Importance of Tetramer Formation by the Nitrogen Assimilation Control Protein for Strong Repression of Glutamate Dehydrogenase Formation in Klebsiella pneumoniae

Abstract

The nitrogen assimilation control protein (NAC) from Klebsiella pneumoniae is a very versatile regulatory protein. NAC activates transcription of operons such as hut (histidine utilization) and ure (urea utilization), whose products generate ammonia. NAC also represses the transcription of genes such as gdhA, whose products use ammonia. NAC exerts a weak repression at gdhA by competing with the binding of a lysine-sensitive activator. NAC also strongly represses transcription of gdhA (about 20-fold) by binding to two separated sites, suggesting a model involving DNA looping. We have identified negative control mutants that are unable to exert this strong repression of gdhA expression but still activate hut and ure expression normally. Some of these negative control mutants (e.g., NAC^86ter and NAC^132ter) delete the C-terminal domain, thought to be required for tetramerization. Other negative control mutants (e.g., NAC^L111K and NAC^L125R) alter single amino acids involved in tetramerization. In this work we used gel filtration to show that NAC^86ter and NAC^L111K are dimers in solution, even at high concentration (NAC^WT is a tetramer). Moreover, using a combination of DNase I footprints and gel mobility shifts assays, we showed that when NAC^WT binds to two adjacent sites on a DNA fragment, NAC^WT binds as a tetramer that bends the DNA fragment significantly. NAC^L111K binds to such a fragment as two independent dimers without inducing the strong bend. Thus, NAC^L111K is a dimer in solution or when bound to DNA. NAC^L111K (typical of the negative control mutants) is wild type for every other property tested: (i) it activates transcription at hut and ure; (ii) it competes with the lysine-sensitive activator for binding at gdhA; (iii) it binds to the same sites at the hut, ure, nac, and gdhA promoters as NAC^WT; (iv) the relative affinity of NAC^L111K for these sites follows the same order as NAC^WT (ure > gdhA > nac > hut); (v) it induces the same slight bend as dimers of NAC^WT; and (vi) its DNase I footprints at these sites are indistinguishable from those of NAC^WT (except for features ascribed to tetramer formation). The only two phenotypes we know for negative control mutants of NAC are their inability to tetramerize and their inability to cause the strong repression of gdhA. Thus, we propose that in order for NAC^WT to exert the strong repression, it must form a tetramer that bridges the two sites at gdhA (similar to other DNA looping models) and that the negative control mutants of NAC, which fail to tetramerize, cannot form this loop and thus fail to exert the strong repression at gdhA.

Under conditions where the preferred nitrogen source, ammonia, is limiting, Klebsiella pneumoniae must obtain nitrogen from other sources. In almost every case, this ability requires a two-component system in which a phosphorylated activator, NTRC∼P, binds to an enhancer and interacts with RNA polymerase bearing the unusual sigma factor σ⁵⁴. The Ntr system activates a set of genes involved in the catabolism of amino acids and other nitrogenous compounds whose degradation products include ammonia or glutamate. The Ntr system also activates the transcription of another transcriptional regulator, the nitrogen assimilation control protein (NAC) (3, 11). NAC regulates a subset of genes that are dependent on RNA polymerase bearing σ⁷⁰ for their transcription. Thus, NAC serves to couple the σ⁵⁴-dependent transcription of the Ntr system with that of σ⁷⁰-dependent catabolic genes.

NAC is a member of the large, LysR family of transcriptional regulators (34). Like all LysR family members, NAC has a helix-turn-helix motif in its N terminus which is involved in DNA binding. Most LysR family members require an allosteric change in order to activate transcription (34). This change is not required for DNA binding but results in a change in the extent of the DNase I footprint, the DNA bend angle induced by the binding, or both. Although DNA binding information is encoded in the N-terminal domain, the allosteric change is induced by the binding of a small molecule to the C-terminal domain of most LysR family proteins or by a chemical modification of that domain. In contrast, NAC does not appear to require any cofactor or modification for its activity (36), and NAC mutants lacking the C-terminal domain are able to activate transcription normally (28).

NAC is a very versatile regulatory protein which uses a variety of different mechanisms to regulate transcription. At the hut and put operons, NAC activates transcription by a mechanism that requires NAC binding to a site centered at −64 relative to the start of transcription (15). Data from other LysR-type regulators suggest by analogy that NAC may interact with the α-subunit of RNA polymerase in such cases (22, 38). At the dad, ure, and cod operons, NAC activates transcription by a mechanism that requires binding to a site centered at −44, −49, and −59, respectively (20, 29; unpublished data). Little is known how this activation occurs. At its own nac promoter, NAC binds to two adjacent sites located between the enhancer and the promoter and alters the flexibility of the DNA, which in turn restricts the ability of the enhancer-bound NTRC∼P to contact RNA polymerase (12). At serA, NAC interferes with the ability of Lrp to activate transcription by an unknown mechanism (D. S. Heikka and R. A. Bender, unpublished results).

At the gdhA gene, NAC represses by two distinct mechanisms. NAC exerts a weak repression, about threefold, by binding to a site centered at −89 (16) and competing with a lysine-sensitive positive effector (16). NAC also exerts a strong repression of 10-fold or greater by a mechanism that requires that NAC bind to two sites, the one centered at −89 and another centered at +57 (16). This arrangement resembles that seen at ara, gal, and lac, where formation of a DNA loop by the repressor is important for repression (10, 17, 30). In an attempt to separate the various regulatory mechanisms of NAC, we isolated a negative control mutant, NAC^L111K, that could activate transcription at hut and ure but could not cause the strong repression at gdhA (18). The data presented here suggest that the only difference between NAC^L111K and wild-type NAC is that NAC^L111K cannot form tetramers and exists only as dimers, a property shared by other negative control mutants of NAC (NAC^NC). Thus, we suggest that the formation of a tetramer is important for the strong repression at gdhA, probably because tetramers are required for DNA loop formation.

MATERIALS AND METHODS

Bacterial strains and plasmids.

Descriptions and genotypes of the bacterial strains, plasmids, and fusions used here are given in Table 1. All Klebsiella pneumoniae strains (previously called K. aerogenes) were derived from strain MK53 (4).

TABLE 1.

Strains and plasmids used in this study

Strain, plasmid, or fusion	Relevant genotype or description	Source or reference
Strains
EB4335	DH5α/pCB1083, pCB1026	This work
EB5221	DH5α/pCB1083, pCB1252	This work
KC4727	Δ[bla]-2 hutC515 dadA1 nac204::λplacMu53	This work
KC4728	Δ[bla]-2 hutC515 dadA1 nac204::λplacMu53 gltD702::Tn5-131	This work
KC4989	Δ[bla]-2 hutC515 dadA1 hutC::attλ zed-2::Tn5-131 nac-1/pCB832 (λRZ-5ΦCB1202)	This work
Plasmids
pACYC184	Medium-copy-number cloning vector	4
pBluescript	High-copy-number cloning vector	Stratagene
pCB634	nac gene expressed constitutively by lacP in pGD103	This work
pCB730	Upstream region of the gdhA promoter from −173 to −23 cloned into pRJ800	This work
pCB832	2.4-kb fragment carrying lamB cloned into pGB2	16
pCB904	Downstream region of the gdhA promoter from −39 to +68 cloned into pRJ800	This work
pCB1026	EcoRI-BamHI fragment, encoding NAC fused with six histidine codons at the C terminus, cloned into pQE70	This work
pCB1083	lacIq cloned into pACYC184	6
pCB1041	Kmr gene from pGD103 replaced with the bla gene from pUC19	This work
pCB1051	EcoRI-HindIII fragment carrying the nac gene from pCB634 cloned into pCB1041	This work
pCB1227	NACL111K version of pCB1051	This work
pCB1235	NAC86ter version of pCB1051	This work
pCB1242	ure promoter region cloned into pBluescript	This work
pCB1252	NACL111K version of pCB1026	This work
pCB1426	nac promoter region cloned into pBluescript	This work
pCB1456	bla gene from pCB1051 replaced with a ClaI-HindIII fragment carrying the Emr gene from pJDC9	This work
pCB1465	NACL125R version of pCB1051	This work
pCB1556	NACL125R version of pCB1456	This work
pCB1557	NAC86ter version of pCB1456	This work
pCB1558	NACL111K version of pCB1456	This work
pGD103	Low-copy-number cloning vector expressing inserted material from lacP	9
pJDC9	Erythromycin-resistant cloning vector	7
pJF200	nac promoter region cloned into pTE103	11
pOS1	hut promoter region cloned into pUC18	31
pQE70	Expression vector	QIAGEN
pRJ800	Promoterless lacZYA reporter	33
pUC18	High-copy-number cloning vector	Gibco-BRL
pUC19	High-copy-number cloning vector	Gibco-BRL
Fusion
ΦCB1202	gdhAp (−116 to +23) fused to lacZ	16

Enzyme assays.

Cultures were grown at 30°C with aeration as described previously (1, 24). Histidase, β-galactosidase, glutamate dehydrogenase, and urease assays were performed on whole cells as described previously (24). Protein levels were determined by the method of Lowry et al. (23). A modified version of the urease assay was developed in order to screen a large number of mutants for elevated (NAC-activated) levels of urease. A 2-μl sample of concentrated cells (approximately 1 mg of cellular protein/ml) was incubated in 88 μl of 50 mM potassium phosphate buffer (pH 7.5) containing 20 μg/ml cetyltrimethylammoniumbromide for 5 min at room temperature. The reaction was initiated by adding 10 μl of 1 M urea and allowed to proceed for 5 min. The reaction was terminated and color developed as described elsewhere (24), except at room temperature.

Selection for NC mutants.

Plasmid pCB1051 was mutagenized by passage through the mutator strain XL-1 Red (Stratagene) following the manufacturer's instructions. The mutagenized plasmid DNA was introduced into strain KC4728 by electroporation using a Bio-Rad GenePulser. Cells were made electrocompetent by washing exponential-phase cells grown in SOB medium (25) twice with 10% glycerol and concentrating 10- to 50-fold. After recovering in SOC medium for 1 hour, cells were washed once in 1% KCl to remove the rich medium and plated on minimal medium (32) supplemented with ampicillin (100 μg/ml). Colonies were purified by single colony isolation on selective medium. Overnight cultures of each transformant were diluted 1:50 in fresh medium and supplemented with 100 μg/ml ampicillin (to maintain selection for the plasmid) and 2 μM nickel sulfate (to supply the requirement for the nickel-containing urease holoenzyme) and grown for approximately 4 to 5 h (optical density at 600 nm, 0.4 to 0.6) with aeration at 30°C in a roller drum. Cells were washed once in an equal volume of 1% KCl and concentrated 10-fold. Each culture was tested for urease activity using a modified version of the urease assay (as described above). Mutants that produced large amounts of urease were further tested by measuring histidase levels. Plasmid DNA from potential negative control mutants (those that grew on minimal medium and retained high levels of urease and histidase) was isolated and reintroduced into strain KC4728. Urease, histidase, and GDH levels were determined on these secondary transformants as described above.

Mutagenesis.

NAC^L111K, NAC^L125R, and NAC^86ter were created by site-directed mutagenesis with the GeneEditor in vitro site-directed mutagenesis system from Promega. The primers that were used resulted in the following mutations: L111K (a trinucleotide change from CTG to AAG at amino acid 111), L125X (a trinucleotide change from CTG to XXX, which would encode all of the possible amino acids at amino acid 125), L125R (a trinucleotide change from CTG to CGA at amino acid 125), and NAC^86ter (a trinucleotide change from CAG to TAG at amino acid 87). Changes in the nucleotides for the corresponding mutations were confirmed by sequencing both strands of the entire nac gene at the University of Michigan Sequencing Core Facility.

Purification.

Histidine-tagged proteins were purified using nickel affinity resin purchased from QIAGEN according to the manufacturer's protocol. One-liter cultures (LB supplemented with 100 μg/ml of ampicillin and 10 μg/ml of tetracycline) were grown to an optical density at 600 nm of 0.6, at which point isopropyl-β-d-thiogalactopyranoside (IPTG) was added to a concentration of 1 mM to induce the production of the protein. After 4 h of incubation, cells were harvested by centrifugation and resuspended in 5 ml of cracking buffer (100 mM sodium phosphate [pH 7.0], 250 mM NaCl, 2.5 mM MgCl₂, 10% glycerol, 10 mM imidazole, and 1 mM β-mercaptoethanol). Cells were disrupted by passage through a French pressure cell (twice), and debris was removed by centrifugation (30,000 × g for 30 min). Five milliliters of lysate was incubated with 1 ml of nickel resin with agitation for 1 hour at 4°C. This mixture was poured onto a column that retained the resin, while the remaining lysate was discarded. The resin was washed with 20 to 30 ml of wash buffer (100 mM sodium phosphate [pH 7.0], 500 mM NaCl, 2.5 mM MgCl₂, 10% glycerol, 90 mM imidazole, and 1 mM β-mercaptoethanol), and the His-tagged proteins were then eluted (using cracking buffer, with imidazole concentration increased to 250 mM). These fractions were pooled and dialyzed overnight in buffer 4 (250 mM NaCl, 100 mM NaH₂PO₄ [pH 7.0], 2.5 mM MgCl₂, 1 mM β-mercaptoethanol) to remove the imidazole. The NAC was concentrated using Centricon filters as per the manufacturer's directions. The protein was either used immediately or stored at −20°C in glycerol to 50%. Protein concentration was determined by the Lowry method (23).

Gel filtration.

About 7.5 μM His-tagged NAC^WT or His-tagged NAC^L111K was prepared in buffer 4 in a total of volume of 250 μl. The protein was applied to a Sephadex S200 gel filtration column (Amersham Pharmacia Biotech) equilibrated with buffer 4. During elution, the protein was detected by monitoring UV absorbance at 280 nm. Samples (200 μl) were collected and saved for Western blot analysis.

Western blotting.

Protein samples (15 μl) were loaded onto a 12% sodium dodecyl sulfate-polyacrylamide gel electrophoresis gel (Invitrogen), and the proteins were separated by electrophoresis at 100 V for 1 to 1.5 h. Separated proteins were transferred to a nitrocellulose membrane. The membrane was washed in TBS buffer (200 mM Tris [pH 7.5], 200 mM NaCl) and blocked with 10% powdered milk in TBS for 1 h. The membrane was washed twice in TBS. The primary antibody, anti-NAC 4E (Cocalico), was added at a dilution of 1:5,000 in milk-TBS buffer for 2 h at room temperature (or at 4°C overnight). The membrane was then washed three times in TBS TT buffer (TBS with 2% Triton X and 0.5% Tween). The secondary antibody, anti-rabbit-horseradish peroxidase (Sigma), was added at a dilution of 1:10,000 in milk-TBS buffer for 1 h at room temperature. The membrane was washed twice in TBS TT buffer, followed by a wash in TBS buffer. Immunoblots were developed using the ECL Western blotting detection reagents as per the manufacturer's instructions (Amersham Biosciences). The blots were exposed to Kodak BioMax film for 5 min.

Gel mobility shifts assay.

Gel mobility shift assays were performed with DNA from the hut, ure, gdhA, and nac promoters from K. pneumoniae. DNA was prepared by PCR with oligonucleotides that would amplify a 150-bp region of each promoter from pOS1 (plasmid containing the hut promoter region), pCB1242 (ure), pCB904 (plasmid containing the upstream region of the gdhA promoter), pCB730 (downstream region of gdhA), and pJF200 (nac). Purified His-tagged NAC^WT and His-tagged NAC^L111K were used for gel mobility shift assays. Protein was incubated with approximately 30 to 45 ng DNA in the presence of bovine serum albumin at room temperature for 20 min. Afterwards, 1.5 μl of loading buffer (40 mM Tris HCl [pH 8.4], 4 mM EDTA, 0.2% bromophenol blue, 0.2% xylene cyanol, 25% glycerol) was added and loaded onto a 4% acrylamide-Tris-acetate-EDTA gel. The samples were subjected to electrophoresis at 150 V for 3 to 4 h at 4°C. Gels were stained with ethidium bromide for 10 min with agitation and washed with water. DNA was visualized by UV fluorescence.

DNA bending assay.

DNA bending assays were performed with PCR-amplified DNA fragments of the nac promoter from the plasmid pJF200. Primers were used that generated a 205-bp DNA fragment which placed the center of the NAC protected region from nac at different positions in five DNA fragments. The center of the NAC-binding region was placed 40, 75, 105, 135, or 180 bp from one end of the DNA. Approximately 50 ng DNA was used for gel mobility shift assays (see above) with 0.82 pmol NAC or 11.0 pmol NAC^L111K. The bend angle, α, is calculated using the equation μ_m/μ_e = cos(α/2), where μ_m is the mobility of the protein bound to the DNA in the middle of the fragment and μ_e is the mobility of the protein bound to the DNA at the end of the DNA (39).

DNase I footprint assay.

DNase I footprint analysis was performed as described by Goss and Bender (15). Essentially, 10 μg of pCB1426 was digested with BamHI and labeled with [α-³²P]dATP (MP Biomedicals) in the presence of Klenow (Promega), followed by a second digest with XbaI. This generated two DNA fragments, one long fragment containing the nac promoter region and a short DNA fragment of less than 25 bp. Approximately 0.31 pmol labeled DNA was incubated with NAC protein for 20 min at room temperature. Afterwards, 1.7 × 10⁻⁴ U of DNase I (Promega) was added and incubated for 3 min at room temperature. Digests were stopped with loading buffer (40 mM Tris HCl [pH 8.4], 4 mM EDTA, 0.2% bromophenol blue, 0.2% xylene cyanol, 25% glycerol, 50% formamide) and boiled for 5 min. Digested DNA fragments were separated by gel electrophoresis on a 6% urea-Tris-borate-EDTA sequencing gel at 1,200 V for 3.5 to 4 h. The gel was dried on Whatman paper and exposed to Kodak BioMax film. An A+G ladder was prepared with 0.31 pmol of labeled DNA as previously described (2).

RESULTS

L111 is a part of a tetramerization domain in NAC^WT.

Most members of the LysR family of transcriptional regulators are tetrameric (34). However, the short footprint of NAC at the hut promoter (15) and other hints led us to suspect that NAC might function as a dimer (3). We therefore examined the oligomeric state of NAC^WT and the NAC^NC mutant, NAC^L111K (both isolated as His-tagged variants) by gel filtration analysis. NAC^WT eluted from a fast-performance liquid chromatography column with an apparent mass of 130 kDa, in close agreement with the predicted mass of a homotetramer, 128 kDa (Fig. 1A). NAC^L111K eluted with an apparent mass of 64 kDa, in agreement with the predicted mass of a homodimer, 64 kDa (Fig. 1A). The elution profiles of both the NAC^WT and NAC^L111K preparations showed a minor peak eluting with a lower apparent mass; however, Western blot analysis showed that this material was not recognized by either anti-NAC antibody (Fig. 1B) or antibody directed against the His tag (not shown). When NAC^WT and NAC^L111K were mixed before gel filtration, both the tetramer peak and the dimer peak were seen (Fig. 1A). Finally, when the NAC^WT and NAC^L111K preparations were diluted 10-fold before gel filtration, NAC again eluted as a tetramer and NAC^L111K eluted as a dimer (data not shown). Thus, we conclude that NAC is able to form tetramers in solution, whereas NAC^L111K cannot and is dimeric.

FIG. 1.

Fast-performance liquid chromatography analysis of the oligomerization state of NAC^WT and NAC^L111K. A. NAC^WT, NAC^L111K, or a mixture of both NAC^WT and NAC^L111K was applied to a Sephadex S200 column and eluted in buffer 4. Protein was detected by the absorbance (mA) at 280 nm. Positions of size standards (adolase [158 kDa], bovine serum albumin [67 kDa], ovalbumin [43 kDa], chymotrypsin [25 kDa], and RNase [16 kDa]) are shown at the top of the graph. B. Western blot analysis of fractions collected from the gel filtration column. Filters were probed with an antibody raised against purified NAC. Fractions where size standards elute are indicated at the top of each blot.

The OxyR protein from Escherichia coli is a LysR family member that is very similar to NAC from K. pneumoniae (24% identical over the entire protein, 37% identical for the N-terminal 86 amino acids) (35). A three-dimensional structure had been determined for OxyR's C-terminal domain (8), the domain that contains I110 of OxyR, the amino acid equivalent to L111 of NAC. In OxyR, I110 and L124 of one subunit of OxyR interact with either F219 (in the reduced form of OxyR) or A232 (in the oxidized form) of another subunit, and this interaction appears important for tetramerization (8). The amino acids near I110 and L124 (corresponding to L111 and L125 of NAC) are generally conserved within the LysR family, and so we reasoned that mutagenesis of codon 125 in NAC might yield a negative control mutant. A pool of site-directed mutants derived by using a randomized trinucleotide at codon 125 was screened as described in Materials and Methods, and a mutant with the negative control phenotype was isolated. This mutant had a single amino acid change at position 125, from a leucine to an arginine. This mutant, NAC^L125R, like NAC^L111K, was able to activate expression of the hut and ure operons nearly as well as NAC^WT but was unable to show the strong repression of gdhA expression (Table 2).

TABLE 2.

Characterization of NC mutants

Plasmid	nac allelea	Sp act (nmol of product formed/min/mg of protein)b
		Histidase	Urease	GDH
pCB1041	No NAC	35	25	673
pCB1051	Wild type	318	779	38
pCB1227	L111K	239	1,080	431
pCB1465	L125R	192	775	329
pCB1235	86ter	228	943	288
pCB1236	133ter	169	907	368

^aThe NAC⁻ strain KC4727 (hutC5151 Δ[bla]-2, nac-204::λplacMu53) contained a plasmid which carried the various nac alleles that were expressed constitutively from a lac promoter.

^bThe medium was W4 minimal salts (32) supplemented with 0.4% glucose, 0.2% ammonium sulfate, 0.2% glutamine, and 100 ug/ml ampicillin. The specific activities of histidase and urease (which requires NAC for activation) and GDH (which requires NAC for repression) were measured. Assay values are reported as specific activities and are the means of at least three independent experiments in which the standard error of the mean was <15%.

The amino acids near F219 and A232 (corresponding to G217 and G231 of NAC) are not conserved within the LysR family. Nevertheless, we attempted to isolate negative control mutants by site-directed mutagenesis of codons G217, E218, L225, A230, and G231 in NAC. We were able to identify many mutations at each codon that were unable to repress gdhA expression. However, none of these mutants was a negative control mutant, since these mutants also failed to activate transcription at the hut and ure operons.

In our initial screens (18), we had previously isolated two additional negative control mutants. One mutant, NAC^86ter, had a stop codon in position 87 and produced a NAC polypeptide containing only the N-terminal 86 amino acids of NAC. The other mutant, NAC^132ter, had a 2-bp deletion in codon 127 and yielded a NAC polypeptide with 132 amino acids, 126 of which were authentic NAC sequence and the C-terminal 6 of which were translated in the wrong reading frame before reaching a stop codon in that frame. Like the other negative control mutants, NAC^86ter and NAC^132ter were able to activate hut and ure expression but failed to show the strong repression of gdhA (Table 2). As expected, gel filtration indicated that NAC^86ter was dimeric in solution (data not shown; NAC^132ter was not tested.) The properties of these truncated forms suggest that the C-terminal domain of NAC is necessary for tetramer formation and for the strong repression at gdhA but not for activation at hut and ure.

NAC binds to the promoter region of the hut and ure promoters as a dimer.

The DNase I footprint of many LysR family members on their cognate DNA sequences is consistent with that expected from the binding of a dimer of dimers. For example, NAC protects two adjacent regions ofthe nac promoter of about 26 bp each, with a region of hypersensitivity between them (12). Similarly E. coli OxyR protects two regions of about 22 bp with either a space of one helical turn of the DNA space between them (reduced form) or no space between them (oxidized form) (40). However, NAC protects the hut and ure promoter regions from DNase I in a single region of about 26 bp (15). This suggested either that NAC binds as a dimer at hut and ure or that only one dimer within a NAC tetramer is able to protect the DNA. To distinguish between these two models, we compared the binding of NAC^WT and NAC^L111K at hut and ure using gel mobility shift analysis.

In order to normalize for the different affinities of the NAC-binding sites at hut and ure, the concentrations used in the mobility shift assays in Fig. 2 were adjusted such that about 50% of the input DNA was shifted. NAC^L111K gave a single shifted species with hut DNA (Fig. 2, lane 3). Because NAC^L111K is a dimer in solution, we believe that the shifted species is a complex of a NAC^L111K dimer bound to the DNA. In some cases it is possible to estimate the size of the protein in a shifted band by estimating the apparent molecular weight of the complex from its mobility (13, 21). The mobility of the shifted complex suggests a mass of 170 kDa, close to the 160 kDa predicted for one fragment of DNA (96 kDa) plus two protomers of NAC (64 kDa). Thus, we refer to the mobility shift seen with NAC^L111K bound to hut DNA as a “dimer shift”.

FIG. 2.

Gel mobility shift assay of NAC-regulated promoters with NAC^WT or NAC^L111K. Approximately 30 to 45 ng (0.3 to 0.45 pmol) DNA containing the NAC-binding site from hut (lanes 2 and 3), ure (lanes 4 and 5), nac (lanes 6, 7, and 8), the downstream NAC-binding site from gdhA (lanes 9, 10, and 11), or the upstream NAC-binding site from gdhA (lanes 12 and 13) was incubated with NAC^WT (lanes 2, 4, 6, 9, 10, and 12 [1.4, 0.6, 1.1, 1.0, 1.9, and 0.8 pmol, respectively], denoted as “WT”) or NAC^L111K (lanes 3, 5, 7, 8, 11, and 13 [7.9, 3.3, 6.9, 31.5, 4.7, and 4.4 pmol, respectively], denoted as “L111K”). Lane 1 is the 1-kb Plus DNA ladder (Invitrogen). Estimated molecular masses of the DNA ladder are marked to the left of the figure. Each binding site had a different affinity for NAC^WT. The NAC^WT protein concentration was adjusted to shift approximately 50% of each DNA fragment. The activity of NAC^L111K is typically 20% that of NAC^WT at all NAC-binding sites tested. Thus, a 5:1 ratio of NAC^L111K to NAC^WT was used in all gel mobility shift assays and the DNase I footprint assay. “Excess L111K” in lane 8 and “excess WT” in lane 10 indicate an excess amount of NAC was used to visualize the ability of these proteins to produce an extra shift at the respective DNA fragments.

NAC^WT also gave a single shifted band at hut under these conditions (Fig. 2, lane 2). This shifted species had the same mobility as the corresponding complexes formed with NAC^L111K. Even when the concentration of NAC was increased to a level where all the free DNA was shifted, this single band was the only shifted species seen (data not shown). At very high concentrations of NAC, both the hut promoter and the vector DNA present in the binding mixture were shifted to very slow mobilities, as expected for the nonspecific binding of a DNA binding protein (not shown). Because both NAC^WT and NAC^L111K produced the same shifted species, we conclude that NAC^WT binds as a dimer at hut.

When DNA containing the ure promoter was used in place of hut DNA, NAC^WT and NAC^L111K again gave identical shifted bands, and the mobility of this band was identical to the dimer shift seen with hut DNA (Fig. 2, lanes 4 and 5).

NAC binds as a tetramer at the nac promoter.

When the NAC-binding site from the nac promoter was used in mobility shift assays, NAC^WT shifted a small amount of the DNA to a position identical to the dimer shift seen with hut and ure DNA (Fig. 2, compare lanes 2, 4, and 6). However, the major bound species had a much-reduced mobility (Fig. 2, lane 6). NAC^WT is known to bend nac DNA (12), and so the mobility of the slowly migrating species is dependent on the position of the NAC-binding site within the DNA fragment (Fig. 3, lanes 1 to 5). This makes it difficult to estimate the mass of the complex from the mobility. The effect of bending on DNA mobility is at a minimum when the binding site is at the end of a DNA fragment (12). By extrapolating to the mobility expected when the binding site is at the end of the DNA, we estimated that the mass of the slow-migrating complex is about 240 kDa, in reasonable agreement with the 228-kDa mass expected for a DNA fragment (96 kDa) plus four protomers of NAC (132 kDa). The degree of bending induced by the binding of NAC^WT to nac DNA can be estimated from the changes in mobility as a function of distance from the end of the fragment (39). The data in Fig. 3 suggest a bend of about 113° for the tetramer shift. The dimer shift suggests a small bend of about 42°, as had been seen for NAC bound at hut (27).

FIG. 3.

DNA bending of the nac promoter by NAC. DNA fragments were PCR amplified with primers to generate 205-bp fragments that positioned the center of the NAC-binding region 40 bp (lanes 1 and 6), 75 bp (lanes 2 and 7), 105 bp (lanes 3 and 8), 135 bp (lanes 4 and 9), and 180 bp (lanes 5 and 10) from one end of the DNA. Approximately 50 ng (0.52 pmol) of DNA was incubated with 0.82 pmol NAC^WT (lanes 1 to 5) or 11.0 pmol NAC^L111K (lanes 6 to 10), and the bound and free DNA fragments were separated by electrophoresis on a 4% polyacrylamide gel as described in Materials and Methods.

When NAC^L111K bound to nac DNA, the predominant species corresponded to the dimer shift (Fig. 2, lane 7). At high concentrations of NAC^L111K, a small amount of a slower-migrating species was seen, but this differed from that seen with NAC^WT in that its mobility was essentially independent of the position of the NAC-binding site within the fragment (Fig. 2, lane 8, and Fig. 3, lanes 6 to 10). Thus, NAC^L111K binds to nac DNA as a dimer. The slight amount of slower-migrating material does not include the bending seen with the true tetramer of NAC^WT and probably represents the weak binding of a second, independent dimer (see Discussion). The mass of this second shift was estimated to be 230 kDa, essentially equivalent to the mass of the DNA plus four protomers of NAC. Thus, we argue that NAC^L111K is a dimer in solution and remains so even when bound to DNA.

The DNase I footprint of NAC bound at the nac promoter supports this conclusion. As we have shown before (12), NAC protects two adjacent regions of DNA with a strong hypersensitive region between them (Fig. 4). There is also a weak hypersensitive site within one of the sites (Fig. 4, region A), similar to the slight hypersensitivity that is sometimes seen when dimeric NAC binds to hut or ure DNA (15). The sequences required for NAC binding at repression sites are not fully understood but seem to require either ATA-N₉-TAT (also required as part of an activating site) or a related sequence, ATC-N₉-TAT, which is equivalent to ATA-N₉-GAT (15, 27). Region A of the protection at nac mediated by NAC^WT contains the sequence ATC-N₉-TAT centered within the protected region (Fig. 5). Region B contains the sequence ATA-N₉-GCT at an equivalent position within the protected region, perhaps suggesting at least a half-site for NAC binding or perhaps a full site with a 1-bp mismatch from consensus.

FIG. 4.

DNase I footprint of NAC^WT and NAC^L111K bound to the nac promoter region. Radioactively labeled pCB1426 DNA was digested with DNase I in the presence of NAC^WT (lanes 2 to 6 [0, 0.9, 1.4, 2.2, and 4.4 pmol, respectively]) or NAC^L111K (lanes 7 to 11 [4.7, 9.5, 12.6, 12.6, and 25.2 pmol, respectively]). Lane 1 is the radiolabeled DNA without any DNase I treatment. Lane G is the G ladder. The solid line on the right of the footprint is the region protected by both NAC^WT and NAC^L111K. Arrowheads indicate the regions of DNase I hypersensitivity by NAC^WT, NAC^L111K, or both.

FIG. 5.

DNA sequence of the nac promoter region from −240 to +1 (relative to the start of transcription). The DNA sequence protected by NAC is indicated by the bold lines (regions A and B) corresponding to regions A and B in Fig. 4, with the hypersensitive sites indicated byarrowheads. Boxed nucleotides represent the NAC consensus sequence containing ATC-N₉-TAT. A proposed second NAC-binding site, ATA-N₉-GCT, is underlined. The σ⁵⁴-dependent promoter of nac is indicated by the dashed line above the DNA sequence.

In contrast to the result with NAC^WT, NAC^L111K protects only region A, with the slight hypersensitive site within that region (Fig. 4). At very high concentrations of NAC^L111K, some protection of region B is seen, but without the strong hypersensitivity between regions A and B seen when NAC^WT was used (Fig. 4). We interpret this to suggest the weak binding of a second, independent dimer of NAC^L111K in region B without the concomitant formation of a tetramer capable of bending the DNA and creating the hypersensitive site that results from the bending. Thus, it appears that NAC^L111K cannot form a tetramer either in solution or when bound to DNA.

Lack of tetramer formation and failure of gdhA repression.

The strong repression of gdhA expression by NAC requires two NAC-binding sites, one centered at −89 and the other at +57 relative to the start of transcription. Elimination of either site abolishes the strong repression (16). There are two broad classes of explanation for the failure of the negative control mutants to exert this strong repression: they might fail to recognize the binding sites that are specific for repression and bind only to the “activation sites,” or they might fail at some stage in the interaction between the NAC-DNA complexes at these two sites.

We have previously determined that NAC^WT and NAC^L111K bind to DNA fragments containing either of the NAC-binding sites from the gdhA promoter (18). The data in Fig. 2 show that both NAC^WT and NAC^L111K caused identical dimer shifts at the upstream site (centered at −89) (lanes 12 and 13) and at the downstream site (centered at +57) (lanes 9 and 11). In addition, NAC^WT caused a distinct tetramer shift at the downstream NAC-binding site when NAC concentrations were higher (Fig. 2, lane 10), but NAC^L111K did not (18). Thus, as was seen earlier (18), NAC^L111K binds as well as NAC^WT to both sites in vitro.

To determine whether NAC^L111K could recognize a NAC binding site from gdhA in vivo, we took advantage of the weak repression of gdhA by NAC. This weak repression requires only the upstream NAC binding site and results from a competition between NAC and a lysine-sensitive positive effector, probably ArgP (T. J. Goss and R. A. Bender, unpublished data), that also binds to this region.

Transcription of gdhA is reduced about two- to threefold when any of these conditions exists: when the lysine-sensitive effector (ArgP) is absent (Goss and Bender, unpublished), when the upstream binding site is mutated or deleted (16), when lysine inactivates the effector (19), or when NAC is bound to the upstream site (16). Thus, we used a gdhA-lacZ transcriptional fusion with only the upstream NAC-binding region to test for the ability of NAC (and the NAC^NC mutants) to compete with the lysine-sensitive effector. In the presence of lysine (but no NAC), gdhA transcription was reduced about two- to threefold (Fig. 6). When NAC^WT was present, transcription was reduced to this lower level, even in the absence of lysine, demonstrating the effective competition by NAC^WT (Fig. 6). When the NAC^NC mutants replaced NAC^WT in the assay, gdhA transcription was reduced to a level comparable to that seen when NAC^WT was used (Fig. 6). Addition of lysine to inactivate the positive effector had little or no additional effect (Fig. 6). Thus, each of the NAC^NC mutants tested was able to compete as effectively as NAC^WT to prevent activation by the effector.

FIG. 6.

Ability of NAC^WT and NAC^NC mutants to block activation by the lysine-sensitive effector at gdhA. Strain KC4989 carries the gdhA promoter region from −116 to +23 (relative to start of transcription) fused to a lacZ reporter and integrated in single copy into the chromosome. The gdhA promoter region contains the upstream NAC-binding site and the binding site for the lysine-sensitive positive effector. When grown in medium lacking lysine, β-galactosidase activity from KC4989 is expressed at its maximum from this promoter construct (nac allele labeled “none” [white bar]), whereas in medium containing 0.01% lysine, the β-galactosidase activity is expressed at its basal level (“none” [gray bar]). Plasmids that constitutively expressed NAC^WT (pCB1456) or the NAC^NC mutants [NAC^L111K (pCB1558), NAC^L125R (pCB1556), or NAC^86ter (pCB1557)] from the lac promoter were introduced into KC4989, and β-galactosidase activity was assayed in each strain grown in media without (white bars) or with (gray bars) lysine. Activities are the averages of at least three assays with a standard error of the mean of less than 15%.

Since NAC represses gdhA expression by two mechanisms (weak and strong repression), it was important to show the effects of the mutations on each mechanism independently. The presence of lysine eliminates the activation by the effector (and hence the weak repression by NAC). When cells are grown in the presence of lysine, only the strong repression by NAC^WT is relevant. As seen in Fig. 7, expression of NAC^WT resulted in a 10- to 20-fold repression of GDH accumulation, whereas expression of the NAC^NC mutants resulted in less than a 30% reduction. Thus, the NAC^NC mutants are fully functional in exerting the weak repression (which requires binding and competing with the lysine-sensitive effector) but seriously defective in exerting the strong repression (which requires binding to two sites and some other step, presumably tetramer formation).

FIG. 7.

The NAC^NC mutants fail to exhibit the strong repression at gdhA. Plasmids that constitutively expressed NAC^WT (pCB1051) or the NAC^NC mutants [NAC^L111K (pCB1227), NAC^L125R (pCB1465), or NAC^86ter (pCB1235)] from the lac promoter were introduced into the NAC⁻ strain KC4727. The plasmid pCB1041 which did not carry the nac gene (labeled as “none”) was also introduced into KC4727. Specific activity of GDH was measured for each strain grown in medium containing 0.01% lysine. The presence of lysine allowed us to examine the repression by NAC^WT or the NAC^NC mutants in the absence of the lysine-sensitive positive effector. Activities are the averages of at least three assays, with a standard error of the mean of less than 15%.

DISCUSSION

The negative control mutants described here were all identified by their failure to cause the strong repression at gdhA. Some of these, like NAC^L111K and NAC^L125R, have nonconservative amino acid changes in their C-terminal domains; others, like NAC^86ter, NAC^132ter, and NAC^100ter (28), lack the entire C-terminal domain. The one feature that all the negative control mutants share is their inability to form tetramers in solution or on DNA.

This failure in tetramerization is the only defect we have detected. NAC^L111K binds to the same sites as NAC^WT at the hut, ure, nac, and gdhA promoters. The ability of the bound dimer of NAC^L111K to induce a slight DNA bend resembles that of NAC^WT (Fig. 3) (27). The DNase I footprints of bound dimers of NAC^L111K and NAC^WT are indistinguishable at hut, ure, gdhA (not shown), and nac (Fig. 4). The ability of NAC^L111K to activate transcription at hut and ure is almost as great as that of NAC^WT. The ability of NAC^L111K to compete with the lysine-sensitive activator at gdhA is as strong as that of NAC^WT. Finally, although the affinity of NAC^L111K for DNA appears to be slightly weaker than that of NAC^WT, its binding specificity is unaltered. In vitro (in gel mobility shift assays), the relative affinities of both NAC^WT and NAC^L111K follow the order ure > upstream gdhA > downstream gdhA > nac > hut. This order parallels that seen in vivo, where NAC production is regulated by increasing IPTG addition (36). The in vivo order is urease activation > GDH repression > histidase activation.

The fact that the truncated NAC^NC mutants (e.g., NAC^86ter) have the same phenotypes as the full-length NAC^NC mutants (e.g., NAC^L111K) suggests two things: (i) that the first 86 amino acids are sufficient for DNA binding, interacting with RNA polymerase, and activating transcription, and (ii) that the C-terminal portion is necessary for the strong repression of gdhA transcription. The crystal structure of a related protein, CbnR, shows that amino acid 86 is the last amino acid in the N-terminal domain and lies at the end of a long helix that seems to be involved in dimer formation of CbnR (26). When we constructed truncations shorter than NAC^86ter, all NAC activities were lost and no negative control mutants were found, presumably because disruption of the dimerization helix traps NAC as monomers or because the fragments are degraded. In fact, Western blot analysis of NAC from chloramphenicol-treated cells suggests that NAC^86ter was degraded more rapidly (and thus less stable) than NAC^WT (data not shown). Moreover, we have isolated a mutant (NAC^I71A) in what should be the dimerization helix of NAC. NAC^I71A is monomeric in solution, and the mutant strain has a Nac⁻ phenotype, despite the accumulation of substantial amounts of NAC polypeptide (Q. Liu and R. A. Bender, unpublished observations). Thus, dimerization appears to be important for NAC to function, and a dimer of NAC^86ter carries most of NAC's functions intact.

All the negative control mutants isolated to date share a single defect: the inability to form tetramers. This strong correlation strongly suggests that tetramer formation is an essential component of the strong repression of gdhA transcription. Thus, our working model for this repression is formation of a DNA loop with one dimer of a NAC tetramer bound to the site at −89 and another dimer bound at +57.

The crystal structures of other LysR proteins suggest that the NAC tetramer is actually a dimer of dimers. The data presented here raise an interesting question about the stability of the tetramer. In gel filtration experiments, NAC elutes as a tetramer when applied at concentrations as low as 4.5 μM (at lower concentrations, NAC elutes as a dimer). Yet upon binding to hut or ure DNA, only one dimer is bound. It appears that the interaction between the N-terminal domain and DNA somehow weakens the interaction between the two dimers in the NAC tetramer. In LysR-type proteins, where the C-terminal domain is involved in signal transduction, the allosteric change induced in the C-terminal domain (caused by binding a small molecule or by chemical modification) can sometimes affect the interaction (affinity) of the N-terminal domain with DNA. So it would not be surprising if an allosteric effect could be transmitted in the opposite direction. Thus, binding of the N-terminal domain to DNA might somehow alter the C-terminal domain so as to weaken the dimer-dimer interactions. (Nor is the notion of DNA acting as an allosteric effector a new one [5, 14, 37].) If so, then how can we reconcile this with the requirement for tetramer formation in the strong repression? There are clear differences between the NAC-binding sites where NAC activates transcription and those where NAC represses. In fact, a properly positioned NAC-binding site from gdhA failed to function in NAC-mediated activation for hut transcription (32). Most sites where NAC activates have within their consensus sequence the pattern ATA-N₉-TAT, whereas the repression sites often contain the pattern ATC-N₉-TAT (or its equivalent, ATA-N₉-GAT). If these and other sequence differences are responsible for the different activities of NAC, we might expect that the conformation of NAC at activating sites favors an asymmetric dimer (32), whereas that at repressing sites is more permissive for the tetramer.