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Journal of Bacteriology, October 2001, p. 5982-5990, Vol. 183, No. 20
Departments of
Microbiology1and
Biochemistry,2 University of Illinois
at Urbana-Champaign, Urbana, Illinois 61801
Received 2 May 2001/Accepted 24 July 2001
In Escherichia coli expression of the genes of fatty
acid degradation (fad) is negatively regulated at the
transcriptional level by FadR protein. In contrast the unsaturated
fatty acid biosynthetic gene, fabA, is positively
regulated by FadR. We report that fabB, a second
unsaturated fatty acid biosynthetic gene, is also positively regulated
by FadR. Genomic array studies that compared global transcriptional
differences between wild-type and fadR-null mutant
strains, as well as in cultures of each strain grown in the presence of
exogenous oleic acid, indicated that expression of fabB
was regulated in a manner very similar to that of fabA
expression. A series of genetic and biochemical tests confirmed these
observations. Strains containing both fabB and fadR mutant alleles were constructed and shown to
exhibit synthetic lethal phenotypes, similar to those observed in
fabA fadR mutants. A fadR strain was
hypersensitive to cerulenin, an antibiotic that at low concentrations
specifically targets the FabB protein. A transcriptional fusion of
chloramphenicol acetyltransferase (CAT) to the fabB
promoter produces lower levels of CAT protein in a strain lacking
functional FadR. The ability of a putative FadR binding site within the
fabB promoter to form a complex with purified FadR
protein was determined by a gel mobility shift assay. These experiments
demonstrate that expression of fabB is positively regulated by FadR.
Bacteria regulate membrane fluidity
by manipulating the relative levels of saturated and unsaturated fatty
acids within the phospholipids of their membrane bilayers (1,
13). There are eight known genes (fab) involved in
fatty acid biosynthesis in Escherichia coli (reviewed in
references 8 and 34). Of these, only
fabA and fabB are specifically required for the
synthesis of unsaturated fatty acids (4, 5, 12, 46).
Likewise, there are at least five separate gene products involved in
the degradation of long-chain fatty acids to acetyl coenzyme A (for a
review, see reference 34). The FadR regulatory protein
negatively controls expression of the genes of the fatty acid
degradation pathway (33, 40) and also functions as a
positive regulator of unsaturated fatty acid synthesis (19, 29,
30, 38).
Only two unique biochemical reactions are required to specifically
produce unsaturated fatty acids in the overall course of fatty acid
biosynthesis in E. coli (4, 5, 12, 46). When the growing acyl chain coupled to acyl carrier protein (ACP) reaches the 3-hydroxydecanoyl-ACP stage, either of two enzymes can carry out
the dehydration reaction to produce trans-2-decenoyl-ACP. In
vitro, the 3-ketoacyl-ACP dehydratases, FabZ and FabA, both have broad
and overlapping ranges of substrate chain length specificity (28). In vivo, FabZ is involved in the dehydration of all
chain lengths of 3-hydroxyacyl-ACPs and seems especially important in lipid A biosynthesis (36). Genetic data argue that the
activity of FabA in vivo is restricted to 10 carbon substrates. The
enzyme not only catalyzes the dehydration of 3-hydroxydecanoyl-ACP but also isomerizes the trans-2-enoyl bond of the ACP-bound
substrate to the cis-3 isomer (3, 28). This
isomerization places the nascent acyl chain in the unsaturated fatty
acid synthetic pathway. The 3-ketoacyl-ACP synthase I (FabB) enzyme
appears to be required for elongation of the cis-3
decenoyl-ACP produced by FabA and is known to be the primary factor in
determining cellular unsaturated fatty acid content (10).
The Claissen-type condensation of malonyl-ACP with cis-3
decenoyl-ACP catalyzed by FabB produces
cis-5-ene-3-ketododecenoyl-ACP, which is competent to
undergo all the subsequent reactions typical of fatty acid
biosynthesis. Ultimately this results in production of cis-9
hexadecenoyl and cis-11 octadecenoyl chains, which are incorporated into phospholipid (34).
The regulation of unsaturated fatty acid biosynthesis is complex. The
fabA gene is known to have a strong promoter that is positively regulated by FadR (19, 29, 30) as well as a
weaker constitutive promoter. The reasons why a regulatory factor for fatty acid degradation is involved in regulating unsaturated fatty acid
biosynthesis remain obscure. A model advanced by Cronan and Subrahmanyam (15) addresses the issue of why it seems
advantageous to have two promoters for fabA but fails to
answer the question of why FadR regulates fabA per se.
DiRusso and Nyström (21) have postulated that FadR
interacts with a number of other regulatory activities to coordinate
lipid biosynthesis and degradation in response to stress and aging.
While this seems an attractive proposal, it still begs the question of
why the synthesis of unsaturated acids in particular, as opposed to
that of saturated fatty acids, is regulated by FadR. Experimental
evidence that both genes involved in unsaturated fatty acid
biosynthesis are regulated similarly would discount the possibility
that FadR regulation of fabA is merely fortuitous or
vestigial in nature. Computer-assisted searches for consensus FadR
recognition sites within the E. coli genome identify
fabB as a potential target of FadR regulation
(45). It should be noted that although several reviews
state that fabB is positively regulated by FadR, neither
these reports (2, 18, 21) nor the specific reference cited
therein (19) contains data supporting this claim. We
report several different lines of evidence showing that FadR positively
regulates fabB transcription.
Bacterial strains and plasmids.
The bacterial strains and
plasmids used in this study are listed in Table
1. Unless otherwise indicated, strains
were obtained from local laboratory stocks or from the E. coli Genetic Stock Center (CGSC) (Yale University, New Haven,
Conn.). Phage transductions and other basic genetic techniques were
generally carried out as previously described in reference
53. Strain CAG18497 is from the ordered Tn10
collection of Singer and coworkers (48). Strains JWC264,
JWC286, and JWC287 were made by P1vir
transduction of the fadR613::Tn10
allele of CAG18497 into strains MG1655, M8, and M5, respectively.
Strain JWC264 was selected on rich broth plates containing tetracycline
at 37°C. Strain JWC276 is a fabB::CAT (chloramphenicol acetyltransferase) transcriptional fusion in the
strain MG1655 background containing a wild-type copy of the fabB gene expressed from the araBAD promoter of
plasmid pARA14 (7). Strain JWC277 was made by transduction
of fadR613::Tn10 from strain CAG18497
into strain JWC276 and selecting for tetracycline resistance at 30°C
on rich broth plates supplemented with 0.01% oleate.
0021-9193/01/$04.00+0 DOI: 10.1128/JB.183.20.5982-5990.2001
Copyright © 2001, American Society for Microbiology. All rights reserved.
Escherichia coli FadR Positively
Regulates Transcription of the fabB Fatty Acid
Biosynthetic Gene
ABSTRACT
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
INTRODUCTION
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
MATERIALS AND METHODS
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
TABLE 1.
Bacterial strains used in this study
(51). All enzymes and
buffers were from Gibco-BRL (Gaithersburg, Md.) or New England Biolabs
(Beverly, Mass.). Recombinants were recovered on rich media containing
25 µg of a combination of clavulanate potassium and ticarcillin
disodium (CPTD) (Timentin; SmithKline Beecham Pharmaceuticals,
Philadelphia, Pa.) per ml. Plasmid DNA was recovered from
CPTD-resistant colonies and screened by restriction digestions for the
presence of the fabB gene. Several constructs were
transformed into M5, and the abilities of the recombinant strains to
grow at 42°C in the presence of arabinose were examined. A plasmid capable of supporting growth at 42°C was retained as
pARAfabB.
The fabB::CAT fusion strains were made by a
modification of the 
Red-mediated recombination method of Datsenko
and Wanner (16). A primer having 41 bases of homology to
the start codon region of fabB at its 5' end was
synthesized. This primer also included an 11-base sequence containing
translation stops (TAA) in all three reading frames immediately
downstream of the fabB homologous sequence, followed by 27 bases of homology to the beginning of the CAT gene of plasmid pKD3
(23). The sequence of this primer was
5'-GAATGAAACGTGCAGTGATTACTGGCCTGGGCATTGTTTCCTAACTAACTAATCAGGAGCTAAGGAAGCTAAAATGGAG-3'. A second primer included 41 bases of homology to the reverse
complement of the stop codon region of fabB at the 5' end,
and 25 bases of homology to the P1 site of pKD3 on the 3' end. The
sequence of this primer was
5'-GAATTAATCTTTCAGCT TGCGCATTACCAGCGTGGCGTTG-GTGTGTAGGCTGGAGCTGCTTC GAAG-3'.
These primers were used to amplify the cat gene of plasmid pKD3 in a standard PCR. The enzyme and buffers in these reactions were
from Gibco-BRL. The purified 1.5-kb PCR product was transformed into
strain BW25113 (16), and the cells were plated onto rich broth agar containing chloramphenicol and supplemented with
0.01% oleate. Chloramphenicol-resistant colonies were isolated, and phage P1vir was grown on 10 ml cultures of these
strains. The resulting lysates were used to transduce MG1655 to
chloramphenicol resistance. The resulting strain, JWC276, contains the
CAT gene, including its ribosome-binding site, downstream of 3 translational stop codons, which are located 12 codons upstream from
the normal fabB start. In this strain, CAT is expressed from
the fabB promoter. Strain JWC277 was made by transducing the
fadR613::Tn10 insertion of CAG18497
into JWC276 and selecting for tetracycline resistance on rich-broth
plates containing 0.01% oleate at 30°C.
Microbial methods. Rich broth contained 10 g of tryptone, 5 g of NaCl and 1 g of yeast extract per liter. Minimal medium was M9 (44). Glucose was added to 0.2%, and acetate was added to 0.4% by weight. Oleic acid was neutralized with KOH, solubilized in Tergitol NP-40, and used at a final concentration of 100 µg/ml. Antibiotics were used at the following concentrations: tetracycline HCl, 12 µg/ml; kanamycin sulfate, 50 µg/ml; and CPTD, 25 µg/ml. CPTD was used for the reasons previously described (51). Chloramphenicol was present at 34 µg/ml, unless otherwise indicated. The detergent, antibiotics, and most bulk chemicals were obtained from Sigma (St. Louis, Mo.). Solid media contained 1.5% (wt/vol) BactoAgar (Difco, Milwaukee, Wis.). The phenotypes of the various fab mutants were verified by testing for a requirement for unsaturated fatty acids (oleate) on rich-broth plates.
Cultures for genomic expression analyses were grown in minimal M9 medium (35) with 0.4% glycerol as carbon source. Exponentially growing cultures (doubling time of about 2 h) were grown aerobically at 37°C with vigorous rotary shaking. The cultures were repeatedly diluted to ensure exponential growth and were harvested at a cell density of about 108 cells/ml (about 1/20 of the maximal cell density attained in this medium). The growth curves of the fadR and wild-type strains were indistinguishable in the exponential phase of growth in all of the media tested, although the wild-type strain reached a slightly higher maximal cell density. Cerulenin was dissolved in chloroform, and various volumes of this solution were placed into empty sterile culture tubes and allowed to evaporate to dryness at room temperature. One milliliter of rich broth containing kanamycin was inoculated 1:200 with overnight cultures of strain JWC288 or strain JWC289 and was added to each tube, and the cultures were incubated for 6 h at 37°C in a roller drum. Cell growth was measured spectrophotometrically at 600 nm.Genomic expression profiling analysis. The Sigma-Genosys (The Woodlands, Tex.) E. coli Panorama array system was used to evaluate the expression of each of the 4,290 open reading frames (ORFs) in the E. coli genome. Experimentally this involved measuring differences in expression in cells grown in M9-glycerol medium with and without supplementation with 0.01% oleic acid. Differential expression was also measured between the reference strain, MG1655, and the isogenic fadR mutant, JWC264, when both strains were grown in M9-glycerol. A third experiment measured the transcriptional differences between the JWC264 (fadR) strain grown with and without oleate supplementation.
RNA isolation and sample handling procedures were those of Tao et al. (52). Briefly, early-log-phase cultures were removed from the shaking incubator and immediately decanted into an equal volume of boiling lysis buffer composed of 2% sodium dodecyl sulfate (SDS), 250 mM sodium acetate (pH 4.5), and 20 mM EDTA. The lysed cells were extracted twice with 60°C phenol equilibrated with 100 mM sodium acetate at pH 4.5 and then extracted once with phenol-chloroform (1:1) at room temperature. Nucleic acids were precipitated in 0.5 volumes of 2-propanol and rinsed with a small volume of ice-cold 70% ethanol. The pellet was air dried for 10 to 15 min and dissolved in a small volume of sterile RNase-free water. Approximately 20 U of RNase inhibitor (Promega, Madison, Wis.) was added. Residual DNA was removed by incubation with 10 U of RQ1 RNase-free DNase I (Promega) at 37°C for about 1 h. The RNA samples were applied to RNeasy columns (Qiagen) and recovered in 30 µl of sterile diethyl pyrocarbonate-treated water. The RNA was quantitated by absorbance at 260 nm against a water blank. Sample purity was determined by the A260/A280 ratio. Total yields of 10 to 15 µg of RNA with A260/A280 ratios of 1.8 to 2.1 were routinely observed. The RNA samples consist of a mixture of the various stable RNAs, including tRNA and rRNA, as well as mRNA. To restrict probe synthesis to mRNA, the Panorama E. coli cDNA labeling and hybridization kit (Sigma-Genosys) was used in a standard cDNA synthesis reaction. This kit consists of an equimolar mixture of each of 4,290 C terminus-specific primers. Sample RNA (1 µg) was added to a solution containing a 0.33 mM concentration each of dATP, dGTP, and dTTP and 4 µl of the Sigma-Genosys primer mix. The reaction mixture was brought to a total volume of 25 µl in first-strand synthesis buffer in a small, thin-walled Eppendorf tube. The reaction mixture was placed into a thermal cycler, heated to 90°C for 2 min, and then linearly cooled to 42°C over a 20-min period. On reaching 42°C, 200 U of Superscript II RNase H
Reverse Transcriptase
(Gibco-BRL) was added along with 20 U of RNase Inhibitor (Promega) and
20 to 30 µCi of [-33P]dCTP (3,000 Ci/mmol)
(NEN Life Science Products, Inc., Boston, Mass.). The reaction mixture
was incubated at 42°C for an additional 3 h. Unincorporated
nucleotides were removed by centrifugation through Sephadex G-25 gel
filtration columns (Boehringer Mannheim, Indianapolis, Ind.).
Incorporation levels of 80 to 90% of the total label were routinely
achieved by this procedure.
Typically, about 10 to 15 ng of cDNA was recovered from the reverse
transcriptase reactions. Estimates of the mRNA content of E. coli range from 3% of total RNA based on pulse-labeling studies
to 1.4% based on hybridization experiments (6, 31). These
estimates place the total amount of mRNA available in the reverse
transcriptase reaction in the neighborhood of 14 to 30 ng. This closely
matches the amount of cDNA recovered from the reaction. A crude
estimate of the molar amounts of mRNA, based on an average mRNA length
of 1 to 2 kb, places the total number of transcripts at about 100 pmol
or approximately 1010 molecules. The available
nucleoside triphosphates in the reaction mixture could theoretically
support the synthesis of about 1011 molecules of
1-kb single-stranded DNA. Under these conditions, with limiting
template concentration, excess nucleotides and primer, and a single
annealing cycle, the cDNA products approximate both the complexity and
the relative abundance of the individual ORFs within the mRNA population.
The hybridization and washing steps were carried out as recommended by
the manufacturer of the array. Hybridization buffer consists of 5×
SSPE (1× SSPE is 0.18 M NaCl, 10 mM
NaH2PO4, and 1 mM EDTA [pH
7.7]), 2% SDS, and 1× Denhardt's reagent supplemented with 100 µg
of sheared and sonicated herring sperm DNA per ml. The membranes were
prehybridized overnight at 65°C in a roller oven. After
prehybridization, the buffer was exchanged for 6 ml of fresh, prewarmed
buffer. The cDNA probe sample was heated to 94°C for a few minutes
and then added to the hybridization bottle. Samples were hybridized for
at least 18 h at 65°C in the roller oven.
When hybridization was complete the membranes were washed twice in 50 ml of a wash solution consisting of 0.5× SSPE and 0.2% SDS at room
temperature. This was followed by two washes at 65°C with the same
wash solution for 20 min each in the hybridization oven. After the
final wash step, the membranes were placed onto blotting paper, wrapped
in plastic food wrap, and placed in a PhosphorImager cassette
(Molecular Dynamics, Sunnyvale, Calif.). The phosphor screen was
exposed for 2 to 3 days prior to quantitation.
The TIFF images of each blot were analyzed for pixel depth at each spot
position on the membrane using Molecular Dynamics ImageQuant software.
The identity of each spot was determined by using a series of ad hoc
Perl and AWK scripts and the final output was imported into a
spreadsheet. The relative order of gene expression was determined by
sorting the data set by ascending ratio value such that the denominator
of each ratio is the control sample value. Relative rank was determined
by vertical position within the sorted spreadsheet in a manner similar
to that of Wei et al. (56). In E. coli, the
gene having the greatest increase in expression relative to the control
had a relative rank of 1 whereas the gene having the greatest decrease
in gene expression had a relative rank of 4,290.
CAT protein assays. Cells to be assayed were grown overnight in rich media containing the appropriate antibiotics and then diluted 1:200 into 10 ml of rich broth with no antibiotics present. The experimental cultures were incubated at 37°C for about 3 h. Cells were concentrated by low-speed centrifugation and washed twice in ice-cold lysis buffer consisting of 40 mM Tris-HCl (pH 7.8), 1 mM EDTA, and 150 mM NaCl. After cells were resuspended in a small volume of lysis buffer, they were sonicated three times for 15 s each with an S&M sonicator at 40% full power (Sonics and Materials, Inc., Danbury, Conn.). The sonicate was centrifuged for 30 min at high speed (approximately 10,000 × g) to remove insoluble material. Protein concentrations were determined by the method of Bradford using the Bio-Rad (Richmond, Calif.) protein assay kit with bovine serum albumin as a standard.
A CAT assay kit, produced by Roche, Inc., was used to measure the mass of CAT protein in cell extracts. The method is based on a sandwich enzyme-linked immunosorbent assay involving antibodies to CAT bound to the surface of microtiter wells. The cells were lysed, and the cell extracts containing CAT enzyme were added to the wells. Following a 1-h incubation at 37°C the sample extract was discarded. The wells were then washed five times with 250 µl of a washing buffer composed of phosphate-buffered saline and Tween 20. A digoxigenin-labeled antibody to CAT was added, and the plate was incubated for another hour at 37°C. The unbound digoxigenin-labeled antibody was removed, and the wells were washed five times as before. A solution containing peroxidase conjugated to an anti-digoxigenin antibody was added to each well, and the plate was incubated for another hour at 37°C. The peroxidase conjugate was removed, and the plate was washed five times again. The final reaction involved adding the chromogenic peroxidase substrate, 2,2'-azino-di-(3-ethylbenzthiazoline sulfonate), and allowing color development for half an hour at room temperature. The absorbance of each well was read at 405 nm, and a reference was taken at 490 nm. A standard activity curve was generated using authentic CAT protein provided by the manufacturer. The standard curve and samples were plotted as A405
A490 by CAT enzyme concentration.
Gel mobility shift analysis.
A 950-bp DNA fragment including
approximately 500 bp of DNA on either side of the fabB start
codon was produced by PCR amplification. In this case a standard
Taq polymerase amplification reaction was carried out in the
presence of a small quantity of [-33P]dCTP
(3,000 Ci/mmol) to generate a labeled DNA product. The primers were
5'-AATGTAAGCGTTGTAATTGC-3' on the 5' side and
5'-TGTGCGGAAGTCGCACACGC-3' on the 3' side. The 950-bp
product was purified on a Qiagen PCR purification column.
-D-thiogalactopyranoside (IPTG) to the culture. The cells were concentrated by low-speed centrifugation and washed twice in a lysis buffer consisting of 50 mM
sodium phosphate (pH 8.0) and 300 mM NaCl. After the cells were
resuspended in a small volume of lysis buffer, they were sonicated for
1 min on ice at 40% of full power. The cell extract was centrifuged
for 30 min at high speed (approximately 10,000 × g) to
remove insoluble material. The fusion protein was prepared by nickel
chelate chromatography of the cell-free lysate. The cleared lysate was
loaded onto a (0.5 by 2 cm) nickel-nitrilotriacetic acid column
(Qiagen, Inc.) and washed with 40 ml of a buffer consisting of 50 mM
sodium phosphate (pH 6.0), 300 mM NaCl, and 10% glycerol. A 30-ml
linear gradient of imidazole, from 0 to 500 mM, was used to elute the
recombinant FadR. The protein-containing fractions were pooled and
dialyzed against a buffer of 20 mM Tris-HCl (pH 7.5), 50 mM NaCl, and 1 mM EDTA containing 1 mM dithiothreitol. Polyacrylamide gel
electrophoresis showed the FadR protein to be substantially pure at
this stage. Glycerol was added to a final concentration of 5 mM, and
the protein was stored in small aliquots at 80°C until use. Protein
concentrations were determined as above.
Gel retardation assays were performed essentially as described by Henry
and Cronan (30). Briefly, the basic binding reaction consisted of adding 10 µg of FadR protein to 100 ng of labeled DNA
(approximately 50,000 cpm) in a buffer composed of 12 mM HEPES-NaOH (pH
7.9), 5 mM Tris-HCl (pH 7.9), 60 mM KCl, 1 mM EDTA, 1 mM
dithiothreitol, 1 µg of poly(dI-dC), 100 µg of bovine serum albumin
and 12% (vol/vol) glycerol. Under these conditions an almost equimolar
ratio of FadR dimer (173 pmol) to DNA (162 pmol) is present. The total volume of the binding assay reaction mixtures was 20 µl. The reaction mixture was allowed to stand at room temperature for 15 to 20 min and
then loaded onto a low-ionic-strength, 4% polyacrylamide gel
(8). The preequilibrated gel was run for approximately 3 h at 40 mA at 4°C with constant buffer recirculation. The gel was removed from the electrophoresis apparatus, placed on blotting paper, and dried at 80°C under vacuum. The dried gel was placed in an
autoradiography cassette and exposed to Kodak (Rochester, N.Y.) XAR5
film overnight.
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RESULTS |
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Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
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Genomic array experiments indicate that fabB is positively regulated by FadR. Genomic array analysis of global differential transcription patterns in bacteria is useful in defining the extents of metabolic regulons in bacteria (44, 52, 56). A transcriptional regulator such as FadR, having both positive and negative regulatory roles, provides a good test of this experimental approach. Data sets of differential transcription analysis from three separate experimental conditions involving various combinations of FadR and long-chain fatty acid supplementation have been assembled and are available online at http://www.life.uiuc.edu/~jwcampbe. Analysis of these data confirms a number of features of the current model of FadR regulation.
The FadR protein is known to bind long-chain fatty acyl coenzyme A (acyl-CoA) at nanomolar concentrations (22, 30). When FadR binds acyl-CoA, the ability of the protein to bind DNA is greatly decreased and it no longer functions as an effective transcriptional repressor or activator (11, 19, 30). Transcription of fadBA (encoding the two subunits of the fatty acid degradation complex) is higher in wild-type strains grown in the presence of oleate than in the same strains grown without fatty acids (9) (Table 2). Conversely, transcription of fabA decreased in the absence of FadR or in the presence of long-chain fatty acids (29, 30) (Table 2).
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-oxidation regulon (23, 40). The third
data set was produced by comparing gene expression patterns in strain
JWC264 grown in the presence and absence of 0.01% oleic acid. Not
surprisingly, the magnitude of the differential transcriptional response of FadR regulated genes was greatest in the data set that
compared the global transcriptional patterns of the FadR null mutant
with the wild-type strain. The data set produced by comparing the
global transcriptional activities in a wild-type FadR strain in the
presence and absence of oleic acid was similar to that observed in the
FadR mutant versus wild type experiment. However, the overall magnitude
of the differential transcription of FadR regulated genes was generally
lower. This was probably due to residual FadR DNA binding activity in
the presence of long-chain acyl-CoAs, versus the complete absence of
FadR in strain JWC264.
The fadBA -oxidation operon is transcribed from a
FadR-regulated promoter (9), and thus, the parallel
behavior of these two genes within the global transcriptional arrays
was expected. However, fabA and fabB are
well-separated transcriptional units (one-fifth of the genetic map).
The major fabA promoter is regulated by FadR and is
downstream of a second weak constitutive promoter. The promoter of
fabB is not as well characterized (32) but is likely to be complex. The similarity of the differential
transcriptional responses of fabB to fabA under
the various experimental conditions suggests that fabB
expression is modulated by a mechanism similar to that of
fabA.
Temperature-sensitive fabB mutations are synthetically lethal with fadR. The first indications that fabA was positively regulated by FadR were the discoveries that fadR strains contained abnormally low levels of cellular unsaturated fatty acid and, more definitively, that fabA(Ts) fadR double mutants were unable to grow even at low temperature unless unsaturated fatty acids were provided (38). The explanation for this finding is that in the presence of functional FadR, sufficient mutant FabA is produced to provide a level of isomerase activity that satisfies cellular unsaturated fatty acid requirements. Indeed, fabA(Ts) strains have been shown to contain abnormally low levels of cellular unsaturated fatty acid at permissive temperatures (38). In contrast, even at the permissive temperature (30°C), fabA(Ts) fadR strains are unable to synthesize enough mutant FabA to provide sufficient unsaturated fatty acid for functional cell membranes (15). Our interpretation of these data is that E. coli can tolerate normal levels of expression of a FabA enzyme of compromised catalytic efficiency or low levels of expression of the wild-type enzyme but cannot survive low levels of expression of a catalytically compromised mutant enzyme.
To test whether fabB mutants might also be synthetically lethal with fadR mutations, the behaviors of fabB(Ts) and fabB(Ts) fadR strains grown under various conditions were examined. Each strain was grown overnight on medium supplemented with 0.01% oleate prior to inoculating experimental cultures. The cultures were incubated at 30 or 42°C overnight and then examined for growth (Table 3). The fabA(Ts) strain M8 grew at 30°C in the presence or absence of oleate, but at 42°C the strain grew only when supplemented with oleate (4, 5). In contrast, at either temperature a fabA(Ts) fadR double mutant grew only in the presence of exogenous oleate (38). A phenotypic behavior that paralleled the fabA case was shown by fabB(Ts) and fabB(Ts) fadR strains. The fabB(Ts) strain M5 required oleate for growth only at 42°C, whereas the fabB(Ts) fadR strain, JWC287, failed to grow at either temperature unless oleate was provided. Thus, the synthetic lethal behavior towards fadR reported for fabA(Ts) mutants was also found for a fabB(Ts) mutant. This result strongly suggests that FadR positively activates expression of fabB, as is the case with fabA. Two other observations support this hypothesis. The first is that the fabB(Ts) fadR strain grew at the permissive temperature on solid media, but the same strain failed to grow in liquid cultures shaken in flasks under standard conditions. However, the fabB(Ts) fadR strain grew in liquid medium when mechanical stress was minimized (in a thin film of liquid with minimal agitation as in Table 3). It therefore seems that the structural integrity of the membranes of these strains was compromised to the point that the cells became very sensitive to mechanical stress (5).
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A fadR strain is hypersensitive to cerulenin.
The behavior of
the fabB(Ts) strains with respect to FadR suggested that
fadR mutants might be more susceptible to the specific 3-ketoacyl-ACP synthase inhibitor cerulenin (39). Although
both 3-ketoacyl-ACP synthases I and II are inhibited by this
antibiotic, synthase I (FabB) is about sevenfold more sensitive than
synthase II (FabF) in vitro (41). In vivo results indicate
similar differential effects on the two enzymes (6, 53).
Therefore we expected that, due to decreased FabB levels,
fadR strains should be more sensitive to cerulenin than were
wild-type strains. This was found to be the case (Fig.
1). The cerulenin concentration that gave one-half the maximal growth of the wild-type strain was between 8 and
16 µg/ml, which agrees well with the value of 12.5 µg/ml reported
by Omura (39). In contrast, the fadR mutant
strain failed to show detectable growth even at low cerulenin
concentrations and almost complete growth inhibition was seen at 1 µg
of cerulenin per mg. Therefore, the fadR strain was at least
10-fold more sensitive to cerulenin than was the wild-type strain,
indicating lower levels of FabB, the primary target of the antibiotic.
Both strains used in this experiment contained plasmid pRC1 which
produces sufficient FabA enzyme to allow growth of fabA(Ts)
mutants at the nonpermissive temperature (14). This
plasmid was introduced to ensure that the effect of cerulenin was due
to changes in the level of FabB and not to possible secondary effects
of the fadR mutation on fabA expression.
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Decreased expression of fabB in fadR strains.
Our genomic
array experiments indicated that fabB was positively
regulated by FadR and that the level of expression was depressed in
cultures of the fadR mutant strain, JWC264. Measuring
fabB expression by assays of 3-ketoacyl-ACP synthase
activity is complicated by the presence of a second 3-ketoacyl-ACP
synthase (FabF), which has a similar level of activity and overlapping
substrate specificity. We therefore constructed a chromosomal
transcriptional fusion in which the fabB promoter was used
to drive expression of a promoterless CAT gene. Since FabB provides at
least one essential cellular function, we provided a functional copy of
fabB in trans to allow growth in the absence of
unsaturated fatty acid supplementation. The plasmid used,
pARAfabB, expresses fabB from the
Salmonella enterica serovar Typhimurium araBAD
promoter. Although this promoter is generally considered to have low
residual activity in the absence of inducer (7, 27), the
basal level of activity from the uninduced promoter provided sufficient
3-ketoacyl-ACP synthase activity to complement loss of the chromosomal
fabB gene (Fig. 2).
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Purified FadR binds to DNA sequences upstream of the fabB 10
promoter region.
Scrutiny of the sequences in the promoter region
of fabB showed that a region about 70 bp upstream of the
fabB start codon had a good match to known FadR binding
sites. Alignment of this sequence with those of the known FadR
binding sites (listed at http://arep.med.harvard.edu/ecoli_matrices/dat/fadR.dat)
is shown in Fig. 3 (data of Robison et
al. [45]). The sequence of the putative fabB
FadR binding site most closely matches those of the two positively
regulated genes iclR and fabA. Both the
iclR- and fabA-associated FadR binding sites are
known to overlap the 35 regions of their respective promoters
(26, 30). The fabB transcriptional start has
been mapped (32), and the upstream region is typical of a
promoter requiring positive activation in that the 10 region has a
good match to the consensus for E. coli promoters, but
there is no obvious 35 consensus sequence (25). As is
the case for both iclR and fabA, the putative
FadR binding site of fabB overlaps the 35 region of the
promoter, which is thought to position FadR protein to act as a
transcriptional activator.
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DISCUSSION |
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Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
|
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The evidence presented indicates that FadR positively regulates fabB transcription. The fact that both fabA and fabB, alone among the fatty acid biosynthetic genes, are directly regulated by FadR indicates that E. coli has a vested interest in regulating unsaturated fatty acid biosynthesis even when exogenous fatty acids are available. In retrospect, coordinated regulation of fabA and fabB might have been anticipated since in Pseudomonas aeruginosa it was recently shown that the two genes are cotranscribed in a two-gene fabA-fabB operon (31), although there is no evidence of transcriptional regulation of the operon. The available partial genomic sequences of Pseudomonas putida and Pseudomonas syringae that fabA and fabB are also adjacent genes in these organisms (data not shown).
The two E. coli long-chain acyl-ACP elongation enzymes, FabB and FabF, play different roles in unsaturated fatty acid synthesis. Both enzymes catalyze the Claissen-like condensation of malonyl-ACP with the growing acyl chain. Although the two enzymes are structurally and functionally very similar, they play different roles in cellular metabolism, as shown by the very different phenotypes of fabB and fabF mutants and of strains carrying recombinant plasmids encoding either of the two enzymes. Null fabF mutants have no growth phenotype, whereas fabB mutants require an exogenous source of unsaturated fatty acids. Likewise, strains carrying recombinant fabB plasmids are well behaved, whereas similar plasmids carrying fabF are prone to deletion and rearrangement (34, 51). Recently, it has been shown that FabF effectively titrates FabD (malonyl coenzyme A:ACP transacylase) and that overproduction of FabD offsets the toxicity of fabF plasmids (51). The exact role of FabB in unsaturated fatty acid biosynthesis has not been directly demonstrated in vitro and currently seems to be a puzzle (41, 55). FabB is generally thought to catalyze elongation of the cis-3-decanoyl-ACP produced by FabA to 3-ketododecanoyl-ACP. This is inferred from the findings that fabF strains have no apparent growth phenotype and that fabB mutants require only unsaturated fatty acids for growth. In vitro tests with purified FabF and FabB proteins show that both enzymes are capable of catalyzing the condensation of malonyl-ACP and cis-3-decenoyl-ACP (24, 55). The discrepancy between the genetic and biochemical observations indicates that the biochemical characterization of these enzymes does not accurately reflect their behavior in vivo. Unfortunately, resolution of these issues is problematic, since double knockouts of fabB and fabF are nonviable and fabF(Ts) strains are not available. Furthermore, double mutant fabF fabB(Ts) strains are nonviable at high temperature despite supplementation with both saturated and unsaturated fatty acids probably due to the inability to provide precursors for lipid A biosynthesis (24).
The levels of FabA activity normally present do not limit the synthetic capacity for unsaturated fatty acid biosynthesis in E. coli. This has been shown by examining the effects of FabA overproduction on cellular fatty acid composition (10). Unexpectedly, the cellular content of unsaturated fatty acids did not change, but the content of saturated fatty acids increased markedly. This effect was reversed by introduction of a second plasmid encoding FabB. These results were interpreted as indicating that the level of FabB governs the overall rate of unsaturated fatty acid biosynthesis. This is supported by studies that showed that overproduction of FabB about 10-fold (without manipulating FabA) results in increased unsaturated fatty acid production (17). Therefore, fabB may be the most effective point at which to regulate flux of this pathway. However, in order to regulate the level of unsaturated fatty acid biosynthesis without the complication of altering the level of saturated fatty acids produced, E. coli may need to simultaneously coordinate changes in expression of both fabA and fabB.
Why does FadR regulate only the synthesis of unsaturated fatty acids in response to exogenous supplementation? One clue may be that although exogenous supplementation can satisfy the entire unsaturated fatty acid requirement of E. coli, supplementation with saturated fatty acids cannot completely replace the function of that branch of the pathway. The reason for the differing efficiencies of supplementation is the fatty acid chains of the essential outer membrane component lipid A, which are derived from the saturated pathway. These fatty acids (mainly 3-hydroxymyristic acid) cannot be effectively supplied in the medium. Although when added to the culture medium 3-hydroxymyristic acid enters E. coli (and can serve as a carbon source), the supplied acid is not incorporated into lipid A (34). This is probably due to lack of conversion of the acid to the required ACP thioester (42). Therefore, if E. coli were to shut down both the saturated and unsaturated fatty acid synthetic pathways in response to exogenous supplementation, the organism would perish from a lack of the acyl chains needed to synthesize lipid A.
In general, the amplitudes of transcriptional regulation which we observed in our microarray analyses are less than those observed in enzyme assays, Northern blot analyses, and gene fusion experiments. To an unknown extent this may be due to the use of ratio values as the output of the current array techniques. If the expression signal of a gene in the control cultures is small and variable (for either technical or physiological reasons), these factors can have very large effects on the calculated ratios. Another possibility is that the differences might reflect mRNA decay. In our protocol we are primarily sampling the 3' end of the mRNAs due to the use of primers that hybridize to the ORF C termini, whereas the other techniques measure the full-length mRNA (either directly or indirectly). Therefore, the systems may differ due to mRNA turnover. Bacterial mRNA turnover is not yet well understood and the patterns of degradation seem to be mRNA specific (49). Recent data argue that in many messages the 3' end is the last segment degraded (49). (This is attributed to blocking of nuclease activity by the RNA hairpin left from factor-independent transcription termination.) Moreover, it is unclear whether or not the level of a given mRNA in E. coli can alter the rate or pattern of degradation of itself or of other mRNAs.
Many improvements to the existing array technologies seem possible. Quantitative tests of the various RNA isolation, priming, blotting, and detection methods have yet to be published. It is our opinion that even when the technology is fully developed this technology might best be viewed as a divining rod to find new regulatory circuitry, rather than as a quantitative tool.
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ACKNOWLEDGMENTS |
|---|
This work was supported by National Institutes of Health grant AI15650.
We thank C. O. Rock for free exchanges of information.
| |
FOOTNOTES |
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* Corresponding author. Mailing address: Department of Microbiology, University of Illinois, B103 Chemical and Life Sciences Laboratory, 601 S. Goodwin Ave., Urbana, IL 61801. Phone: (217) 333-7919. Fax: (217) 244-6697. E-mail: j-cronan{at}life.uiuc.edu.
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Journal of Bacteriology, October 2001, p. 5982-5990, Vol. 183, No. 20
0021-9193/01/$04.00+0 DOI: 10.1128/JB.183.20.5982-5990.2001
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