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Appl Environ Microbiol, February 1998, p. 479-485, Vol. 64, No. 2
0099-2240/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
Adaptive Changes in Membrane Lipids of Barophilic
Bacteria in Response to Changes in Growth Pressure
Yutaka
Yano,*
Akihiko
Nakayama,
Kenji
Ishihara, and
Hiroaki
Saito
Marine Biochemistry Division, National
Research Institute of Fisheries Science, Yokohama, Kanagawa 236, Japan
Received 2 September 1997/Accepted 3 December 1997
 |
ABSTRACT |
The lipid compositions of barophilic bacterial strains which
contained docosahexaenoic acid (DHA [22:6n-3]) were examined, and the
adaptive changes of these compositions were analyzed in response to
growth pressure. In the facultatively barophilic strain 16C1,
phosphatidylethanolamine (PE) and phosphatidylglycerol (PG) were major
components which had the same fatty acid chains. However, in PE,
monounsaturated fatty acids such as hexadecenoic acid were major
components, and DHA accounted for only 3.7% of the total fatty acids,
while in PG, DHA accounted for 29.6% of the total fatty acids. In
response to an increase in growth pressure in strain 16C1, the amounts
of saturated fatty acids in PE were reduced, and these decreases were
mainly balanced by an increase in unsaturated fatty acids, including
DHA. In PG, the decrease in saturated fatty acids was mainly balanced
by an increase in DHA. Similar adaptive changes in fatty acid
composition were observed in response to growth pressure in obligately
barophilic strain 2D2. Furthermore, these adaptive changes in response
were also observed in response to low temperature in strain 16C1. These
results confirm that the general shift from saturated to unsaturated
fatty acids including DHA is one of the adaptive changes in response to
increases in pressure and suggest that DHA may play a role in
maintaining the proper fluidity of membrane lipids under high pressure.
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INTRODUCTION |
Lipids, in particular,
phospholipids, are the main components of cell membranes, in which many
important biological functions occur. The dynamic states of lipids,
such as the fluidity and the order, are known to be closely related to
the functions of biological membranes (9). The dynamic
states are dependent on factors such as temperature and pH
(16). Thus, it is well known that a number of bacteria,
plants, and poikilotherms regulate the lipid compositions of their
membranes in response to changes in the environmental temperature so as
to maintain the membrane lipid fluidity necessary for proper biological
function (9). This phenomena is known as homeoviscous
adaptation (25). For example, a decrease in growth
temperature generally results in an increase in the level of
unsaturation and a decrease in the average chain length of fatty acids
in bacterial lipids (16, 24). Furthermore, in cold-adapted
species, the levels of unsaturated fatty acids or polyunsaturated fatty
acids (PUFAs) are known to be relatively high (9, 31).
Barophiles are defined as organisms which grow optimally or
preferentially at pressures greater than atmospheric pressure (0.1 MPa). Barophilic bacteria have been isolated from various deep-sea
environments and have been shown to grow rapidly at low temperatures
and high pressures (11, 35, 36). High pressure and low
temperature in deep-sea environments theoretically decrease the
fluidity of lipids and possibly depress the functions of biological membranes (9, 14). Thus, barophiles seem to have some
mechanism which allows their lipids to adapt to deep-sea environments.
In fact, in a barophilic strain, it was shown that an increase in growth pressure resulted in an increase in the level of unsaturated fatty acids in the lipids, suggesting a homeoviscous adaptation in
response to pressure (3). In general, bacteria have been considered to be unable to produce PUFAs (5, 7). A great number of bacterial strains isolated from terrestrial and shallow-sea environments are reported to produce fatty acids which have chain lengths of less than 20 (19, 20, 30). However, marine
bacteria such as deep-sea isolates and barophilic isolates have been
reported to contain PUFAs such as docosahexaenoic acid (DHA
[22:6n-3]) and eicosapentaenoic acid (EPA [20:5n-3]) in their
lipids (4, 12, 22). PUFAs have relatively low melting points
(16), and so they may assist in maintaining the proper
fluidity of membrane lipids that the marine bacteria require to adapt
to deep-sea environments. Furthermore, it was found that the relative
amounts of PUFAs were higher during both medium- and high-pressure
incubations, than during low-pressure incubations (4, 32).
From these studies, we assumed that PUFAs may be involved in the
adaptation of barophiles to deep-sea environments. However, the details
of the lipid compositions remain unknown in bacteria containing PUFAs,
and the adaptive changes of lipid compositions in barophilic bacteria
have been reported only for the fatty acid composition of total lipids.
In a previous study (33), we showed that the barophilic
strains isolated from the intestinal contents of deep-sea fish
contained DHA and EPA. Furthermore, we found that bacteria containing
DHA were generally and abundantly distributed in the intestines of deep-sea fish, compared with shallow-sea poikilothermic animals (34), suggesting the involvement of DHA in the adaptation to deep-sea environments. Thus, in the present study, we initially examined the lipid compositions of a barophilic strain which contained DHA in its lipids. Next, we examined the effect of growth pressure on
the phospholipid and the fatty acid compositions in facultatively and
obligately barophilic strains containing DHA.
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MATERIALS AND METHODS |
Bacterial strains and medium.
Strain 16C1 was facultatively
barophilic and was originally isolated from the intestinal contents of
the deep-sea fish Coryphaenoides armatus, which was
retrieved from a depth of 3,100 m (34). Strain 2D2 was
obligately barophilic and was isolated from the intestinal contents of
the deep-sea fish Coryphaenoides yaquinae, which was retrieved from a depth of 6,100 m (18). These strains were
maintained in retortable pouches (nylon-CPP; Sanei Chemical industry,
Tokyo, Japan) containing marine broth (Difco, Detroit, Mich.) at 5°C and at in situ pressures (strain 16C1, 41.1 MPa; strain 2D2, 62.1 MPa)
by using pressure vessels (18). Growth characteristics of
strains 16C1 and 2D2 in relation to pressure were examined by
epifluorescence microscopy with 4',6-diamidino-2-phenylindole (DAPI)
(21), as described previously (18).
Fatty acid composition of strain 16C1.
For analyses of the
fatty acid composition of strain 16C1, the culture was grown in
defatted marine broth. Defatted marine broth consisted of (per liter)
5 g of Bacto Peptone (Difco), 1 g of Bacto yeast extract
(Difco), 0.1 g of ammonium ferric citrate, 20 mM MOPS
(morpholinepropanesulfonic acid [pH 7.0]), and 30.0 g of
artificial seawater salts (Senju, Osaka, Japan). Before the preparation
of the medium, peptone and yeast extract were extracted with
chloroform-methanol (2:1 [vol/vol]) to remove the lipids. The culture
of strain 16C1 was grown in the medium at 5°C and 41.4 MPa. After a
7-day incubation period which led the strain to the stationary phase,
bacterial cells were harvested by centrifugation at 9,000 × g for 15 min under 4°C, immediately frozen at 
20°C, freeze-dried, and used for lipid analyses.
Total lipids were extracted from dried cells with chloroform-methanol
(2:1 [vol/vol]) by the method of Folch et al. (6). The
organic phase was filtered and washed with one-quarter its volume of
distilled water. After removal of the solvent by evaporation, the total
lipid extracts were redissolved in a known volume of chloroform-methanol (2:1 [vol/vol]) and stored at 20°C under nitrogen for further analysis. The total lipids were added to 2.5%
methanolic HCl and heated at 85°C for 2.5 h for preparation of
fatty acid methyl esters (FAMEs). The resulting FAMEs were extracted
three times with n-hexane and stored at 20°C under nitrogen for further analysis. The analyses of the FAMEs were performed
with a G-5000 gas chromatograph (Hitachi, Tokyo, Japan) equipped with
an Omega wax 320 capillary column (30 m by 0.32 mm [inside diameter];
Supelco, Bellefonte, Pa.) and a flame ionization detector. The oven
temperature was programmed from 170 to 215°C at a rate of 1°C per
min. Helium was used as the carrier gas, and the injector and the
detector were maintained at 255 and 260°C, respectively. The samples
were further analyzed with an SP-2330 capillary column (30 m by 0.25 mm
[inside diameter]; Supelco) with the oven temperature programmed as
described above. The FAMEs were identified by comparison with authentic
standards and measured with a recording integrator attached to the gas
chromatograph.
The unsaturated nature of fatty acids was confirmed by hydrogenation
with platinum oxide and reanalysis with the gas chromatograph
(
2). The FAMEs were dissolved with methanol in a test tube,
and PtO
2 was added. The tube was flushed with hydrogen to
remove
any air, and then was vigorously shaken and maintained for
2 h.
The solvent was then evaporated, and the resulting saturated
FAMEs
were taken up in
n-hexane. The resulting saturated
FAMEs were
analyzed for gas chromatography and compared with the
original
FAMEs.
Positions of double bonds in fatty acids were determined by gas
chromatography-mass spectrometry (GC-MS) according to the
procedure
(the pyrrolidine method) of Anderson et al. (
1,
2).
The
FAMEs were dissolved with pyrrolidine in a test tube, and
acetic acid
was added. The mixture was heated at 100°C for 30
min. The resulting
derivatives were taken up in dichloromethane
and washed with 2 N HCl
and then with water. GC-MS analyses of
derivatized FAME samples were
performed on a JMS-DX303 instrument
(JEOL, Tokyo, Japan) equipped with
an Omega wax 320 capillary
column (30 m by 0.32 mm [inside diameter];
Supelco). The oven
temperature was 260°C. Electron impact mass
spectra were measured
at 70 eV.
For analysis of constituent fatty acids, known amounts of the total
lipids were spotted on thin-layer chromatography plates
and separated
with chloroform-methanol-water (65:25:4 [vol/vol/vol])
as a mobile
phase. Developed chromatograms were sprayed with 0.01%
(wt/vol)
primurin reagent (Tokyo Kasei, Tokyo, Japan) and viewed
under UV light.
Bands containing phosphatidylethanolamine (PE)
and phosphatidylglycerol
(PG) were identified by reference to
the standard, scraped from the
plates, and then subjected to transesterification
as described above. A
known amount of 23:0 methyl ester was added
as an internal standard.
After cooling, the resulting FAMEs were
extracted and stored at
20°C under nitrogen until GC analyses.
GC analyses were performed
under the conditions described above.
Phospholipid composition was
determined from the weight of the
constituent fatty acids by using the
internal standard.
Effect of pressure and temperature on lipid compositions.
Chilled marine broth (1,000 ml) was inoculated with 1.0 ml of strain
cultures which grew at 5°C and at in situ pressures for a week. This
was distributed to four retortable pouches (250 ml each) and then
heat-sealed. Next, the pouches were placed in pressure vessels and
incubated at 5°C and at various pressures. After incubation, bacterial cells were harvested in early stationary phase in a manner
similar to that described above. That is, in the case of strain 16C1,
bacteria cells were harvested after 4 days from both the 20.7- and
41.4-MPa incubations and after 5 days from the 0.1-MPa incubation. In
the case of strain 2D2, harvesting was conducted after 4 days from the
41.4-MPa incubation and after 5 days from both the 20.7- and 62.1-MPa
incubations. Bacterial cells were immediately frozen at 20°C,
freeze-dried, and used for lipid analyses as described above.
Furthermore, in the case of strain 16C1, in order to examine the effect
of growth temperature on fatty acid composition, the pouches containing
the medium were incubated at 1°C and atmospheric pressure (0.1 MPa)
at the same time with the pressure experiment. After a 5-day
incubation, the bacterial cells were harvested, frozen at 20°C,
freeze-dried, and used for lipid analyses as described above. In all of
these experiments, the cell numbers of the cultures were counted by epifluorescence microscopy at the harvesting time.
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RESULTS |
Growth characteristics.
Figure 1
shows the cell numbers of strains 16C1 and 2D2 after a 2-day incubation
period when the cells were grown at 5°C and at various pressures.
Strain 16C1 optimally grew at 20.4 MPa. Because the strain was capable
of growing at 0.1 MPa, it was considered to be facultatively
barophilic. Strain 16C1 grew to the early stationary phase for 4 days
at both 20.7 and 41.4 MPa, and for 5 days at 0.1 MPa (data not shown).
Strain 2D2 optimally grew at 41.4 MPa, as shown in the previous study
(18). Furthermore, the strain was confirmed to be obligately
barophilic, because it was not capable of growing at 0.1 MPa. Strain
2D2 grew to the early stationary phase for 4 days at 41.4 MPa and for 5 days at both 20.7 and 62.1 MPa (data not shown).
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FIG. 1.
Growth of barophilic strains 16C1 ( ) and 2D2 ( ) at
different pressures and 5°C. Markers show the cell numbers of the
cultures at 2 days of incubation, respectively. The cell numbers at the
beginning of incubation were 4.2 × 104 cells/ml in
strain 16C1 and 1.7 × 105 cells/ml in strain 2D2.
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Fatty acid composition of strain 16C1.
The representative
fatty acid compositions of total lipids, PE, and PG in strain 16C1 are
shown in Table 1. In the total lipids of
strain 16C1, the major fatty acids were tetradecanoic acid (14:0),
tetradecenoic acid (14:1), hexadecanoic acid (16:0), hexadecenoic acid
(16:1), and DHA, according to the comparison of retention time with the
standards on the gas chromatogram. Octadecanoic acid (18:0) and
octadecenoic acid (18:1) were minor components. The PUFAs of
C18 (the carbon length is 18; this notation is also used
below) and C20 were also found as minor fatty acids in
total lipids, but those of C22, except for DHA, were not
detected. After hydrogenation, all peaks which were regarded as
unsaturated fatty acids disappeared on the gas chromatogram, and the
resulting fatty acid composition was almost the same as the composition calculated from the fatty acid composition before hydrogenation (data
not shown). Furthermore, on the mass spectrum of DHA, the characteristic peaks of m/z 381, 366, 352, 338, and 326 were
found (Fig. 2), and the spectrum was the
same as that of the authentic 22:6n-3 (data not shown). The positions
of double bonds in monounsaturated fatty acids of C14 and
C16 and 20:5n-3 were also confirmed (data not shown).
In PE, the fatty acid composition was, in general, similar to that of
the total lipids; however, it is notable that 16:1n-7
was present at
higher levels (55.6% of the total fatty acids)
and that DHA amounted
to only 3.7% of the total fatty acids. In
contrast, 16:1n-7 was
present at a lower level (35.9% of the total
fatty acids), and DHA
accounted for 29.6% of the total fatty acids
in PG.
Effect of pressure on phospholipid composition.
Phospholipid
compositions of strain 16C1 and obligately barophilic strain 2D2, grown
at different pressures, are shown in Table
2. In the case of strain 16C1, the
proportion of PE was greater with higher pressure incubations than that
with the 0.1-MPa incubation. However, this proportion was not different
between the 20.7- and 41.4-MPa incubations. Phospholipids in strain 2D2 were also generally PE and PG, and the change in composition was not
distinctly observed among cells grown at any given pressure, while PE
appeared to increase slightly in cells grown at higher pressures.
Effect of pressure on fatty acid composition.
The fatty acid
compositions of total lipids, PE, and PG from strain 16C1, grown at
different pressures, are shown in Table 3. The major fatty acids were 14:0,
14:1n-7, 16:0, 16:1n-9, 16:1n-7, and DHA, regardless of the growth
pressures. In total lipids, the change in response to growth pressure
was observed in the C14 and C16 acids. The
proportions of 14:0 and 14:1n-7 in the cells grown at 0.1 MPa were 26.9 and 19.2% of the total fatty acids, respectively, and these
proportions decreased in the cells grown at 20.7 MPa (10.0 and 6.2% of
total fatty acids, respectively) and 41.4 MPa (9.0 and 5.8% of total
fatty acids, respectively). This decrease was largely balanced by an
increase in 16:0 and 16:1. DHA accounted for 6.6% of the total fatty
acids at 0.1 MPa and increased to 8.4% at 20.7 MPa. However, this
change in fatty acid composition was not distinct between the cultures
at 20.7 and 41.4 MPa. Thus, in the total lipids, the proportions of
saturated fatty acids decreased with an increase in growth pressure,
while the proportions of monounsaturated fatty acids increased. In
addition, the proportions of PUFAs slightly increased.
The change in fatty acid composition of PE was generally similar to
that observed in the total lipids. With increasing growth
pressure, the
proportions of saturated fatty acids decreased,
while the proportions
of monounsaturated fatty acids and PUFAs
increased.
In PG, the change in fatty acid composition in response to growth
pressure was most markedly observed in the level of 14:0
and DHA. The
proportion of 14:0 was 23.6% of the total fatty acids
at 0.1 MPa, and
this decreased to 10.2% at 20.7 MPa. DHA accounted
for 12.5% of the
total fatty acids at 0.1 MPa and increased to
28.6% at 20.7 MPa. The
proportion of 16:1 was generally constant.
The change was not distinct
between the culture at 20.7 MPa and
the culture at 41.4 MPa. Thus, the
decrease in saturated fatty
acids in PG associated with increasing
growth pressure was balanced
by an increase in PUFAs.
The change in fatty acid composition of the obligately barophilic
strain 2D2 in response to growth pressure is shown in Table
4. In this strain, the fatty acid
compositions of total lipids,
PE, and PG were generally similar to
those of strain 16C1, and
the major fatty acids were 14:0, 14:1, 16:0,
16:1, and DHA. However,
the fatty acid compositions in strain 2D2 were
notably different
from those of strain 16C1, in that strain 2D2
contained a lower
proportion of 16:1 and a far higher proportion of
DHA. In particular,
DHA accounted for about 50% of the fatty acids of
PG. The change
in fatty acid composition of the 2D2 strain was, in
general, similar
to that in strain 16C1. In the total lipids and PE,
the proportions
of 14:0 and 14:1n-7 decreased and those of 16:0 and
16:1 increased,
with increasing growth pressure. Thus, with increasing
growth
pressure, the proportions of saturated fatty acids decreased,
while those of monounsaturated fatty acids and PUFAs increased.
In PG,
the proportion of 14:0 decreased and that of DHA increased
with
increasing growth pressure. The degree of change in fatty
acid
composition, however, was not as great as that observed in
the PG of
strain 16C1. The proportion of 16:1 remained constant.
Thus, the
proportions of saturated fatty acids decreased in PG,
while those of
PUFAs increased.
Effect of temperature on fatty acid composition.
The fatty
acid compositions of total lipids, PE, and PG from strain 16C1 grown at
1°C and 0.1 MPa are shown in Table 5.
For the convenience of comparison, the table includes the data for cells grown at 5°C and 0.1 MPa, previously shown in Table 3. The
major fatty acids of the total lipids were 14:0, 14:1, 16:0, 16:1 and
DHA, regardless of the respective growth temperature. In response to an
increase in growth temperature, changes were observed in
C14 acids, C16 acids, and DHA. These changes,
in general, corresponded with the changes caused by growth pressure for
the same fatty acids. Thus, with decreasing growth temperature, the proportions of saturated fatty acids decreased and those of unsaturated fatty acids increased.
| |
DISCUSSION |
Production of DHA in procaryotes was first reported in deep-sea
isolates, including barophilic bacteria (4). The present study showed that the DHA observed in the barophilic strain 16C1 was
22:6n-3, which contained methylene-interrupted double bonds and was
normally observed in lipids of eucaryotes (17, 26). Furthermore, it was found that the double-bond positions of
monounsaturated fatty acids were n-9 and n-7 in the strain. Generally,
unsaturated fatty acids in bacteria have been considered to be produced
by two possible mechanisms: an oxygen-independent (anaerobic) pathway and an oxygen-dependent (aerobic) pathway (7). In the
anaerobic pathway, which is catalyzed by a fatty acid synthetase,
palmitoleic acid (16:1n-7) and cis-vaccenic acid (18:1n-7)
are produced. In the aerobic pathway, which involves fatty acid
desaturation analogous to that of eucaryotes, palmitoleic acid
(16:1n-7) and oleic acid (18:1n-9) are commonly produced from saturated
fatty acids by position-specific desaturase. A production mechanism for
PUFA, such as DHA, however, is unknown in bacteria. In procaryotic
algae producing 18:4 (28) and eucaryote-synthesized DHA
(17), it was reported that C18 and
C20 PUFAs of the n-3 and n-6 series were present in their
lipids. Also in strain 16C1, those PUFAs were shown to be contained in
the lipids, although no previous reports have been made of the presence
of C20 PUFAs, even EPA, in bacteria containing DHA (4,
8). Furthermore, according to the known elongation system of
fatty acid, it is difficult to imagine that 22:6n-3 containing
methylene-interrupted double bonds is formed by means of an anaerobic
pathway. Thus, we suppose that DHA in the bacterial strain 16C1 may be
also synthesized via C18 and C20 PUFAs of the
n-3 and n-6 series, as is the case in eucaryotes. However,
C22 PUFAs (with the except of DHA) were not detected in
strain 16C1 or in other bacteria containing DHA. In fish, DHA is
considered to be formed by the addition of C2 to EPA,
followed by desaturation (13). However, in rat hepatocytes, it has been reported that elongation of 22:5n-3 to 24:5n-3 is followed
by desaturation to 24:6n-3, and then this is metabolized, via
-oxidation, to 22:6n-3 (27). Thus, further investigations are needed to clarify the synthetic pathway of DHA in bacteria.
Phospholipids in strain 16C1 were mainly PE and PG, as described
previously (34). PE was found to be abundant in
monounsaturated fatty acids (74% of total fatty acids) and lacking in
PUFAs, including DHA (5.4% of total fatty acids), compared with PG. In
PG, monounsaturated fatty acids were present at a lower level (47% of
total fatty acids) and DHA accounted for 29.6% of the total fatty
acids. This tendency was also observed in strain 2D2, as shown in Table
4, and the proportion of DHA in the PG accounted for about 50% of the
total fatty acids. Although there have been no reports on the fatty
acid composition of phospholipids in bacteria containing DHA, it has
been reported that PG has higher levels of EPA than PE in two
EPA-producing bacterial strains isolated from freshwater fish and
shallow-sea fish (10, 29).
The change in phospholipid composition in relation to growth pressure
was not distinct, except that the proportion of PE became high when
strain 16C1 underwent pressurized incubation. It is well known that the
phospholipid compositions of bacteria and the changes related to
incubation conditions vary with bacterial species (24). In a
bacterium containing EPA, which has PE and PG as the major lipids, the
proportion of PE was reported to increase at a lower growth temperature
(10), although what the change meant was unknown.
With increasing growth pressure, DHA levels increased in the fatty
acids of the total lipids in strains 16C1 and 2D2. This indicates that
DHA was more necessary at high pressures and confirms that DHA plays
some roles in the growth and biological functions of barophilic
bacteria at high pressures, suggested by DeLong and Yayanos
(4). Furthermore, the present study found that the increases
in the level of DHA occurred in phospholipids, especially PG. This
finding suggests that the role of DHA may be closely related to the
functions of the membrane.
In PE, in the present study, the proportions of 14:0 and 14:1 were
reduced with an increase in pressure, and these decreases were mainly
balanced by a increase in 16:1. In PG, the decrease of 14:0 was mainly
balanced by an increase in DHA. Generally speaking, with increasing
growth pressure, saturated fatty acids decreased and unsaturated fatty
acids, including DHA, increased in major phospholipids of barophilic
strains. It is well known that the increase in the level of
unsaturation of fatty acids occurs with the lowering of growth
temperature in bacteria and poikilotherms (15, 23). In the
present study also, the lowering of growth temperature caused the
increase in unsaturated fatty acids. These results suggest that the
barophilic strains compensate for pressure increases through
homeoviscous adaptation in a fashion similar to the response to
lowering of temperature. Thus, the present study further confirms the
suggestion of DeLong and Yayanos (3, 4). That is to say, one
of the roles of DHA may be in maintaining the fluidity of lipids at
high pressure.
In the present study, the changes in fatty acid composition in
phospholipids were slight or ambiguous between medium pressure and high
pressure. It has also been reported that in the barophilic strain MT41,
there was a decrease in the relative amount of DHA at the highest
growth pressure compared with that at the medium pressure
(4). If these strains always regulate their lipid states
when pressure increases, it is difficult to understand slight or no
change in the fatty acid composition at high pressure, as observed in
the present study. Furthermore, although it is known that a shortening
of carbon length in fatty acids is observed at lower temperatures
(9, 24), the proportions of 14:0 and 14:1 were reduced at
higher pressures in the present study. These results may suggest that
there were changes in the lipid composition which were not detected by
the analysis of fatty acid composition. This indicates the need for
more detailed analysis of the molecular species of phospholipids.
| |
ACKNOWLEDGMENTS |
We thank Y. Ezura, K. Takahashi, H. Shinano, and K. Yoshida for
valuable advice.
This study was partially supported by grant BRP-97-I-A-4 from the
Ministry of Agriculture, Forestry, and Fisheries.
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FOOTNOTES |
*
Corresponding author. Mailing address: National
Research Institute of Fisheries Science, 2-12-4, Fukuura, Kanazawa-ku,
Yokohama, Kanagawa 236, Japan. Phone: 45-788-7669. Fax: 45-788-5001. E-mail: yanoya{at}nrifs.affrc.go.jp.
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Appl Environ Microbiol, February 1998, p. 479-485, Vol. 64, No. 2
0099-2240/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
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Abstract
Introduction
