophorohexadecaoses isolated from X. campestris pv. campestris. Carbohydr. Res. 340, 673-677 https://doi.org/10.1016/j.carres.2004.12.030
  17. Talaga, P., Stahl, B., Wieruszeski, J. M., Hillenkamp, F., Tsuyumu, S., Lippens, G. and Bohin, J. P. (1996) Cell-associated glucans of Burkholderia solanacearum and Xanthomonas campestris pv. citri: a new family of periplasmic glucans. J. Bacteriol. 178, 2263-2271 https://doi.org/10.1128/jb.178.8.2263-2271.1996
  18. York, W. S. (1995) A conformational model for cyclic $\beta$- (1→2)-linked glucans based on NMR analysis of the $\beta$-glucans produced by Xanthomonas campestris. Carbohydr. Res. 278, 205-225 https://doi.org/10.1016/0008-6215(95)00260-X
  19. Cho, E., Lee, S. and Jung, S. (2007) Benzoate methanolysis catalyzed by $\alpha$-cyclosophorohexadecaose isolated from Xanthomonas oryzae. Carbohydr. Polym. 70, 174-180 https://doi.org/10.1016/j.carbpol.2007.03.013
  20. Amemura, A. and Cabrera-crospo, J. (1986) Extracellular oligosaccharides and low-Mr polysaccharides containing (1$\rightarrow$2)-b-D-glucosidic linkages from strains of Xanthomonas, Escherichia coli and Klebsiella pneumonia. J. Gen. Microbiol. 132, 2443-2452
  21. Cho, E., Lee, S. and Jung, S. (2008) Novel acetylated $\alpha$-cyclosophorotridecaose produced by Ralstonia solanacearum. Carbohydr. Res. 343, 912-918 https://doi.org/10.1016/j.carres.2008.01.023
  22. Wieruszeski, J. M., Bohin, A., Bohin, J. P. and Lippens, G.(2001) In vivo detection of the cyclic osmoregulated periplasmic glucan of Ralstonia solanacearum by high-resolution magic angle spinning NMR. J. Magn. Reson. 151, 118-123 https://doi.org/10.1006/jmre.2001.2348
  23. Talaga, P., Cogez, V., Wieruszeski, J. M., Stahl, B., Lemoine, J., Lippens, G. and Bohin, J. P. (2002) Osmoregulated periplasmic glucans of the free-living photosynthetic bacterium Rhodobacter sphaeroides. Eur. J. Biochem. 269, 2464-2472 https://doi.org/10.1046/j.1432-1033.2002.02906.x
  24. Lopez-Lara, I. M., Orgambide, G., Dazzo, F. B., Olivares, J. and Toro, N. (1993) Characterization and symbiotic im portance of acidic extracellular polysaccharides of Rhizobium sp. strain GRH2 isolated from acacia nodules. J. Bacteriol. 175, 2826-2832 https://doi.org/10.1128/jb.175.10.2826-2832.1993
  25. Zevenhuizen, L. P., van Veldhuizen, A. and Fokkens, R. H. (1990) Re-examination of cellular cyclic β-1,2-glucans of Rhizobiaceae: distribution of ring sizes and degrees of glycerol-1-phosphate substitution. Antonie Leeuwenhoek 57, 173-178 https://doi.org/10.1007/BF00403952
  26. Geiger, O., Weissborn, A. C. and Kennedy, E. P. (1991) Biosynthesis and excretion of cyclic glucans by Rhizobium meliloti 1021. J. Bacteriol. 173, 3021-3024 https://doi.org/10.1128/jb.173.9.3021-3024.1991
  27. Cho, E. and Jung, S. (2009) Novel acetylated linear periplasmic glucans isolated from Pseudomonas syringae. Bull. Korean Chem. Soc. 30, 1-4 https://doi.org/10.5012/bkcs.2009.30.10.2433
  28. Roset, M. S., Ciocchini, A. E., Ugalde, R. A. and de Iannino, N. I. (2006) The Brucella abortus cyclic $\beta$-1,2-glucan virulence factor is substituted with O-ester-linked succinyl residues. J. Bacteriol. 188, 5003-5013 https://doi.org/10.1128/JB.00086-06
  29. Lacroix, J. M., Loubens, I., Tempête, M., Menichi, B. and Bohin, J. P. (1991) The mdoA locus of Escherichia coli consists of an operon under osmotic control. Mol. Microbiol. 5, 1745-1753 https://doi.org/10.1111/j.1365-2958.1991.tb01924.x
  30. Jackson, B. J., Bohin, J. P. and Kennedy, E. P. (1984) Biosynthesis of membrane derived oligosaccharides: characterization of opgB mutants defective in phosphoglycerol transferase I activity. J. Bacteriol. 160, 976-981
  31. Lanfroy, E. and Bohin, J. P. (1993) Physical map location of the Escherichia coli gene encoding phosphoglycerol transferase I. J. Bacteriol. 175, 5736-5737 https://doi.org/10.1128/jb.175.17.5736-5737.1993
  32. Lacroix, J. M., Lanfroy, E., Cogez, V., Lequette, Y., Bohin, A. and Bohin, J. P. (1999) The mdoC gene of Escherichia coli encodes a membrane protein that is required for succinylation of osmoregulated periplasmic glucans. J. Bacteriol. 181, 3626-3631
  33. Lequette, Y., Odberg-Ferragut, C., Bohin, J. P. and Lacroix, J. M. (2004) Identification of mdoD, an mdoG paralog which encodes a twin-arginine-dependent periplasmic protein that controls osmoregulated periplasmic glucan backbone structures. J. Bacteriol. 186, 3695-3702 https://doi.org/10.1128/JB.186.12.3695-3702.2004
  34. Goldberg, D. E., Rumley, M. K., and Kennedy, E. P. (1981) The biosynthesis of membrane-derived oligosaccharides: a periplasmic phosphoglycerol transferase. Proc. Natl. Acad. Sci. U.S.A. 78, 5513-5517 https://doi.org/10.1073/pnas.78.9.5513
  35. Jackson, B. J. and Kennedy, E. P. (1983) The biosynthesis of membrane-derived oligosaccharides: a membrane-bound phosphoglycerol transferase. J. Biol. Chem. 258, 2394-2398
  36. Bohin, J. P. and Kennedy, E. P. (1984) Regulation of the synthesis of membrane-derived oligosaccharides in Escherichia coli. Assay of phosphoglycerol transferase I in vivo. J. Biol. Chem. 259, 8388-8393
  37. Lequette, Y., Lanfroy, E., Cogez, V., Bohin, J. P. and Lacroix, J. M. (2008) Biosynthesis of osmoregulated periplasmic glucans in Escherichia coli: the membrane-bound and the soluble periplasmic phosphoglycerol transferases are encoded by the same gene. Microbiology 154, 476-483 https://doi.org/10.1099/mic.0.2007/013169-0
  38. Mukhopadhyay, P., Williams, J. and Mills, D. (1988). Molecular analysis of a pathogenicity locus in Pseudomonas syringae pv. syringae. J. Bacteriol. 170, 5479-5488 https://doi.org/10.1128/jb.170.12.5479-5488.1988
  39. Page, F., Altabe, S., Hugouvieus-Cotte-Pattat, N., Lacroix, J. M., Robert-Baudouy, J. and Bohin, J. P. (2001) Osmoregulated periplasmic glucans synthesis is required for Erwinia chrysanthemi pathogenicity. J. Bacteriol. 183, 3134-3141 https://doi.org/10.1128/JB.183.10.3134-3141.2001
  40. Zorreguieta, A., Geremia, R. A., Cavaignac, S., Cangelosi, G. A., Nester, E. W. and Ugalde, R. A. (1988) Identification of the product of an Agrobacterium tumefaciens chromosomal virulence gene. Mol. Plant-Microbe Interact. 1, 121-127 https://doi.org/10.1094/MPMI-1-121
  41. Ielpi, L., Dylan, T., Ditta, G. S., Helinski, D. R. and Stanfield. S. W. (1990) The ndvB locus of Rhizobium meliloti encodes a 319-kDa protein involved in the production of $\beta$-(1,2)-glucan. J. Biol. Chem. 265, 2843-2851
  42. Zorreguieta, A. and Ugalde, R. A. (1986) Formation in Rhizobium and Agrobacterium spp. of a 235-kilodalton protein intermediate in $\beta$-D(1,2)-glucan synthesis. J. Bacteriol. 167, 947-951 https://doi.org/10.1128/jb.167.3.947-951.1986
  43. Breedveld, M. W., Hadley, J. A. and Miller, K. J. (1995) A novel cyclic-$\beta$-1,2-glucan mutant of Rhizobium meliloti. J. Bacteriol. 177, 6346-6351 https://doi.org/10.1128/jb.177.22.6346-6351.1995
  44. Bahgwat, A. A., Mith$\ddot{o}$fer, A., Pfeffer, P. E., Kraus, C., Spickers, N., Hotchkiss, A., Ebel, J. and Keister, D. L. (1999) Further studies of the role of cyclic $\beta$-glucans in symbiosis. An ndvC mutant of Bradyrhizobium japonicum synthesizes cyclodecakis-(1$\rightarrow$3)-$\beta$-glucosyl. Plant Physiol. 119, 1057-1064 https://doi.org/10.1104/pp.119.3.1057
  45. Cogez, V., Evgueni Gak, E., Puskas, A., Kaplan, S. and Bohin, J. P. (2002) The opgGIH and opgC genes of Rhodobacter sphaeroides form an operon that controls backbone synthesis and succinylation of osmoregulated periplasmic glucans. Eur. J. Biochem. 269, 2473-2484 https://doi.org/10.1046/j.1432-1033.2002.02907.x
  46. Minsavage, G. V., Mudgett, M. B., Stall, R. E. and Jones, J. B. (2004) Importance of opgHXcv of Xanthomonas campestris pv. vesicatoria in host-parasite interactions. Mol. Plant-Microbe Interact. 17, 152-161 https://doi.org/10.1094/MPMI.2004.17.2.152
  47. Salanoubat, M., Genin, S., Artiguenave, F., Gouzy, J., Mangenot, S., Arlat, M., Billault, A., Brottier, P., Camus, J. C., Cattolico, L., Chandler, M., Choisne, N., Claudel-Renard, C., Cunnac, S., Demange, N., Gaspin, C., Lavie, M., Moisan, A., Robert, C., Saurin, W., Schiex, T., Siguier, P., Thebault, P., Whalen, M., Wincker, P., Levy, M., Weissenbach, J. and Boucher, C. A. (2002) Genome sequence of the plant pathogen Ralstonia solanacearum. Nature 415, 497-502 https://doi.org/10.1038/415497a
  48. Brown, D. G. and Allen, C. (2004) Ralstonia solanacearum genes induced during growth in tomato: an inside view of bacterial wilt. Mol. Microbiol. 53, 1641-1660 https://doi.org/10.1111/j.1365-2958.2004.04237.x
  49. Okada, Y., Horiyama, S. and Koizumi, K. (1985) Studies on inclusion complexes of non-steroidal anti-inflammatory agents with cyclosophoraose-A. Yakugaku Zasshi 106, 240-247
  50. Koizumi, K., Okada, Y., Horiyama, S., Utamura, T., Higashiura, T. and Ikeda, M. (1984) Preparation of cyclosophoraose-A and its complex-forming ability. J. Incl. Phenom. 2, 891-899 https://doi.org/10.1007/BF00662259
  51. Morris, V. J., Brownsey, G. J., Chilvers, G. R., Harris, J. E., Morris, V. J., Brownsey, G. J., Chilvers, G. R., Harris, J. E., Gunning, A. P. and Stevens, B. H. J. (1991) Possible biological roles for Rhizobium leguminosarum extracellular polysaccharide and cyclic glucans in bacteria-plant interactions for nitrogen-fixing bacteria. Food Hydrocoll. 5, 185-188 https://doi.org/10.1016/S0268-005X(09)80312-3
  52. Lee, S., Seo, D., Park, H., Choi, Y. and Jung, S. (2003) Solubility enhancement of a hydrophobic flavonoid, luteolin by the complexation with cyclosophoraoses iso lated from Rhizobium meliloti. Antonie van Leeuwenhoek 84, 201-207 https://doi.org/10.1023/A:1026075215921
  53. Kang, S., Lee, S., Kwon, C. and Jung, S. (2006) Solubility enhancement of flavonoids by cyclosophoraose isolated from Rhizobium meliloti 2011. J. Microbiol. Biotechnol. 16, 791-794
  54. Lee, S., Seo, D., Kim, H. and Jung, S. (2001) Investigation of inclusion complexation of paclitaxel by cyclohenicosakis-(1→2)-($\beta$-D-glucopyranosyl), by cyclic-(1→2)-$\beta$-D-glucans (cyclosophoraoses), and by clomaltoheptaoses ($\beta$-cyclodextrins). Carbohydr. Res. 334, 119-126 https://doi.org/10.1016/S0008-6215(01)00178-1
  55. Lee, S., Kwon, C., Choi, Y., Seo, D., Kim, H., and Jung, S. (2001) Inclusion complexation of a family of cyclosophoraoses with indomethacin. J. Microiol. Biotechnol. 11, 463-468
  56. Park, H., Choi, Y. Kang, S. Lee, S. Kwon, C. and Jung, S. pH-Dependent inclusion complexation of carboxymethylated cyclosophoraoses to N-acetyl phenylalanine. Carbohydr. Polym. 64, 85-91
  57. Park, H. and Jung, S. (2005) pH-Dependent on-off inclusion complexation of carboxymethylated cyclosophoraoses with neutral red. Bull. Korean Chem. Soc. 26, 675-678 https://doi.org/10.5012/bkcs.2005.26.4.675
  58. Lee, S., Park, H., Seo, D., Choi, Y. and Jung, S. (2004) Synthesis and characterization of carboxymethylated cyclosophoraose, and its inclusion complexation behavior. Carbohydr. Res. 339, 519-527 https://doi.org/10.1016/j.carres.2003.11.011
  59. Kwon, C., Choi, Y., Kim, N., Yoo, J., Yang, C., Kim, H. and Jung, S. (2000) Complex forming ability of a family of isolated cyclosophoraoses with erogosterol and its Monte Carlo docking comutatational analysis. J. Incl. Phenom Macro. Chem. 36, 55-65 https://doi.org/10.1023/A:1008050432556
  60. Cho, E., Jeon, Y. and Jung, S. (2009) Chiral separation of hesperetin and hesperetin-O-glycoside in capillary electrophoresis using microbial $\beta$-1,2-glucans. Bull. Korean Chem. Soc. 30, 1870-1872 https://doi.org/10.5012/bkcs.2009.30.8.1870
  61. Park, H. and Jung, S. (2005) Separation of some chiral flavonoids by microbial cyclosophoraoses and their sulfated derivatives in micellar electrokinetic chromatography. Electrophoresis 26, 3833-3838 https://doi.org/10.1002/elps.200500194
  62. Jung, Y., Lee, S., Paik, S. R. and Jung, S. (2004) Cyclosophoraose as a novel chiral stationary phase for enantioseparation. J. Microbiol. Biotechnol. 14, 1338-1342
  63. Park, H., Lee, S., Kang, S., Jung, Y. and Jung, S. (2004) Enantioseparation using sulfated cyclosophoraoses as a novel chiral additive in capillary electrophoresis. Electrophoresis 25, 2671-2674 https://doi.org/10.1002/elps.200405971
  64. Lee, S., Choi, Y. Lee, S., Jeong, K. and Jung, S. (2004) Chiral recognition based on enantioselective interactions of propranolol enantiomers with cyclosophoraoses isolated from Rhizobium meliloti. Chirality 16, 204-210 https://doi.org/10.1002/chir.20010
  65. Lee, S. and Jung, S. (2003) Enantioseparation using cyclosophoraoses as a novel chiral additive in capillary electrophoresis. Carbohydr. Res. 338, 1143-1146 https://doi.org/10.1016/S0008-6215(03)00083-1
  66. Lee, S. and Jung, S. (2002) $^1^3{C}$ NMR spectroscopic analysis on the chiral discrimination of N-acetylphenylalanine, catechin and propranolol induced by cyclic-(1,2)-β-D-glucans (cyclosophoraoses). Carbohydr. Res. 337, 1785-1789 https://doi.org/10.1016/S0008-6215(02)00286-0
  67. Park, H., Kang, L. and Jung, S. (2008) Methanolysis of 7-acetoxy-4-methylcoumarin catalyzed by cyclosophoraoses. Bull. Korean Chem. Soc. 29, 228-230 https://doi.org/10.5012/bkcs.2008.29.1.228
  68. Park, H. and Jung, S. (2008) Methanolysis of ethyl esters of N-acetyl amino acids catalyzed by cyclosophoraoses isolated from Rhizobium meliloti. Carbohydr. Res. 343, 274-281 https://doi.org/10.1016/j.carres.2007.10.033
  69. Lee, S. and Jung, S. (2004) Cyclosophoraose as a catalytic carbohydrate for methanolysis. Carbohydr. Res. 339, 461-468 https://doi.org/10.1016/j.carres.2003.11.004
  70. Lee, S., Kwon, C., Park, B. and Jung, S. (2009) Synthesis of selenium nanowires morphologically directed by Shinorhizobial oligosaccharides. Carbohydr. Res. 344, 1230-1234 https://doi.org/10.1016/j.carres.2009.04.014
  71. Choi, Y., Yang, C., Kim, H. and Jung, S. (2000) Molecular dynamics simulations cyclohenicosakis-[(1→2)-$\beta$-D-gluco-henicosapyranosyl], a cyclic (1→2)-$\beta$-D-glucan (a cyclosophoraose') of DP 21. Carbohydr. Res. 326, 227-234 https://doi.org/10.1016/S0008-6215(00)00050-1
  72. Kwon, C., Choi, J., Lee, S., Park, H. and Jung, S. (2007) Chiral separation and discrimination of catechin by microbial cyclic $\beta$-(1$\rightarrow$3),(1$\rightarrow$6)-glucans isolated from Bradyrhizobium japonicum. Bull. Korean Chem. Soc. 28, 347-350 https://doi.org/10.5012/bkcs.2007.28.2.347
  73. Cho, E., Jeon, Y. and Jung, S. (2009) Chiral separation of hesperetin and hesperetin-O-glycoside in capillary electrophoresis using microbial $\beta$-1,2-glucans. Bull. Korean Chem. Soc. 30, 1870-1872 https://doi.org/10.5012/bkcs.2009.30.8.1870
  74. Lee, S., Cho, E., Kwon, C. and Jung, S. (2007) Cyclosophorohexadecaose and succinoglycan monomers as catalytic carbohydrates for the Strecker reaction. Carbohydr. Res. 342, 2682-2687 https://doi.org/10.1016/j.carres.2007.07.006
  75. Lippens, G., Wieruszeski, J. M., Horvath, D., Talaga, P. and Bohin, J. P. (1998) Slow dynamics of the cyclic osmoregulated periplasmic glucan of Ralstonia solanacearum as revealed by heteronuclear relaxation studies. J. Am. Chem. Soc. 120, 170-177 https://doi.org/10.1021/ja970960u
  76. Wieruszeski, J. M., Bohin, A., Bohin, J. P. and Lippens, G. (2001) In vivo detection of the cyclic osmoregulated periplasmic glucan of Ralstonia solanacearum by high-resolution magic angle spinning NMR. J. Magn. Reson. 151, 118-123 https://doi.org/10.1006/jmre.2001.2348
  77. Kim, H., Jeong, K., Cho, K. W., Paik, S. R. and Jung, S. (2006) Molecular dynamics simulations of a cyclic-$\beta$-(1→2) glucancontaining an a-(1→6) linkage as a 'molecular alleviator' for the macrocyclic conformational strain. Carbohydr. Res. 341, 1011-1019 https://doi.org/10.1016/j.carres.2006.02.025
  78. Choi, Y. and Jung, S. (2005) The $\alpha$-(1→6) glycosidic linkage as a novel conformational entropic regulator in osmoregulated periplasmic α-cyclosophorohexadecaose. Carbohydr. Res. 340, 2550-2557 https://doi.org/10.1016/j.carres.2005.08.020

Cited by

  1. Characterization and applications of cyclic β-(1,2)-glucan produced from R. meliloti vol.4, pp.22, 2014, https://doi.org/10.1039/c3ra47073c
  2. New insights into the biological role of the osmoregulated periplasmic glucans in pathogenic and symbiotic bacteria vol.7, pp.5, 2015, https://doi.org/10.1111/1758-2229.12325
  3. Swarm and swim motilities of Salmonella enterica serovar Typhimurium and role of osmoregulated periplasmic glucans vol.3, pp.1, 2015, https://doi.org/10.7243/2052-6180-3-3
  4. Cyclic β-(1, 2)-glucan production by Rhizobium meliloti MTCC 3402 vol.48, pp.12, 2013, https://doi.org/10.1016/j.procbio.2013.08.024
  5. Synthesis and Characterization of Cyclic β-(1, 2)-Glucan from Agrobacterium Tumefaciens 2013, https://doi.org/10.12720/jolst.1.1.44-46
  6. Identification and Characterization of Novel Salmonella Mobile Elements Involved in the Dissemination of Genes Linked to Virulence and Transmission vol.7, pp.7, 2012, https://doi.org/10.1371/journal.pone.0041247
  7. Resistance and survival strategies of Salmonella enterica to environmental stresses vol.45, pp.2, 2012, https://doi.org/10.1016/j.foodres.2011.06.056
  8. Characterization of cyclic β-glucans of Bradyrhizobium by MALDI-TOF mass spectrometry vol.346, pp.13, 2011, https://doi.org/10.1016/j.carres.2011.05.015
  9. Generation of Free Oligosaccharides from Bacterial Protein N-Linked Glycosylation Systems vol.99, pp.10, 2013, https://doi.org/10.1002/bip.22296
  10. Phosphoethanolamine Transferase LptA in Haemophilus ducreyi Modifies Lipid A and Contributes to Human Defensin Resistance In Vitro vol.10, pp.4, 2015, https://doi.org/10.1371/journal.pone.0124373
  11. The Rcs Signal Transduction Pathway Is Triggered by Enterobacterial Common Antigen Structure Alterations in Serratia marcescens vol.193, pp.1, 2011, https://doi.org/10.1128/JB.00839-10
  12. Cyclic β-glucans at the bacteria-host cells interphase: One sugar ring to rule them all vol.20, pp.6, 2018, https://doi.org/10.1111/cmi.12850
  13. Structural characterization and applications of a novel polysaccharide produced by Azospirillum lipoferum MTCC 2306 vol.35, pp.1, 2019, https://doi.org/10.1007/s11274-019-2588-y