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1 Institute of Genetics and Biophysics ‘Adriano Buzzati Traverso’, via P. Castellino 111, 80131 Naples, Italy
2 Dipartimento di Scienze Cliniche e Biologiche, Ospedale S. Luigi, Regione Gonzole 10, Orbassano (TO), Italy
3 Dipartimento di Chimica Organica e Biochimica, Università Federico II di Napoli, Via Cinthia, 80126 Napoli, Italy
Correspondence
Roberto Defez
defez{at}igb.cnr.it
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ABSTRACT |
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The GEO accession number for the array studies reported in this paper is GSE4941.
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INTRODUCTION |
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). It was recently reported that nitric oxide mediates the IAA response through the activation of soluble guanylate cyclase and the following transient increase of cyclic GMP (Pagnussat et al., 2003). IAA also plays a key role in the regulation of ‘stringent control’, a process mediated by the bacterial alarmone ppGpp, which is produced in the chloroplasts of plant cells in response to stressful conditions (Takahashi et al., 2004). Soil bacteria (such as species of Pseudomonas, Azospirillum, Agrobacterium and Rhizobium) involved in intimate relationship with plants synthesize IAA as a communication system with the host plant, and many of them use IAA in pathogenic interactions (tumours, hairy roots, etc.). These bacterial systems mainly synthesize IAA from tryptophan via the indoleacetamide (Lemcke et al., 2000) or indole-3-pyruvate (Zimmer et al., 1998) biosynthetic pathways.
IAA is also involved in the morphogenetic development of Saccharomyces cerevisiae. At high concentration, IAA blocks the growth of yeast cells, whereas at lower concentration it induces filamentation and adhesion, factors that are important in plant infection by fungi. These events are mediated by a family of transporters and by the fungal-specific transcription factor Yap1 (Prusty et al., 2004). In mammals IAA is a metabolite of the neurotransmitter serotonin (Artigas et al., 1985). In enteric bacteria IAA is able to compensate for the absence of cAMP, restoring some specific pathways. It is able to replace cAMP in the regulation of the arabinose operon (Ebright & Beckwith, 1985) and the acetohydroxy acid synthase gene (Williams, 1986) in Escherichia coli. The mechanism of IAA action on the regulation of these genes is unclear. Experiments with two separate classes of small molecules, cAMP and indole derivatives, support the hypothesis of a local denaturation of the DNA helix (Ebright & Wong, 1981). Thus, IAA appears to trigger, in different biological systems, changes in gene regulation leading to different developmental programmes. However, although IAA's role as the main auxin plant hormone has been known for 70 years, its function in eukaryotes and prokaryotes is still largely unexplored. In the present study, we investigated the effect of IAA on the transcript, enzyme and metabolic profile of E. coli, a bacterium not usually exposed to IAA in its natural environment. We demonstrate that IAA activates the tricarboxylic acid (TCA) cycle and the glyoxylate shunt, while it reduces the NADH/NAD+ ratio.
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METHODS |
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for another E. coli K-12 strain. At least three independent experiments were performed for all the results presented in this work. When more than three repeated experiments were done, the number of repetitions is indicated in the table footnote or figure legend.
RNA isolation.
Total RNA was purified from cells grown in M9 arabinose medium using the RNeasy Mini-Kit from Qiagen and following the manufacturer's instructions. The isolated RNA was incubated at 37 °C with 5 U DNase I (Promega) and 40 U RNasin ribonuclease inhibitor (Promega) for 30 min, extracted with phenol/chloroform, precipitated with 2.5 vols ethanol and then dissolved in diethylpyrocarbonate-treated water. After purification, the RNA concentration was determined by absorbance at 260 nm and the RNA was stored at –20 °C until further use.
Probe synthesis.
33P-labelled probes were prepared using E. coli gene-specific primers (Sigma-Genosys) following the protocol provided by the manufacturer, except that the reaction mixture contained 40 µCi (1.48 MBq) [
-33P]dCTP (1000–3000 Ci mmol–1; Amersham Pharmacia), and 200 U Superscript II (Promega) in a 30 µl volume. Unincorporated radiolabelled nucleotides were removed by applying the reaction mixture to a Sephadex G-25 gel filtration spin column.
Array hybridization.
The DNA arrays (Panorama E. coli gene arrays) used in the hybridization experiments were produced by Sigma-Genosys Biotechnologies. Each DNA array consists of a 12x24 cm positively charged nylon membrane on which 10 ng each of all 4290 PCR-amplified ORF-specific DNA fragments are robotically printed in duplicate. The hybridization and washing steps were carried out as described by the manufacturer. Briefly, the nylon membranes were pre-hybridized in 5 ml hybridization buffer [5x SSPE (1x SSPE is 0.18 M NaCl, 10 mM NaH2PO4 and 1 mM EDTA, pH 7.7), 2 % SDS, 1x Denhardt's reagent and 100 µg denatured, sonicated salmon sperm DNA per ml]. After incubation for 2 h at 65 °C, the pre-hybridization solution was replaced with 3 ml hybridization buffer containing the denatured radioactively labelled cDNA probes. Filter hybridizations were carried out for 16 h in a hybridization oven. The filters were then washed three times, for 5 min each, at room temperature with 50 ml wash solution (0.5x SSPE, 0.2 % SDS) and then three times at 65 °C with 100 ml warmed wash solution. The washed filters were blotted between two pieces of Whatman 3 MM paper for 5 min and then wrapped in Saran Wrap. For quantification, filters were exposed to a phosphorimager screen. The blots were stripped at 100 °C with 1 % SDS in Tris/EDTA buffer as specified by the manufacturer.
Data analysis.
The exposed phosphorimager screens were scanned at 100 µm resolution using a STORM 445 PhosphorImager (Molecular Dynamics). The resulting files were analysed to determine the signal intensities for each spot by summing the value of each pixel within the boundaries of the spot using ImageQuant (version 3.0) software (Molecular Dynamics). Background values were measured in the surrounding region of the four corner grids containing E. coli genomic DNA, as a positive control, and empty spots. The intensity values for each spot were exported from ImageQuant to Microsoft Excel. Each ORF-specific spot was present in duplicate, and the mean of the normalized intensity values of the duplicate spots of each gene was used for further analysis. To avoid extreme intensity ratio for genes close to or below the detection limit, signal intensity values corresponding to a signal-to-background (S/B) ratio <2.0 were scaled up to a value corresponding to the normalized background means. To identify ORFs whose expression was altered by addition of IAA, the two growth conditions (with and without IAA addition) were compared by determining the ratio of the corresponding intensities of each pair of ORF-specific spots on two blots. These ratios represent the relative transcript levels of each E. coli ORF under the same growth condition. Ratios were calculated such that the log2 of the absolute value of the expression ratio was positive for intensities that were higher in IAA-treated cells and negative for intensities that were higher in control cells. A gene was considered significantly regulated when the relative intensity was least two times greater than the background expression threshold and the log2 expression ratio was 
0.8 (up-regulated) or 
–0.8 (down-regulated) and more than 4 times standard deviation from the mean.
Real-time PCR studies.
Total RNA was isolated from cells grown in M9 arabinose medium using a RNeasy Mini Kit (Qiagen) and following the manufacturer's protocol. Residual DNA present in the RNA preparations was removed by RNase-free DNase I treatment (Epicentre Technologies). cDNAs were synthesized with the StrataScript reverse transcription reagents (Stratagene) and random hexamers as primers. One ‘no RT’ control (without reverse transcriptase) for each RNA sample and one ‘no RNA’ control (replacing RNA with distilled H2O) for each primer and probe set were also performed. Specific primer pairs were designed using the Primer3 software. Primers for rrsA of the 16S rRNA gene were also designed, and this gene was included in all the Q-RT-PCR analyses for data normalization. Real-time PCR was performed with each specific primer pair by using the DyNamo HS SYBR Green qPCR kit (Finnzymes). The reactions were performed with the DNA Engine OPTICON 2 system (MJ Research). RT-PCR amplification for each cDNA sample was performed in triplicate wells. During the reactions the fluorescence signal due to SYBR Green intercalation was monitored to quantify the double-stranded DNA product formed in each PCR cycle. Results were recorded as relative gene expression changes after normalizing for rrsA gene expression and computed using the comparative CT method (2–
CT) described in detail by Livak & Schmittgen (2001).
Metabolite determination.
For this analysis the cells were grown in M9 arabinose medium. The level of NADH and NAD+ was measured by first extracting the nucleotides from cells harvested at different growth stages and then assaying for the two metabolites as described by Zhang et al. (2000) by using alcohol dehydrogenase to convert NAD+ to NADH and lactate dehydrogenase to convert NADH to NAD+. For intracellular acetyl-CoA determination the extraction and assay was based on the method described by Pruss & Wolfe (1994) with some modifications. E. coli cells (40 ml) were centrifuged, resuspended in 4 ml washing buffer [10 mM sodium phosphate (pH 7.5), 10 mM MgCl2 and 1 mM EDTA] and treated with 0.8 ml 3 M ice-cold HClO4. After rapid vortexing, the mixture was kept in ice for 30 min. Cell debris was removed from the mixture by centrifugation at 2800 g for 5 min. The HClO4 in the supernatant was neutralized immediately with 1.3 ml of 3 M KHCO3. The resulting HClO4 precipitate was removed by further centrifugation at 8000 g for 10 min. A 0.5 ml sample of this neutralized extract was assayed for acetyl-CoA by using a coupled enzyme assay with malate dehydrogenase and citrate synthase (Pruss & Wolfe, 1994), and acetyl-CoA was measured indirectly by monitoring the reduction of NAD+ to NADH. The intracellular acetyl-CoA concentration was expressed in mM using a measured cell density of 5x1011 cells per l per OD600 unit and an approximated cell volume of 3x10–15 l per cell (Chohnan et al., 1998). The extracellular levels of acetate and pyruvate were measured by a coupled reaction with acetate kinase and phosphotransacetylase (Pruss & Wolfe, 1994) and by a colorimetric assay (Patnaik et al., 1992), respectively. The concentrations were expressed in mM.
Preparation of cell extracts.
Harvested cells were dissolved in extraction buffer (20 mM Tris/HCl, pH 8.0, containing 1 mM EDTA, 20 mM KCl, 1 mM DTT, 10 %, v/v, glycerol and 1 mM PMSF) and then disrupted by sonication (MSE 150 W) for 2 min (in 30 s periods). Cell debris was removed by centrifugation at 16 000 g (Eppendorf 5415 D) for 10 min; the resulting crude cell-free extracts were immediately used for determination of specific enzyme activities or stored at –20 °C. Protein concentrations were determined by the Bradford assay, using BSA as a standard.
Enzyme activity measurement.
The activity of 34 enzymes was monitored in M9 arabinose medium for control and IAA-treated cells cultured at the same growth stage as for mRNA analysis. Three of these enzymes, involved in the TCA cycle and the glyoxylate shunt (citrate synthase, isocitrate dehydrogenase and malate synthase), were selected to assay their activity in cells grown with five different carbon sources (arabinose, glucose, mannitol, lactose and glycerol).
Previously published methods were used to assay the activity of the overall Entner–Doudoroff (E-D) pathway enzymes [including 6-phosphogluconate dehydratase (Edd) and 2-keto-3-deoxy-6-phosphogluconate aldolase (Eda)] (Alleanza & Lessie, 1982), ribose phosphate isomerase (RpiA) (Hove-Jensen & Maigaard, 1993), transketolase (TktA) (Josephson & Fraenkel, 1969), transaldolase (TalA) (Tchola & Horecker, 1966), phosphotransacetylase (Pta) (Brown et al., 1977), acetyl-CoA synthetase (Acs) (Brown et al., 1977), acetate kinase (Ack) (Skarstedt & Silverstein, 1976), alcohol dehydrogenase (AdhE) (Clark & Cronan, 1980), lactate dehydrogenase (LdhA) (Kochhar et al., 1992), phosphoglucose isomerase (Pgi) (Reithel, 1966), phosphofructokinase (Pfk) (Ling et al., 1969), fructose bisphosphate aldolase (Fba) (Rutter et al., 1969), triosephosphate isomerase (TpiA) (Eikmanns, 1992), glyceraldehyde-3-phosphate dehydrogenase (GAPDH) (Eikmanns, 1992), phosphoglycerate kinase (Pgk) (D'alessio & Josse, 1966), phosphoglyceromutase (PgmA) (D'alessio & Josse, 1966), enolase (Eno) (Westhead, 1966), pyruvate kinase (Pyk) (Valentine & Tanaka, 1966), fructose-1,6-bisphosphatase (Fbp) (Donahue et al., 2000), phosphoenolpyruvate carboxylase (Ppc) (Wohl & Markus, 1972), pyruvate dehydrogenase (AceE) (Schartz et al., 1968), phosphoenolpyruvate synthetase (PpsA) (Cooper & Kornberg, 1966), malic enzyme (SfcA) (Stols & Donnelly, 1997), citrate synthase (GltA) (Weitzman, 1969), aconitase (AcnA) (Fansler & Lowenstein, 1969), isocitrate dehydrogenase (IcdA) (Muro-Pastor & Florencio, 1992), 2-oxoglutarate dehydrogenase (SucAB) (Reed & Mukherjee, 1969), succinyl-CoA synthetase (SucCD) (Bridger et al., 1969), succinate dehydrogenase (SdhAB) (King, 1969), fumarase (FumA) (Hill & Bradshaw, 1969), malate dehydrogenase (Mdh) (Kitto, 1969), isocitrate lyase (AceA) (McFadden, 1969), malate synthase (AceB) (Dixon & Kornberg, 1969), and acetohydroxy acid synthase (IlvB) (De Felice et al., 1988).
Each reaction was started by the addition of enzyme extract and the activity was measured spectrophotometrically in a Beckman Coulter recording spectrophotometer. The wavelength and the millimolar absorption coefficients for NAD+, NADH, NADP+ and NADPH were 340 nm and 6.22 cm–1 mM–1, respectively. One unit of specific enzyme activity is defined as the amount of enzyme required to convert 1 µmol substrate per min per mg protein under the published reaction conditions. The control cultures were used as a reference for comparison.
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). The mRNA levels of 41 genes increased and that of 9 genes decreased in IAA-treated cells as compared to controls.
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) we found that 50 % of the up-regulated genes were associated with energy metabolism and amino acid biosynthesis.
Carbon and energy metabolism
Several genes involved in the TCA cycle (gltA, sucA, sucD, sdhB and sdhC) were up-regulated in IAA-treated cells. In addition, the expression of the aceA gene, encoding isocitrate lyase, involved in the isocitrate catabolic pathway at the branch point between the glyoxylate shunt and the TCA cycle, and of the cyoD gene, encoding a subunit of the aerobic respiratory system, cytochrome bo terminal oxidase complex, was higher in treated cells.
The expression of several carbon-transport genes, such as the arginine ABC transport gene artQ and the ribose-uptake gene rbsC, which are regulated by catabolite repression, was affected by IAA treatment. In addition, the mlc gene, which encodes the global regulator (repressor) Mlc, involved in glucose uptake (Shin et al., 2001), and glpK, encoding a glycerol kinase involved in the utilization of glycerol as carbon source (Weissenborn et al., 1992), were also induced in IAA-treated cells. In contrast, the expression level of the uhpA gene, which regulates the induction of the sugar-phosphate transport system (Insland et al., 1992), and of the adhE gene, encoding the fermentative enzyme alcohol dehydrogenase, induced under anaerobic conditions, was severely down-regulated in IAA-treated cells. Gene members of several operons (cyoABCDE, ilvGMEDA, sucABCD, sdhCBAD, proBA, dnaKJ and leuABCD) were regulated in a coordinate way; however, not all the genes of these operons were included in Table 1 either because of a high standard deviation or because of a log2 expression ratio below ±0.8. For the above reasons we did not find the araBAD operon and the ilvB gene among the genes significantly regulated, although they were both slightly induced. The ara operon and the ilvB gene were previously described as induced by IAA treatment (Ebright & Beckwith, 1985; Williams, 1986).
Biosynthetic pathways
Genes involved in amino acid biosynthesis, in particular those involved in the branched-chain amino acid biosynthetic pathways, were significantly induced in IAA-treated cells. Among them we found two genes of the ilvGMEDA operon, which encodes the enzymes for isoleucine and valine synthesis, and two genes of the leuABCD operon, the products of which are required for leucine biosynthesis. In addition, expression of the proA gene, which encodes 
-glutamyl phosphate reductase, involved in the biosynthesis of proline, was up-regulated in IAA-treated cells.
Confirmation of the macroarray results by real-time PCR.
A mixture of six upregulated and downregulated metabolic genes (sucA, sucD, gltA, aceA, adhE and galR), was selected for real-time PCR studies. The results of this analysis confirmed those obtained with macroarray hybridizations, although the absolute values of fold changes were different (Table 2), probably due to the different sensitivity of these two techniques.
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, Table 4 and Fig. 1). We found that most of the TCA cycle activities were significantly induced following IAA treatment. At the important control point of the junction between the TCA cycle and the glyoxylate shunt, isocitrate lyase (AceA) and malate synthase (AceB) were also induced in IAA-treated cells. The activities of several glycolytic enzymes (Pgi, GapA, Pgk, Eno, Pyk and the anaplerotic Ppc) were only slightly increased or unregulated following IAA treatment. Simultaneously, the activities of the gluconeogenic enzymes PpsA and Fbp were reduced and unaffected, respectively. In addition, the pyruvate dehydrogenase (aceEF operon product) and malic enzyme (sfcA gene product) activities were both induced in IAA-treated cells. We could not find a significant alteration of the enzyme activities involved in the oxidative pentose phosphate pathway, including ribose-5-phosphate isomerase (RpiA), transketolase (TktA) and transaldolase (TalA), and the E-D pathway, including 6-phosphogluconate dehydratase (Edd) and 2-keto-3-deoxy-6-phosphogluconate aldolase (Eda). Among the fermentative enzymes, the activity of lactate dehydrogenase (LdhA) was not influenced by IAA treatment, while the activity of alcohol dehydrogenase (AdhE) was reduced. The activity of the enzymes Pta and AckA, involved in acetate synthesis, was unregulated and induced, respectively, whereas that of Acs, involved in acetate consumption, was reduced. Finally, the activity of acetohydroxy acid synthase (IlvB), catalysing the common first step of isoleucine and valine biosynthesis, increased up to 40 % after IAA treatment (Table 3).
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). When the cells were grown in arabinose minimal medium the ratio of enzyme activities in IAA-treated versus control cells was more consistently increased. This increased ratio might be explained by the high specificity of IAA for the araBAD operon as previously reported (Ebright & Beckwith, 1985).
We found an excellent correlation between gene induction (gltA, aceA, sdhC, sdhB, sucA) and the activation of the corresponding enzymes. Similarly adhE was repressed along with the corresponding enzyme AdhE. However, there was a major exception in the consistency among gene and enzyme regulation concerning the sucD gene (up-regulated) and the relative enzyme activity of succinyl-CoA synthetase (down-regulated). The sucD gene is the last member of the PsdhsdhCDAB-PsucsucAB-sucCD cluster. The transcription of the sucABCD genes, which encode subunits of 2-oxoglutarate dehydrogenase (sucAB) and succinyl-CoA synthetase (sucCD), occurs from two distinct promoter elements, Psdh and Psuc. The promoter located near the start site of the sucA gene provides a relatively low constitutive level of sucABCD expression. The second promoter transcribes both the sdhCDAB and the sucABCD operon. It was demonstrated that the transcription of the sucABCD genes is primarily initiated and regulated at the upstream sdh promoter, which, unlike the suc promoter, harbours a cAMP receptor protein (CRP)-binding site (Cunningham & Guest, 1998). All the eight genes of this cluster are up-regulated in IAA-treated cells and we hypothesize that this is due to the ability of IAA to mimic cAMP action (Kline et al., 1980) even at this CRP-binding site. The induction of the sucD gene is not converted into a higher enzyme activity probably because this section of the TCA cycle is not required to be more active in IAA-treated cells. The metabolic product of SucD, succinate, can also be produced by AceA, which is activated in these cells; this might cause a feedback inhibition of SucD.
Effect of IAA treatment on metabolite and cofactor levels
To determine the effects of the IAA treatment on intermediary carbohydrate metabolism, we measured the intracellular levels of acetyl-CoA and the extracellular level of acetate and pyruvate. The concentrations of acetyl-CoA (1.0±0.1 mM for control cells, 2.7±0.3 mM for IAA-treated cells) and acetate (0.480±0.021 mM for control cells, 0.692±0.060 mM for IAA-treated cells) increased while the level of pyruvate excreted (0.066±0.003 mM for control cells, 0.046±0.002 mM for IAA-treated cells) decreased. In addition, the determination of the steady-state internal redox state, as reflected by the NADH/NAD+ ratio, showed that IAA treatment led to a 1.3-fold decrease of this ratio after 30 min (Fig. 2). This effect was much higher after 2 h, when the NADH/NAD+ ratio was even lower (3.2-fold decrease). The total dinucleotide pool also increased up to 40 % in IAA-treated cells (data not shown).
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The determination of metabolite levels revealed that IAA-treated cells synthesized higher acetyl-CoA levels as compared to the control cells. In response to this perturbation, treated cells conveyed acetyl-CoA to the phosphotransacetylase (Pta)-acetate kinase (Ack) pathway (Table 3, Fig. 1), producing and excreting more acetate in the medium. The increased acetate production was accomplished by excreting less pyruvate into the medium, in agreement with the well-known increased pyruvate production in mutants with a defective Pta-AckA pathway (Chang et al., 1999).
The higher acetate excretion observed in treated cells is an effect caused by the imbalance between the uptake of the substrate and the demand for biosynthesis and energy production. This adjustment functions as a safety valve, as proposed by Holms (1996) from stoichiometric flux analysis of the central metabolic pathways in E. coli. The accumulation of acetate observed in the extracellular milieu of IAA-treated cells might be of evolutionary advantage: it could increase the metabolic flexibility of these cells during adaptation to a changing environment. In particular, during the transition to stationary phase, the cells might well undergo a metabolic switch: instead of excreting acetate, they could reabsorb it, activate it to acetyl-CoA by means of Acs, and utilize the acetate to generate energy and biosynthetic components via the TCA cycle and the glyoxylate shunt, respectively. Most of the metabolic genes differentially expressed in IAA-treated cells are catabolite-sensitive genes: their transcription is subject to cAMP-dependent catabolite repression. Among them we found the TCA cycle genes gltA, sdhB, sucA and sucD, in the gltA-sdhCDAB-sucAB-sucCD gene cluster, and the cyoD gene, encoding one component of the low-affinity terminal oxidase cytochrome bo3, the respiratory system that is derepressed in aerobic conditions.
It is known that the cytochrome bo3 respiratory system, as well as cytochrome bd, is necessary to regenerate NAD+ from NADH and to eliminate excess reducing equivalents (Garcia-Horsman et al., 1994). Accordingly, we found that the NADH/NAD+ ratio after IAA treatment dropped to values roughly threefold lower than those found for control cells. This effect plays an important role in IAA-treated cells avoiding repression of NADH-producing enzymes, specially two of the TCA cycle's rate-controlling enzymes (IcdA and SucAB), subject to a product inhibition mechanism (de Graef et al., 1999). The lower NADH/NAD+ ratio correlates with the reduced production of the less energy-efficient fermentative gene adhE and the repression of the alcohol dehydrogenase activity. Indeed, Leonardo et al. (1996), by directly measuring the NADH-NAD+ levels and by monitoring the expression of adhE, demonstrated that this E. coli gene is induced under high-NADH/NAD+ conditions and repressed under low-NADH/NAD+ conditions. The decrease of NADH/NAD+ also prevents the inactivation of another enzyme subject to catabolite repression, the AceE enzyme in the pyruvate dehydrogenase complex (required for the oxidative carboxylation of pyruvate to produce CO2 and acetyl-CoA). The induction of the pyruvate dehydrogenase complex observed in treated cells correlated with the reduced conversion of pyruvate through phosphoenolpyruvate synthase (Pps) (Table 3).
In addition, IAA-treated cells induced the synthesis of genes encoding protective molecules such as molecular chaperones (yrfI, dnaK, yfhE and htpG gene products) to prevent the damage caused to many biological molecules by reactive oxygen species, such as hydrogen peroxide, produced during aerobic respiration.
Finally, the up-regulation of the two central backbones of metabolism, the TCA cycle and the glyoxylate shunt, provides cofactor regeneration and building blocks for amino acid biosynthesis (Akashi & Gojobori, 2002). We found that 10 % of the genes showing altered expression levels in IAA-treated cells were involved in amino acid biosynthesis: they were all up-regulated. Among them we found genes encoding enzymes required for the biosynthesis of valine and leucine, derived from intermediates produced in late steps of glycolysis, and for isoleucine and proline, derived from oxalacetate and 2-oxoglutarate, two precursor metabolites produced in the TCA cycle. We thus hypothesize that the increased expression of the amino acid biosynthetic genes is indicative of a nitrogen-sufficient condition.
Taken globally, these results suggest that IAA mimics cAMP action in the catabolite derepression of many metabolic genes (i.e. artQ, rbsC, mlc, glpK, TCA cycle genes and cyoD). It has been described that the ilvB gene (Williams, 1986) and the araBAD operon (Ebright & Beckwith, 1985) were specifically induced by IAA. Those reports and our data confirm that IAA cannot replace cAMP at all known sites of action, as we did not find that all catabolite-repressed genes are induced. Further studies to investigate a physical interaction between IAA and the ara operon transcriptional regulators might explain this aspect. We cannot say if the catabolite gene activator protein CAP (the crp gene product) is directly involved in the modulation of the catabolite repression observed in IAA-treated cells. Regulation, exerted by the cAMP–CRP complex in catabolite repression, is well established. However, several lines of evidence indicate that the cAMP–CRP complex may not be the only regulator involved in catabolite repression (Crasnier, 1996). In addition, the induction of the arabinose operon in cya mutant strains does not require CRP, as shown by Ebright & Beckwith (1985). Thus IAA might act through a different mechanism, for example via interaction with a new mediator.
IAA functions as a regulator of cellular metabolism in E. coli: it co-ordinates changes in carbon and nitrogen metabolism that are used by the cells to maintain metabolic homeostasis and to optimize energy production during aerobic life.
De Melo et al. (2004) found that human tumour cells treated with IAA showed activated flux of substrates through the TCA cycle and a decreased NADH/NAD+ ratio, showing a striking similarity with our data, in a quite distant system. The alterations triggered by IAA in the main metabolic pathways of bacterial, yeast and human cells leave an open question. Can some of the effects observed in higher organisms be explained by the action of IAA on cytoplasmic organelles? Does IAA act in E. coli as a hormone, as a second messenger or via physical interaction with DNA (Ebright & Wong, 1981)?
Our results provide the first extensive analysis of the action of IAA on the bacterial metabolism, allowing similar kinds of questions to be more precisely addressed.
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ACKNOWLEDGEMENTS |
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Received 13 December 2005;
revised 12 April 2006;
accepted 27 April 2006.
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