Journal of General Microbiology (1984), 130, 1 137-1 150. Printed in Great Britain
1137
Distribution of Hopanoid Triterpenes in Prokaryotes
By M I C H E L R O H M E R , ’ * P I E R R E T T E B O U V I E R - N A V E Z
AND G U Y OURISSON3
EcoIe Nationale Supkrieure de Chimie de Mulhouse, UniversitP de Haute Alsace, 3 rue Alfred
Werner, 68093 Mulhouse Cedex, France
Institut de Botanique, Universitk Louis Pasteur, 28 rue Goethe, 67083 Strasbourg Cedex,
France
Centre de Neurochimie, Universitg Louis Pasteur, 5 rue Blaise Pascal, 67084 Strasbourg Cedex,
France
(Received 3 October 1983; revised 4 January 1984)
Pentacyclic triterpenoids of the hopane family were found in about half of some 100 strains of
prokaryotes belonging to diverse taxonomic groups, such a wide distribution indicating the
biological significance of these compounds. Hopanoids were found in almost all the
cyanobacteria and obligate methylotrophs examined, in all the purple non-sulphur bacteria
studied and in many taxonomically diverse Gram-negative or Gram-positive chemoheterotrophs. They were absent in all archaebacteria and purple sulphur bacteria examined as well as
in various other Gram-positive or Gram-negative genera. The C30hopanoids, diploptene and
diplopterol, are present in almost all hopanoid-containing prokaryotes. The major compounds
are always the C35 bacteriohopanepolyols, which are present at a level of 0-1-2 mg per g dry
weight, the most common one being bacteriohopanetetrol. Because of their structural
characteristics and their influence on the properties of biological membrane models, these
compounds might be the structural equivalents of the sterols found in eukaryotes.
INTRODUCTION
Of the many diverse families of pentacyclic triterpenes (Devon & Scott, 1972)most have been
isolated from widely scattered taxonomic groups. For example, derivatives of a given triterpene
skeleton have been found in isolated plants, in the secretion of one family of trees, in some ferns,
mosses or lichens, or in the cuticle of some grasses. Such triterpenes have been considered, like
many secondary plant metabolites, to be ‘evolutionary molecular frills’ possibly with ecological,
but without obvious physiological significance. This was precisely the case, until recently, for
the hopane family of triterpenes (Forster et al., 1973; Ourisson et al., 1979a, 6).
We have shown that ‘hopanoids’ (see Figs 1 and 2) are among the most widespread of all
complex natural products (Van Dorsselaer et al., 1977; Ourisson et al., 1979a, b), and that they
are probably essential constituents of many prokaryotes (Rohmer & Ourisson, 1976b ; Rohmer et
al., 1979). The first indication of their significance was the discovery of the ubiquity of their
molecular fossils in all sediments, pointing to a wide distribution over space and time, and
suggesting a bacterial origin (Ourisson et al., 1979a). The simplest hopanoid, diploptene (Fig. 2,
IV) had been found previously in three cyanobacteria (Gelpi et al., 1970)and two other bacteria
(Bird et al., 1971 ;De Rosa et al., 197l), and complex hopanoids, the bacteriohopanepolyols, had
been isolated and identified in ‘Acetobacter xylinum’ (Forster et al., 1973; Rohmer & Ourisson,
1976a) and in Bacillus acidocaldarius (Langworthy et al., 1976; Langworthy & Mayberry, 1976).
Hopanoids are structurally similar to sterols in their molecular dimensions and amphiphilic
character. Their biosynthesis is similar to that of sterols, from which it diverges by being fully
anaerobic (Rohmer et al., 1979) and by involving apparently a more primitive cyclase to form
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M. ROHMER, P . BOUVIER-NAVE A N D G . OURISSON
the polycyclic structure (Anding et al., 1976; Rohmer et al., 1980a, b; Bouvier et al., 1980).
Despite repeated reports of sterols in bacteria, it must be recognized that these derivatives are
generally absent in prokaryotes (Asselineau, 1962; Goldfine, 1972; Bouvier, 1978), at least in
amounts comparable to those found in eukaryotes. In two cases only, Methylococcus capsulatus
(Bird et al., 1971; Bouvier et al., 1976; Rohmer et al., 1980b) and Nannocystis exedens (Kohl et
al., 1983), have larger amounts of sterols been detected, and their de novo biosynthesis
unambiguously demonstrated.
We have postulated that hopanoids are phylogenetic sterol ancestors, acting as membrane
reinforcers, as sterols do in the membranes of eukaryotes (Rohmer et al., 1979; Ourisson &
Rohmer, 1982). For instance, tetrahymanol, a hopanoid-like triterpene, can replace the sterols
in the membranes of the protozoon Tetrahymenapyriformis, depending on the culture' conditions
(Conner et al., 1968).The growth of the sterol-requiring Mycoplasma mycoidessubsp. capriis also
supported by a hopanoid, diplopterol (Kannenberg & Poralla, 1982). Various hopanoids have
been shown to induce an effect similar to that of cholesterol on artificial phospholipid
membrane models (Poralla et al., 1980; Bisseret, 1982; Bisseret et al., 1983). Furthermore
bacteriohopanetetrol and diplopterol induce an orientation of the cellulose microfibrils
produced by 'Acetobacter xylinum' (Haigh et al., 1973).
Hopanoids therefore represent a major lipid family of probable physiological significance;
this paper provides details of the analysis of about 100 strains of prokaryotes for the possible
presence of hopanoids.
METHODS
Strains and growth ofbacteria. Details of the bacteria studied are included in Tables 1 and 2. Some were grown in
our laboratory and others received as freeze-dried samples. Names of organisms not on the Approved Lists of
Bacterial Names (Skerman et a f . , 1980) are shown in inverted commas.
Analytical methods. Gas-liquid chromatography (GLC), gas-liquid chromatography/mass spectrometry (GLC/
MS) and the radioactivity measurements were carried out as described by Rohmer et al. (1980a). The quantity of
each product, down to the pg level, was measured by GLC by comparing the peak areas with that of an internal
standard of n-dotriacontane in the case of diplopterol trimethylsilyl ether and bacteriohopane derivatives, or of nhexatriacontane in the case of diploptene. Diplopterol (Fig. 2, V) was silylated using N,O-bis(trimethylsily1)trifluoroacetamide (20 pl) in dry pyridine (20 pl) for 30 min at room temperature.
Isolation and pur9cation ofhopanoids. Freeze-dried cells (0.5-2 g, depending on the experiment) were extracted
twice using chloroform/methanol(50 ml, 2 : 1, v/v) under reflux for 1 h. The combined extracts were evaporated to
dryness and treated with periodic acid (H5106)by one or both of the following procedures.
In procedure 1, the chloroform/methanol extract was hydrolysed for 1 h under reflux in 6% (w/v) methanolic
KOH ( 5 ml). After addition of water (10 ml), the non-saponifiable lipids were extracted three times with diethyl
ether (10 ml). The solution was dried over anhydrous Na,SO, and evaporated to dryness under vacuum after
filtering off the drying agent. To convert the bacteriohopanepolyols into convenient derivatives, the nonsaponifiable lipids were treated for 1 h at room temperature with a solution (0.5-2 ml) of periodic acid (100 mg) in
tetrahydrofuran/water (5 ml, 95 :5, v/v). The ice-cold reaction mixture was then added dropwise to a suspension of
NaBH, (100 mg) in ethanol (2 ml) under vigorous stirring and cooling at 0 "C, and stirred for 1 h at room
temperature. After addition of water (10 ml), the hopanoids were extracted three times with petroleum ether
(10 ml, b.p. 40-60 "C).As this first procedure was time-consuming and resulted in poor yields in some cases (e.g.
Rhodospirilfumrubrum or cyanobacteria of the LPP group: Table 3) because of the basic hydrolysis, the following
modified procedure was developed.
In procedure 2, the crude chloroform/methanol extract was directly treated by stirring at room temperature for
1 h with a solution of H5106(300 mg) in tetrahydrofuran/water (3 ml, 8 :1, v/v). After addition of water (10 ml),
the lipids were extracted three times with petroleum ether (10 ml), and the solution was dried over anhydrous
Na,SO, and evaporated to dryness. The residue was reduced by stirring for 1 h at room temperature with an excess
of NaBH, (100 mg) in ethanol (3 ml). After addition of a solution of KH2PO4(15 ml, 100 mM), the hopanoids were
extracted as previously described with petroleum ether.
The reaction mixture obtained after either of the above H510,/NaBH, treatments was separated by TLC using
a double development with dichloromethane on Merck H F 254 (0.25 mm) silica gel plates into hydrocarbons
containing diploptene (Fig. 2, IV) and squalene (RF = 0.79), diplopterol (V, RF = 0.26), (22 S)-bacteriohopane
derivatives (VIII, RF = 0.19) and (22 R)-bacteriohopane derivatives (IX-XIV, RF = 0.15). After spraying with a
0.1 % alcoholic solution of berberin chlorhydrate, the bands were visualized under UV light (366 nm) and scraped
off; the triterpenoids were recovered from silica gel using dichloromethane.
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Hopanoids in prokaryotes
1139
The hydrocarbons were further purified by TLC using cyclohexane as eluent into diploptene (IV, RF = 0.53)
and squalene (AF= 0.25). Primary alcohols were acetylated overnight at room temperature using acetic anhydride
(50 pl) in dry pyridine (50 pl), dry toluene (up to 200 pl) being sometimes added to solubilize the hopanoids
completely. Excess reagent was removed with a stream of NZ,and the acetates were purified by TLC
(cyclohexane/ethyl acetate, 90 : 10, RF = 0.52).
RESULTS
Choice of an analytical procedure and structure determinations
The analytical method had to be rapid and sensitive, but it also had to give incontrovertible
proof of the presence of hopanoids by actual isolation and characterization. No particular
difficulty was encountered with the simplest hopanoids (Fig. 2) diploptene (IV) and diplopterol
(V) as they are readily extracted from freeze-dried cells by organic solvents, and can be
characterized by GLC and by combined GLC/MS. This is not the case for the more complex
hopanoids, the bacteriohopanepolyols (Fig. 1, I-111), whose extraction and TLC analysis is
hampered by their high polarity and low solubility in any organic solvents. The high molecular
weights of these latter compounds make it difficult to analyse them by GLC or GLC/MS. In five
cases, Acetobacter aceti ssp. xylinum, Nostoc muscorum, Methylobacterium organophilum,
Rhodopseudomonas acidophila and Bacillus acidocaldarius, we have isolated the polyols and
determined their structures by unambiguous chemical and physical methods (Rohmer, 1975;
Rohmer & Ourisson, 1976a, b, c; M. Rohmer & J. M. Renoux, unpublished results).
In all other cases our standard procedure was the degradation of the polyhydroxylated sidechains: H5106oxidation gave aldehydes which were reduced by NaBH, into primary alcohols
(Fig. 1). These alcohols (Fig. 2, VI-XV) were readily isolated and purified by TLC and the
corresponding acetates analysed by GLC and GLC/MS. In many analyses performed by
procedure 2 the H5IO6/NaBH, degradation releases large amounts of unidentified aliphatic
alcohols which cannot be separated by TLC from the primary triterpenic alcohols. However, the
presence of these compounds does not interfere with the analysis of the hopanoids by GLC, as
their retention times are much shorter. It must be remembered, however, that the
characterization of hopanoids after the oxidation-reduction sequence described does not allow
the detection of any variation in the side-chain. For instance, the isolation of the C32primary
alcohol (VIII) does not distinguish between the initial presence of C35tetrol isolated from the
strains studied in detail and mentioned above, and the possible presence in other strains of
higher homologues or conjugates. For our present purpose the limitations of the method are
unimportant, as it gives, with a good sensitivity (detection limit < 1 pg), positive proof for the
presence of the hopane skeleton. It is therefore assumed that all C32primary alcohols (VIII, IX,
XII, XIV) are derived from a tetrol identical or similar to the tetrols isolated by us from
Acetobacter aceti subsp. xylinum (Rohmer & Ourisson, 1976a) and the C31 alcohols (VII, XI,
XIII) from one of the pentols isolated from Nostoc muscorum (Rohmer, 1975; Rohmer &
Ourisson, 19766). The assignment of the C-22 configuration is tentatively based upon NMR
spectroscopy and steric considerations (Rohmer & Ourisson, 1976a). The two diastereoisomers
(VIII and IX) are separated both by TLC on silica gel and by GLC; these two methods were used
to determine this configuration. In a few cases (methylotrophs, purple non-sulphur bacteria), the
results have been confirmed by proton NMR spectroscopy. The presence of C30-typehopanoids
(X and XIII), obtained after H5IO6/NaBH, treatment of extracts of Methylococcus capsulatus or
type I methylotrophs, is unusual since they are not present in the intact chloroform/methanol
extract and are only released after degradation of a polyhydroxylated precursor of unknown
structure.
Distribution of hopanoids in prokaryotes
The bacteria containing no detectable hopanoids are listed in Table 1, and the hopane
producers in Table 2. The hopanoid compositions of the hopanoid-containing strains are shown
in Table 3 ; those of the Acetobacter strains containing very complex and peculiar
bacteriohopanepolyol mixtures are listed separately in Table 4. Hopanoids have been identified
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M. ROHMER, P. BOUVIER-NAVE A N D G . OURISSON
OH OH
____)
d
=
r
O
"
VIII, IX, XII. XIV
-
I1
?OH
VII, XI, XI11
OH
OH OH OH
Fig. 1. Structural variations in the bacteriohopanepolyol side-chains and their degradation to
simplified products (see Fig. 2).
by oiher authors in a few other strains: diploptene (IV) has been found in the cyanobacteria
Chroococcus,Lyngbya and Nostoc (Gelpi et al., 1970) and in Methylococcus capsulatus (Bird et al.,
1971), bacteriohopanetetrol (I) and diploptene (IV) in Bacillus acidocaldarius (De Rosa et al.,
1971; Langworthy et al., 1976) and in other thermoacidophilic bacilli (Hippchen et al., 1981),
diploptene (IV) and diplopterol (V) in a Pseudomonas sp. (Natori et al., 19Sl), and
methylhopanols of uncertain structures in Rhodomicrobium vannielii (Howard & Chapman,
1981). The most widespread bacteriohopanepolyol side-chain contained four hydroxyl groups (I)
but the pentols (11) have been found in most cases, at least as minor components. Extra methyl
groups were found at C-3 in most Acetobacter species and in Methylococcus capsulatus (Rohmer
& Ourisson, 1976c), or in a yet unknown position on rings A or B of the bacteriohopanepolyols
from the cyanobacteria of the Nostoc group or methyldiplopterol from Methylobacterium
organophilum. Additional double bonds were found at C-6 in some scattered strains, especially
in the Acetobacter strains, and at C-6 and/or C-11 (Rohmer & Ourisson, 19766; Rohmer, 1975).
Usually only one diastereoisomer at C-22 was present; the (22R)-diastereoisomer was the most
abundant, with substantial amounts of the other diastereoisomer isolated only from the
Acetobacter species. Structural features that are still unresolved and are currently under
investigation include the stereochemistry of the polyhydroxylated side-chain, the location of the
extra methyl group in the Nostoc and Methylobacterium hopanoids and the structures of the
hopanoids from Methylococcus capsulatus.
Hopanoid biosyn thesis
Labelling experiments were performed in vivo with selected prokaryotes using various
precursors. The incorporation rates were rather modest, but they prove unambiguously for the
first time that prokaryotes are capable of synthesizing polycyclic triterpenoids de novo (Table 5).
[ 1- 4C]Acetate was incorporated into the hopanoids of Nostoc rnuscorum and Methylococcus
capsulatus and [5-3H]riboseor [methyl-14C]methionineinto those of Acetobacter pasteurianus. In
the last case the position of the labelling has not been determined, but we assume that it is on the
methyl group at C-3.
DISCUSSION
From the present screening it appears that hopanoids are present in about 50 of the
approximately 100 examined. They were found in many prokaryotes belonging to various
taxonomic groups, and this wide distribution emphasizes the importance of this triterpene
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Hopanoids in prokaryotes
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M. ROHMER, P. BOUVIER-NAVE A N D G . OURISSON
Table 1. Bacteria containing no detectable hopanoids
Organism*
A. Cyanobacteria
Spirulina sp.t
Synechococcus sp. ATCC 27144t
Synechococcus sp. L 1402-17
B. Purple sulphur and green sulphur bacteria
Amoebobacter roseus 661 1
Chlorobium limicola
Chlorobium limicola var. ‘thiosulfatophilum’6230
Chromatium vinosum D
Ectothiorhodospira mobilis 8 115
Ectothiorhodospira shaposhnikovii Moskau N 1
Thiocapsa roseopersicina 63 11
Thiocystis violacea 231 1
C. Gram-negative chemoautotrophs and
chemoheterotrop hs
Agrobacterium tumefaciens CIP 671
Caulobacter crescentus CIP 7715
Erwinia herbicola NCIB 9680
Escherichia coli CIP K 1212000
Flexithrix QQ-1
‘Methylomonas Clara’ Hoechst
‘Methylophilus rnethylotrophus’ ICI
‘Moraxella displex non liquefaciens’ CIP 5545
‘Moraxella lwoffi’ CIP 5382
Paracoccus denitrificans DSM 38 1-65b
Proteus mirabilis CIP A235
Pseudomonas acidovorans ATCC 17046
Pseudomonas aeruginosa ATCC 15692
Pseudomonas chlororaphis ATCC 9446
Pseudomonas diminuta CIP 7129
Pseudomonas jlmrescens
‘Pseudomonas maltophilia’ ATCC 17445
Pseudomonas stutzeri ATCC 17588
Rhizobium lupini CIP 6357
Thiobacillus A2
Thiobacillus thioparus NCIB 8370
Xanthomonas campestris CIP 7423
D. Gram-positive bacteria
Actinoplanes brasiliensis ATCC 25844
Bacillus subtilist
Brevibacterium linens CIP 6372
Clostridium paraputrificum ATCC 25780
Desulfovibrio desulfuricans NCIB 8310
‘Micrococcusjlavus’ CIP 53160
Micromonospora sp. Roche 2207-85
Streptococcusfaecalis CIP 761 17
‘Sporosarcina lutea’ CIP 5345
Source and growth conditions
J. P. Van de Casteele, Institut Franqais du Pktrole, Rueil
Malmaison, France
R. Y. Stanier, Institut Pasteur, Paris, France (Stanier &
Cohen-Bazire, 1977)
W. Koch, Pflanzenphysiologisches Institut, Gottingen,
FRG; grown in medium of Kratz & Myers (1955)
according to Brandt et al. (1970)
N. Pfennig, Universitat Konstanz, FRG ; grown according
to Pfennig (1977)
R. Y. Stanier, Institut Pasteur, Paris, France
As for A . roseus
As for A . roseus
H. Truper, Universitat Bonn, FRG
As for E. mobilis
As for A . roseus
As for A . roseus
Grown for 24 h at 30 “C in Difco bactotryptone (10 g 1- l ) ,
Difco yeast extract (5 g 1- l ) and NaCl(5 g 1- l ) with initial
pH 7.3
Grown for 24 h at 28 “C in Difco bactopeptone (2 g 1- l ) ,
Difco yeast extract (1 g 1-l) and MgS0,.7H20 (0.2 g 1-l)
with initial pH 7.3
Grown for 24 h at 30 “C in Difco bactopeptone (10 g 1-I)
and Difco yeast extract (10 g 1-l) with initial pH 7.3
Growth conditions as for Agrobacterium
R. Levin, University of California, San Diego, USA
P. Prave, Hoechst AG, Frankfurt/Main, FRG
J. McNairney, Imperial Chemical Industries, Billingham,
UK
Growth conditions as for Agrobacterium
Growth conditions as for Agrobacterium
P. Vignais, Universitt de Grenoble, France
Growth conditions as for Agrobacterium
N. Palleroni, Hoffmann-La Roche, Nutley, NJ, USA
J. M. Meyer, Universitt Louis Pasteur, Strasbourg, France
As for P . aeruginosa
Growth conditions as for Agrobacterium
As for P . aeruginosa (strain isolated by B. Wurtz, UniversitC
Louis Pasteur, Strasbourg, France)
As for P . aeruginosa
As for P . aeruginosa
Growth conditions as for Agrobacterium
Grown on acetate medium by J. C. Gottschal, Biologisch
Centrum, Groningen, The Netherlands
Grown for 72 h at 30 “C in chemoautotrophic thiobacilli
medium of Adachi & Suzuki (1977)
Growth conditions as for Agrobacterium
N. Palleroni, Hoffmann-La Roche, Nutley, NJ, USA
Strain from B. Wurtz, Universitt Louis Pasteur, Strasbourg,
France ; growth conditions as for Agrobacterium
A. Fauve, Universitt de Clermont-Ferrand, France
As for B. linens
J. Le Gall, CNRS, Marseille, France
Growth conditions as for Agrobacterium
As for A . brasiliensis
Growth conditions as for Agrobacterium
Growth conditions as for Agrobacterium
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Hopanoids in prokaryotes
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Table 1. (continued)
Organism*
Source and growth conditions
E. Archaebacteria
Halobacterium cutirubrum NRCCC 54001
Methanobacterium thermoautotrophicum
‘Sarcina littoralis’ NRCCC 16006
SulJolobus acidocaldarius 98-3
Thermoplasma acidophilum 122-1B2
M. Kates, University of Ottawa, Canada
R. S. Wolfe, University of Illinois, Urbana, USA
As for H. cutirubrum
D. Searcy, University of Massachusetts, Amherst, USA ;
type strain isolated by D. Brock
As for S . acidocaldarius
* Abbreviations : ATCC, American Type Culture Collection; CIP, Collection de 1’Institut Pasteur ; DSM,
Deutsche Sammlung fur Mikroorganismen; NCIB, National Collection of Industrial Bacteria; NRCCC,
National Research Council of Canada Collection.
t Results obtained by procedure 1 (see Methods). All other results were obtained by procedure 2.
Table 2. Bacteria containing hopanoids
Source and growth conditions
Organism*
A. Cyanobacteria
Anabaena variabilis
Calothrix sp. ATCC 27914
Fischerella sp. ATCC 29174
LPP ATCC 27902
LPP ATCC 27984
Nostoc sp. ATCC 27985
Nostoc muscorum B 1452-12b
Oscillatoria sp. ATCC 27935
Scytonema sp. ATCC 29171
Synechocystis ATCC 27 170
Synechocystis ATCC 27 178
B. Purple non-sulphur bacteria
Rhodomicrobiurn vannielii RM5
Rhodopseudomonas acidophila 7050
Rhodopseudomonas acidophila 10050
Rhodopseudomonas palustris
Rhodospirillum rubrum Ha
Rhodospirillum rubrum
C. Methylotrophs
Type I ‘Methylomonas albus’ BG8
‘Methylomonas rnethanica’ SI
‘Methylomonas’ sp. N 1D 1
Type I1 ‘Methylocystisparvus’ OBBP
‘Methylosinus sporium’ 5
‘Methylosinus trichosporium’ PG
‘Methylosinus trichosporiurn’ OB3b
Type X Methylococcus capsulatus MC
Methylococcus capsulatus TRMC
D. Gram-negative chemoautotrophs
Nitrosomonas europaea
1
Strain (Kratz & Myers, 1955) from J. G. Carr, University
of Bristol, UK; grown according to Brandt et al. (1970)
in medium of Kratz & Myers (1955)
R. Y. Stanier, Institut Pasteur, Paris, France
W. Koch, Pflanzenphysiologisches Institut, Tubingen,
FRG; grown according to Brandt et al. (1970) in
medium of Kratz & Myers (1955)
R. Y. Stanier, Institut Pasteur, Paris, France
R. Whittenbury, University of Warwick, UK ; strain
grown according to Whittenbury & Dow (1977)
N. Pfennig, Universitat Konstanz, FRG
R. Y. Stanier, Institut Pasteur, Paris, France
R. Whittenbury, University of Warwick, UK ; grown
according to Whittenbury et al. (1970)
Strain (Hazeu, 1972) obtained from W. Hazeu, Delft
University of Technology, The Netherlands
R. Whittenbury, University of Warwick, UK ; grown
according to Whittenbury et al. (1970)
I. H. Higgins, University of Canterbury, UK
R. Whittenbury, University of Warwick, UK ; grown
according to Whittenbury et al. (1970)
Strain from N. Walker, Rothamsted Experimental
Station, Harpenden, UK; grown at 28 “C for 5 d in a
200 1 fermenter by Hoffmann-La Roche, Basel,
Switzerland, using ATCC medium Nb 221 to yield 1 g
dry cells
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M. ROIIMER, P . BOUVIER-NAVE A N D G . OURISSON
Table 2 (continued)
Organism*
E. Gram-negative chemoheterotrophs
Acetobacter aceti subsp. aceti NCIB 8621
Acetobacter aceti subsp. liquefaciens NCIB 941 8
Acetobacter aceti subsp. xylinum NCIB 41 12
Acetobacter aceti subsp. xylinum Roche 2277
Acetobacter pasteurianus subsp. estunensis
NCIB 8935
Acetobacter pasteurianus subsp. lovaniensis
NCIB 8620
Acetobacter pasteurianus subsp. 'orleanensis'
NCIB 6426
Acetobacter pasteurianus subsp. pasteurianus
NCIB 6429
Acetobacter pas teurianus subsp. pasteurianus
NCIB 8856
Acetobacter peroxydans NCIB 8087
Acetobacter peroxydans NCIB 8618
Azotobacter chroococcum CIP
Azotobacter vinelandii CCM 289
Gluconobacter oxydans subsp. oxydans
NCIB 9013
Hyphomicrobium sp. X
Methylobacterium organophilum
'Pseudomonas cepacia' Berkeley 382
F. Gram-positive chemoheterotrophs
Bacillus acidocaldarius Brock 104-1A
Streptomyces chartreusis NRLL 3882
Streptomyces sp. Hoechst G 1815
Streptomyces sp. Hoechst 4-6609
Source and growth conditions
Grown for 24 h at 30 "C in the medium of Hestrin &
I
J
A. Kaiser, Hoffmann-La Roche, Basel, Switzerland
Grown for 24 h at 30 "C in the medium of Hestrin &
Schramm (1954)
Grown for 96 h at 28 "C on modified Burck medium with
(NH&S04 as nitrogen source (Winogradski, 1949)
J. M. Meyer, Universite Louis Pasteur, Strasbourg,
France
Grown for 24 h at 30 "C in the medium of Hestrin &
Schramm (1954)
Grown on dimethylamine medium by J . C. Gottschal,
Biologisch Centrum, Groningen, The Netherlands
Type strain (Patt & Hanson, 1978)from R. Whittenbury,
University of Warwick, UK; grown according to
Whittenbury et al. (1970) in AMS medium containing
0.5% (v/v) methanol
N. Palleroni, Hoffmann-La Roche, Nutley, NJ, USA
K. Poralla, Universitat Tubingen, FRG
A. Faure, Universite de Clermont-Ferrand, France
}P.
Prave, Hoechst AG, Frankfurt/Main, FRG
* Abbreviations not defined in Table 1 : CCM, Czechoslovak Collection of Micro-organisms; LLP, LyngbyaPhormidium-Plectonema, a provisional group of cyanobacteria of uncertain affinity (Stanier & Cohen-Bazire,
1977); NRRL, Northern Regional Research Laboratory.
family. As bacteriohopanepolyols have never been described so far in any eukaryote, even in
C,,-hopanoid-containing eukaryotes like ferns or lichens (P. Bouvier-Nave & M. Rohmer,
unpublished results), they have to be considered as typical prokaryotic metabolites. We have
proved that these compounds are synthesized de nouo by the micro-organisms, for instance from
acetate. Furthermore we have shown for the first time that cell-free systems prepared from the
bacteria Acetobacter pasteurianus and Methylococcus capsulatus catalyse the cyclization of
squalene into diploptene (IV) and diplopterol (V) (Anding et al., 1976; Rohmer et al., 1980a, b).
In most cases the hopanoid content of bacteria is of the same order of magnitude (0.1-2 mg
per g dry weight) as the sterol content of eukaryotic cells. This conclusion requires qualification
since the yields varied depending on which of the two analytical procedures was employed
(Tables 3 and 4). The first method, which involves a basic hydrolysis before the H5106oxidation
proved satisfactory for the Acetobacter strhi'ns, but led to very poor yields in the case for instance
of Rhodospirillum rubrum or cyanobacteria. In the second method the crude extract is directly
oxidized with H510,, and the hopanoid yield is 20-fold greater, starting from the same freezedried cells of Rhodospirillum rubrum or o f the cyanobacterium of the LPP group (Table 3). This
means that more than 9 0 x of the hopanoids are lost during the first procedure. Acetobacters
apparently contain mostly free polyols, but in other prokaryotes bacteriohopanepolyols may be
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Hopanoids in prokaryotes
1145
Table 3 . Hopanoid composition of hopanoid-containing strains (other than acetic acid bacteria)
d,
m
a
s
.e
22
v)
3 5
m a
(Pg g-’)
Organism
A. Cyanobacteria
Anabaena cariabilist
Calothrir sp. ATCC 279147
Fischerella sp. ATCC 19174t
LPP ATCC 27902t#
LPP ATCC 27984t#
LPP ATCC 27984#
Nostoc ATCC 27985
Nostoc muscorum B 1453-12btg
Oscillatoria sp. ATCC 27935t
Scytonema sp. ATCC 291711
Synechocystis ATCC 27 1707
Synechocystis ATCC 27 178t
B. Purple non-sulphur bacteria
Rhodomicrobium vannielii
Rhodopseudomonas acidophila 7050
Rhodopseudomonas acidophila 10050
Rhodopseudomonas palustris
Rhodospirillum rubrum?
Rhodospirillum rubrum Hat
Rhodospirillum rubrum Ha
C. Methylotrophs
Type I ‘Methylomonas albus’ BG8
‘Methylomonas methanica’ SI
‘Methylomonas’ N 1D 1
Type I1 ‘Methylocystisparous’ OBBP
‘Methylosinus sporium’ 5
‘Methylosinus trichosporium’ PG
‘Methylosinus trichosporium’ OB3bt
Type X Methylococcus capsulatus MC
Methylococcus capsulatus TRMC
D. Gram-negative chemoautotrophs
Nitrosomonas europaeat
E. Gram-negative chemoheterotrophs
Methylobacterium organophilum
Hyphomicrobiurn sp. X
‘Pseudomonas cepacia’ Berkeley 382
Azotobacter chroococcum CIP I
Azotobacter vinelandii CCM 289q
F. Gram-positive bacteria
Bacillus acidocaldarius Brock 104-1A
Streptomyces chartreusis N RLL 3882
Streptomyces sp. G 1815
Streptomyces sp. 4-6609
40
14
55
5
35
5
5
0.5
1
140
20
10
20
290
100
170
35
240
20
200
15
45
1
4800
1400
2000
2600
50
104
1650
120
15
100
230
1600
500
1
6
110
20
25
95
3000
5800
3400
1300
4200
4700
1200
1800
2900
10
350
+
+
185
20
50
3
f
22s
VIII
VI
VII
19
+
36:
19$
1
1
5
4
10
93
95
74
+
64
69
7
4
20
40
16
31
64
1
2
2500
55
20
9
900
45
2
60
1
55
96
100
81
100
100
100
64$
77$
100
100:
100
100
A6-IX X
4
4
100
99
99
95
100
96
90
+
1
7
60
84
69
36
3
1
97
100
88
3
35
10
4
IX
11
+
32
29
100
15
5 1001
20
1
3
22 R
+
140
30
60
4
3
65
250
130
3
110
60
110
110
10
50
10
5
35
20
Percentage composition of the
bacteriohopanepolyol fraction*
4
65
98
100
100
99
* Bacteriohopanepolyols are listed according to the alcohols obtained after H,IO,/NaBH, treatment (see Fig. 2 for structures).
The hopanoid containing a double bond at C-6 is listed under A6-IX.
t Results obtained by procedure 1 (see Methods). All other results were obtained by procedure 2.
$ Mixture of the normal hopanoid and its x-methyl homologue. These two compounds are not separated by GLC and were
detected by GLC/MS. In the case of Methylobacterium no diplopterol has been identified, but only x-methyldiplopterol.
0 This Nostoc contains a significant amount (35 pg g-l) of a pentol (111) giving a diol (XV) after H510,/NaBH, treatment.
II The polyol fraction was not analysed in Azotobacter chroococcum.
7 The hopanoid content was not estimated in Azotobacter vinelandii: as this micro-organism was grown in presence of CaC03,it
was impossible to determine the dry weight of the cells.
# The distinctions among genera of this subgroup of cyanobacteria are not clear. They have been placed in a provisional
category, the LPP (Lyngbya-Phormidium-Plectonema)group (Stanier & Cohen-Bazire, 1977).
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NCIB 901 3
180
40
140
40
85
10
25
165
20
30
30
290
40
40
620
200
100
540
40
170
200
60
60
280
550
720
1210
1250
3350
410
1680
1430
1490
4530
3100
1370
16
27
17
16
30
6
10
23
10
20
27
23
+
6
2
8
1
2
4
1
1
3
3
2
+
A6VII
+
1
1
5
+
1
1
4
2
+
1
1
VII
All-
+
VII
A6,11-
A
+
+
+
36 40
31 39
2 4 4 8 +
20 50
32 38
46 15
32 33
40 18
43 17
31 37
40 35
20 56
A6- A l l IX IX IX
2
+
5
+
IX
A6.11-
XI1
1
2
1
5
2
+ 2 0
5
10
5
XI
All-
3
10
2
6
1
4
2
4
2
XI1
A6-
Percentage composition of the bacteriohopanepolyol fraction*
2
1
+
1
+
3
1
XI1
8 1 1-
+
4
XI1
\
A6.1 I -
Bacteriohopanepolyolsare listed according to the primary alcohols obtained after HsIO,/N~BH, treatment (see Fig. 2 for structures). The hopanoids containing a double bond at C-6,
C-11 or C-6 and C-11 are listed using the symbols A6, A l l or A6,11.The structures of these compounds have been previously reported (Rohmer, 1975; Rohmer & Ourisson, 19766).
A . aceti subsp. aceti NCIB 8621
A . aceti subsp. Iiquefaciens NCIB 9418
A . aceti subsp. xylinum NCIB 41 12
A . aceti subsp. xylinum R 2277
A . pasteurianus subsp. estunensis NCIB 8935
A . pasteurianus subsp. lovaniensis NCIB 8620
A . pasteurianus subsp. ‘orleanensis’ NCIB 6426
A . pasteurianus subsp. pasteurianus NCIB 6429
A . pasteurianus subsp. pasteurianus NCIB 8856
A . peroxydans NCIB 8087
A . peroxydans NCIB 8618
Gluconobacter oxydans subsp. oxydans
Organism
Bacteriohopane- r
Diploptene Diplopterol
polyol
(pgg-1) (pgg-1)
(pgg-1)
VIII VI VII
Table 4. Hopanoid composition of acetic acid bacteria
U
Z
0
v,
v,
Y
C
P
0
36
m
w
K
c
C-L
Hopanoids in prokaryotes
1147
Table 5. In vivo incorporation of labelled precursors into prokaryotic hopanoids
Organism
Methylococcus capsulatus
Nostoc muscorum
Acetobacter pasteurianus
subsp. pasteurianus
Labelled
precursor
Specific
Total
activity
activity Incubation
(mCi mmol-I) (mCi)
(h)
[ 1 -I4C]Acetate
55
[ lJ4C]Acetate
45
0.30
24
[ 1-l4C]Acetate
49
0.50
18
1
3
~-[methyI-'~C]Methionine
9.05
0.10
24
~-[merhyl-'~C]Methionine
9.05
0-04
24
0.20
18
D-( +)-[5-3H]Ribose
50
Labelled
triterpenoids
Recovered
activity
(d.p.m.)
Squalene*
Diploptene*
Squalene*
Diploptene*
Squalene
Diploptene
Bacteriohopanepolyolst
I$
11s
111s
Bacteriohopanepolyolst
43 200
15800
91 300
21 000
5 200
3 800
106000
145 000
8 600
49 400
200 800
Diploptene
32 000
Diplopterol
28 000
Bacteriohopanepolyolss 23 1 000
* The radioactivity of the corresponding fraction was checked by preparative GLC and counting of the trapped triterpenoid by
liquid scintillation spectrometry.
t The radioactivity of the bacteriohopanepolyol fraction was checked after HSI06/NaBH4cleavage and TLC on silver nitrate
impregnated silica gel of the acetates of the primary alcohols obtained.
$ The tetraacetate of (I) and the pentaacetates of (11) and (111) were isolated by TLC and analysed separately.
8 The radioactivity of this fraction was checked by preparationof: tetraacetates,dicarbonatesobtained by phosgen treatment in
dry pyridine, and acetates of alcohols produced by H5106/NaBH4cleavage.
present as more water-soluble forms which cannot be extracted with diethyl ether after basic
hydrolysis. Such very polar derivatives have already been described by Langworthy et al. (1976)
in Bacillus acidocaldarius where bacteriohopanetetrol is linked through a glycosidic bond to N acylglucosamine. Recently we have isolated, from the facultative methylotroph Methylobacterium organophilum and from the purple non-sulphur bacterium Rhodomicrobiurn vannielii, new
series of much more polar bacteriohopane derivatives whose structures are currently under
investigation and which represent the quasi-totality of the bacteriohopanepolyol content of the
cells (J. M. Renoux, S. Neunlist & M. Rohmer, unpublished results). At the moment the direct
H5106 oxidation of the crude extract (procedure 2) is seen as the best method for a rapid
screening of the bacteriohopanepolyol content of a prokaryote as it includes the free polyols as
well as the known polar derivatives.
About half the strains analysed do not contain detectable hopanoids. There are several
possible explanations for this. (i) These prokaryotes may be unable to synthesize hopanoids
because they lack the required enzymes. (ii) Synthesis of hopanoids may be possible, but it does
not occur under the growth conditions used. Modification of some growth parameters might
induce the appearance of these compounds. (iii) Hopanoids may be in fact present, but are not
detected using our analytical procedure. The last hypothesis refers to a few puzzling cases where
only traces of hopanoids have been detected. For instance in a Pseudomonas aeruginosa strain
very small amounts of pure 3-methyldiplopterol (0.3 pg per g dry weight) have been found by
GLC/MS. Most probably this very rare hopanoid, which is found as a minor component
together with diplopterol, does not arise from accidental contamination, but rather indicates the
capacity of this micro-organism to synthesize hopanoids.
At the moment it is impossible to draw clear taxonomic conclusions from the distribution of
hopanoids in prokaryotes, since information on hopanoid content has been obtained for
relatively few organisms. A few clear-cut differences are already observable, however : for
example archaebacteria are devoid of hopanoids, and the purple non-sulphur bacteria can be
readily distinguished by their high hopanoid content from the purple sulphur bacteria, which
lack hopanoids. The hopanoid-containing strains are scattered through all taxonomic groups,
and even if some taxa appear to be homogeneous as to the presence of hopanoids (e.g. the
cyanobacteria, the purple non sulphur-bacteria, the obligate methylotrophs or the acetobacters),
most of the other hopanoid-containing strains were found in various taxonomic groups which
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1148
M. R O H M E R , P. B O U V I E R - N A V E A N D G . O U R I S S O N
apparently lack any polycyclic triterpenoid. Why are the two cyanobacterial genera
Synechococcus and Synechocystis, which are physiologically and biochemically very similar
(Stanier & Cohen-Bazire, 1977),different in hopanoid content? The former lacks any detectable
hopanoid, whereas the latter contains the usual bacteriohopanepolyols. Why is ‘Pseudomonas
cepacia’ the only hopanoid-containing pseudomonad? According to the Pseudomonas
classification of Palleroni et al. (1973) ‘Pseudomonas cepacia’ belongs to a peculiar subgroup
different from those of the other Pseudomonas strains. It would be interesting to analyse other
Pseudomonas species of the same subgroup to see if this difference is relevant. From the present
results it is evident that none of the bacteriohopanepolyols characterizes a strain or a group. It is
also evident that the composition of a bacteriohopanepolyol fraction in a given micro-organism
is dependent on the growth conditions since we had difficulties in ensuring the reproducibility of
the composition of the bacteriohopanepolyol fraction in some bacteria. This was particularly
true in the case of some Acetobacter strains which contain the most complex mixtures of
bacteriohopanepolyols. For instance we have found that the content of the 3-methylhopanoids
in Acetobacterpasteurianus can be markedly increased by addition of L-methionine to the culture
medium, and that the concentration of the A1 l-bacteriohopanepentol is apparently dependent
on the oxygenation of the cultures of Acetobacter aceti subsp. xylinum. Similarly the proportion of
the x-methylhopanoids in Nosroc muscorum varied from 0 to 60% for unknown reasons. The
values given in Tables 3 and 4 have to be considered as preliminary; the composition of a
particular bacteriohopanepolyol fraction will be more significant when the physiological role
of these compounds is better understood.
The hopanoids, whose importance was first revealed by the wide distribution of their
molecular fossils, appear now as characteristic and important constituents of numerous
prokaryotes belonging to the most varied taxonomic groups. We have postulated that these
compounds might be sterol surrogates in prokaryotes and even their phylogenetic ancestors
(Rohmer et al., 1979; Ourisson & Rohmer, 1982). Hopanoid biosynthesis in prokaryotes is the
most primitive triterpenoid biosynthetic route observed to date. Indeed the following
characteristics of this biosynthetic pathway can be considered as primitive compared to those of
sterol biosynthesis. (i) In hopanoid biosynthesis, squalene, a simpler substrate, is cyclized
instead of (3S)-squalene epoxide in the biosynthesis of other triterpenoids. (ii) The squalene
cyclization implies the involvement of the most favourable all pre-chair conformation of the
polyene which is thermodynamically less constrained than that required for the formation of
lanosterol, which must be partly in a pre-boat conformation. (iii) Hopanoid formation implies
only a simple cyclization without rearrangement or further oxidative degradation as in sterol
biosynthesis. (iv) The three squalene cyclases studied so far (Rohmer et al., 1980a, b ; Bouvier et
al., 1980) are not highly substrate specific: they cyclize squalene as well as the two enantiomers
of squalene epoxide, whereas the eukaryotic cyclases act specifically on (3s)-squalene epoxide.
(v) Hopanoid biosynthesis is completely independent from molecular oxygen since no oxidation
step is required as in sterol biosynthesis; it is therefore compatible with an ancient prebiotic
atmosphere.
The role of the hopanoids as membrane reinforcers is already supported by experiment;
hopanoids induce a condensing effect on artificial phospholipid monolayers or bilayers, much
like cholesterol (Poralla et al., 1980; Bisseret et al., 1983). Numerous problems concerning the
hopanoids are still unresolved : structural, biosynthetic and functional studies are currently
being pursued. Furthermore, like hopanoids, which can be regarded as ‘molecular coelacanths’,
other families of biolipids are so far only known from their molecular fossils, examples being CZ9
hopanoids, isoarborinol, tricyclopolyprenol derivatives and various, possibly archaebacterial,
lipid ethers. We have postulated that these compounds are molecular fossils of prokaryotic
membrane constituents (Ourisson et al., 1982); their identification in living organisms is still
awaited.
We acknowledge gratefully our debt to the late Professor R. Y. Stanier for the very helpful discussions which
initiated this work and for his constant interest for this screening. We are grateful to all our colleagues who
supplied us with freeze-dried material or living strains and helped in the growth of difficult micro-organisms.
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Hopanoids in prokaryotes
1149
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