Previous Article | Next Article 
Journal of Bacteriology, August 2000, p. 4288-4294, Vol. 182, No. 15
0021-9193/00/$04.00+0
Copyright © 2000, American Society for Microbiology. All rights reserved.
Metabolic Instability of Escherichia
coli Cyclopropane Fatty Acid Synthase Is Due to
RpoH-Dependent Proteolysis
Ying-Ying
Chang,1
Johannes
Eichel,2 and
John E.
Cronan Jr.1,3,*
Departments of
Microbiology1 and
Biochemistry,3 University of Illinois,
Urbana, Illinois 61801, and Hans-Knöll Institute for
Natural Products Research, 07745 Jena, Germany2
Received 6 March 2000/Accepted 10 May 2000
 |
ABSTRACT |
Cyclopropane fatty acids (CFAs) are generally synthesized as
bacterial cultures enter stationary phase. In Escherichia
coli, the onset of CFA synthesis results from increased
transcription of cfa, the gene encoding CFA synthase.
However, the increased level of CFA synthase activity is transient; the
activity quickly declines to the basal level. We report that the loss
of CFA activity is due to proteolytic degradation dependent on
expression of the heat shock regulon. CFA synthase degradation is
unaffected by mutations in the lon, clpP, and
groEL genes or by depletion of the intracellular ATP pools.
It seems likely that CFA synthase is the target of an unidentified
energy-independent heat shock regulon protease. This seems to be the
first example of heat shock-dependent degradation of a normal
biosynthetic enzyme.
| |
INTRODUCTION |
Cyclopropane fatty acids (CFAs) are
very widely distributed among the bacteria and are formed by addition
of a methylene group, derived from the methyl group of
S-adenosyl-L-methionine (AdoMet), across the
carbon-carbon double bond of unsaturated fatty acids (UFAs)
(11). Methylene addition does not involve free fatty acids
or intermediates of phospholipid biosynthesis, but rather, uses mature
phospholipid molecules previously incorporated into and functioning
within membrane bilayers. CFAs are typically produced at the onset of
stationary phase in bacterial cultures, and CFA formation can thus be
considered a conditional, postsynthetic modification of bacterial
membrane lipid bilayers. This modification is catalyzed by a soluble
enzyme, despite the fact that one of the substrates, the UFA double
bond, is normally sequestered deep within the hydrophobic interior of
the phospholipid bilayer. This and other properties imply topologically
novel protein-lipid interactions in the biosynthesis of CFAs. No
physiological role for CFA formation had been ascribed until we
recently reported that Escherichia coli strains lacking CFA
survive poorly when exposed to pH 3 (2).
Only a small fraction of UFAs are converted to CFAs in exponentially
growing cultures of E. coli and other bacteria
(11). However, as cultures reach stationary phase, the UFAs
of the membrane phospholipid bilayers are rapidly and almost
quantitatively converted to CFA. Wang and Cronan (28) showed
that the growth-phase dependence of CFA synthesis is due to activation
of an RpoS-dependent cfa promoter. The cfa gene
encodes CFA synthase, the 43-kDa enzyme catalyzing the modification
(29), and is transcribed from two promoters, P1 and P2
(28). Promoter P1 is a standard RpoD (
70)
promoter and is active throughout the growth curve, whereas function of
promoter P2 requires the stationary-phase sigma factor RpoS
(38). The onset of stationary phase results in efficient
conversion of phospholipid UFA moieties to CFAs by two mechanisms, the
accumulation of RpoS gives increased cfa expression, and the
decreased phospholipid synthetic rate allows CFA synthesis to catch up
to phospholipid synthesis (CFA synthase must no longer cope with a
continually increasing substrate pool) (28). However, these
studies also gave the unexpected result that CFA synthase activity did
not plateau at the higher level but instead rapidly decreased to the level seen in exponentially growing cultures. This abrupt decrease in
CFA synthase activity suggested that CFA synthase is extremely sensitive to proteolysis, and consistent with this suggestion, a CFA
fusion protein carrying a C-terminal tag was found to be very unstable
in vivo (28). However, the presence of the abnormal fusion
junction could have destabilized the CFA synthase portion of the fusion
protein, and thus these results might not be a valid indication of CFA
synthase degradation. We have now examined the stability of E. coli CFA synthase by immunoprecipitation of the radioactively
labeled protein. We report that native CFA synthase is a short-lived
protein in vivo and its degradation is dependent on expression of the
heat shock regulon.
| |
MATERIALS AND METHODS |
Bacterial strains and media.
All bacterial strains are
derivatives of E. coli K-12 and are listed in Table
1. Genetic markers were transferred among
the strains by phage P1vir transduction. Plasmid pAYW19,
which carries the cfa gene, was constructed previously
(29). The liquid medium used was minimal medium E
(27) plus either glucose (0.4%) or glycerol (0.4%) as
carbon source. L-Methionine (45 µg/ml) was added to the
media of methionine auxotrophs. The concentrations of antibiotics and
amino acids used in media (in milligrams per liter) were tetracycline
hydrochloride, 10; sodium ampicillin, 100; chloramphenicol, 30;
L-tryptophan, 20.5; L-leucine, 50;
L-cystine, 60; and L-threonine, 50. L-Methionine supplementation was as given below.
Pulse-chase experiments.
These experiments were carried out
using two different protocols. In protocol A, an overnight culture was
diluted 1:20 into methionine-supplemented minimal medium E plus either
glucose or glycerol as carbon source (the overnight cultures were grown
on the same medium). Ampicillin was added when required to select plasmid maintenance. The cultures were grown to mid-exponential phase
(A600 of 0.5 to 0.6) at the temperature given,
centrifuged, and washed once with medium E, and the cells were
resuspended in the same medium except that the methionine concentration
was decreased to 2.5 µg/ml to permit efficient radioactive labeling. A mixture of L-[35S]methionine and
L-[35S]cysteine (70:25 molar ratio; American
Radiolabeled Chemicals, Inc.) was added to 50 µCi/ml (specific
activity of 1,175 Ci/mmol) when the labeling was performed on strains
carrying plasmids or 100 µCi/ml when strains carrying a single
cfa gene copy were labeled. The labeling period was usually
30 to 40 min. Labeling was terminated by addition of nonradioactive
L-methionine (45 µg/ml), and samples were taken following
growth for various lengths of time. In protocol B, radioactive labeling
was done as in protocol A except that L-cystine (60 µg/ml) was included in all media. Labeling was halted by centrifuging
the cells, washing once with minimal medium E, and resuspension in
minimal medium plus glucose, L-methionine (45 µg/ml), and
L-cystine (60 µg/ml).
The samples were processed by harvesting the cells by centrifugation
followed by washing once with 0.1 M potassium phosphate
buffer (pH
7.5). The cell pellets were then suspended in 1/10
of the original
sample volume of phosphate buffer, lysed by sonication,
and centrifuged
at 30,000 ×
g for 1 h. The resulting supernatants
were treated with preimmune serum to decrease the nonspecific
binding
before immunoprecipitation with a polyclonal rabbit antiserum
prepared
against purified
E. coli CFA synthase. Usually, 0.1 ml
of
the cell extracts was treated with 2 µl of the CFA synthase
antiserum
and incubated for 1 h at 0°C, and 16 µl of protein A-Sepharose
CL-4B beads (Pharmacia Inc.) was added. The protein A-Sepharose
CL-4B
beads were prepared as a 1:1 (vol/vol) suspension in bead
buffer
containing 10 mM Tris-HCl (pH 8.0), 10 mM EDTA, 150 mM
NaCl, 1% bovine
serum albumin, and 0.4% (wt/vol) Triton X-100.
After incubation for 30 min on ice, the Sepharose beads were washed
twice with the bead buffer,
once with 0.1 M Tris-HCl (pH 8.0),
and finally once with 0.01 M
Tris-HCl (pH 8.0). To the washed
beads, 20 µl of sodium dodecyl
sulfate-polyacrylamide gel electrophoresis
(SDS-PAGE) sample buffer was
added, and the sample was boiled
for 2 min. The samples were
centrifuged, and the supernatants
were loaded on SDS-10%
polyacrylamide gels. Following SDS-PAGE,
the gels were dried and
scanned with a PhosphorImager (Molecular
Dynamics, Inc.). The software
for data analyzing and storage was
ImageQuaNT (Molecular Dynamics). The
gels were also exposed to
X-ray film for permanent records. Although we
used equal amounts
of sample radioactivity for each
immunoprecipitation, the total
radioactivity in each gel lane varied
somewhat, probably due to
the multiple steps of the procedure.
Therefore, we normalized
the radioactivity present in the CFA synthase
band to the total
radioactivity present in the lane. A strain carrying
a
cfa null
mutation (strain YYC1317) was used to determine
the background
radioactivity in the gel section where CFA synthase
migrated.
The anti-CFA synthase antibody nonspecifically precipitated numerous
other proteins, and thus the CFA synthase band was identified
by use of
the
cfa null mutant plus an
35S-labeled CFA
synthase standard prepared from strain YYC1321,
which expresses CFA
synthase from a phage T7 promoter. This strain
was grown to mid-log
phase, and CFA synthase production was induced
(0.4 mM
isopropyl-
-
D-thiogalactopyranoside) for 30 min at 37°C
followed by addition of rifampin (200 µg/ml) to block expression
of
chromosomal genes. After 5 min, the culture was labeled with
radioactive methionine-cysteine, and the proteins were extracted
as
described above. Samples of the extracts were loaded on the
gels
without immunoprecipitation to serve as a CFA synthase marker.
As
expected from the work of Studier and Moffatt (
25), CFA
synthase
was the major labeled
protein.
Antibody preparation.
CFA synthase was purified as described
previously (29). The purified CFA synthase was further
purified by on a preparative SDS-polyacrylamide gel (BioRad model 491 Prep Cell; 14-cm length and 2.8-cm diameter; 10% running and 4%
stacking gel). Fractions containing only CFA synthase were pooled and
dialyzed against 0.1 M potassium phosphate buffer (pH 7.5). The
antiserum was prepared by injection of rabbits with 150 µg of the
protein dispersed in complete Freund's adjuvant and boosted after 2 and 4 weeks with 50 µg of CFA synthase dispersed in incomplete
Freund's adjuvant.
| |
RESULTS |
E. coli CFA synthase is unstable in vivo.
In
preliminary studies of the degradation of E. coli CFA
synthase, we tested several different growth and labeling protocols. We
first attempted to label cultures in early stationary phase, since that
is the time CFA synthase levels are maximal. However, labeling under
these conditions was very inefficient such that CFA synthase, a minor
protein, could not be detected. We therefore labeled mid-log-phase
cultures and allowed the culture to enter stationary phase during the
chase period. Glycerol-grown cultures of E. coli strain
YYC1314, which contains only the chromosomal copy of cfa,
were first examined. The intensity of the labeled CFA synthase band
decreased with time during the chase with unlabeled methionine (Fig.
1). After chase for 40 or 80 min with
excess unlabeled methionine, only 63 or 35%, respectively, of the
original radioactivity of CFA synthase remained. Similar results
obtained when this strain was grown with glucose as carbon source. The half-life of the protein calculated from these experiments was in the
range of 40 to 60 min (the doubling time for this strain under these
conditions was about 60 min). Strain YYC1318, which contains a
high-copy-number plasmid encoding cfa (Fig. 1), was also
examined. About half of the radioactivity in CFA synthase disappeared
after 30 min of chase at 37°C (Fig. 1), and most of the remaining
label was subsequently lost. In several repeats of this experiment in
glucose medium, the CFA synthase half-life varied within 30 to 60 min.
Therefore, the half-life of the CFA synthase encoded by multiple copies
of the cfa gene was similar to that obtained for a strain
having one copy of the gene. All subsequent experiments used strains
carrying this plasmid to take advantage of the increased sensitivity
resulting from increased cfa expression.
View larger version (48K):
[in this window]
[in a new window]
|
FIG. 1.
Proteolytic degradation of CFA synthase. Autoradiograms
show SDS-polyacrylamide gel separations of immunoprecipitates of
pulse-chase experiments with various strains. Three independent gels
are shown (lanes 1 to 4, 5 to 9, and 10 to 14). Lanes 1 to 3 are
extracts of strain YYC1314 (which contains only the chromosomal
cfa gene) growing with glycerol as carbon source. Lane 1, extracts from the strain labeled for 40 min without chase; lanes 2 and
3, the same as lane 1, but chases were for 40 and 80 min, respectively.
Lane 4 is strain YYC1168 carrying plasmid pAYW19 and was used as a CFA
synthase standard (Materials and Methods). Lanes 5 to 9 are extracts of
strains grown with glucose as carbon source. Lane 5, cfa
null mutant, YYC1317; lanes 6 to 9, extracts of the cfa
plasmid-containing strain YYC1318 labeled for 30 min and chased for 0, 30, 60, and 90 min, respectively. Lanes 11 to 14 are extracts from
experiments done on strain YYC1318 grown with glucose and labeled for
30 min followed by chase periods of 0, 60, 90, and 120 min,
respectively. Lane 10 is a CFA synthase marker in which CFA synthase
was labeled in the presence of rifampin in strain YYC1321 (Materials
and Methods). The arrows indicate the protein band corresponding to CFA
synthase (note that the standard in lane 4 differs from that used in
lane 10 and elsewhere in this study). The experiments were done with
protocol A (Materials and Methods). Note that residual labeling with
cysteine occurred during the chase (see text). Abbreviations: bgrd,
background (cfa null mutant); std, standard.
|
|
In the early experiments, comparison with the null mutant background
control showed that some radioactivity remained in the
CFA synthase
band even after extended chase periods (e.g., 90
or 120 min). It should
be noted that the
35S-labeled methionine source used was a
mixture containing 70%
methionine and 25% cysteine plus 5%
miscellaneous amino acids.
We had not added cysteine (or cystine) to
our chase medium because
we expected that endogenous cysteine synthesis
would provide an
effective chase. To test this assumption, the method
was modified
by including exogenous cystine (cysteine is toxic to
E. coli K-12
at high concentrations) in all of the media.
With this modification,
the half-life (15 min) of CFA synthase obtained
was shorter than
that measured in the absence of a source of exogenous
cysteine
(30 to 60 min), and the CFA synthase band completely
disappeared
upon prolonged chase (Fig.
2). We therefore added cystine to all
the
media used in subsequent experiments. These results were obtained
from
cultures grown at 37°C beginning from mid-exponential phase
(
A600 of 0.5 to 0.6) (Fig.
2). Similar results
were obtained for
late-log-phase cultures (
A600
of 1.0), whereas stationary-phase
cultures could not be sufficiently
labeled to detect CFA synthase.
View larger version (39K):
[in this window]
[in a new window]
|
FIG. 2.
Effect of temperature on degradation of CFA synthase.
(A) Autoradiograms of SDS-polyacrylamide gel separations of
immunoprecipitates of pulse-chase experiments with various strains are
shown. The cfa plasmid carrying strain YYC1318 was grown on
glucose minimal medium supplemented with methionine and cysteine and
labeled as described in Materials and Methods. In each case, the
cultures were labeled with radioactive methionine for 30 min at 37°C
before initiation of the chase period. In lanes 2 and 3, the chase
experiments were done at 30°C. Lane 1, no chase; lane 2, 30-min
chase; lane 3, 60-min chase. Lanes 4 and 5 were chased at 37°C for 30 and 60 min, respectively. Lane 6 was chased for 5 min at 42°C and
then for 30 min at 37°C. Lane 7 is the CFA synthase marker. Arrows
indicate the CFA synthase band. std, standard. (B) Turnover of CFA
synthase of strain YYC1318 at 30°C (open circles) and 37°C (filled
circles). Each point represented the result from one lane of an
experiment such as that of panel A. The experiments were done
with protocol B (Materials and Methods).
|
|
The effect of temperature on CFA synthase degradation was also tested.
When strain YYC1318 was grown at 30°C, the half-life
of CFA synthase
was 30 min (Fig.
2), about twice that obtained
at 37°C, whereas at
42°C the half-life was too rapid to measure
accurately but was
considerably less than 15 min. Therefore,
E. coli CFA
synthase was metabolically unstable and the rate of degradation
increased with
temperature.
Degradation of E. coli CFA synthase depends on
induction of the heat shock regulon.
The temperature-dependent
degradation of E. coli CFA synthase suggested that heat
shock proteases may be involved in the instability of this enzyme. For
example, Lon and the Clp family of proteases are well-known members of
the heat shock regulon (9, 12). The genes of this regulon
are specifically transcribed by RNA polymerase directed by the RpoH
sigma factor (32). Therefore, the effect of a mutation
in rpoH on degradation of E. coli CFA synthase
was tested by use of strain K165 (21), which carries an
rpoH nonsense mutation plus a temperature-sensitive suppressor tRNA. The temperature-sensitive suppressor tRNA is inactive
at 42°C and suppresses poorly at lower temperatures (1, 5). Strain K165 grows fairly well at 30°C, whereas higher
temperatures block cell division and are lethal. To avoid these
complications, we chose to examine cells deficient in (rather than
devoid of) this transcription factor (1, 5). We first
transduced strains K165 and its parent, strain SC122, to methionine
auxotrophy followed by introduction of the cfa plasmid
pAYW19 to give strains YYC1325 and YYC1324, respectively. These strains
were then grown at 30°C, and the half-lives of CFA were determined
for both strains (Fig. 3 and Table
2). In the parental strain the CFA
synthase half-life was about 45 min, whereas no detectable loss of
radioactive CFA synthase occurred in the RpoH-deficient strain during a
90-min chase. Therefore, it seems that degradation of the CFA synthase of E. coli requires high level expression of RpoH.
View larger version (48K):
[in this window]
[in a new window]
|
FIG. 3.
Effect of mutations in rpoH, lon,
clpP, or groEL on degradation of CFA synthase.
Autoradiograms of SDS-polyacrylamide gel separations of
immunoprecipitates of pulse-chase experiments with various strains are
shown. Three gels are shown (lanes 1 to 5, 6 to 12, and 13 to 16).
Lanes 2 and 3 are extracts of strain YYC1324 (wild type [wt]) labeled
with [35S]methionine for 40 min at 30°C and then chased
for 0 (lane 2) or 90 (lane 3) min at 30°C. Lanes 4 and 5 are
identical to lanes 2 and 3, respectively, except that the strain
labeled was YYC1325 (rpoH). Lanes 6 and 7 are strain YYC1328
(wild type), lanes 8 and 9 are strain YYC1329 (lon), and
lanes 10 and 11 are strain YYC1332 (lon clpP). In each case,
the strain was labeled for 30 min at 37°C followed by no chase (lanes
6, 8, and 10) or by chase for 60 min at 37°C (lanes 7, 9, and 11). In
lanes 13 to 16, the experiments were done at 30°C. Lanes 13 and 14 were strain YYC1336 (wild type), whereas lanes 15 and 16 were from
strain YYC1338 (groEL). In each case, the odd-numbered lane
received no chase period, whereas the even-numbered lane was chased for
60 min (labeling was done for 30 min). Lanes 1 and 12 contain the CFA
synthase marker, and the arrows denote the CFA synthase protein band.
The experiments were done using protocol A (Materials and Methods).
std, standard.
|
|
The lack of CFA synthase turnover in the RpoH-deficient strain should
result in higher levels of the protein, and if this
protein is fully
functional, higher levels of CFA should be present
in this strain than
in the wild-type strain. To test this implication,
we isolated
phospholipids from mutant strain K165 and parental
strain SC122 grown
at 30°C. The lipids were analyzed by collision-induced
dissociation
electrospray mass spectroscopy (
26) (Fig.
4). We
found that the levels of synthesis
of the C17 and C19 CFAs in
the RpoH-deficient strain were eight- and
fivefold higher, respectively,
than the levels of these acids present
in the parental strain.
Moreover, CFA synthesis began earlier in the
growth curve of the
RpoH-deficient strain than in the parental strain
(e.g., at an
optical density of 1.3, the RpoH-deficient strain
contained about
threefold more CFA than the wild-type strain). These
results indicated
that the
32 deficiency stabilized CFA
synthase in vivo and the protein was
enzymatically active. The low
levels of CFA in these strains can
be attributed to the
relA1 and
spoT1 mutations present in these
strains. These mutations decrease RpoS levels and thereby inhibit
CFA
accumulation (
4). This was of advantage in the present
experiments because the supply of UFA moieties did not become
limiting.
View larger version (21K):
[in this window]
[in a new window]
|
FIG. 4.
Effects of RpoH deficiency on cellular CFA content. (A)
Acyl chain region of the spectrogram obtained by collision-induced
dissociation electrospray mass spectroscopy of phospholipids of the
wild-type strain SC122; (B) parallel analysis of the phospholipids of
the RpoH-deficient strain K165. Stationary-phase cultures (optical
density of 3) grown at 30°C were analyzed. Fatty acids: 14:0,
myristic acid; 16:0, palmitic acid; 16:1, palmitoleic acid; 17 ,
cis-9,10-methylenehexadecanoic acid (C17 CFA); 18:1,
cis-vaccenic acid; 19,
cis-11,12-methyleneoctadecanoic acid (C19 CFA). CFAs are
formed by methyleneation of the UFAs palmitoleic and
cis-vaccenic acids.
|
|
Mutations in lon, clpP, or
groEL do not affect the degradation of E. coli
CFA synthase.
To determine if a known heat shock protease was
involved in the CFA synthase degradation, we first tested the Lon
protease, which plays an important role in general protein degradation
(7-9, 12, 18). We constructed an isogenic pair of strains
in which strain YYC1328 carried the wild-type lon allele
whereas strain YYC1329 carried a lon null mutation. The
strains were methionine auxotrophs and contained the high-copy-number
cfa plasmid pAYW19 (Table 1). The strains were labeled with
radioactive methionine and then chased with nonradioactive methionine
for 60 min at 37°C (Fig. 3). The rates of CFA synthase degradation
were the same in both strains (Fig. 3 and Table 2), and thus we
concluded that Lon protease was not responsible for CFA synthase degradation.
We then proceeded to test
clpP and
groEL mutants.
Both ClpP and GroEL are heat shock proteins. ClpP (
7-9,
12,
18) is
an active protease when complexed with either ClpA or
ClpX, whereas
GroEL is a chaperone that plays an important role in
protein folding
and assembly but can also facilitate proteolysis of
certain proteins
(
14,
15). We constructed strain YYC1332
carrying both
lon and
clpP null mutations and
found that CFA synthase turnover in
this strain was the same as in the
isogenic wild-type strain (YYC1328)
and in the single
lon
mutant strain YYC1329 (Fig.
3 and Table
2). Likewise, we tested the
groEL strain YYC1338 and found that
the rate of CFA synthase
turnover was identical to that of the
isogenic wild-type strain YYC1336
(Fig.
3 and Table
2). Therefore,
CFA synthase degradation does not
require the function of any
of these heat shock
proteins.
Degradation of CFA synthase is not energy dependent.
The
function of the known heat shock proteins involved in proteolysis is
energy dependent since ATP is required for activity of these proteases
and chaperones (8, 9). We tested the energy dependence of
the protease responsible for the CFA synthase turnover by depleting the
intracellular ATP pool during the chase period. ATP depletion was done
by carbon source starvation alone or together with cyanide treatment
(17, 24). To ascertain that the decrease in intracellular
ATP levels was sufficient to inhibit ATP-dependent proteolysis, we
examined the degradation of the incomplete proteins formed by premature
translation termination in the presence of puromycin. Degradation of
such abnormal peptides is strongly (albeit not completely) dependent on
ATP-dependent proteases, with Lon playing the major role (5, 9,
24). Cultures were treated with puromycin (0.3 mg/ml) for 15 min
at 37°C and then labeled with [35S]methionine-cysteine
for 5 min at 37°C. After labeling, the cells were washed free of
[35S]methionine-cysteine and suspended either in minimal
medium containing glucose or in minimal medium lacking glucose and
supplemented with 1 mM potassium cyanide (KCN) for 30 min at either 37 or 30°C. We then measured the radioactivity present in the
trichloroacetic acid-soluble supernatants (a measure of the proteolytic
degradation of the puromycyl-modified proteins) at both temperatures.
At 37°C the presence of KCN inhibited the degradation of puromycyl
protein fragments by 67%, whereas an inhibition of 44% was seen at
30°C. The energy dependence of CFA synthase degradation was then
examined under these conditions. We found that the rate of CFA synthase degradation was unchanged by the KCN-plus-glucose starvation treatment (Fig. 5). Therefore, ATP-dependent
proteases such as those we have tested and the newly described
HslU-HslV (ClpQ-ClpY) protease (16, 19, 22) are not involved
in CFA synthase degradation. The lack of energy dependence also argues
that an ATP-dependent chaperone such as DnaK is not required to present
CFA synthase to a protease responsible for degradation.
View larger version (106K):
[in this window]
[in a new window]
|
FIG. 5.
Degradation of E. coli CFA synthase is
independent of energy. Autoradiograms of SDS-polyacrylamide gel
separations of immunoprecipitates of pulse-chase experiments are shown.
Lane 1 is the CFA synthase marker, lanes 2 to 5 were from strain
YYC1318, and lane 6 is the cfa null strain YYC1317. Strain
YYC1318 was pulse-labeled for 30 min without chase (lane 2) or chased
for 30 min in the presence of glucose (lane 3). Lanes 4 and 5 were the
same as lane 3 except that the chase was done in the absence of glucose
without or with 1 mM KCN, respectively. Arrows denote the CFA synthase
protein band. The experiments were done with protocol B (Materials and
Methods). Abbreviations are as in Fig. 1.
|
|
| |
DISCUSSION |
Our results show that the loss of CFA synthase activity early in
stationary phase is due to proteolysis rather than to another process
that inactivates the enzyme (e.g., covalent modification or tight
binding of an inhibitory molecule). Our finding that CFA synthase
degradation is dependent on induction of the heat shock regulon was
unexpected since the known targets of heat shock-dependent protein
degradation are abnormal proteins or proteins that play regulatory
rather than metabolic roles (8, 9, 18).
The first question raised by these results is, Why is CFA synthase
rapidly degraded while most other E. coli proteins are stable? Preferential proteolysis of a normally folded protein in
E. coli has often been found to play a regulatory role, for example, the Lon-dependent degradation of the cell division inhibitor SulA during recovery from the SOS response (18). However,
since we know of no regulatory role for either CFA synthase or CFA, it
seems more likely that CFA synthase degradation is of some physiological advantage to E. coli. Note that the decrease
in CFA synthase activity occurs after virtually all of the membrane phospholipid UFA moieties have been converted to CFA (28),
and since cyclopropanation is not reversible (3),
degradation of CFA synthase does not affect the level of CFA formation.
The most straightforward rationale for degradation would be to prevent high levels of CFA formation when the cells reenter exponential growth
and resume synthesis of UFA-containing phospholipids. This rationale
predicts that cells containing CFA-modified phospholipids might grow
more poorly than those containing UFA-containing lipids. However,
strains carrying cfa plasmids have normal exponential-phase growth rates despite efficient conversion of the phospholipid UFA
moieties to CFA (10). Moreover, UFA-auxotrophic strains supplemented with exogenous CFAs (in place of UFAs) grow well (3). Therefore, CFA-containing phospholipids do not prevent rapid growth. Another possibility arises from the fact that CFA synthesis is the major consumer of AdoMet in E. coli. Each
cyclopropanation reaction consumes a molecule of AdoMet, and an
E. coli cell contains 22 million phospholipid molecules,
each of which contains at least one UFA to be converted to CFA. In
contrast, the total consumption of AdoMet in the synthesis of other
molecules (chiefly in methylation of nucleic acids and in the synthesis
of spermidine) is about 10-fold less (calculated from the data of
reference 19). Thus, degradation of CFA synthase may
redirect AdoMet to the synthesis of the other molecules during
exponential growth. Note that synthesis of these nonlipid molecules is
essentially complete when cells enter stationary phase, and thus there
is little or no competition for AdoMet at the time when CFA synthesis
is maximal. Therefore, we propose that degradation of CFA synthase is a
cellular optimization strategy designed for efficient use of a
metabolically expensive intermediate (three ATPs are expended in the
synthesis of each AdoMet molecule). The delay of CFA synthesis until
entry into stationary phase seems appropriate since the only
physiological function of CFA formation presently known is to protect
bacteria from acid shock (2), and it is in stationary phase
where cells are most likely to encounter an acid environment in nature
(13, 23).
It is not clear why CFA synthase degradation is part of the heat shock
response. However, it should be noted that RpoH null mutants grow
poorly even at very low growth temperatures, and this phenotype
indicates that some level of expression of the heat shock regulon is
present at all growth temperatures (12). Heat shock is not a
general stress response and has not been reported to be induced by
entry into stationary phase or by acid shock (12, 13, 23).
One possibility is that the presence of CFA in membrane phospholipids
may hinder growth at high temperatures, but this does not appear to be
the case (see above). However, high-level production of misfolded
proteins induces heat shock in exponentially growing cells
(6), and if misfolded proteins accumulate during entry to
stationary phase, some induction of the heat shock regulon could occur.
Nevertheless, we have no evidence that the rate of CFA synthase
degradation increases as cultures enter stationary phase, although
given the inherently imprecise nature of decay measurements (the
decrements at successive time points are small), only large changes in
degradation rates would be detected. The most straightforward
interpretation of our data is that a protease transcribed from an
RpoH-dependent promoter degrades CFA synthase. We favor this
interpretation because degradation of the protein is not energy
dependent. The lack of dependence on cellular ATP argues against other
interpretations such as a heat shock chaperone being required to fold a
constitutively expressed protease or to present CFA synthase to a
constitutive protease.
The timing of CFA synthesis is controlled by two mechanisms involving
sigma factors, the RpoS-dependent P2 promoter and RpoH-dependent proteolysis. Can these two elements adequately explain the spike in CFA
synthase activity observed as cultures enter stationary phase? In many
cases low levels of a protein are maintained by a rate of degradation
that closely matches the rate of synthesis (7, 18). Upon an
environmental stimulus, the rate of synthesis of the protein is
increased and exceeds the capacity of the degradation apparatus. As
pointed out by Miller (18), this regulatory strategy has
several advantages. The protein can be maintained at a very low level,
but when the synthetic rate is increased such that it no longer matches
the rate of degradation, the levels of protein can increase
dramatically. Moreover, when the protein has completed its task and the
rate of synthesis has decreased, proteolysis results in a rapid return
to normal levels of the protein without the need for dilution by cell
growth. In the case of CFA synthase, the rate of degradation seems
closely matched to the level of enzyme produced from the constitutive
RpoD (70)-dependent P1 promoter. Upon entry into
stationary phase, the large increase in RpoS levels gives activation of
the strong P2 promoter (28), which increases the synthetic
rate of CFA synthase above the rate of degradation, thus producing the
large increase in activity observed. The enzyme then proceeds to modify
the membrane lipids and can subsequently be degraded. Although
transcription of cfa continues in stationary phase
(28), we were unable to radioactively label the CFA synthase
protein in stationary-phase cultures (see above). It therefore seems
likely that degradation plus the generally low translation rate in
stationary-phase cells is responsible for the observed loss of CFA
synthase activity.
| |
ACKNOWLEDGMENTS |
This work was supported by National Institutes of Health grant
AI15650 and by the Deutsche Forschungsgemeinschaft (SFB197, project A7).
| |
FOOTNOTES |
*
Corresponding author. Mailing address: B103 Chemical
and Life Sciences Laboratory, 601 South Goodwin Ave., Urbana, IL 61801. Phone: (217) 333-0425. Fax: (217) 244-6697. E-mail:
j-cronan{at}life.uiuc.edu.
| |
REFERENCES |
| 1.
|
Baker, T. A.,
A. D. Grossman, and C. A. Gross.
1984.
A gene regulating the heat shock response in Escherichia coli also affects proteolysis.
Proc. Natl. Acad. Sci. USA
81:6779-6783[Abstract/Free Full Text].
|
| 2.
|
Chang, Y. Y., and J. E. Cronan, Jr.
1999.
Membrane cyclopropane fatty acid content is a major factor in acid resistance of Escherichia coli.
Mol. Microbiol.
33:249-259[CrossRef][Medline].
|
| 3.
|
Cronan, J. E., Jr.,
W. D. Nunn, and J. G. Batchelor.
1974.
Studies on the biosynthesis of cyclopropane fatty acids in Escherichia coli.
Biochim. Biophys. Acta
348:63-75[Medline].
|
| 4.
|
Eichel, J.,
Y. Y. Chang,
D. Riesenberg, and J. E. Cronan, Jr.
1999.
Effect of ppGpp on Escherichia coli cyclopropane fatty acid synthesis is mediated through the RpoS sigma factor (S).
J. Bacteriol.
181:572-576[Abstract/Free Full Text].
|
| 5.
|
Goff, S. A.,
L. P. Casson, and A. L. Goldberg.
1984.
Heat shock regulatory gene htpR influences rates of protein degradation and expression of the lon gene in Escherichia coli.
Proc. Natl. Acad. Sci. USA
81:6647-6651[Abstract/Free Full Text].
|
| 6.
|
Goff, S. A., and A. L. Goldberg.
1985.
Production of abnormal proteins in E. coli stimulates transcription of lon and other heat shock genes.
Cell
41:587-595[CrossRef][Medline].
|
| 7.
|
Gottesman, S.
1996.
Proteases and their targets in Escherichia coli.
Annu. Rev. Genet.
30:465-506[CrossRef][Medline].
|
| 8.
|
Gottesman, S.,
S. Wickner,
Y. Jubete,
S. K. Singh,
M. Kessel, and M. Maurizi.
1995.
Selective, energy-dependent proteolysis in Escherichia coli.
Cold Spring Harbor Symp. Quant. Biol.
60:533-548[Abstract/Free Full Text].
|
| 9.
|
Gottesman, S.,
S. Wickner, and M. R. Maurizi.
1997.
Protein quality control: triage by chaperones and proteases.
Genes Dev.
11:815-823[Free Full Text].
|
| 10.
|
Grogan, D. W., and J. E. Cronan, Jr.
1984.
Cloning and manipulation of the Escherichia coli cyclopropane fatty acid synthase gene: physiological aspects of enzyme overproduction.
J. Bacteriol.
158:286-295[Abstract/Free Full Text].
|
| 11.
|
Grogan, D. W., and J. E. Cronan, Jr.
1997.
Cyclopropane ring formation in membrane lipids of bacteria.
Microbiol. Mol. Biol. Rev.
61:429-441[Abstract].
|
| 12.
|
Gross, C. A.
1996.
Function and regulation of the heat shock proteins, p. 1382-1399.
In
F. C. Neidhardt, R. Curtiss III, J. L. Ingraham, E. C. C. Lin, K. B. Low, B. Magasanik, W. S. Reznikoff, M. Riley, M. Schaechter, and H. E. Umbarger (ed.), Escherichia coli and Salmonella typhimurium: cellular and molecular biology, 2nd ed., vol. 1. American Society for Microbiology, Washington, D.C.
|
| 13.
|
Hengge-Aronis, R.
1996.
Regulation of gene expression during entry into stationary phase, p. 1497-1512.
In
F. C. Neidhardt, R. Curtiss III, J. L. Ingraham, E. C. C. Lin, K. B. Low, B. Magasanik, W. S. Reznikoff, M. Riley, M. Schaechter, and H. E. Umbarger (ed.), Escherichia coli and Salmonella typhimurium: cellular and molecular biology, 2nd ed., vol. 1. American Society for Microbiology, Washington, D.C.
|
| 14.
|
Kandror, O.,
L. Busconi,
M. Sherman, and A. L. Goldberg.
1994.
Rapid degradation of an abnormal protein in Escherichia coli involves the chaperones GroEL and GroES.
J. Biol. Chem.
269:23575-23582[Abstract/Free Full Text].
|
| 15.
|
Kandror, O.,
M. Sherman, and A. Goldberg.
1999.
Rapid degradation of an abnormal protein in Escherichia coli proceeds through repeated cycles of association with GroEL.
J. Biol. Chem.
274:37743-37749[Abstract/Free Full Text].
|
| 16.
|
Kessel, M.,
W. Wu,
S. Gottesman,
E. Kocsis,
A. C. Steven, and M. R. Maurizi.
1996.
Six-fold rotational symmetry of ClpQ, the E. coli homolog of the 20S proteasome, and its ATP-dependent activator, ClpY.
FEBS Lett.
398:274-278[CrossRef][Medline].
|
| 17.
|
Maurizi, M. R.,
P. Trisler, and S. Gottesman.
1985.
Insertional mutagenesis of the lon gene in Escherichia coli: lon is dispensable.
J. Bacteriol.
164:1124-1135[Abstract/Free Full Text].
|
| 18.
|
Miller, C. G.
1996.
Protein degradation and proteolytic modification, p. 938-954.
In
F. C. Neidhardt, R. Curtiss III, J. L. Ingraham, E. C. C. Lin, K. B. Low, B. Magasanik, W. S. Reznikoff, M. Riley, M. Schaechter, and H. E. Umbarger (ed.), Escherichia coli and Salmonella typhimurium: cellular and molecular biology, 2nd ed., vol. 1. American Society for Microbiology, Washington, D.C.
|
| 19.
|
Missiakas, D.,
F. Schwager,
J. M. Betton,
C. Georgopoulos, and S. Raina.
1996.
Identification and characterization of HsIV HsIU (ClpQ ClpY) proteins involved in overall proteolysis of misfolded proteins in Escherichia coli.
EMBO J.
15:6899-6909[Medline].
|
| 20.
|
Neidhardt, F. C., and H. E. Umbarger.
1996.
Chemical composition of Escherichia coli, p. 13-16.
In
F. C. Neidhardt, R. Curtiss III, J. L. Ingraham, E. C. C. Lin, K. B. Low, B. Magasanik, W. S. Reznikoff, M. Riley, M. Schaechter, and H. E. Umbarger (ed.), Escherichia coli and Salmonella typhimurium: cellular and molecular biology, 2nd ed., vol. 1. American Society for Microbiology, Washington, D.C.
|
| 21.
|
Neidhardt, F. C.,
R. A. VanBogelen, and E. T. Lau.
1983.
Molecular cloning and expression of a gene that controls the high- temperature regulon of Escherichia coli.
J. Bacteriol.
153:597-603[Abstract/Free Full Text].
|
| 22.
|
Rohrwild, M.,
O. Coux,
H. C. Huang,
R. P. Moerschell,
S. J. Yoo,
J. H. Seol,
C. H. Chung, and A. L. Goldberg.
1996.
HslV-HslU: a novel ATP-dependent protease complex in Escherichia coli related to the eukaryotic proteasome.
Proc. Natl. Acad. Sci. USA
93:5808-5813[Abstract/Free Full Text].
|
| 23.
|
Slonczewski, J. L., and J. W. Foster.
1996.
pH-regulated genes and survival at extreme pH, p. 1539-1549.
In
F. C. Neidhardt, R. Curtiss III, J. L. Ingraham, E. C. C. Lin, K. B. Low, B. Magasanik, W. S. Reznikoff, M. Riley, M. Schaechter, and H. E. Umbarger (ed.), Escherichia coli and Salmonella typhimurium: cellular and molecular biology, 2nd ed., vol. 1. American Society for Microbiology, Washington, D.C.
|
| 24.
|
Straus, D. B.,
W. A. Walter, and C. A. Gross.
1988.
Escherichia coli heat shock gene mutants are defective in proteolysis.
Genes Dev.
2:1851-1858[Abstract/Free Full Text].
|
| 25.
|
Studier, F. W., and B. A. Moffatt.
1986.
Use of bacteriophage T7 RNA polymerase to direct selective high-level expression of cloned genes.
J. Mol. Biol.
189:113-130[CrossRef][Medline].
|
| 26.
|
Sweetman, G.,
M. Trinei,
J. Modha,
F. P. Kuselm,
J. I. Fishov,
D. Joseleau-Petit,
C. Redman,
P. Farmer, and V. Norris.
1996.
Electrospray ionization mass spectrometric analysis of phospholipids of Escherichia coli.
Mol. Microbiol.
20:233-234[CrossRef][Medline].
|
| 27.
|
Vogel, H. J., and D. M. Bonner.
1956.
Acetyl-ornithinase of Escherichia coli: partial purification and some properties.
J. Biol. Chem.
218:97-106[Free Full Text].
|
| 28.
|
Wang, A. Y., and J. E. Cronan, Jr.
1994.
The growth phase-dependent synthesis of cyclopropane fatty acids in Escherichia coli is the result of an RpoS(KatF)-dependent promoter plus enzyme instability.
Mol. Microbiol.
11:1009-1017[CrossRef][Medline].
|
| 29.
|
Wang, A. Y.,
D. W. Grogan, and J. E. Cronan, Jr.
1992.
Cyclopropane fatty acid synthase of Escherichia coli: deduced amino acid sequence, purification, and studies of the enzyme active site.
Biochemistry
31:11020-11028[CrossRef][Medline].
|
Journal of Bacteriology, August 2000, p. 4288-4294, Vol. 182, No. 15
0021-9193/00/$04.00+0
Copyright © 2000, American Society for Microbiology. All rights reserved.
This article has been cited by other articles:
-
Boissier, F., Bardou, F., Guillet, V., Uttenweiler-Joseph, S., Daffe, M., Quemard, A., Mourey, L.
(2006). Further Insight into S-Adenosylmethionine-dependent Methyltransferases: STRUCTURAL CHARACTERIZATION OF Hma, AN ENZYME ESSENTIAL FOR THE BIOSYNTHESIS OF OXYGENATED MYCOLIC ACIDS IN MYCOBACTERIUM TUBERCULOSIS. J. Biol. Chem.
281: 4434-4445
[Abstract]
[Full Text]
-
Hartig, C., Loffhagen, N., Harms, H.
(2005). Formation of trans Fatty Acids Is Not Involved in Growth-Linked Membrane Adaptation of Pseudomonas putida. Appl. Environ. Microbiol.
71: 1915-1922
[Abstract]
[Full Text]
-
Kim, B. H., Kim, S., Kim, H. G., Lee, J., Lee, I. S., Park, Y. K.
(2005). The formation of cyclopropane fatty acids in Salmonella enterica serovar Typhimurium. Microbiology
151: 209-218
[Abstract]
[Full Text]
-
Weichart, D., Querfurth, N., Dreger, M., Hengge-Aronis, R.
(2003). Global Role for ClpP-Containing Proteases in Stationary-Phase Adaptation of Escherichia coli. J. Bacteriol.
185: 115-125
[Abstract]
[Full Text]
-
Kremer, L., Guerardel, Y., Gurcha, S. S., Locht, C., Besra, G. S.
(2002). Temperature-induced changes in the cell-wall components of Mycobacterium thermoresistibile. Microbiology
148: 3145-3154
[Abstract]
[Full Text]
Top
Abstract
Introduction
