- Email: [email protected]
Flash pyrolysis – a rapid method for screening bacterial species for the presence of bacteriohopanepolyols
Organic Geochemistry Organic Geochemistry 36 (2005) 975–979 www.elsevier.com/locate/orggeochem
Note
Flash pyrolysis – a rapid method for screening bacterial species for the presence of bacteriohopanepolyols Mark A. Sugden, Helen M. Talbot *, Paul Farrimond School of Civil Engineering and Geosciences, University of Newcastle, Newcastle Upon Tyne NE1 7RU, UK Received 9 November 2004; accepted 7 December 2004 (returned to author for revision 24 November 2004) Available online 22 February 2005
Abstract Flash pyrolysis–gas chromatography–mass spectrometry (Py–GC–MS) was applied to freeze dried cells of eleven bacterial species. Six species known to synthesise bacteriohopanepolyols (BHPs) were found to produce easily identifiable C27–C35 hopenes and hopanes upon flash pyrolysis. These compounds could not be observed in the pyrolysis products of five species, two of which are known not to produce BHPs. In the search for BHP-containing bacteria this rapid screening approach prevents unnecessary time-consuming laboratory work to characterise species that would produce negative results. 2005 Elsevier Ltd. All rights reserved.
1. Introduction Bacteriohopanepolyols (BHPs), such as bacteriohopane-32,33,34,35-tetrol, are triterpenoids synthesised by certain types of bacteria, including gram-negative bacteria, methanotrophs, cyanobacteria, acetic acid bacteria, nitrogen fixers and purple non-sulfur bacteria (Rohmer et al., 1984; Farrimond et al., 1998, and references therein). An ever increasing range of BHP structures, with a complex biological functionality at C-35, are being found (e.g., Rohmer, 1993; Talbot et al., 2003a,b), and these can vary significantly among bacterial groups/species. The ubiquitous occurrence of BHPs and their diagenetic products (e.g., hopanols, hopanoic acids, hopenes and hopanes) in recent sediments (e.g., Innes et al., 1997; Talbot et al., 2003c) shows the impor* Corresponding author. Tel.: +44 191 246 4886; fax: +44 191 246 4961. E-mail address: [email protected] (H.M. Talbot).
tance of the bacterial contribution to the biomarker pool in the sedimentary record (Ourisson and Rohmer, 1992). BHPs in sediments are a record of bacterial activity and can therefore provide an indication of the palaeoenvironmental conditions at the time of deposition (e.g., Farrimond et al., 2000). To fully understand this record it is necessary to establish which species of the bacterial population contain BHPs and the specific composition of these BHPs. To date, they have been found in approximately 50% of the bacteria examined specifically for their presence, but they are not equally distributed across the various bacterial taxa (Farrimond et al., 1998). The highly functionalised and amphiphilic BHPs are not readily amenable to analysis by conventional GC–MS (Innes et al., 1997). However, a new technique involving atmospheric pressure chemical ionisation liquid chromatography/ion trap mass spectrometry (APCI-LC/MSn) has recently been used to identify them in a range of bacteria and modern sediment samples (Talbot et al., 2003a,b). This work requires dual stage
0146-6380/$ - see front matter 2005 Elsevier Ltd. All rights reserved. doi:10.1016/j.orggeochem.2004.12.003
976
M.A. Sugden et al. / Organic Geochemistry 36 (2005) 975–979
solvent extraction techniques followed by derivatisation using acetic anhydride and pyridine (Talbot et al., 2003a). In order to better constrain the occurrence and composition of BHPs in bacteria it is necessary to perform this time consuming laboratory work on a diverse range of species from different taxa, with the knowledge that hopanoids would likely not be detected in a significant proportion of the species tested. Thus, a rapid technique to screen for the presence of hopanoids is needed. A catalytic hydropyrolysis method for the screening of microbial cultures has recently been shown to produce a series of C27–C35 hopanes and hopenes from hopanoid-containing bacteria (Love et al., 2005). Here, we report the results from the application of pyrolysis–gas chromatography–mass spectrometry (Py–GC–MS) to a range of bacterial species. The aim was to establish a rapid low-cost method, using facilities that are available worldwide, to detect whether a particular species contained BHPs or not. This knowledge, prior to solvent extraction for analysis using APCI-LC/MSn, prevents unnecessary laboratory work on species which do not contain hopanoids.
2. Experimental 2.1. Cultures The species, strain, phylum or class and family of bacterial cells subjected to Py–GC–MS are summarised in Table 1, together with their measured BHP concentrations. 2.2. Py–GC–MS Py–GC–MS was performed on freeze dried cells using a resistance-heated CDS 120 Pyroprobe unit, fitted
with a platinum coil pyroprobe, coupled to a HewlettPackard 5890GC with split/splitless injector (300 C), linked to a Hewlett-Packard 5972MSD (electron voltage 70 eV, filament current 200 lA, source temperature 180 C, multiplier voltage 2000 V, interface temperature 300 C). The acquisition was controlled with a HP Vectra 486 Chemstation computer, in either full scan (range m/z 50–700 amu/s), or selected ion mode (m/z 191, 250 ms dwell). The sample (<1 mg) was placed in a quartz tube plugged with pre-extracted silica wool at both ends. Pyrolysis was performed at 510 C for 10 s, after which the split was opened. Various pyrolysis temperatures were tested (360, 410, 460, 510, 560, 610 and 660 C), 510 C being found to give a consistently high hopanoid response for Rhodopseudomonas palustris. R. palustris was also subjected to Py–GC–MS at 510 C with TMAH, with no obvious benefits. The oven temperature was programmed from 50 C (2 min) to 350 C at 10 C/ min and then held for 10 min. He was used as carrier gas (flow 1 mL/min, pressure of 50 kPa, split at 30 mL/min). The pyrolysis products were separated on a fused silica column (15 m · 0.25 mm i.d.) coated with a 5% phenylmethylsilicone bonded stationary phase (DB5-HT, film thickness 0.1 lm). Hopanes and hopenes were identified from their mass spectra (R. palustris and Methylosinus trichosporium) with reference to published spectra and spectra obtained from standard oil samples.
3. Results and discussion Fig. 1a shows the m/z 191 partial mass chromatogram of the hopanoid degradation products released after flash pyrolysis of R. palustris at 510 C for 10 s. The distribution (Table 2) is dominated by C27–C32 hopenes and hopanes, although the largest peak is actually a
Table 1 Bacterial species subjected to Py–GC–MS Species
Strain
Phylum or class
Family
BHPs
References
Methylobacterium fujisawaense Methylosinus trichosporium Rhodopseudomonas palustris Nitrosomona europaea ÔAnacystis montana fo. minorÕ
SAL-7 OB3b
Alphaproteobacteria Alphaproteobacteria Alphaproteobacteria Betaproteobacteria Cyanobacteria
Methylobacteriaceae Methylocystaceae Bradyrhizobiaceae Nitrosomonadaceae None
5.6 mg/g 4.0 mg/g 2 mg/g >3.68 mg/g 220 lg/g
Knani et al. (1994) Neunlist and Rohmer (1985) Neunlist et al. (1988) Seemann et al. (1999) Herrmann et al. (1996)
Cyanobacteria
None
600 lg/g
Talbot, unpublished results
Cyanobacteria Actinobacteria Actinobacteria Actinobacteria Actinobacteria
Prochlorococcaceae Nocardiaceae Nocardiaceae Nocardiaceae Nocardiaceae
None None Unknown Unknown Unknown
Talbot, unpublished results Talbot, unpublished results
Synechocystis sp. Prochlorococcus marinus Rhodococcus rhodochrous Rhodococcus rhodnii Rhodococcus equi Rhodococcus erythropolis a b
CCAP 1405/3a CCAP 1480/4a MIT9313b
CCAP = Culture Collection of Algae and Protozoa (Scottish Association for Marine Science Research Services Ltd.). MIT = Massachusetts Institute of Technology.
M.A. Sugden et al. / Organic Geochemistry 36 (2005) 975–979 2
27
(a) R. palustris 3
9
14
6
10
4 8 57
1
14
18
3 13
10 6 9 13 11
24
11 12 15 16
23
17 19
21
4
13 6 9 11 14
26
4
24 27
3 46
3
27 22 or 23
6
16
14
9 11 16 17 19
22
24
22 or 23
3
19
(g) R. erythropolis
26
9 6 10 11 4
20
22
24
26
17 16 26
19
(i) P. marinus
(h) R. rhodochrous
28
27
2
27
4
26 24
(f) M. fujisawaense
14 10 14
20
16 17 19
(e) Synecocystis sp.
2
22 27 25
16 17 19
2
10 9 11
(c) M. trichosporium
2
(d) A. montana
20
10
2 3
(b) N. europaea
977
28
20
22
24
26
28
Retention time Fig. 1. m/z 191 Partial mass chromatograms for labelled bacterial species. Peak assignments are given in Table 2. Table 2 Identified hopanoid products from Py–GC–MS of R. palustris (Numbers refer to peaks in Fig. 1) Peak no.
Name
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27
C27 Hopene C27 Hop-17(21)-ene C27 17b Hopane C29 Hop-17(21)-ene C29 Hopene C29 17b, 21a Hopane C30 Hopene C29 Hopene C29 17b, 21b Hopane C29 Hopene C29 Hopene C30 Hopene C30 17b, 21b Hopane C30 17b, 21b Hop-22(29)-ene (diploptene) C30 17b, 21b Hop-21(22)-ene C31 17b, 21b Hopane C32 17b, 21b Hopane C30 17b, 21b Hopan-30-ol (diplopterol) C33 Hop-17(21)-ene C32 Hopan-31-one C35 Hopanefuran (I)a C34 Hopanefuran (II)a C33 Ene-ol or C33 hopanone Unknown nitrogen-containing hopanoid C35 Hopanefuran (isomer of 21) C35 Arylhopane (III)a C35 Hopanefuran (isomer of 21)
a
Structure – see Fig. 2.
C35 hopane furan (peak 27). On the basis of their mass spectra peaks 14 and 18 have been identified as diploptene and diplopterol (e.g., Philp, 1985), respectively, which are likely to have been volatilised on to the column during pyrolysis, as R. palustris is known to biosynthesise these components (Rohmer et al., 1984). The remaining C27–C30 hopenes and hopanes may be generated from the breakdown or structural rearrangement of diploptene and diplopterol or alternatively, together with the C31–C35 hopenes and hopanes, they may be degradation and rearrangement products of BHPs. Figs. 1b–i show the m/z 191 partial mass chromatograms for a further eight bacterial species subjected to Py–GC–MS. Some of these species were only analysed in SIM mode and therefore it has not been possible to identify all the hopane and hopene components in each case; however, many peaks have been identified by matching relative retention times to those of R. palustris or M. trichosporium (Figs. 1a, c and Fig. 2; Table 2). It was possible to identify hopane and hopene peaks in all of the hopanoid-containing bacteria (Figs. 1b–f), even those with relatively low hopanoid contents such as the cyanobacteria ÔAnacystis montanaÕ and Synechocystis sp. (Table 1; Figs. 1d–e). Five further species, two of which were known not to contain BHPs (R. rhodochrous and P. marinus; Talbot, unpublished results) from APCI-LC/MSn analysis following traditional solvent extraction and derivatisation (e.g., Talbot et al., 2003a), were also subjected to Py– GC–MS and analysed in both SIM and SCAN modes.
978
M.A. Sugden et al. / Organic Geochemistry 36 (2005) 975–979
32
O
31
35
I
O
34
II
III Fig. 2. Tentative structures of novel components identified in Table 2 based on interpretation of mass spectra.
No hopane or hopene components could be found in their m/z 191 partial mass chromatograms (three representative examples are shown in Fig. 1g–i). The unidentified compounds in the mass chromatograms of these non-hopanoid producing bacteria could not be matched (by either relative retention times or mass spectra) to the hopanes and hopenes generated by R. palustris or M. trichosporium (Fig. 1a), or to any of the other hopanoidcontaining bacteria (Fig. 1b–f). 4. Conclusions Flash pyrolysis of bacteria that synthesise BHPs generates hopanes and hopenes that can be easily identified in m/z 191 mass chromatograms, even for those strains of bacteria (e.g., ÔA. montanaÕ and Synechocystis sp.) with relatively low hopanoid contents. No hopanes or hopenes could be detected in the corresponding m/z 191 partial mass chromatograms of non-hopanoid producing bacteria. Py–GC–MS is therefore an efficient method for screening bacterial strains for hopanoid synthesis, and prevents unnecessary laboratory work on bacterial strains that lack hopanoids. Acknowledgements M.A.S. and H.M.T. thank the NERC for supportive grants. Professors M. Rohmer, C.J. Murrell and R.E. Summons, and Drs. J. Day and A. Ward and also Allison Coe are thanked for kindly providing the bacterial strains. Paul Donohoe (University of Newcastle) is thanked for valuable technical assistance with the Py– GC–MS as are Professor J.R. Maxwell and an anonymous reviewer for valuable comments. Associate Editor—J.R. Maxwell
References Farrimond, P., Fox, P.A., Innes, H.E., Miskin, I.P., Head, I.M., 1998. Bacterial sources of hopanoids in recent sediments: improving our understanding of ancient hopane biomarkers. Ancient Biomolecules 2, 147–166. Farrimond, P., Head, I.M., Innes, H.E., 2000. Environmental influence on the biohopanoid composition of recent sediments. Geochimica et Cosmochimica Acta 64, 2985–2992. Herrmann, D., Bisseret, P., Connan, J., Rohmer, M., 1996. Relative configurations of carbapseudopentose moieties of hopanoids of the bacterium Zymomonas mobilis and the cyanobacterium ÔAnacystis montanaÕ. Tetrahedron Letters 37, 1791–1794. Innes, H.E., Bishop, A.N., Head, I.M., Farrimond, P., 1997. Preservation and diagenesis of hopanoids in recent lacustrine sediments of Priest Pot, England. Organic Geochemistry 26, 565–576. Knani, M., Corpe, W.A., Rohmer, M., 1994. Bacterial hopanoids from the pink-pigmented facultative methylotrophs (PPFMs) and from green plant surfaces. Microbiology 140, 2755–2759. Love, G.D., Bowden, S.A., Jahnke, L.L., Snape, C.E., Campbell, J.G., Day, J.G., Summons, R.E., 2005. A catalytic hydropyrolysis method for the rapid screening of microbial cultures for lipid biomarkers. Organic Geochemistry 36, 63–82. Neunlist, S., Rohmer, M., 1985. The hopanoids of ÔMethylosinus trichosporiumÕ: Aminobacteriohopanetriol and Aminobacteriohopanetetrol. Journal of General Microbiology 131, 1363–1367. Neunlist, S., Bisseret, P., Rohmer, M., 1988. The hopanoids of the purple non-sulfur bacteria Rhodopseudomonas palustris and Rhodopseudomonas acidophila. European Journal of Biochemistry 171, 245–252. Ourisson, G., Rohmer, M., 1992. Geohopanoids: the most abundant natural products on earth. Accounts of Chemical Research 25, 398–402. Philp, R.P., 1985. Fossil Fuel Biomarkers: Applications and Spectra. Elsevier, Amsterdam, p. 293. Rohmer, M., 1993. The biosynthesis of the triterpenoids of the hopane series in the Eubacteria: a mine of new
M.A. Sugden et al. / Organic Geochemistry 36 (2005) 975–979 enzyme reactions. Pure and Applied Chemistry 65, 1293– 1298. Rohmer, M., Bouvier-Nave, P., Ourisson, G., 1984. Distribution of hopanoid triterpenes in prokaryotes. Journal of General Microbiology 130, 1137–1150. Seemann, M., Bisseret, P., Tritz, J.-P., Hooper, A.B., Rohmer, M., 1999. Novel bacterial triterpenoids of the hopane series from Nitrosomonas europaea and their significance for the formation of the C35 bacteriohopane skeleton. Tetrahedron Letters 40, 1681–1684. Talbot, H.M., Squier, A.H., Keely, B.J., Farrimond, P., 2003a. Atmospheric pressure chemical ionisation reversed-phase
979
liquid chromatography/ion trap mass spectrometry of intact bacteriohopanepolyols. Rapid Communications in Mass Spectrometry 17, 728–737. Talbot, H.M., Summons, R., Jahnke, L., Farrimond, P., 2003b. Characteristic fragmentation of bacteriohopanepolyols during atmospheric pressure chemical ionisation liquid chromatography/ion trap mass spectrometry. Rapid Communications in Mass Spectrometry 17, 2788– 2796. Talbot, H.M., Watson, D.F., Pearson, E.J., Farrimond, P., 2003c. Diverse biohopanoid compositions of non-marine sediments. Organic Geochemistry 34, 1353–1371.
Note
Flash pyrolysis – a rapid method for screening bacterial species for the presence of bacteriohopanepolyols Mark A. Sugden, Helen M. Talbot *, Paul Farrimond School of Civil Engineering and Geosciences, University of Newcastle, Newcastle Upon Tyne NE1 7RU, UK Received 9 November 2004; accepted 7 December 2004 (returned to author for revision 24 November 2004) Available online 22 February 2005
Abstract Flash pyrolysis–gas chromatography–mass spectrometry (Py–GC–MS) was applied to freeze dried cells of eleven bacterial species. Six species known to synthesise bacteriohopanepolyols (BHPs) were found to produce easily identifiable C27–C35 hopenes and hopanes upon flash pyrolysis. These compounds could not be observed in the pyrolysis products of five species, two of which are known not to produce BHPs. In the search for BHP-containing bacteria this rapid screening approach prevents unnecessary time-consuming laboratory work to characterise species that would produce negative results. 2005 Elsevier Ltd. All rights reserved.
1. Introduction Bacteriohopanepolyols (BHPs), such as bacteriohopane-32,33,34,35-tetrol, are triterpenoids synthesised by certain types of bacteria, including gram-negative bacteria, methanotrophs, cyanobacteria, acetic acid bacteria, nitrogen fixers and purple non-sulfur bacteria (Rohmer et al., 1984; Farrimond et al., 1998, and references therein). An ever increasing range of BHP structures, with a complex biological functionality at C-35, are being found (e.g., Rohmer, 1993; Talbot et al., 2003a,b), and these can vary significantly among bacterial groups/species. The ubiquitous occurrence of BHPs and their diagenetic products (e.g., hopanols, hopanoic acids, hopenes and hopanes) in recent sediments (e.g., Innes et al., 1997; Talbot et al., 2003c) shows the impor* Corresponding author. Tel.: +44 191 246 4886; fax: +44 191 246 4961. E-mail address: [email protected] (H.M. Talbot).
tance of the bacterial contribution to the biomarker pool in the sedimentary record (Ourisson and Rohmer, 1992). BHPs in sediments are a record of bacterial activity and can therefore provide an indication of the palaeoenvironmental conditions at the time of deposition (e.g., Farrimond et al., 2000). To fully understand this record it is necessary to establish which species of the bacterial population contain BHPs and the specific composition of these BHPs. To date, they have been found in approximately 50% of the bacteria examined specifically for their presence, but they are not equally distributed across the various bacterial taxa (Farrimond et al., 1998). The highly functionalised and amphiphilic BHPs are not readily amenable to analysis by conventional GC–MS (Innes et al., 1997). However, a new technique involving atmospheric pressure chemical ionisation liquid chromatography/ion trap mass spectrometry (APCI-LC/MSn) has recently been used to identify them in a range of bacteria and modern sediment samples (Talbot et al., 2003a,b). This work requires dual stage
0146-6380/$ - see front matter 2005 Elsevier Ltd. All rights reserved. doi:10.1016/j.orggeochem.2004.12.003
976
M.A. Sugden et al. / Organic Geochemistry 36 (2005) 975–979
solvent extraction techniques followed by derivatisation using acetic anhydride and pyridine (Talbot et al., 2003a). In order to better constrain the occurrence and composition of BHPs in bacteria it is necessary to perform this time consuming laboratory work on a diverse range of species from different taxa, with the knowledge that hopanoids would likely not be detected in a significant proportion of the species tested. Thus, a rapid technique to screen for the presence of hopanoids is needed. A catalytic hydropyrolysis method for the screening of microbial cultures has recently been shown to produce a series of C27–C35 hopanes and hopenes from hopanoid-containing bacteria (Love et al., 2005). Here, we report the results from the application of pyrolysis–gas chromatography–mass spectrometry (Py–GC–MS) to a range of bacterial species. The aim was to establish a rapid low-cost method, using facilities that are available worldwide, to detect whether a particular species contained BHPs or not. This knowledge, prior to solvent extraction for analysis using APCI-LC/MSn, prevents unnecessary laboratory work on species which do not contain hopanoids.
2. Experimental 2.1. Cultures The species, strain, phylum or class and family of bacterial cells subjected to Py–GC–MS are summarised in Table 1, together with their measured BHP concentrations. 2.2. Py–GC–MS Py–GC–MS was performed on freeze dried cells using a resistance-heated CDS 120 Pyroprobe unit, fitted
with a platinum coil pyroprobe, coupled to a HewlettPackard 5890GC with split/splitless injector (300 C), linked to a Hewlett-Packard 5972MSD (electron voltage 70 eV, filament current 200 lA, source temperature 180 C, multiplier voltage 2000 V, interface temperature 300 C). The acquisition was controlled with a HP Vectra 486 Chemstation computer, in either full scan (range m/z 50–700 amu/s), or selected ion mode (m/z 191, 250 ms dwell). The sample (<1 mg) was placed in a quartz tube plugged with pre-extracted silica wool at both ends. Pyrolysis was performed at 510 C for 10 s, after which the split was opened. Various pyrolysis temperatures were tested (360, 410, 460, 510, 560, 610 and 660 C), 510 C being found to give a consistently high hopanoid response for Rhodopseudomonas palustris. R. palustris was also subjected to Py–GC–MS at 510 C with TMAH, with no obvious benefits. The oven temperature was programmed from 50 C (2 min) to 350 C at 10 C/ min and then held for 10 min. He was used as carrier gas (flow 1 mL/min, pressure of 50 kPa, split at 30 mL/min). The pyrolysis products were separated on a fused silica column (15 m · 0.25 mm i.d.) coated with a 5% phenylmethylsilicone bonded stationary phase (DB5-HT, film thickness 0.1 lm). Hopanes and hopenes were identified from their mass spectra (R. palustris and Methylosinus trichosporium) with reference to published spectra and spectra obtained from standard oil samples.
3. Results and discussion Fig. 1a shows the m/z 191 partial mass chromatogram of the hopanoid degradation products released after flash pyrolysis of R. palustris at 510 C for 10 s. The distribution (Table 2) is dominated by C27–C32 hopenes and hopanes, although the largest peak is actually a
Table 1 Bacterial species subjected to Py–GC–MS Species
Strain
Phylum or class
Family
BHPs
References
Methylobacterium fujisawaense Methylosinus trichosporium Rhodopseudomonas palustris Nitrosomona europaea ÔAnacystis montana fo. minorÕ
SAL-7 OB3b
Alphaproteobacteria Alphaproteobacteria Alphaproteobacteria Betaproteobacteria Cyanobacteria
Methylobacteriaceae Methylocystaceae Bradyrhizobiaceae Nitrosomonadaceae None
5.6 mg/g 4.0 mg/g 2 mg/g >3.68 mg/g 220 lg/g
Knani et al. (1994) Neunlist and Rohmer (1985) Neunlist et al. (1988) Seemann et al. (1999) Herrmann et al. (1996)
Cyanobacteria
None
600 lg/g
Talbot, unpublished results
Cyanobacteria Actinobacteria Actinobacteria Actinobacteria Actinobacteria
Prochlorococcaceae Nocardiaceae Nocardiaceae Nocardiaceae Nocardiaceae
None None Unknown Unknown Unknown
Talbot, unpublished results Talbot, unpublished results
Synechocystis sp. Prochlorococcus marinus Rhodococcus rhodochrous Rhodococcus rhodnii Rhodococcus equi Rhodococcus erythropolis a b
CCAP 1405/3a CCAP 1480/4a MIT9313b
CCAP = Culture Collection of Algae and Protozoa (Scottish Association for Marine Science Research Services Ltd.). MIT = Massachusetts Institute of Technology.
M.A. Sugden et al. / Organic Geochemistry 36 (2005) 975–979 2
27
(a) R. palustris 3
9
14
6
10
4 8 57
1
14
18
3 13
10 6 9 13 11
24
11 12 15 16
23
17 19
21
4
13 6 9 11 14
26
4
24 27
3 46
3
27 22 or 23
6
16
14
9 11 16 17 19
22
24
22 or 23
3
19
(g) R. erythropolis
26
9 6 10 11 4
20
22
24
26
17 16 26
19
(i) P. marinus
(h) R. rhodochrous
28
27
2
27
4
26 24
(f) M. fujisawaense
14 10 14
20
16 17 19
(e) Synecocystis sp.
2
22 27 25
16 17 19
2
10 9 11
(c) M. trichosporium
2
(d) A. montana
20
10
2 3
(b) N. europaea
977
28
20
22
24
26
28
Retention time Fig. 1. m/z 191 Partial mass chromatograms for labelled bacterial species. Peak assignments are given in Table 2. Table 2 Identified hopanoid products from Py–GC–MS of R. palustris (Numbers refer to peaks in Fig. 1) Peak no.
Name
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27
C27 Hopene C27 Hop-17(21)-ene C27 17b Hopane C29 Hop-17(21)-ene C29 Hopene C29 17b, 21a Hopane C30 Hopene C29 Hopene C29 17b, 21b Hopane C29 Hopene C29 Hopene C30 Hopene C30 17b, 21b Hopane C30 17b, 21b Hop-22(29)-ene (diploptene) C30 17b, 21b Hop-21(22)-ene C31 17b, 21b Hopane C32 17b, 21b Hopane C30 17b, 21b Hopan-30-ol (diplopterol) C33 Hop-17(21)-ene C32 Hopan-31-one C35 Hopanefuran (I)a C34 Hopanefuran (II)a C33 Ene-ol or C33 hopanone Unknown nitrogen-containing hopanoid C35 Hopanefuran (isomer of 21) C35 Arylhopane (III)a C35 Hopanefuran (isomer of 21)
a
Structure – see Fig. 2.
C35 hopane furan (peak 27). On the basis of their mass spectra peaks 14 and 18 have been identified as diploptene and diplopterol (e.g., Philp, 1985), respectively, which are likely to have been volatilised on to the column during pyrolysis, as R. palustris is known to biosynthesise these components (Rohmer et al., 1984). The remaining C27–C30 hopenes and hopanes may be generated from the breakdown or structural rearrangement of diploptene and diplopterol or alternatively, together with the C31–C35 hopenes and hopanes, they may be degradation and rearrangement products of BHPs. Figs. 1b–i show the m/z 191 partial mass chromatograms for a further eight bacterial species subjected to Py–GC–MS. Some of these species were only analysed in SIM mode and therefore it has not been possible to identify all the hopane and hopene components in each case; however, many peaks have been identified by matching relative retention times to those of R. palustris or M. trichosporium (Figs. 1a, c and Fig. 2; Table 2). It was possible to identify hopane and hopene peaks in all of the hopanoid-containing bacteria (Figs. 1b–f), even those with relatively low hopanoid contents such as the cyanobacteria ÔAnacystis montanaÕ and Synechocystis sp. (Table 1; Figs. 1d–e). Five further species, two of which were known not to contain BHPs (R. rhodochrous and P. marinus; Talbot, unpublished results) from APCI-LC/MSn analysis following traditional solvent extraction and derivatisation (e.g., Talbot et al., 2003a), were also subjected to Py– GC–MS and analysed in both SIM and SCAN modes.
978
M.A. Sugden et al. / Organic Geochemistry 36 (2005) 975–979
32
O
31
35
I
O
34
II
III Fig. 2. Tentative structures of novel components identified in Table 2 based on interpretation of mass spectra.
No hopane or hopene components could be found in their m/z 191 partial mass chromatograms (three representative examples are shown in Fig. 1g–i). The unidentified compounds in the mass chromatograms of these non-hopanoid producing bacteria could not be matched (by either relative retention times or mass spectra) to the hopanes and hopenes generated by R. palustris or M. trichosporium (Fig. 1a), or to any of the other hopanoidcontaining bacteria (Fig. 1b–f). 4. Conclusions Flash pyrolysis of bacteria that synthesise BHPs generates hopanes and hopenes that can be easily identified in m/z 191 mass chromatograms, even for those strains of bacteria (e.g., ÔA. montanaÕ and Synechocystis sp.) with relatively low hopanoid contents. No hopanes or hopenes could be detected in the corresponding m/z 191 partial mass chromatograms of non-hopanoid producing bacteria. Py–GC–MS is therefore an efficient method for screening bacterial strains for hopanoid synthesis, and prevents unnecessary laboratory work on bacterial strains that lack hopanoids. Acknowledgements M.A.S. and H.M.T. thank the NERC for supportive grants. Professors M. Rohmer, C.J. Murrell and R.E. Summons, and Drs. J. Day and A. Ward and also Allison Coe are thanked for kindly providing the bacterial strains. Paul Donohoe (University of Newcastle) is thanked for valuable technical assistance with the Py– GC–MS as are Professor J.R. Maxwell and an anonymous reviewer for valuable comments. Associate Editor—J.R. Maxwell
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