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Genetics. Aug 2007; 176(4): 2165–2176.
PMCID: PMC1950622

Genetic Analysis of the Histidine Utilization (hut) Genes in Pseudomonas fluorescens SBW25

Abstract

The histidine utilization (hut) locus of Pseudomonas fluorescens SBW25 confers the ability to utilize histidine as a sole carbon and nitrogen source. Genetic analysis using a combination of site-directed mutagenesis and chromosomally integrated lacZ fusions showed the hut locus to be composed of 13 genes organized in 3 transcriptional units: hutF, hutCD, and 10 genes from hutU to hutG (which includes 2 copies of hutH, 1 of which is nonfunctional). Inactivation of hutF eliminated the ability to grow on histidine, indicating that SBW25 degrades histidine by the five-step enzymatic pathway. The 3 hut operons are negatively regulated by the HutC repressor with urocanate (the first intermediate of the histidine degradation pathway) as the physiological inducer. 5′-RACE analysis of transcriptional start sites revealed involvement of both σ54 (for the hutU–G operon) and σ70 (for hutF); the involvement of σ54 was experimentally demonstrated. CbrB (an enhancer binding protein for σ54 recruitment) was required for bacterial growth on histidine, indicating positive control of hut gene expression by CbrB. Recognition that a gene (named hutD) encoding a widely distributed conserved hypothetical protein is transcribed along with hutC led to analysis of its role. Mutational and gene fusion studies showed that HutD functions independently of HutC. Growth and fitness assays in laboratory media and on sugar beet seedlings suggest that HutD acts as a governor that sets an upper bound to the level of hut activity.

PSEUDOMONAS fluorescens strain SBW25 is a common saprophytic bacterium that activates expression of a suite of amino acid uptake and degradation pathways when growing in the plant environment (Rainey 1999; Gal et al. 2003). The histidine uptake and utilization pathway (the hut locus) confers on SBW25 the ability to utilize histidine as a sole source of carbon, nitrogen, and energy (Zhang et al. 2006).

In bacteria, catabolism of histidine occurs via either a four or a five-step enzymatic pathway (Coote and Hassall 1973a; Magasanik 1978). The first three steps, from histidine to urocanate, to imidazolone propionate (IPA), to formiminoglutamate (FIGLU), are catalyzed by the gene products of hutH, hutU, and hutI, respectively, and are common to both the four-step and the five-step pathways (outlined in Figure 1A). Breakdown of FIGLU differs among organisms. Enteric bacteria, e.g., Salmonella typhimurium and Klebsiella aerogenes (Magasanik 1978), and the gram-positive bacterium Bacillus subtilis (Chasin and Magasanik 1968) hydrolyze FIGLU directly to form glutamate and formamide. However, P. putida (Hu et al. 1987) and Streptomyces coelicolor (Kendrick and Wheelis 1982) are known to employ two enzymes (FIGLU iminohydrolase encoded by hutF and formylglutamase encoded by hutG) to convert FIGLU into glutamate plus formate with formylglutamate (FG) as an intermediate. Notably, no ATP is generated in this process; thus the energy (and most building blocks) required for growth on histidine must be derived from further degradation of glutamate.

Figure 1.
The histidine degradation pathways (A) and structure of the P. fluorescens SBW25 hut locus (B) with details of the promoter regions of PhutF and PhutU (C). (A) The five-step histidine degradation pathway of Pseudomonas (Hu et al. 1987) is shown by solid ...

Regulation of hut expression is complex and not fully understood. In gram-negative bacteria, hut is negatively regulated by the product of hutC (Magasanik 1978; Allison and Phillips 1990). Repression by HutC is relieved by urocanate, the first intermediate of the histidine degradation pathway (Lessie and Neidhardt 1967; Newell and Lessie 1970), which interacts with the HutC repressor (Hu et al. 1989). In enteric bacteria, expression of hut requires derepression (of HutC) and also requires activation by additional positive regulation factors. When histidine is utilized as a source of carbon, hut transcription is activated by catabolite-activating protein (CAP) charged with cAMP. Thus, like the lac operon, expression is subject to control by catabolite repression exerted by glucose (Magasanik 1978). However, when histidine is a nitrogen source, hut expression is activated by the nitrogen assimilation control protein (NAC), whose transcription is controlled by the NtrBC two-component system in response to nitrogen starvation (Pomposiello et al. 1998).

Little is known about positive regulation of hut in Pseudomonas species, although succinate-provoked carbon catabolite repression has been observed (Lessie and Neidhardt 1967; Coote and Hassall 1973b; Phillips and Mulfinger 1981), an effect thought to be attributable to the inhibitory effects of succinate on urocanase (Hug et al. 1968). Recently, Nishijyo et al. (2001) showed that expression of the hut enzymes in P. aeruginosa requires the two-component system CbrAB, but just how CbrAB regulates hut transcription is unclear.

Here we report a genetic analysis of the hut locus from P. fluorescens SBW25. The study builds upon a substantive body of mainly enzymatic data acquired during the 1980s by Phillips and co-workers who focused on P. putida ATCC 12633 (Hu et al. 1987, 1989; Hu and Phillips 1988; Allison and Phillips 1990). Our work began with the unannotated whole-genome sequence of SBW25. Initially on the basis of the analysis of this sequence, but confirmed using genetics, we show that the hut locus is composed of 13 genes organized in 3 transcriptional units. We reveal the identity of the gene encoding FIGLU iminohydrolase (hutF), show that one copy of hutH is nonfunctional, and confirm negative regulation via HutC (and the role of urocanate as the specific inducer). In addition, we identify positive regulators (σ54 and the enhancer-binding protein CbrB) and attribute a phenotype to the widely distributed and highly conserved protein encoded by pflu0360 (hutD).

MATERIALS AND METHODS

Bacterial strains, plasmids, and growth conditions:

Escherichia coli DH5αλpir was used for gene cloning and conjugative transfer into P. fluorescens. P. fluorescens strains and plasmids are listed in Table 1. Pseudomonas and E. coli strains are routinely grown in Luria–Bertani medium (LB) at 28° and 37°, respectively (Sambrook et al. 1989). Where indicated, Pseudomonas strains were also cultivated in minimal-M9 medium (Sambrook et al. 1989) supplemented with glucose (0.4% or 22.2 mm) and NH4Cl (1 mg ml−1 or 18.7 mm). When histidine or urocanate were used as sole carbon and nitrogen sources, they replaced the glucose and ammonia of the M9 medium and were added at a final concentration of 15 mm. When necessary, antibiotics were included at the following concentrations: tetracycline (Tc), 10 μg ml−1; kanamycin (Km), 50 μg ml−1; spectinomycin (Sp), 100 μg ml−1; nitrofurantoin (Nf), 100 μg ml−1. M9 plates containing half-strength cetrimide, fucidin, and cephalosporin (CFC) supplement from Oxoid were used to select for P. fluorescens recovered from the sugar beet seedlings. Glutamine (1 mm) was added to minimal-M9 medium to support growth of the rpoN mutant (PBR808).

TABLE 1
Bacterial strains and plasmids

Growth kinetics of P. fluorescens SBW25 and the derived mutant strains were determined in microtiter plates using a VersaMax microtiter plate reader with SOFTmax PRO software (Molecular Devices). To ensure that all bacteria were physiologically equivalent, strains were inoculated from cells stored in a −80° freezer. They were first grown in LB broth (24 hr) and then subcultured once in M9 broth (24 hr) before use in assay conditions. Absorbance at the wavelength of 450 nm was determined every 5 min over a period of 48 hr.

Mutational analysis and complementation:

Site-directed and insertional mutagenesis was performed using standard DNA manipulation techniques (Sambrook et al. 1989). Gene mutations were achieved by SOE–PCR (splicing by overlapping extension using the polymerase chain reaction; Horton et al. 1989) in conjunction with a two-step allelic-exchange strategy using the suicide-integration vector pUIC3 (Rainey 1999). Details of oligonucleotide primers are available on request. All PCR-generated fragments were checked by sequencing prior to exchange into the chromosome.

Cycloserine enrichment was used to enrich for strains that had lost the chromosomally integrated pUIC3-based vector. Strains were grown overnight in 20 ml LB broth; 400 μl of the overnight culture was inoculated into 20 ml prewarmed LB broth and cultivated at 28° with shaking (150 rpm) for 30 min. Tetracycline was added at the final concentration of 10 μg ml−1 to inhibit the growth of cells that had lost pUIC3. After growth for 2 hr, cycloserine was added at 800 μg ml−1 and growth was continued for another 4 hr (during this step the growing TcR cells are killed). The cells were then washed in sterile water, diluted, and inoculated onto LB plus X-Gal plates. SBW25 strains carrying polar mutations were generated by SOE–PCR and allelic replacement as described above, but an Ω-Sp cassette [retrieved from plasmid pHP45Ω (Fellay et al. 1987)] was inserted in place of the deleted gene.

Complementation of the hutH genes was performed by cloning the PCR-amplified coding region of hutH1 or hutH2 into pME6010 (Heeb et al. 2000) at the EcoRI site. The plasmid was introduced into PBR801 (ΔhutH1/H2) by conjugation with the help of pRK2013 (Tra+).

Construction of lacZ transcriptional fusions and assay for β-galactosidase activity:

lacZ reporter fusions were generated by cloning an ~800-bp fragment in front of the promoterless ‘lacZ carried on the integration vector of pUIC3 (Rainey 1999). The resulting plasmid was mobilized into P. fluorescens SBW25 or derived mutant strains by conjugation with the help of pRK2013. Integration into the genome by insertion–duplication was selected on LB plates supplemented with Nf and Tc.

Expression of ‘lacZ fusions was measured by β-galactosidase assay using 4-methylumbelliferyl-β-d-galactoside (4MUG) as the enzymatic substrate. The product (7-hydroxy-4-methylcoumarin, 4MU) was detected using a Hoefer DyNA Quant 200 fluorometer (Pharmacia Biotech) following the manufacturer’s instructions. The reaction was monitored at 460 nm with an excitation wavelength of 365 nm. Cell density was determined by measuring the absorbance of the culture at 600 nm. The enzyme activity was expressed as “aM 4MU min−1 cell−1” (1 aM = 10−18 mol).

Rapid amplification of cDNA 5′-ends:

Transcriptional start sites of the hut operons were determined using the rapid amplification of cDNA 5′-ends (5′-RACE) system (Invitrogen, Carlsbad, CA). Total RNA was isolated by using the TRIzol RNA extraction reagent (Invitrogen) from P. fluorescens SBW25 cells grown in M9 salt medium supplemented with histidine. Primers PhutU5 (5′-TGAGGACCCAACGGCCTTTG-3) and PhutU1 (5′-TCTGGCCGTACATGGCTAG-3′) were used for the cDNA synthesis and the subsequent nested PCR amplification of the hutU transcript. To identify the hutF transcriptional start, primers PhutC1 (5′-GAGTGGAAGCACAGGCCCA-3′) and PhutF1 (5′-TGATCTGACGCGACAGTTC-3′) were used. The final 5′-RACE products were purified in agarose gel and extracted by using QIAGEN's QIAquick gel extraction kit before being cloned into pCR8/GW/TOPO (Invitrogen). Ten randomly chosen colonies were analyzed by DNA sequencing.

Assessment of bacterial fitness in laboratory media and in planta:

Performance of mutant strains growing in laboratory media and in planta was examined by direct competition with the lacZ-marked wild-type strain of P. fluorescens SBW25 (SBW25-lacZ). Competition experiments were initiated with a 1:1 ratio of each strain [acclimated for 48 hr prior to initiating competition (see above)]. The initial frequency was determined by dilution plating onto LB plus X-Gal plates. For performance in laboratory medium, 5 μl of the bacterial suspension was inoculated into 5 ml of the tested medium in a 20-ml plastic tube. After growth for 24 hr, it was subcultured at a 1000× dilution. After two transfers (~20 generations), final ratios were determined by counting colonies on LB plus X-Gal plates. Relative fitness was expressed in terms of the selection rate constant (SRC) (Lenski 1991). A plant competitive colonization assay was performed as previously described (Zhang et al. 2006).

RESULTS

Genetic identification of pflu0358 as hutF:

Interrogation of the genome sequence of P. fluorescens SBW25 using the amino acid sequence of proteins known to be involved in histidine metabolism revealed a cluster of 12 genes (hutChutG) (Figure 1, Table 2). In addition to the core degradative enzymes (HutHUIG), the SBW25 hut operon harbors a second copy of the histidase gene (hutH) along with five open reading frames (ORFs) that are predicted to play a role in uptake: plfu03630365 are likely to encode an ABC-type transport system and pflu0362 and pflu0368 encode predicted permeases [pflu0368 is required for histidine utilization (Zhang et al. 2006)].

TABLE 2
The P. fluorescens SBW25 hut genes, predicted functions and similarity with homologs from P. putida ATCC 12633, KT2440, and P. aeruginosa PAO1

The ORFs immediately adjacent to the hut locus were examined to see whether any might be assigned a role in histidine uptake or metabolism. ORF pflu0358 encodes a predicted protein of 454 amino acids (Figure 1). It contains a conserved amidohydrolase domain (Pfam01979, E-value 8e−5) and belongs to a group of metallo-dependent hydrolases with unknown function (cd01313, E-value 1e−141). The possibility that this gene might encode FIGLU iminohydrolase (HutF) was tested by construction of a mutant PBR800 (pflu0358::Ω). PBR800 grew normally on minimal-M9 medium with glucose and ammonia as carbon and nitrogen sources, but was incapable of growing when histidine (or urocanate) was the sole carbon and nitrogen source. Next, a lacZ transcriptional fusion was constructed to pflu0358 and integrated into the genome of SBW25 in such a way as to ensure that pflu0358 function was not affected. The resulting fusion strain (PBR811) was grown in minimal-M9 medium (with glucose and ammonia) supplemented with histidine or urocanate and β-galactosidase activity was measured. Results showed that pflu0358 transcription was elevated 12-fold by the addition of either histidine or urocanate (mean aM 4MU min−1 cell−1 ± standard error): 0.49 ± 0.18 in M9, 5.67 ± 0.83 in M9 plus histidine, and 8.57 ± 3.22 in M9 plus urocanate.

A search of sequenced Pseudomonas genomes shows that in each genome hutC lies adjacent to a homolog of pflu0358, but oriented in the opposite direction. Furthermore, a homolog of pflu0358 exists in S. coelicolor A3(2), a bacterium, which like P. putida (Table 2), has a five-enzyme histidine degradation pathway (Kendrick and Wheelis 1982; Consevage et al. 1985). No homologs were identified in the genomes of bacteria known to possess the four-enzyme degradation pathway, for example, S. typhimurium, K. pneumoniae (Magasanik 1978), or B. subtilis (Chasin and Magasanik 1968). Together these data are consistent with the genetic organization of hutF reported in P. putida ATCC 12633 (Consevage et al. 1985) and indicate that pflu0358 is hutF. Further evidence was provided by the HutF amino acid sequence from P. putida ATCC 12633 (A. Phillips, unpublished data) and P. aeruginosa PAO1 (Marti-Arbona et al. 2006), which show 84 and 76% sequence identity, respectively, to the deduced amino acid sequence of hutF from P. fluorescens SBW25.

Functional characterization of the hutH-like genes:

The hut locus of SBW25 harbors two copies of hutH-like genes (hutH1 and hutH2), which encode proteins that show 36 and 84% sequence identity with the biochemically characterized HutH from P. putida (Table 2). Interestingly two copies of HutH-like genes are also found within the hut locus of all the genome-sequenced Pseudomonas strains (P. syringae DC3000, 1448A, and B728a; P. aeruginosa PAO1; and P. fluorescens Pf0-1 and Pf-5) with the exception of P. putida KT2440. To investigate functionality, both copies of hutH were deleted from the SBW25 genome. The mutant strain PBR801 (ΔhutH1/H2) was unable to grow on histidine, but could grow on urocanate, which confirms the histidase (histidine ammonia-lyase) function of hutH1 and/or hutH2. Next, single-deletion mutants PBR802 (ΔhutH1) and PBR803 (ΔhutH2) were produced: deletion of hutH1 had no effect; however, deletion of hutH2 resulted in a strain unable to grow on histidine. Additionally, complementation of PBR801 was achieved by introduction of a cloned copy of hutH2, but not hutH1. Analysis of the deduced amino acid sequences of hutH1 and hutH2 showed that HutH1 lacks the conserved Ala–Ser–Gly active site residues (Schwede et al. 1999) consistent with the genetic data that indicate that hutH2 (but not hutH1) encodes a functional histidase.

In P. putida ATCC 12633, urocanate, rather than histidine, is the physiological inducer of hut genes (Allison and Phillips 1990). If this also holds for SBW25, then histidine should be incapable of activating hut transcription in a mutant of SBW25 lacking histidase function. To this end, induction of hutF transcription was examined in the double hutH1/H2 mutant (PBR801) using the chromosomally integrated hutF-‘lacZ fusion (fusion strain PBR820). Histidine was no longer capable of inducing hutF expression, whereas urocanate induction was not significantly affected (mean aM 4MU min−1 cell−1 ± standard error): 0.29 ± 0.01 in M9, 0.51 ± 0.17 in M9 plus histidine, and 3.59 ± 0.27 in M9 plus urocanate. Therefore, both histidine and urocanate induce hut expression, but urocanate is the direct inducer.

Transcriptional organization of the hut genes:

hutF must be transcribed as a single gene unit from a promoter between hutF and hutC (PhutF). However, the transcriptional organization of the remaining 12 hut genes was uncertain (Figure 1B). To test whether the 12 hut genes from hutC to hutG are transcribed as a single mRNA, a lacZ fusion was made to the first (hutC) and the last (hutG) of the 12 genes (to give fusion strains PBR812 and PBR814, respectively), and their response to urocanate determined. In the wild-type background, both hutC and hutG were induced by the presence of urocanate (Table 3, PBR812 and PBR814). This result confirms the existence of a urocanate-inducible promoter in the front of hutC (PhutC).

TABLE 3
Urocanate-induced hut gene expression in strains that carry polar mutations of the hut loci

To test for the existence of additional promoter(s) downstream of hutC, a hutC polar mutant, PBR804, was generated (hutC::Ω). In both hutC::Ω and wild-type backgrounds, hutG expression was elevated in the presence of urocanate compared to the wild-type hutG-‘lacZ fusion grown in M9 (Table 3, fusion strains PBR818 and PBR814). The fact that hutG transcription was not abolished by the hutC polar mutation indicates the presence of a second transcriptional unit. To localize the predicted additional hut promoter(s), the intergenic regions located downstream of hutC were examined. The TGA stop codon of hutC overlaps the ATG start codon of the adjacent gene (pflu0360), suggesting cotranscription. Between pflu0360 and hutU lies a 405-nucleotide region that could potentially define a promoter. To test this possibility, a lacZ fusion to hutU was constructed in hutC::Ω and expression was measured by β-galactosidase assay. In both the hutC::Ω and wild-type backgrounds, hutU transcription was elevated in the presence of urocanate (Table 3, fusion strains PBR813 and PBR817). The hutC polar mutation did not abolish hutU expression in the presence of urocanate, indicating the existence of a second transcriptional unit initiated from a promoter immediately upstream of hutU (PhutU).

To determine the existence of additional promoter(s) between hutU and hutG, a hutU polar mutation was generated (PBR815). Unlike PBR804 (hutC::Ω), PBR815 (hutU::Ω) was unable to grow on either histidine or urocanate (hutU encodes urocanase; in addition the omega cassette has polar effects). As shown in Table 3, hutG-lacZ expression was not induced by urocanate in the hutU::Ω genetic background (fusion strain PBR819) while the positive control (hutU-‘lacZ) remained urocanate inducible (fusion strain PBR832). This shows that there is no functional urocanate-inducible promoter between hutU and hutG.

Taken together, the genetic data show that the hut locus is organized into three transcriptional units: hutF, hutCD, and hutUG (Figure 1B).

Data on the effect of hutC inactivation on the transcription of hutC, hutU and hutG (Table 3), and hutF (not shown) are consistent with HutC from SBW25 also functioning as a repressor of hut gene transcription (Hu et al. 1989). To determine whether urocanate is the direct inducer of hutCD and hutU–G (in addition to hutF), histidine and urocanate-induced expression of hutCD and hutU–G was determined in the ΔhutH1/H2 genetic background using ‘lacZ fusions to hutC and hutU (fusion strains PBR821 and PBR822, respectively). Expression was stimulated by urocanate, but not by histidine (mean aM 4MU min−1 cell−1 ± standard error): hutC expression was 0.41 ± 0.2 in M9, 0.38 ± 0.04 in M9 plus histidine, and 1.76 ± 0.15 in M9 plus urocanate; hutU expression was 2.08 ± 0.22 in M9, 2.23 ± 0.67 in M9 plus histidine, and 16.7 ± 0.51 in M9 plus urocanate. Urocanate is therefore the physiological inducer of all three hut operons.

Determination of the hut transcriptional start sites:

To identify the promoter sequences of PhutF and PhutU (the two putative promoters that control expression of hut structural genes) 5′-RACE was used to identify the 5′-end sequence of each transcript from cells grown on minimal-M9 medium with histidine as the sole source of carbon and nitrogen (Figure 1C). Immediately upstream of the hutF transcriptional start site resides a sequence of nucleotides (TTACCG N16 TATATG) that is similar to the σ70-promoter consensus (TTGACA N16–18 TATAAT). In addition, a putative HutC binding site (Hu et al. 1989) overlaps the −10 region of PhutF (Figure 1C).

When the nucleotide sequence upstream of the hutU transcriptional start was examined, a motif (TGGCCG N5 TTGCA) was identified that shows strong similarity to the σ54-promoter −24/−12 consensus (TGGCAC N5 TTGCW). However, the start site is the 14th nucleotide downstream from the conserved C residue of the −12 element (Figure 1C), whereas transcription from such promoters is typically initiated at the 12th nucleotide downstream from the conserved C residue of the −12 element. This atypical spacing has been reported for other σ54-dependent promoters (Barrios et al. 1999). Also present in the vicinity of the hutU transcriptional start is a putative HutC binding site (Figure 1C) (Hu et al. 1989).

Transcriptional activation of hut requires σ54 and CbrB:

The observation that hut genes in the hutUG operon are controlled by a σ54-dependent promoter suggested the involvement of both σ54 and a σ54 activator (Wosten 1998). In Pseudomonas, σ54 is encoded by rpoN; thus a rpoN deletion mutant (PBR808, a gift from Jake Jones and Gail Preston) was examined for its ability to utilize histidine and urocanate. PBR808 grew normally in minimal-M9-salt medium supplemented with glucose and ammonia, but was unable to grow in minimal-M9-salt medium with histidine (or urocanate) as the sole carbon and nitrogen source. This demonstrates the requirement of σ54 for utilization of histidine and urocanate. It also shows that derepression of HutC (by plating cells in the presence of either histidine or urocanate) is a necessary, but not sufficient, requirement for activation of hut transcription.

Next we sought the identity of the σ54 activator. A possible candidate was indicated by Nishijyo et al. (2001) who showed that in P. aeruginosa PAO1 a two-component regulatory system, CbrAB, was associated with hut regulation. Significantly, the response regulator CbrB contains a σ54-type regulator output domain in addition to a signal-receiver domain. In silico analysis of the P. fluorescens SBW25 genome revealed a two-gene locus (pflu5236 and pflu5237) that shows significant similarity to the CbrAB genes of P. aeruginosa PAO1. The genetic organization of this locus in SBW25 (hereafter referred to as SBW25 cbrAB) mirrors that found in PAO1: it is composed of two genes, orientated in the same direction, and separated by 22 nucleotides.

To test the role of CbrAB in hut regulation a cbrA and a cbrB mutant were constructed (PBR809 and PBR810, respectively) and their ability to utilize histidine and urocanate as sole carbon and nitrogen sources was determined. Both PBR809 and PBR810 grew normally in M9 salt medium supplemented with glutamate (the end product of the histidine degradation pathway), but were unable to grow in M9 salt medium supplemented with histidine or urocanate. This result, in conjunction with the in silico analysis data, implicates CbrB as a σ54-enhancer binding protein required for transcriptional activation of the hutU–G operon.

Functional investigation of the hypothetical protein Pflu0360 (HutD):

The TGA stop codon of hutC overlaps the ATG start codon of a downstream ORF (pflu0360). This organization suggests that transcription of pflu0360 is coordinated with hutC and that pflu0360 may play a role in hut regulation, possibly in conjunction with HutC. We designate this open reading frame hutD (Figure 1) on the basis of the functional characterization described below.

Interrogation of the NCBI protein database with the deduced amino acid sequence of hutD provided no clues as to its function: it belongs to a group of uncharacterized proteins that are highly conserved in bacteria (Pfam0596; DUF886, COG 3758). A recently determined crystal structure of HutD (PA5104) from P. aeruginosa PAO1 (PDB.1yll) reveals no insight into function. In Pseudomonas, hutD is always located downstream of hutC in an overlapped manner, whereas in other gram-negative bacteria, e.g., Burkholderia cenocepacia, Mesorhizobium loti, Yersinia pestis, and Serratia marcescens, the hutD homolog is located in the hut locus but not adjacent to hutC.

To investigate the role of hutD and possible interactions with hutC, three in-frame deletion mutants were generated: PBR805 (ΔhutC), PBR806 (ΔhutD), and PBR807 (ΔhutCD). The three mutant strains were subjected to phenotypic assays to determine: first, effects on transcription of the hutU–G operon; second, ability to grow on histidine (and urocanate) as a sole source of carbon and nitrogen; and third, the contribution of each mutation to fitness in laboratory media.

Expression of the hutU–G operon was measured using a chromosomally integrated hutU-‘lacZ fusion in the genetic background of wild-type SBW25 (as a control; PBR813) and the hutC (fusion strain PBR823), hutD (fusion strain PBR824), and hutCD mutants (PBR825). β-Galactosidase was assayed for cells growing in minimal-M9 medium supplemented with glucose and ammonia, histidine, or urocanate. Results are shown in Figure 2. Consistent with previous findings (see above), the hutU–G operon was constitutively expressed in a ΔhutC background (fusion strain PBR823). The operon was also constitutively expressed in ΔhutCD (fusion strain PBR825), whereas in ΔhutD (fusion strain PBR806) and in the wild-type background (PBR813), the hut operon remained histidine and urocanate inducible.

Figure 2.
Expression of hutU in wild-type SBW25 and the hutCD mutants. β-Galactosidase activities (aM 4MU min−1 cell−1) were measured in hutU-‘lacZ fusion strains PBR813 (wild type), PBR823 (ΔhutC), PBR824 (ΔhutD ...

In M9 salt medium supplemented with glucose and ammonia, the basal level of hutU–G expression was negligible in the wild-type background and was not affected by the hutD deletion. However, a statistically significant increase in hutU–G transcription (attributable to hutD) was detected when hutU–G expression was compared in the wild-type and the hutD mutant strains (grown on M9 salt medium plus histidine or urocanate, fusion strains PBR813 and PBR824). A similar effect, also attributable to hutD, was evident when hutU–G transcription was compared in hutC vs. hutCD mutant strains (see Figure 2, PBR823 and PBR825). From these data we conclude that HutC is the sole hut repressor; HutD appears to limit the upper level of transcriptional induction.

Next we examined the growth properties of the mutants in minimal-M9 medium supplemented with glucose and ammonia, histidine, or urocanate (Figure 3 and Table 4). The three mutant strains showed similar growth characteristics to wild-type SBW25 when grown on minimal-M9 medium supplemented with glucose and ammonia, although PBR805 (ΔhutC) and PBR807 (ΔhutCD) mutants showed slower growth, a likely consequence of constitutive expression of enzymes in an environment where they are not required (Savageau 1989, 1998).

Figure 3.
Growth dynamics of the hutCD deletion mutants. Growth was measured for wild-type SBW25 (open circles) and mutants PBR805 (ΔhutC, solid triangles), PBR806 (ΔhutD, open triangles), and PBR807 (ΔhutCD, open squares) in M9 (M9 salts ...
TABLE 4
Maximum growth rate (μmax) of the hutCD mutants when growing in laboratory media

When grown on either histidine or urocanate as the sole carbon and nitrogen source, growth of the hutD mutant strain PBR806 was significantly impaired: the lag time was extended by ~4 hr and the maximum-growth rate (μmax) was ~20% lower than the wild type (Table 4). The opposite effect was observed in the hutC mutant (PBR805): this genotype showed more rapid growth than the wild type, presumably because there is no delay in activating hut gene expression (Savageau 1989). PBR807 (ΔhutCD) displayed an intermediate phenotype (Figure 3).

To further analyze the growth phenotypes, particularly the slow growth of PBR806 (ΔhutD) on histidine or urocanate, the competitive ability of these mutants relative to the wild-type ancestor was determined in three shaken-broth cultures containing minimal-M9 medium and supplemented with (1) glucose and ammonia, (2) histidine, or (3) urocanate. The mutant was mixed 1:1 with a lacZ-marked “wild-type” strain (SBW25-lacZ) and the bacterial mixture was inoculated into test media. After growing in competition for ~20 generations the ratio of mutant to the wild-type competitor was determined by plating onto LB plates supplemented with X-Gal. As a control for any possible effects due to the lacZ marker, ancestral wild-type SBW25 was competed in parallel against the SBW25-lacZ. Results are shown in Figure 4A and closely parallel the trends revealed from the analysis of the growth of individual strains (Figure 3, Table 4). However, when histidine or urocanate was the sole carbon and nitrogen source, the fitness of PBR806 (ΔhutD) and PBR807 (ΔhutCD) was drastically impaired relative to wild type. The fitness of PBR805 (ΔhutC) was not impaired; indeed, PBR805 (ΔhutC) was more fit than wild type on minimal-M9-salt medium supplemented with histidine. The fact that deletion of hutD produced a phenotype equivalent to wild type in the histidine-free environment (and distinct from ΔhutC and ΔhutCD), but equivalent to ΔhutCD in the histidine-containing environment indicates that hutC and hutD are not functionally interdependent.

Figure 4.
Fitness of the hutCD deletion mutants relative to lacZ-marked P. fluorescens SBW25 in laboratory media. (A) Fitness of SBW25 (wild type), PBR805 (ΔhutC), PBR806 (ΔhutD), and PBR807 (ΔhutCD) relative to SBW25-lacZ grown in M9 medium, ...

The observed fitness effects are consistent with the known repressor function of HutC and further suggest that HutD may function to govern the upper level of hut activation. To examine these effects more closely, the fitness of each of the three mutants was determined on M9 salt medium supplemented with 5 mm glutamate (the end product of histidine degradation and itself a source of carbon and nitrogen), but containing different concentrations of histidine (from 0.5 mm to 15 mm). If HutD functions as a governor that limits the upper level of hut expression, then the fitness cost associated with deletion of hutD ought to increase with increasing histidine concentration. Conversely, the fitness cost associated with derepression of hut (brought about by deletion of hutC) should decrease with increasing histidine concentration (Savageau 1989, 1998). Assuming that hutC and hutD function independently of one another and given opposing fitness effects associated with deletion of each gene, then fitness of a ΔhutCD mutant should be low at both low and high concentrations of histidine. The results shown in Figure 4B are fully consistent with these predictions.

Ecological significance of hut regulatory genes in plant environment:

The competitive ability of each mutant [PBR805 (ΔhutC), PBR806 (ΔhutD), and PBR807 (ΔhutCD)] relative to the lacZ-marked “wild-type” strain (SBW25-lacZ) was determined during the course of colonization of sugar beet seedlings. After a 2-week period of competitive colonization, bacteria were recovered from the shoot and rhizosphere and the fitness of each mutant was determined relative to wild type.

The fitness of PBR805 (ΔhutC) and PBR807 (ΔhutCD) was significantly impaired in the plant environment, whereas PBR806 (ΔhutD) was not affected (Figure 5). Drawing upon the data shown in Figure 4 (and its interpretation) the data of fitness in the plant environment indicate that histidine is scarce in this environment—a finding consistent with previous measures that showed histidine to be present in the rhizosphere at ~3 μm (Zhang et al. 2006). The fitness of PBR815 (hutU::Ω) was indistinguishable from wild type (Figure 5), which indicates that neither histidine nor urocanate are important sources of carbon or nitrogen in the plant environment.

Figure 5.
Fitness of the hutCD deletion mutants relative to lacZ-marked P. fluorescens SBW25 on sugar beet seedlings. Data are shown in an order of strains: ancestral SBW25 (wild type), PBR805 (ΔhutC), PBR806 (ΔhutD), and PBR807 (ΔhutCD ...

DISCUSSION

Amino acids are a significant source of carbon and nitrogen in many terrestrial and marine environments (Jaeger et al. 1999; Phillips et al. 2004). Not surprisingly then, bacteria possess specific pathways dedicated to the uptake and degradation of specific amino acids. The histidine uptake and utilization pathway of SBW25 is typical of these pathways.

Our genetic analysis has confirmed previously known aspects of the function of the hut pathway, but has also extended understanding of gene composition, organization, and regulation. Of value has been the bringing together of various aspects of hut genetics within a single study to lay the foundations for further work. Indeed, our original goal in focusing attention on hut had been to study the function of the five predicted uptake components (Zhang et al. 2006), but this proved impossible without the genetics of hut first achieving a contemporary position.

In terms of hut metabolic genes the work reported here makes two new contributions. The first concerns the identity and genomic location of hutF, a gene long presumed to exist within organisms that degrade histidine by the five-step enzymatic pathway (Consevage et al. 1985), but that had escaped identification because the link between DNA sequence, amino acid sequence, and enzyme activity had not been made (Janiyani and Ray 2002; Rietsch et al. 2004).

Recently, Marti-Arbona et al. (2006) characterized three proteins from P. aeruginosa (PA5106, PA5091, and PA3175) with predicted roles in the breakdown of formiminoglutamate. Chemical analysis showed that PA5106 and PA5091 are HutF and HutG, which catalyze the last two steps of the five-enzyme pathway: the steps formiminoglutamate to formylglutamate (and ammonia), and formylglutamate to glutamate (and formate), respectively (Figure 1A). Interestingly, the gene product of PA3175 is able to catalyze the hydrolysis of formiminoglutamate to glutamate and formamide, the end reaction of the four-step hut pathway (Marti-Arbona et al. 2006). This finding, based solely on in vitro assays of enzyme activity, led to the suggestion that P. aeruginosa PAO1 may be capable of degrading histidine by both the four- and the five-step pathways (Marti-Arbona et al. 2006). Our mutant analyses suggest that this is unlikely. The hut operons of PAO1 and SBW25 are highly similar (Table 1); furthermore, SBW25 contains a homolog of PA3175 (Pflu4510: 42% amino acid similarity). If Pflu4510 is expressed, and functional, then PBR800 (hutF::Ω) ought not be compromised in its ability to grow on histidine. The fact that PBR800 (hutF::Ω) is unable to grow on histidine indicates that SBW25 degrades histidine, at least under the conditions of the assay, solely by the five-step pathway. By extension, the same is likely to be true of P. aeruginosa.

The second contribution stems from analysis of the two copies of hutH, both of which are predicted to encode histidine ammonia lyase or histidase. Our data show that hutH1 is incapable of complementing a hutH1/H2 double mutant even when expressed from a constitutive promoter. This indicates that the protein is not a functional histidine ammonia lyase. The fact that the hut locus from six of the seven genome-sequenced Pseudomonas species/strains also carries an additional copy of hutH is puzzling; moreover, the high degree of conservation of the hutH1 homologs [all lack the exact same three amino acid residues (Ser–Gly–Asp) that span the active site and share between 76 and 86% amino acid identity (X.-X. Zhang and P. B. Rainey, unpublished data)] adds to the mystery. A number of evolutionary scenarios can be envisaged, perhaps the most plausible being an ancient duplication or gene-acquisition event (which sits comfortably with the fact that HutH1 and HutH2 share only 37% sequence identity); although given the significant similarity among HutH1 homologs it would be necessary to invoke a single inactivation event in a single ancestral strain. An additional possibility is that hutH1 has an as yet undetermined functional role in the metabolism of histidine, for example, as a protein that binds (but does not metabolize) histidine. Such a role may be related to the need to control the upper limit of hut activity (see below).

In terms of regulation, hut is more complex and interesting than anticipated (Phillips and Mulfinger 1981). In all respects our work supports the previous claims that HutC is a repressor of hut (importantly, we experimentally demonstrated the involvement of HutC in the expression of each of the three operons) and that urocanate is the immediate inducer of expression (of all three operons); however, identification of the promoter sequences of PhutF and PhutU provided evidence of additional control by σ70 and σ54, respectively. Involvement of σ54 led to a search for a likely σ54-activator protein, the most obvious candidate being CbrB, a response regulator with a σ54-type regulator output domain (Nishijyo et al. 2001). While DNA binding studies are required to demonstrate the direct connection between CbrB and hut transcription, the evidence gathered (experimental and in silico) implicates CbrB as the σ54 enhancer binding protein for transcriptional activation of the hutU–G operon. The fact that hutU–G is regulated by both σ54 and HutC (plus urocanate) provides a rare example of a σ54-dependent operon that is subject to both general and specific regulation (Reitzer and Schneider 2001).

Analysis of hutCD demonstrates a further complexity to hut regulation: inactivation of hutD shows that it is required for efficient utilization of histidine (and urocanate) as a sole carbon and nitrogen source. The fact that hutD is translationally coupled to hutC, combined with the fact that HutD shows no homology to known histidine metabolic genes, suggests that HutD plays a regulatory role.

The combined hutC, hutD, and hutCD analyses show that across the range of growth and fitness assays the ΔhutCD genotype expressed a phenotype that was not consistently that of either the ΔhutC or the ΔhutD genotypes. In fact the hutC mutation was dominant to the hutD mutation in environments without histidine, but the reverse was observed in environments replete with histidine. This suggests that HutD acts independently of HutC. Independent action is consistent with an early report that showed that the protein encoded by the hutD homolog (orf2) from P. putida ATCC12633 was not required for binding of HutC to the operator site in front of hutU (Allison and Phillips 1990). Such a finding is also consistent with the fact that HutD lacks a helix-turn-helix domain and thus it is unlikely to act by binding to DNA.

In addition to evidence of independent action, the transcriptional, growth, and fitness assays suggest a more specific role for HutD, possibly as a governor of hut transcription. Such a role would be analogous to, although mechanistically distinct from, the “governor” site at the glnAp2 promoter of glutamine synthase (GS) in E. coli, which serves to limit the maximum activity of the GS promoter (Atkinson et al. 2002). The precise function of HutD remains to be determined but it is possible that HutD may limit hut activation by controlling the intracellular concentration of the hut inducer, urocanate, perhaps by binding this compound and thus preventing the intracellular concentration of inducer from exceeding a critical threshold level. Computational analysis based on the deposited crystal structure of HutD (PA5104) from P. aeruginosa PAO1 (PDB.1yll) indicates that urocanate, but not histidine, docks with the active site of HutD (V. Arcus, unpublished data). The biological need for a governor may relate to the potentially harmful effects that could result from an excess of intracellular ammonia, a likely consequence of too high a rate of histidine metabolism.

The complexity of hut regulation suggests that the metabolism of histidine is of central metabolic importance to Pseudomonas. Drawing upon Savageau's demand theory of regulatory control (Savageau 1974, 1989, 1998), negative regulation indicates that the hut operon is infrequently used and that histidine is scarce in the natural environment of this bacterium. Indeed, the scarcity of histidine in the plant rhizosphere has previously been shown (Zhang et al. 2006) and is also indicated here. Puzzling then is the positive control by CbrB because this suggests that histidine is frequently encountered in the environment. We cautiously suggest that the apparent paradox can be explained by envisaging a more general role for CbrAB, for example, as a sensor of a range of different amino acids and as a positive activator for a number of specific amino acid degradation pathways, a proposal for which there is some published evidence (Nishijyo et al. 2001; Knight et al. 2006) and for which we have additional data (X.-X. Zhang, D. G. Brown and P. B. Rainey, unpublished data).

Acknowledgments

We thank Allen Phillips for comments on this work and access to the unpublished hutF sequence of P. putida ATCC 12633. We also thank Dieter Haas for helpful discussions, Jake Jones and Gail Preston for providing the rpoN deletion mutant, Darby Brown and Mike McDonald for comments on the manuscript, and Annabel Gunn for technical support. This work was supported by the Marsden Fund Council from government funding administered by the Royal Society of New Zealand.

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