FEMS Microbiology Ecology 12 (1993) 39-50
© 1993 Federation of European Microbiological Societies 0168-6496/93/$06.00
Published by Elsevier
39
FEMSEC 00454
A sensitive method for quantification of aceticlastic
methanogens and estimation of total methanogenic
cells in natural environments based on an analysis
of ether-linked glycerolipids
Sadami Ohtsubo a, Mitsuyoshi K a n n o
and Isao Miura ~
a Hiroyoshi Miyahara a Shuhei Kohno a Yosuke Koga b
a Water Treatment Section, Department of Biotechnology and Water Treatment, TOTO Ltd., Kitakyushu, and
b Department of Chemistry, University of Occupational and Environmental Health, Kitakyushu, Japan
(Received 13 November 1992; revision received 2 February 1993; accepted 3 February 1993)
Abstract." A highly sensitive method for the quantification of methanogens in anaerobic digestor sludges was developed, basekt on
an analysis of ether-linked glycerolipids. Core lipids were prepared from total lipids by HF treatment and mild methanolysis, and
these core lipids were quantified as the corresponding 9-anthroyl derivatives by high-performance liquid chromatography with
fluorescence detection. The amounts, in terms of cell carbon content, of Methanosaeta and Methanosarcina were proportional to
the amounts of a-hydroxyarchaeol and fl-hydroxyarchaeol, respectively. Moreover, the total amount of core lipids was well
correlated with the cell mass of aceticlastic and H 2/CO2-consuming methanogens. The limit of detection for Methanosaeta concilii
was 17 ng of cell carbon when the signal/noise ratio was 3. This method allowed us to quantitate aceticlastic methanogens with
high accuracy and to make a rough estimate of total methanogenic cells without any interference by the multifarious impurities that
are present in anaerobic sludges. These results suggest that the present method will be a useful tool for investigations of
methanogenic ecosystems.
Key words: Methanogen quantification; Ether-linked glycerolipid; Aceticlastic methanogen; High-performance liquid chromatography; Fluorescence detection; Methanogenic ecosystem
Introduction
Methane is a biogenic gas produced in anaerobic environments, such as rice paddies, wetlands
and the rumen of livestock. Increases in atmoCorrespondence to: S. Ohtsubo, Water Treatment Section,
Department of Biotechnology and Water Treatment, TOTO
Ltd., Kokurakita-ku, Kitakyushu 802, Japan.
spheric methane are considered to be significant
contributors to 'greenhouse' warming [1], and an
understanding of the biogeochemical processes of
methane cycling is clearly necessary at this time.
In addition, methane fermentation is a major
method for biological digestion of waste materials. Thus, methanogens, the only group of microorganisms that can produce methane, play a
considerable role in systems of environmental
40
importance. Quantification of methanogens in
anaerobic habitats provides basic information
about the ecology of methanogens.
Previously, we proposed a method for measurement of total methanogenic cells by quantification of the alkylglycerol ether portions (core
lipids; Fig. 1) of ether-linked polar lipids by highperformance liquid chromatography (HPLC) [2].
Recently, core lipids with a 3-hydroxyphytanyl
chain (hydroxyarchaeols) have been found in several methanogen species, Methanosarcina barkeri,
Methanosarcina mazeii, Methanosaeta concilii
(= ' Methanothrix soehngenii' = ' Methanothrix concilii'), Methanococcus vannielii, Methanococcus
voltaei, Methanococcus thermolithotrophicus, Methanosphaera stadtmaniae, Methanohalophilus
mahii and Methanolobus tindarius [3-5]. Because
the previous method does not allow detection of
hydroxyarchaeols, this method has a disadvantage
in that it gives underestimates of total numbers of
methanogenic cells in environmental samples that
contain large amounts of hydroxyarchaeols, such
as sludges from anaerobic waste digestors. Aceticlastic methanogens (Methanosaeta and Methanosarcina) catalyze acetate splitting, which is the
rate-limiting reaction in the anaerobic digestion
of organic compounds, and these bacteria play a
key role in methane fermentation [6]. A method
that allows specific quantification of aceticlastic
methanogens, as well as of total methanogenic
cells, would be a very useful tool for investigations of methanogenic ecosystems. In this report,
we describe a new, improved method for quantification of aceticlastic methanogens and for mak<'"
Caldarchaeol
Archaeol
uol ° ~ ' ~ x "
ec.Hydroxyarchaeol
13-Hydroxyarchaeoi
Fig. 1. Structures of representative core lipids in methanogens.
ing a rough estimate of total methanogenic cells.
The method is based on HPLC analysis of hydroxyarchaeols and other core lipids.
The nomenclature of core lipids proposed by
Nishihara et al. [7] is used in this report.
Materials and methods
Chemicals
1,2-Di-O-hexadecyl-rac-glycerol (dihexadecylglycerol) and 4-dimethylaminopyridine (DMAP)
were purchased from Sigma Chemical Co. (St.
Louis, USA). 9-Anthroylnitrile was obtained from
Wako Pure Chemical Industries, Ltd. (Osaka,
Japan). Because the purchased preparation of
9-anthroylnitrile contained fluorescence-positive
impurities, 9-anthroylnitrile was purified by chromatography on a column (1 cm i.d. × 30 cm) of
Silica gel 60 (E. Merck AG, Darmstadt, Germany) with a mixture of tetrahydrofuran/nhexane (20: 80, v/v) as the mobile phase.
Dichloromethane and n-hexane were of HPLC
grade.
Core lipid standards were prepared from
Methanobacterium thermoautotrophicum, Methanosarcina barkeri and Methanosaeta concilii as described previously [3,7,8].
Sources of strains
Methanobacterium thermoautotrophicum AH
( - D S M 1053), Methanospirillum hungateii JF1
(--- DSM 864), Methanobacterium formicicum MF
(= DSM 1535), Methanobre~ibacter arboriphilicus
DH1 ( = DSM 1125), Methanosarcina mazeii $6
( = DSM 2053), Methanosarcina barkeri strains
MS ( = DSM 800) and Jiirich (= DSM 2948) were
obtained from Deutsche Sammlung von Mikroorganismen und Zellkulturen GmbH (DSM),
Braunschweig, Germany. Methanosaeta concilii
Opfikon (= DSM 2139) was a gift from A.J.B.
Zehnder (Agricultural University, Wageningen,
The Netherlands). Methanosaeta concilii GP6 (=
OCM 69) was from the Oregon Collection of
Methanogens (Beaverton, USA). Methanosaeta
concilii MTKO (= DSM 6752 = OCM 252) was
from our collection.
41
Growth of strains
Methanosaeta concilii, Methanobacterium therrnoautotrophicurn, Methanospirillum hungateii and
Methanosarcina barkeri were grown as previously
described [9,10]. Methanosarcina mazeii was
grown in medium no. 120 described in the DSM
catalog 1989 [11], under a nitrogen atmosphere
without shaking. Methanobrevibacter arboriphilicus and Methanobacterium formicicum were
grown in medium No. 119 (DSM catalog 1989)
[11]. Cells were harvested by centrifugation and
suspended in distilled water. Suspensions of cells
were stored at -40°C until extraction of lipids.
Preparation of core lipids
Figure 2 shows a schematic representation of
the procedure of this method.
Lipid extraction. In a 10-ml screw-capped glass
tube, total lipids were extracted from 800/zl of a
Sample (Methanogensand
Other microorganisms)
l Acid Bllgh & Dyer extraction
Total Lipids (Ether polar lipids, Esterpolarlipids
and Non-polarlipids)
HF degradation
Bligh & Dyer extraction
1
1
CHCI3fraction
MethanoI-H:,Ofraction
(Corelipids, Non-polar llpids, (Polar headgroups)
Etherand ester polarlipids)
|
I Mild methanolysis
Discard
| Extraction with
petroleum ether/water (1:2, vol/vol)
1
1
CHCI 3 fraction
MethanoI-H20 fraction
(Corelipids, Non- polar lipids) (Polarhead groups)
Derivatization by 9-AN
Discard
9-AN-core liplds
Non-polar lipids
I
HPLC
9-AN-core lipids
Fig. 2. Schematic representation of the procedure for the
quantitation of core lipids.
cell suspension (approx. 1-200 /zg of methanogenic cell carbon) by the acidic procedure of
Bligh and Dyer, using trichloroacetic acid, as
modified by Nishihara and Koga [12]. When necessary, 0.2 /zg of dihexadecylglycerol were added
as the internal standard. The total lipid fraction
was transferred to a 7-ml screw-capped Teflon
centrifuge tube (Nalge Company, Rochester,
USA) and dried under a stream of nitrogen.
HF degradation. Total lipids were hydrolyzed
with 1 ml of 46% hydrofluoric acid and 0.5 ml of
chloroform (unless otherwise stated, see below)
at 4°C for 30 h. After incubation, lipids were
recovered by partitioning with chloroform/
methanol/water (10:10:9, v/v; Bligh and Dyer
solvent [13]) and dried under a stream of nitrogen.
Core lipids were acetylated by incubation at
100°C for 2 h with 0.2 ml of pyridine and 0.2 ml
of acetic anhydride, and they were quantitated by
gas-liquid chromatography (GLC). GLC was performed on a model GC-9A gas chromatograph
(Shimadzu Corp., Kyoto, Japan) equipped with a
column packed with Dexsil 300GC on Chromosorb W. The column temperature was increased from 200°C to 340°C at a rate of
20°C/min. Hexacosane was used as an internal
standard for the quantification of core lipids.
Acetylated archaeol and acetylated hydroxyarchaeols could not be separated under these
conditions.
Mild methanolysis. Lipids were hydrolyzed by
incubation with 1 ml of 5% methanolic HC1/
chloroform (1:27, v/v) at 50°C for 24 h. After
cooling to room temperature, core lipids were
recovered by partitioning with petroleum ether/
water (1 : 2, v/v).
Derivatization of core lipids with 9-anthroylnitrile
In the standard method, the solution of core
lipids was transferred to a 1.0-ml amber ReactiVial (Pierce, Rockford, USA) and dried under a
stream of nitrogen. Then 100 tzl of a solution of
DMAP (300 izmol/ml in chloroform) were added,
and the mixture was dried. Next, 50 /zl of a
solution of 9-anthroylnitrile (40/z mol/ml in CCI 4)
were added and the mixture was incubated at
75°C for 4 h. After this reaction, 200/zl of ace-
42
tonitrile were added and the mixture was applied
to a SepPak C18 cartridge (Millipore Corp., Bedford, USA). 9-Anthroyl derivatives of core lipids
on the cartridge were washed with 30 ml of
acetonitrile and recovered by elution with 5 ml of
chloroform. The eluate was dried, and the residue
was dissolved in 100/xl of n-hexane.
To examine the influence of the reaction solvent on the derivatization of caldarchaeol, 9-anthroyl derivatives of caldarchaeol were analyzed
by thin-layer chromatography (TLC). TLC was
performed on a silica gel 60 plate (Art. 5721; E.
Merck AG.) with n-hexane/tetrahydrofuran
(80:20, v/v) as the mobile phase. Spots on the
TLC plate were visualized under UV light (254
nm) and by subsequent acid charring with 50%
H z S O 4 (V/V) at 150°C.
of pure-cultured cells of Methanosaeta concilii
GP6 (20.7 /xg of cell carbon), Methanosarcina
barkeri MS (8.9 /xg of cell carbon) and
Methanobacterium formicicum MF (30.0 /xg of
cell carbon) was added to each environmental
sample, and the recoveries of these methanogens
were examined. Anaerobic sludges were collected
from two kinds of anaerobic fixed-bed digestor
maintained in our laboratory, and a soil sample
was a sediment from the bed of the Murasaki
River in Kitakyushu City, Japan.
The dry weight of samples was determined
gravimetrically, and cellular carbon contents were
determined with a model TOC-500 total organic
carbon analyzer (Shimadzu Corp.).
Results
HPLC
9-Anthroyl derivatives of core lipids were separated and quantified by HPLC using a Waters
600E Multisolvent Delivery System (Millipore
Corp.) equipped with a model RF-535 fluorescence detector (excitation, 370 nm; emission, 470
nm; Shimadzu Corp.). A column of TSKgel NH 260 (4.6 mm i.d. x 250 mm; TOSOH Corp., Tokyo,
Japan) was used for the separation. The solvents
for elution of 9-anthroyl derivatives were nhexane/dichloromethane (96 : 4, v/v) for the first
15 min, n-hexane/dichloromethane (77 : 23, v/v)
from 15 to 26 min, and n-hexane/dichloromethane (96 : 4, v/v) from 26 to 40 min. The flow
rate was 1.2 ml/min. From 5 to 30/~1 of solution
was injected on the column with a model 231-401
auto-sampling injector (Gilson Medical Electronics, Inc., Middleton, USA). The data were processed at a Waters Maxima 825J chromatography
workstation (Millipore Corp.).
Quantification of core lipids from methanogenic
cells and environmental samples
A 800-/xl suspension of methanogenic cells or
an environmental sample was dispersed by three
5-rain cycles of sonication at 5-min intervals. Lipid
extraction and preparation, derivatization and
quantification of core lipids were performed as
described above. To examine the applicability of
this method to environmental samples, a mixture
Preparation of core lipids
Core lipids were prepared from total lipids of
Methanosaeta concilii GP6 by HF degradation as
described by Sprott et al. [3]. Preparation of core
lipids was incomplete under these conditions, and
large amounts of intact polar lipids were detected
by TLC (data not shown). Under the conditions
of Sprott et al., HF degradation occurs at the
interface between aqueous HF and polar lipids,
which cannot dissolve in an aqueous solvent. The
nature of this reaction should be one of the
reasons for incomplete degradation by HF.
Therefore, we examined various co-solvents for
improvement of HF degradation (Table 1). Addition of chloroform improved the efficiency of the
reaction, and we chose to add 500 /xl of chloroform to reaction mixture. However, despite this
improvement, the preparation of core lipids was
still insufficient after HF degradation alone, as
described by Sprott et al. [3]. Mild methanolysis
was performed after HF degradation for the
preparation of core lipids. Because the solubility
of lipids in the reaction solvent is also supposed
to influence the reactivity during mild methanolysis, this reaction was performed in 5% methanolic
HCl/chloroform (1:27, v/v) instead of 0.18%
methanolic HC1 used by Sprott et al. When core
lipids were prepared by the combination of HF
degradation and mild methanolysis from total
43
Table 1
The enhancement of HF degradation of total lipids from
Methanosarcina barkeri by the addition of co-solvent
Co-solvent
Relative production
of core lipids (%) a
None (control)
Chloroform
20/xl
200/xl
500/~1
1000/~1
Acetone
20/.d
200 ~1
400 #1
Chloroform/acetone
(1 : 1, v/v)
400/xl
100
Acetonitrile
20 Izl
89
104
168
144
78
83
98
102
39
a The amount of released core lipids was defined as the area
under the peak that contained acetylated archaeol and
acetylated/3-hydroxyarchaeoldivided by the area under the
peak of acetylated internal standard, as determined by
gas-liquid chromatography.
chaeol, from the conditions for TLC. The spots
after TLC were identified as follows. The compounds corresponding to each spot were purified
by TLC and derivatized with 9-anthroylnitrile in
C C l 4 a s the reaction solvent. The compound with
a low Rf value was converted to the compound
with the high Rf value, but the mobility of the
latter did not change by the second reaction (data
not shown). These results indicated that the compound with the high Rf value was di-9-anthroyl
caldarchaeol. The major product of derivatization
in acetonitrile, the original solvent for the preparation of 9-anthroyl derivatives [14], was mono-9anthroyl caldarchaeol, and a large fraction of
caldarchaeol remained (Fig. 3). Best results were
obtained with CC14. Most of the caldarchaeol was
converted to the di-9-anthroyl derivative in this
solvent.
Figure 3 also shows the presence of some
fluorescent spots in addition to those of the 9-anthroyl derivatives of caldarchaeol. These spots
were found after TLC of the 9-anthroylnitrile
purchased from Wako Pure Chemical Industries
and they interfered with the detection of 9-an-
lipids of Methanosaeta and Methanosarcina, any
unreacted polar lipids were not detected, as
judged by TLC. Nevertheless, some of the polar
lipids from Methanobacterium thermoautotrophicum were not converted to core lipids. These
remaining polar lipids were largely glycolipids of
which the core portion was caldarchaeol.
a,
Front
qIF
,?
(e
Preparation of 9-anthroyl derivatives of core lipids
Core lipids were reacted with 9-anthroylnitrile,
a derivatising reagent for alcoholic hydroxy
groups, essentially by the method of Ramesha et
al. [14]. Archaeol and hydroxyarchaeols were
completely reacted under these conditions, but
conversion of caldarchaeol was incomplete (data
n o t shown). Thus, conditions for the reaction of
caldarchaeol were examined. Figure 3 shows the
influence of the reaction solvent on the derivatization of caldarchaeol. Two spots were observed
as 9-anthroyl derivatives of ealdarchaeol, and the
spot that had the lower R f value was considered
to be mono-9-anthroyl caldarchaeol, an intermediate in the formation of di-9-anthroyl caldar-
o
~b
,/5)
4.3
:'
o
~
o
q
•
(t
~-
0
i
1
2
3
4
Di-9-anthroyl
caldarchaeol
Mono-9-anthroyl
caldarchaeol
Caldarchaeol
(~
: Fluorescence positive
O
: Acid-charring positive
Fig. 3. Examination by TLC of the effect of the reaction
solvent on the 9-anthroyl derivatization of caldarchaeol. In a
5-ml screw-capped glass tube, 10 gg of caldarchaeol were
incubated at 70°C for 5 h with 1.5/xmol of 9-anthroylnitrile
and 1.5 /zmol of DMAP in 100 /xl of reaction solvent. The
TLC was developed with n-hexane/tetrahydrofuran (80:20,
v/v), and spots were visualized under UV light (254 nm) and
by subsequent acid charring. Lanes: 1, acetonitrile; 2, carbon
tetrachloride; 3, heptane; 4, dimethyl sulfoxide.
44
throyl core lipids by HPLC (data not shown).
Therefore, we purified 9-anthroylnitrile from the
commercial preparation as described in Materials
and methods.
The level of DMAP in reaction mixtures
markedly affected the derivatization of core lipids
(Fig. 4). A 10 /.~mol of D M A P was sufficient for
the derivatization of dihexadecylglycerol, but
more than 25 /.~mol of DMAP was required for
complete conversion of caldarchaeol to the di-9anthroyl derivative. Finally, the reaction conditions were set as described in Materials and
methods. Under these conditions, the completeness of the derivatization of all core lipids was
assessed by TLC, and there was no indication of
any unreacted core lipids after TLC.
Since di-9-anthroyl caldarchaeol has two 9-anthroyl groups, di-9-anthroyl caldarchaeol must
emit twice as much fluorescence per molecule as
9-anthroyl dihexadecylglycerol which has one 9anthroyl group. However, Fig. 4 indicates that the
peak area, in other words the fluorescence intensity, of di-9-anthroyl caldarchaeol was 5-fold lower
than the expected value. Consequently, we decided to normalize the amount of caldarchaeol by
multiplying the peak area of di-9-anthroyl caldarchaeol in HPLC analysis by 5, so that the peak
%
5
10
15
20
=
25
30
35
,2
L.
5
10
15
20
Time (rain)
25
30
35
Fig. 5. H P L C chromatograms of 9-anthroyl derivatives of core
lipids. (A) A chromatogram after H P L C of the mixture of
purified 9-anthroyl core lipids (the amounts of core lipids,
approx. 40-100 ng). (B) A chromatogram after H P L C of
9-anthroyl core lipids prepared from environmental samples.
9-Anthroyl core lipids prepared from sludge A to which a
mixture of pure cultures of m e t h a n o g e n s had been added (see
Table 2) were dissolved in 100 pA of n-hexane, and a 30-~1
sample was subjected to HPLC.
area of di-9-anthroyl caldarchaeol appeared to be
twice that of the 9-anthroyl dihexadecylglycerol.
Figure 5A shows the profile after HPLC of a
mixture of 9-anthroyl derivatives of purified core
lipids. Under these conditions, the peaks of 9-anthroyl core lipids were separated completely from
one another.
2
0 -
A
5
10
15
20
DMAP (pmol)
25
30
Fig. 4. T h e influence of the level of D M A P on the derivatization of caldarchaeol (10 /~g) and dihexadecylglycerol (1 /~g).
Samples were derivatized with 2.0 /~mol of 9-anthroylnitrile
and the indicated concentration of D M A P at 75°C for 5 h.
The a m o u n t s of derivatives were taken from peak areas after
HPLC. Symbols: ©, 9-anthroyl dihexadecylglycerol; o, di-9-anthroyl caldarchaeol; A, mono-9-anthroyl caldarchaeol.
The relationship between core lipid content and the
biomass of pure cultures of methanogenic strains
Significant amounts of hydroxyarchaeols have
been found in Methanosaeta concilii, two Methanosarcina species ( M. barkeri and M. mazeii ),
Methanosphaera stadtmaniae, three Methanococcus species and a few halophilic methylotrophic
methanogens [3-5]. These methanogens other
than Methanosphaera stadtmaniae and aceticlastic species (Methanosaeta and Methanosarcina)
require NaCl for growth and inhabit marine or
high-salt environments. Methanosphaera stadtma-
45
niae was originally isolated from human feces and
may be present in an anaerobic digestor. Nevertheless, the hydroxyarchaeol-containing methanogens except aceticlastic species were not detected
in anaerobic digestors, as analyzed by immunological techniques [15,16]. These results suggest
that the levels of hydroxyarchaeols in a sludge
from anaerobic sewage digestor reflect the
amounts of aceticlastic methanogens, and hydroxyarchaeols of Methanosaeta and Methanosarcina can be distinguished by the position of glycerol at which a hydroxyphytanyl chain is linked,
sn-3 in Methanosaeta and sn-2 in Methanosarcina
(Fig. 1) [3,4]. If we could show that the levels of
hydroxyarchaeols are correlated with the biomass
of these aceticlastic methanogens, aceticlastic
methanogens could be quantitated. Figures 6 and
7 show the correlation between hydroxyarchaeol
content and cell mass for Methanosaeta and Methanosarcina, respectively. The level of hydroxyarchaeol is represented as the peak size in HPLC
analysis. The level of a-hydroxyarchaeol was proportional to the cell mass of each of three strains
of Methanosaeta concilii (r = 0.97; Fig. 6). Similar
results were obtained for the relationship be-
0.5
0.4
0.3
o A O ~
.
e~
t=
0.2
0.1
0.0
0
2
4
6
8
Cell carbon content (lug)
10
Fig. 6. Correlation between the relative amount (peak response) of a-hydroxyarchaeol and the amount of Methanosaeta cells. Peak response was defined as the area under the
peak of 9-anthroyl a-hydroxyarchaeol divided by the area
under the peak of 9-anthroyl dihexadecylglycerol. The correlation curve was obtained by the least-squares regression
method using all data points. Symbols: o, M. concilii strain
MTKO; o, M. concilii strain Opfikon; zx, M. concilii strain
GP6.
1.0
r=O.9J
A
0.8
~
0.6
o.4
0.2
0.0
0
'
'
1
'
'
2
'
'
3
'
4
5
Cell carbon content (lug)
Fig. 7. Correlation between the relative amount (peak response) of fl-hydroxyarchaeol and the amount of Methanosarcina cells. Peak response was defined as the area under the
peak of 9-anthroyl fl-hydroxyarchaeol divided by the area
under the peak of 9-anthroyl dihexadecylglycerol. The correlation curve was obtained by the least-squares regression
method using the data from strains MS and Jiirich. Symbols:
©, M. barkeri strain MS; e, M. barkeri strain Jiirich; zx, M.
mazeii strain $6.
tween the level of fl-hydroxyarchaeol and the cell
mass of two strains of Methanosarcina barked
(r = 0.99; Fig. 7). Though the level of fl-hydroxyarchaeol was linearly correlated with the cell mass
of Methanosarcina mazeii, the correlation curve
was different from that of Methanosareina barkeri.
The relative amount of total core lipids (RCL)
was defined as the sum of peak areas of 9-anthroyl core lipids (the peak area of di-9-anthroyl
caldarchaeol was corrected by multiplying by 5 as
described above) divided by the peak area of
9-anthroyl dihexadecylglycerol. We examined the
possibility of estimating total methanogenic
biomass by using RCL values. Figure 8A shows
the correlation between RCL values and cell mass
of aceticlastic methanogens. While the molecular
species of hydroxyarchaeol and its relative level
in Methanosaeta concilii were different from those
in Methanosarcina barkeri [3,8], RCL values of
these methanogens showed the same relationship
to cell mass (r = 0.97). In the case of Methanosarcina mazeii, however, the correlation curve was
different from that for the other species. Figure
8B shows the results for H2/COz-consuming
46
3.0
A
3.0
o
B
/""
2.5
2.5
• 0~or j=t .97
2.0
2.0
,d 1.5
.d 1.5
l.O
1.0
0.5
0.5
.
0.0 0
1
.
.
.
.
.
.
.
.
.
2
3
4
5
6
7
Cell carbon content (lug)
0.0 0
,,'"
1
2
3
4
5
6
7
Cell carbon content (lug)
Fig. 8. Correlation between the values of the relative amounts of total core lipids (RCL) and methanogenic cells. The RCL value
was defined at the sum of areas under the peaks of 9-anthroyl core lipids divided by the area under the peak of 9-anthroyl
dihexadecylglycerol. The peak area of di-9-anthroyl caldarchaeol was corrected by multiplying by 5 as a correction factor (see text)
(A) Correlation for aceticlastic methanogens. The correlation curve was obtained by the least-squares regression method using all
data except those for M. mazeii $6. Symbols: o, M. concilii strain Opfikon; *, M. concilii strain GP6; t3, M. concilii strain MTKO;
Ill, M. barkeri strain MS; z~, M. barkeri strain Jiirich; A, M. mazeii strain $6, (B) Correlation for H2/CO2-consuming
methanogens. Dashed line shows the correlation curve for aceticlastic methanogens obtained in (A) of this figure. The correlation
curve (solid line) was obtained by the least-squares regression method using all data points for H 2 / C O z-consuming methanogens.
Symbols: o, M. formicicum; *, M. thermoautotrophicum ; t3, M. hungateii; z~, M. arboriphilicus.
Table 2
Recovery of methanogens from environmental samples
Sample
Species
Added methanogen
concerned
(/zg of cell carbon) (A)
Amount of methanogens in
environmental samples a
(/zg of cell carbon)
Recovery (%) f
Minus pure
cultures (B)
Plus pure
cultures (C)
29.0 _+0.2 e
9.9 -+0.4
54.5 _+0.5
74.8 _+0.7
96.1
91.0
78.2
106.7
Sludge A b
Methanosaeta
Methanosarcina
Total (I) c
Total (II) d
20.6
8.9
59.6
59.6
9.2
1.8
7.9
11.2
Sludge B b
Methanosaeta
Methanosarcina
Total (I) e
Total (II) a
20.6
8.9
59.6
59.6
6.0
0.7
6.8
9.7
25.0 _+1.6
9.3 _+0.1
54.4 _+2.1
74.6 _+2.9
92.2
96.6
79.9
108.9
Soil b
Methanosaeta
Methanosarcina
Total (1) c
Total (lI) a
20.6
8.9
59.6
59.6
1.3
0.6
3.4
5.1
18.1 _+0.4
8.8 _+0.3
48.6 _+1.6
66.7 _+2.2
81.6
92.1
75.8
103.4
a A mixture of Methanosaeta concilii GP6 (20.7/zg of cell carbon), Methanosarcina barkeri MS (8.9 /zg of cell carbon) and
Methanobacterium formicicum MF (30.0/zg of cell carbon) was added to each environmental sample.
b The amounts of environmental samples subjected to the determination of methanogens were as follows: sludge A, 266 /zg dry
weight (ca. 100/zl); sludge B, 489/.~g dry weight (ca. 40 ~1); soil, 38.2 mg.
c Values were calculated from the standard curve for aceticlastic methanogens (see Fig. 8A).
d Values were calculated from the standard curve for H 2 / C O 2 consuming methanogens (see Fig. 8B).
c Values are means +_standard deviations (n = 3).
f Recovery was calculated as ([(C)-(B)]/(A)}x 100 (%).
47
methanogens ( Methanobacterium formicicum,
Methanospirillum hungateii, Methanobacterium
thermoautotrophicum and Methanobrevibacter arboriphilicus). One correlation curve fitted all of
the data from these species very well (r = 0.98),
regardless of the differences in polar lipids composition and the relative levels of core lipids [2].
Nevertheless, the correlation curve did not coincide with that for aceticlastic methanogens. This
difference should be caused by the incomplete
preparation of caldarchaeol from the total lipids
of H2/CO2-consuming methanogens, as indicated in Methanobacterium thermoautotrophicum.
Application of the method to natural samples
This method was applied to the quantification
of methanogenic cells in anaerobic sludges and a
soil sample. Figure 5B shows the HPLC profile of
9-anthroyl core lipids from an anaerobic sludge to
which a mixture of pure-cultured cells of methanogens was added. Only a few peaks derived from
impurities are present in the chromatogram, and
it was confirmed that cellular components from
Escherichia coli did not interfere with the detection of the peaks of 9-anthroyl core lipids (data
not shown). The mixture of pure cultures of three
methanogens, Methanosaeta concilii GP6, Methanosarcina barkeri MS and Methanobacterium
forrnicicum MF, was added to the anaerobic sampies, and the recoveries of these methanogens
were examined (Table 2). The amounts of
Methanosaeta and Methanosarcina were determined from the levels of ot-hydroxyarchaeol and
/3-hydroxyarchaeol, respectively. The correlation
curve for Methanosarcina barked was used in the
determination of Methanosarcina. Methanosaeta
and Methanosarcina were considered to be accurately determined in environmental samples because the added aceticlastic methanogens were
recovered with high yield (81-96%). Total
methanogenic content was estimated by using two
kinds of correlation curve, the curve for aceticlastic methanogens and the curve for H 2 / C O 2consuming methanogens (Fig. 8). Total methanogenic cells were underestimated (75.8-79.9%)
when the correlation curve for aceticlastic methanogens was used, but by contrast total me-
thanogenic cells were slightly overestimated
(103.4-108.9%) when the correlation curve for
Hz/CO2-consuming methanogens was used. Methanobacterium formicicum is H2/CO2-consuming methanogens, and the preparation of core
lipids from this bacterium is incomplete by our
method, as mentioned above. This is the reason
for the difference of recovery depended on the
standard curve. From these results, it is indicated
that our method can quantitate aceticlastic
methanogens and provide a rough estimate (with
a range of + 10-30% error) of the total methanogenic biomass in environmental samples.
Discussion
We have described a new method for quantifying aceticlastic methanogens and estimating the
total mass of methanogenic cells. Previously, we
reported a method for quantitating methanogenic
cells that was based on the analysis of core lipids
by HPLC [2]. This method involved the preparation of core lipids by a combination of acetolysis
and methanolysis, the derivatization of core lipids
with 3,5-dinitrobenzoyl chloride (DNBC) and the
quantification of these derivatives by HPLC with
UV detection. This earlier method allowed us to
estimate total methanogenic cells with high sensitivity. However, it had a disadvantage that it
could not detect hydroxyarchaeols, which caused
an underestimation of methanogenic cells in natural samples.
In the present study, we made two alterations
in our previous method to permit quantification
of hydroxyarchaeols and to enhance its sensitivity.
First, the combination of HF degradation and
mild methanolysis was introduced for the preparation of core lipids to allow us to obtain intact
hydroxyarchaeols, which are acid-labile. Sprott et
al, used this method for screening hydroxyarchaeols in methanogenic bacteria [3]. They reported
that HF degradation alone released head groups
from about 55% of total polar lipids of Methanosarcina barked and that up to 80% of core lipids
were recovered by the combination of HF degradation and mild methanolysis. We enhanced the
48
rate of generation of core lipids without degradation of hydroxyarchaeols by addition of chloroform to the reaction solvent. Under our conditions, polar lipids from Methanosaeta and Methanosarcina were completely converted, as
checked by TLC.
The second modification was the introduction
of 9-anthroylnitrile as the labeling reagent for
detection in HPLC analysis, with the intention of
increasing sensitivity and simplifying conditions
for derivatization of hydroxyarchaeols, a-Hydroxyarchaeol accounts for only one-third of the
core lipids from Methanosaeta concilii [8]. Thus,
higher sensitivity is required for practical quantification of a-hydroxyarchaeol, as compared with
the quantification of total methanogenic cells.
9-Anthroylnitrile is a fluorescent marker and can
provide higher sensitivity than DNBC, which was
used in previous method. The limit of detection
for Methanosaeta concilii was 17 ng of cell carbon
when the signal/noise ratio was 3. 9-Anthroylnitrile was synthesized for the determination of
hydroxysteroids by Goto et al. [17]. They examined reactivities of various hydroxysteroids with
9-anthroylnitrile and indicated that tertiary hydroxy groups were not reacted with this reagent.
Thus, 9-anthroylnitrile would not react with the
hydroxy group on the phytanyl moiety of hydroxyarchaeol (Fig. 1). The derivatization conditions
for hydroxyarchaeols would be simpler than those
for caldarchaeol in which two hydroxy groups
participate in derivatization (Fig. 1). Moreover,
unreacted hydroxy groups on phytanyl moieties
might be favorable for the separation of a- and
/3-hydroxyarchaeols by HPLC. In fact, it was
proven by Fourier-transform infrared spectroscopy that the hydroxy group on the phytanyl
moiety was inert to the reaction with 9-anthroylnitrile (data not shown) and derivatization of
hydroxyarchaeols proceeded more effectively than
that of, caldarchaeol. The peaks of ~-hydroxyarchaeol and /3-hydroxyarchaeol were also separated completely from one another by HPLC
(Fig. 5A and B).
Although two hydroxy groups in caldarchaeol
were reacted with 9-anthroylnitrile, the intensity
of fluorescence from this derivative was much
lower than expected. The reason for this discrep-
ancy is not clear, but it was reported by Bayliss et
al. [18] that the fluorescence intensity of diethylene glycol (DEG) 9-anthroyl diester was nine
times lower than that of DEG 9-anthroyl monoester. This result suggests that the fluorescent
characteristics of 9-anthroyl diesters are very different from those of typical monoesters. We calculated 5 as the correction factor for the amounts
of di-9-anthroyl caldarchaeol. We compared the
ratio of caldarchaeol to archaeol in core lipids of
Methanobacterium thermoautotrophicum from an
analysis of dinitrobenzoyl derivatives, prepared
by our previous method [2], with that obtained
from an analysis of 9-anthroyl derivatives. The
ratio obtained from 9-anthroyl derivatives was
4.7-fold lower than that from dinitrobenzoyl
derivatives. These results indicate that 5 is reasonable as a correction factor.
The cell mass of Methanosaeta concilii and
Methanosarcina barkeri was proportional to the
amount of hydroxyarchaeol (Figs. 6 and 7). The
difference in strains did not affect this relationship. These results suggest that the present
method should allow the quantification of these
aceticlastic methanogens in an anaerobic digestor
sludge. However, the slope of the correlation
curve for Methanosarcina mazeii was less steep
than that for Methanosarcina barkeri (Fig. 7). The
genus Methanosarcina forms aggregates of cells
with an outer layer, composed of heteropolysaccharides [19]. Our method is based on a presumption that lipid content reflects the mass of individual cells, and the presence of the outer layer
has not been taken into consideration. The difference between correlation curves for Methanosarcina barkeri and Methanosarcina mazeii may be
caused by differences related to the nature of
heteropolysaccharide layer.
In the experiment for which results are shown
in Table 2, the added mixture of methanogenic
species included 50.3% Hz/COz-consuming
methanogens and 49.7% aceticlastic methanogens. With this distribution ratio, the amount of
total methanogenic cells was estimated with a
precision of 75.8-108.9%, depending on the standard curve. Because acetielastic methanogens are
determined from the levels of hydroxyarchaeols,
independently of the estimation of total methano-
49
genic cells, the accuracy of estimation of total
methanogenic cells could be increased by selecting an appropriate standard curve. That is to say,
the standard curve for aceticlastic methanogens is
preferable for samples that contain large amounts
of aceticlastic methanogens, and the standard
curve for H2/CO2-consuming methanogens is
suitable for samples that contain small amounts
of aceticlastic methanogens.
Several methods have been used for quantification of methanogens, such as detection of autofluorescence caused by coenzyme F410 [20], the
most probable number (MPN) technique [21] and
immunological detection [15,16]. Each of these
techniques has grave disadvantages for quantification of methanogens in environmental samples.
Methanosaeta contains trace amounts of coenzyme F420, and this aceticlastic methanogen cannot be detected by autofluorescence [22]. Counting of cells by the MPN method is strongly dependent on the growth medium, and this method
counts an aggregate of cells, which are generally
formed in culture of Methanosaeta and Methanosarcina, as a single cell. Immunological methods
can identify and quantify methanogenic strains in
natural samples. However, these methods require
antisera against many reference methanogens.
The preparation of many kinds of antiserum entails a great deal of effort, and laboratories that
can use these techniques are restricted in number. Our new method allows separate quantification of aceticlastic and H2/CO2-consuming
methanogens without a requirement for special
instrumentation and reagents. Moreover, results
are not influenced by the morphology of natural
samples, dispersed or aggregated samples, or the
biological characteristics of the methanogens.
Therefore, though our method cannot identify
methanogenic strains at the species level, unlike
immunological techniques, it should be very useful for analyzing methanogenic ecosystem, that
contains many kinds of methanogen.
Acknowledgments
We thank to K. Demizu for helpful discussions
and Y. Akahoshi for skillful technical assistance.
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