The genome sequence has been submitted to the EMBL database under accession number FN869568.
A blueprint of ectoine metabolism from the genome of the industrial producer Halomonas elongata DSM 2581T
Version of Record online: 16 SEP 2010
© 2010 Society for Applied Microbiology and Blackwell Publishing Ltd
Thematic Issue: Extremophiles. Guest Editors: Ricardo Cavicchioli, Ricardo Amils, Dirk Wagner, Terry McGenity
Volume 13, Issue 8, pages 1973–1994, August 2011
How to Cite
Schwibbert, K., Marin-Sanguino, A., Bagyan, I., Heidrich, G., Lentzen, G., Seitz, H., Rampp, M., Schuster, S. C., Klenk, H.-P., Pfeiffer, F., Oesterhelt, D. and Kunte, H. J. (2011), A blueprint of ectoine metabolism from the genome of the industrial producer Halomonas elongata DSM 2581T. Environmental Microbiology, 13: 1973–1994. doi: 10.1111/j.1462-2920.2010.02336.x
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- Issue online: 21 AUG 2011
- Version of Record online: 16 SEP 2010
- Received 13 January, 2010; accepted 3 August, 2010.
Fig. S1. Species-level assignment of H. elongata proteins by MEGAN analysis. The plot indicates the number of proteins assigned to the named species out of a set of 1672 proteins for which such an assignment was successful. The number of assigned proteins is given in parenthesis after the name of the species. Only 8% of the proteins are not assigned to C. salexigens.
Fig. S2. Alignment of the H. elongata and C. salexigens chromosomes. The chromosomes of H. elongata and C. salexigens were aligned using MUMmer software (Kurtz et al., 2004) and show a strong X-alignment. Such X-alignments have been described for several interspecies comparisons and attributed to genome inversions around the replication origin (Eisen et al., 2000). A prominent X-alignment probably indicates that the number of such genome rearrangement events was small, which is astonishing for species with such an evolutionary distance that they are classified into distinct genera.
Fig. S3. Utilization of acetate as carbon source by H. elongata strains in the absence and presence of ectoine. H. elongata wild type, mutant strains KB41 (ΔdoeA), KB42 (ΔdoeB) and KB47 (ΔdoeD) were incubated for 3 days at 30°C on mineral salt medium (0.51 M NaCl) containing 40 mM acetate (A) and 40 mM acetate plus 10 mM ectoine (B) respectively. All strains were able to grow on acetate as sole carbon source (A). In the presence of ectoine, mutant KB47 failed to grow with acetate, while KB41 and KB42 are still able to utilize acetate (B). The inability of doeD mutant KB47 to grow on acetate in the presence of ectoine explains why KB47 also fails to grow with ectoine alone although acetate should be still provided due to the deacetylase activity of DoeB (Fig. 3).
Fig. S4. RT-PCR and RACE-PCR analysis of the doeABXC region. (A) Genetic and physical organization of the doeABX locus. The position of the σ70-dependent promoter is indicated. Primer binding sites for RT-PCR are marked by triangles. Black triangles with dotted lines show successful generation of PCR products. For the pair of white triangles, no PCR product was obtained. Reverse primers were used for both, RT reaction and PCR. (B) RT-PCR analysis proving that doeABX is organized as one operon. A 1450 bp PCR product was amplified from cDNA and separated by agarose-gel electrophoresis (lane 1), which matched the size of the calculated doeABX PCR product (1473 bp). A corresponding doeX–doeC product could be amplified from genomic DNA with the same primer pair (positive control, lane 2). No PCR product could be amplified from cDNA using a primer pair to doeX and doeC, proving that doeC is not part of the doeABX operon (lane 3). A corresponding doeX–doeC product could be amplified from genomic DNA with the same primer pair (positive control, lane 4). A PCR product could be amplified with primers that both bind within the doeC ORF (control RT reaction, data not shown, product indicated in A). (C) Nucleotide sequence of the doeA promoter region. Arrows indicate the transcription initiation site (+ 1), which was mapped by RACE-PCR. The −35 and −10 sequences of the σ70-dependent promoter upstream of doeA are written in bold.
Fig. S5. Maximum ectoine yield as a function of the ATP load and turnover of ectoine in the synthesis degradation cycle (A) and flux distributions for maximum conversion into ectoine (B). (A) Points I to IV mark the flux distributions described in panel B. Horizontal axes represent total consumption of ATP by processes outside the model and the flux circulating through the ectoine synthesis/degradation cycle. The units are arbitrary, normalized for a glucose uptake of 100 (e.g. molecules). (B) I) One of the admissible flux distributions without ATP demand. II) Unique solution when total ATP demand is less than or equal to one ATP per glucose III) Example of solutions for higher ATP demands, superposition of II and maximum ATP production distribution. IV) Example of solutions for a turnover of ectoine, identical to III everywhere except the ATP load and cycle.
Table S1. The 20 COGs with the highest occupancy.
Table S2. Organisms used for the search for high salt COGs.
Table S3. Average pI difference between proteins from H. elongata and the indicated organisms.
Table S4. Enzymatic reactions used in the metabolic models.
Table S5. Bacterial strains and plasmids used in this study.
Table S6. Locus tag of genes similar to ect genes, ask, and doe genes depicted in Figs 2, 4 and 7, and accession data of genome sequences of corresponding organisms.
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