HflD, an Escherichia coli protein involved in the λ lysis–lysogeny switch, impairs transcription activation by λCII

Archives of Biochemistry and Biophysics 493 (2010) 175–183

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HflD, an Escherichia coli protein involved in the k lysis–lysogeny switch, impairs transcription activation by kCII Pabitra K. Parua, Avisek Mondal, Pradeep Parrack * Department of Biochemistry, Bose Institute, P-1/12, C.I.T. Scheme VIIM, Kolkata 700 054, India

a r t i c l e

i n f o

Article history: Received 6 August 2009 and in revised form 16 October 2009 Available online 22 October 2009 Keywords: Lysis–lysogeny decision Bacteriophage lambda kCII CII–HflD interaction Transcription inhibition

a b s t r a c t The CII protein of bacteriophage lambda is the key regulator for the lytic–lysogenic choice of the viral lifecycle. An unstable homotetrameric transcription activator of the three phage promoters pE, pI and paQ, kCII is stabilized by kCIII and destabilized by the host protease, Escherichia coli HflB (FtsH). In addition, other E. coli proteins HflK, HflC and HflD also influence lysogeny by acting upon CII. Among these, HflD (22.9 kDa), a peripheral membrane protein that is exposed towards the cytoplasm, interacts with CII and decreases the frequency of lysogenization of k by stimulating the degradation of CII. In this study, we show that in addition to helping CII degradation, HflD inhibits the DNA binding by CII, thereby inhibiting CII-dependent transcription activation. From biochemical, biophysical and modelling studies we also suggest that HflD–CII interaction takes place through the Cys31-accessible surface area of monomeric HflD, which binds to tetrameric CII as a 1:1 complex. Ó 2009 Elsevier Inc. All rights reserved.

Introduction When the temperate coliphage k infects Escherichia coli, it can follow either the lytic or the lysogenic pathway, depending upon the conditions of infection and conditions of growth [1–3]. The choice of pathways is influenced by several phage proteins as well as host proteins [4,5]. The most important regulator for the lytic/ lysogenic decision is kCII, an unstable homotetrameric transcription factor that activates the three k promoters pE, pI and paQ, leading to products that help establish the lysogenic pathway [6–13]. kCII is a small (4  97 aa) protein that assumes an all-helix structure with each monomer comprising four a helices (a1–a4) and a disordered C-terminal end (16 residues) [14–17]. The a4 helix (residues 64–77) from each subunit is responsible for the tetramerization of the protein, formed by a four-helix bundle [14,15]. The disordered C-terminal tail is the target for the proteolysis of CII by E. coli HflB (also known as FtsH) and makes CII unstable, a property essential for its function [18–20]. The three-dimensional crystal structure of the CII–DNA complex has revealed that CII binds DNA as a tetramer (dimer of dimers), where monomer A of the AB dimer and monomer C of the CD dimer make nearly identical, sequence-specific interactions with every base pair of the TTGC direct repeats of DNA via the recognition helix (a3) [15].

* Corresponding author. Address: Department of Biochemistry, Bose Institute, P1/12, C.I.T. Scheme VIIM, Kolkata 700 054, West Bengal, India. Fax: +91 33 23553886. E-mail address: [email protected] (P. Parrack). 0003-9861/$ - see front matter Ó 2009 Elsevier Inc. All rights reserved. doi:10.1016/j.abb.2009.10.010

The cellular concentration of CII determines which pathway the phage will follow. The level of CII depends upon kCIII, which stabilizes CII by its antiproteolytic action against the host protease HflB [21–24]. It also depends upon other host factors, viz. HflK, HflC and HflD [5,25–26], which destabilize CII, thereby affecting the lysis– lysogeny decision [4]. Among the latter, HflD (213 amino acids) is a peripheral membrane protein of E. coli that is exposed to the cytosol [26]. The 640 bp hflD gene, previously known as ycfC [27], is located at 23.7 min in the E. coli genome, upstream of purB and downstream of asuE (trmU) [28]. The purB gene encodes succinyl-AMP (S-AMP) lyase required for purine metabolism. Despite its location within the pur operon, ycfC was not found to play any role in purine biosynthesis, neither is it essential for the growth of E. coli [28]. Random mutational studies identified some point and deletion mutations within this gene that increased the lysogenization frequency of k [26]. Subsequently, ycfC was named hflD, a new hfl (high frequency of lysogenization) gene. Its gene product HflD interacts with CII both in vivo and in vitro [26] and stimulates the in vivo degradation of CII, thereby decreasing the lysogenization frequency of k [26]. However, HflD inhibits the HflB-mediated degradation of CII in vitro [26]. The reason behind these contradictory roles of HflD is not known. It was proposed that HflD sequesters CII from its target promoters (pE, pI and paQ) and recruits it to membrane-bound HflB for rapid degradation [26]. Nevertheless, no information is available about the regions of CII or of HflD that are involved in CII–HflD interaction. The mechanism of action of HflD remains unresolved. The three-dimensional structure of HflD (YcfC), deposited in the protein databank in 2004 (PDB ID: 1QZ4 and 1SDI) remains unpublished.

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In this study, we have explored how HflD interacts with CII, and examined the effect of HflD on CII-activated transcription. Through biochemical and biophysical experiments supplemented with modelling studies we provide some newer aspects of CII–HflD interactions, which may be helpful in understanding the mechanism of action of HflD. Materials and methods Bacterial strains and plasmids Commonly used E. coli strains and plasmids were obtained from commercial sources as mentioned in the text. All other strains, plasmids and primers used in this study are shown in Table 1. Construction of plasmids For expression of HflD with a 6-histidine tag (His6-HflD), the hflD gene was removed from pKH449 [26] by digestion with BamHI and cloned into pET28a vector (Novagen) at the BamHI site. The resulting plasmid was named pKP509. pKP07 was prepared to replace the T7 promoter of pET28a by lac promoter. The lac promoter-operator region (lacP/O; 122 bp) was PCR amplified using LacF and LacR as the forward and the reverse primers, and pUC19 (Fermentas) as the template, and was cloned into pET28a at the BglII and XbaI sites. pKP109 (carrying lacZ under the control of pE) was constructed by cloning pE (298 bp) upstream of lacZ at the XbaI site into pSD5B [29] vector in the correct orientation. pKP219 (contains lacZ under pE and cII under lacP) and pKP419 (contains lacZ under pE and cIIA, encoding residues 1–82 of cII, under lacP) were constructed for in vivo transcription assays. First the cII (or cIIA) gene was PCR amplified using CIIF and CIIR (or CIIF and

CIIAR) as forward and reverse primers and pAB305 [30] as template, and cloned into pKP07 at the NdeI and BamHI sites downstream of lacP/O, to obtain pKP106 (or pKP108). The lacP/O-cII (500 bp) (or lacP/O-cIIA, 450 bp) cassette was then removed from pKP106 (or pKP108) by digestion with BglII and BamHI and cloned into the pKP109 vector at the BglII site. pKP917 (carrying hflD under lacP/O) was constructed for in vivo transcription assays. hflD was first removed from pKH449 by digestion with BamHI and cloned into BamHI-digested pKP07 under lacP/O, to obtain pKP107. The lacP/O-hflD cassette (830 bp) was then removed from pKP107 by digestion with BglII and BamHI and cloned into pQE30 (Qiagen) at the BamHI site. Prior to sub-cloning of the lacP/O-hflD cassette into pQE30, the plasmid was modified by removing its existing T5 promoter-lacO region by digestion with XhoI and EcoRI followed by end-filling with Klenow polymerase and self-ligation. The pKP917 was digested by BamHI and self-ligated. The resultant plasmid that contained only the lacP/O region (without any downstream gene) was named pKP908 and was used as the empty control vector for in vivo studies. Mutation and cloning of the hflD gene The specific mutation (W181A) within the hflD gene was carried out by PCR-directed site specific mutagenesis [31a]. In the primary step, two separate PCRs were performed taking pKP509 as template. In one, PDF was used as the forward primer and PRW181A as the reverse primer. In the other, RD and PFW181A were used as the reverse and the forward primers, respectively. To amplify the complete mutated gene, secondary PCR was carried out using the two overlapping fragments obtained from the primary PCR as the template and PDF and RD as the forward and the reverse primers. The resulting 640 bp PCR product was then cloned into pKP07 at the NdeI and XhoI sites and named

Table 1 Bacterial strains, plasmids and primers used. Strains and plasmids

Genotype/description

Protein expressed

Reference/Source

E. coli strains AK525

Dpro-lac thi/F0 lacIq ZM15 Y+ pro+ zgj-460::Tn5 zgj-525::IS1A

–

Kihara et al. [42]

Plasmids pKP07 pKP109 pKP219 pKP917 pKP908 pKP804 pKP509 pAB305 pSD5B pKH449 pKP108 pKP419 pAB412

lacP/O pE-lacZ pE-lacZ & lacP-cII lacP-hflD lacP lacP/O-hflD (W181 is substituted by A) T7P-hflD T7P-cII containing lacZ gene and p15A replicon carrying gst-hflD under the tac promoter lacP/O-cIIA (expressing truncated CII containing 1–82 residues) pE-lacZ & lacP-cIIA T7P-cII

– – His6-CII His6-HflD – His6-HflDW181A(mHflD) – CII (tagless) – GST-HflD His6-CIIA His6-CIIA His6-CII

This study This study This study This study This study This study This study Datta et al. [30] Jain et al. [29] Kihara et al. [26] This study This study Halder et al. [24]

Primers (restriction sites are underlined, mutation sites are in bold) PDFB: 50 -GCAGGGATCCGTGGCAAAGAATTAC-30 (BamHI) RD: 50 -CGCCCTCGAGTCACAACTCCGGGG-30 (XhoI) PDF: 50 -CGGCATATGGTGGCAAAG-30 (NdeI) PFW181A: 50 -GCCGCCGTGCTCGCGCACCAGGTCGGC-30 (Mutation forward primer) PRW181A: 50 -GCCGACCTGGTGCGCGAGCACGGCGGC-30 (Mutation reverse primer) LacF: 50 -GCGGAGATCTGCGCAACGCAATTAATG-30 (BglII) LacR: 50 -GTGGTCTAGAAGCTGTTTCCTGTGTG-30 (XbaI) PEF: 50 -GCGGTCTAGAGGGCATCAAATTAAACC-30 (XbaI) PER: 50 -TATTTCTAGACGCGCCAATCGAGCCATG-30 (XbaI) CIIF: 50 -CAGGCCGTCATATGGTTCGTGCAAAC-30 (NdeI) CIIR: 50 -GCTAACGGGATCCTCAGAACTCCATC-30 (BamHI) CIIAR: 50 -GGCGGGATCCTTTTTTATTGGTGAGAATCGC-30 (BamHI)

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pKP804, which was used for overexpression of His6-HflDW181A, henceforth called mHflD. Overexpression and purification of proteins Escherichia coli BL21 (DE3) (Invitrogen) cells carrying pKP509 or pKP804 were grown in a 750 ml fresh Luria–Bertani medium in presence of kanamycin (50 lg/ml) at 37 °C till A590 of the culture was between 0.4 and 0.5. The culture was then induced by 300 lM isopropyl-b-D-thiogalactopyranoside (IPTG) at 16 °C for 20 h. Bacterial cells were recovered by centrifugation at 5000g for 7 min at 4 °C. The cell pellet was suspended in lysis buffer (20 mM Tris–HCl, 500 mM NaCl, 5 mM MgCl2, 10% glycerol, 10 mM imidazole, 0.5% nonidet P-40, 10 mM b-mercaptoethanol, 1 mM phenylmethylsulfonyl fluoride and 200 lg/ml lysozyme, pH 8.0). The cells were then lysed by sonication followed by centrifugation at 17,400g for 20 min at 4 °C. The supernatant was loaded onto a Ni2+–NTA column (pre-equilibrated with the lysis buffer). After loading, the column was consecutively washed with wash buffers I (20 mM Tris–HCl, 500 mM NaCl, 5% glycerol and 25 mM imidazole, pH 8.0) and II (100 mM NaP, 100 mM NaCl, 5% glycerol, pH 8.0). Elution of the protein (His6-HflD or mHflD) was accomplished with the elution buffer (100 mM NaP, 100 mM NaCl, 5% glycerol and 250 mM imidazole, pH 8.0). Native CII (without a histidine tag) was obtained by overexpressing CII from the recombinant plasmid pAB305 in BL21 (DE3) cells and purification by two steps of ion-exchange chromatography as described earlier [30]. GST-HflD, His6-CII and its truncated variant His6-CIIA required for in vitro interaction studies were overexpressed and purified by the following method. BL21 (DE3) cells carrying pKH449 were induced by 500 lM IPTG at A600  0.5–0.6 for 4 h at 37 °C. Cells were harvested by centrifugation at 5000g at 4 °C for 7 min and suspended in the lysis buffer (1 phosphate buffered saline, 10% glycerol, 10 mM b-mercaptoethanol, 1 mM phenylmethylsulfonyl fluoride, 0.5% nonidet P-40 and 200 lg/ml lysozyme, pH 7.4). Cells were lysed by sonication followed by centrifugation at 17,400g for 20 min at 4 °C to collect the clear supernatant, which was applied to a Glutathione SepharoseTM 4B column (GE Healthcare Bio-Sciences AB, Sweden) pre-equilibrated with the lysis buffer. The column was washed with 10 times the bead volume of wash buffer (1 phosphate buffered saline, 10% glycerol, 0.5% nonidet P-40, pH 7.4). GST-HflD was eluted from the column with elution buffer (100 mM NaP, 100 mM NaCl, 5% glycerol, 10 mM reduced glutathione, pH 7.0). His6-CII was overexpressed within E. coli BL21 (DE3) cells from pAB412 and purified as described in Parua et al. [31b]. His6-CIIA was overexpressed and purified in an identical fashion, from pKP108 in BL21 (DE3) cells. Size-exclusion chromatography Size-exclusion chromatography was carried out to identify the oligomeric state of HflD. Here, 1–2 mg/ml of the recombinant His6-HflD was applied onto a SuperdexÒ75 column. Prior to sample loading, the column was pre-equilibrated with buffer D (100 mM NaP, 100 mM NaCl, 5% glycerol, 1 mM EDTA, pH 8.0). The flow rate was maintained at 0.5 ml/min. Glutaraldehyde crosslinking Glutaraldehyde crosslinking was used to further assess the oligomeric state of HflD as well as the stoichiometry of interaction between CII and HflD, as described in [23]. 0.1% of glutaraldehyde was used to crosslink 8 lM of His6-HflD in buffer D (see above) at 4 °C for 20 min. Reactions were terminated by the addition of

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200 mM of glycine, followed by the addition of 1 SDS–PAGE gel-loading buffer and boiling. The protein samples were electrophoresed on a 13.5% SDS–PAGE followed by electroblotting onto a polyvinylidene difluoride (PVDF; Schleicher and Schuell) membrane. Proteins on the membrane were visualized by immunoblotting with mouse anti-His monoclonal IgG. For measurement of stoichiometry, 8 lM each of CII and His6HflD was incubated together for 10 min at 25 °C followed by the application of 0.1% glutaraldehyde to the reaction mixture. Crosslinking was carried out at 4 °C for the specified time interval (2, 4, 8 and 12 min) as mentioned above. Reactions were terminated and subjected to 12–15% gradient SDS–PAGE followed by electroblotting onto PVDF membranes and were visualized by immunoblotting with anti-CII polyclonal antibody (obtained from Keith Shearwin, University of Adelaide, Australia). Cysteine accessibility of HflD The number of surface-accessible cysteine residues of His6-HflD was determined following the DTNB (5,50 -dithiobis-2-nitrobenzoic acid) method mentioned in [32]. Briefly, 100 ll of DTNB solution was added to 900 ll of protein solution (10 lM) in 100 mM Naphosphate buffer (pH. 8.0) at room temperature (25 °C) with a final protein:DTNB molar ratio of 1:30. Absorbance at 412 nm was measured after the OD value reached saturation. The number of free cysteine residue(s) was determined from the changes in absorbance at 412 nm as the number of TNB (2-nitro-5-thiobenzoate) molecules released due to the reaction between accessible –SH group(s) of the protein and DTNB, using the following equation:

n ¼ ðA412 =TNB Þ=½P where n is the number of surface-accessible cysteine residues, A412 is the measured absorbance at 412 nm, [P] is the concentration of protein and eTNB is the molar extinction coefficient of TNB, for which a value of 13,650 M1cm1 [33] was used. Tryptophan fluorescence measurement The tryptophan fluorescence of CII in the presence of mHflD was measured by titrating 4 lM of CII with increasing concentrations of mHflD (up to 180 nM) in buffer P (20 mM Tris–HCl, 200 mM NaCl, 1 mM EDTA, pH 8.0), in a Hitachi F-3000 spectrofluorimeter. kex = 295 nm and kem = 343 nm were used, with bandwidths of 5 nm on both sides. In vivo b-galactosidase assays The extent of transcription activation by CII or CIIA in the presence or absence of HflD was quantified by measuring the b-galactosidase expressed from a lacZ reporter gene, according to Miller [34]. Escherichia coli AK525 cells carrying a pair of plasmids expressing b-galactosidase and either HflD, CII, CIIA or both CII/ CIIA and HflD (for details, see Table 2) were used as hosts. The cells were allowed to grow at 37 °C in 5 ml Luria–Bertani medium supplemented with 100 lg/ml ampicillin and 50 lg/ml kanamycin, followed by induction by 300 lM IPTG at A590  0.6–0.7 for 60 min and immediate transfer onto ice. The absorbance at 590 nm was recorded. 0.5 ml of these cultures were separately added to 0.9 ml of Z buffer (60 mM Na2HPO4, 40 mM NaH2PO4, 10 mM KCl, 1 mM MgSO4, 50 mM b-mercaptoethanol, pH adjusted to 7.0). The cells were lysed by the addition of 10 ll of 1% SDS, 20 ll of chloroform and vortexing for 10 s. The tubes containing the reaction mixtures were then placed in a 28 °C water bath for 5 min. Reactions were initiated by the addition of 0.2 ml of ortho 2-nitrophenyl-b-D-galcatopyranoside (ONPG) (4 mg/ml, dissolved in Z buffer) at 28 °C. Reactions were stopped by the addition of

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Miller Units ¼ 1000  ðA420  1:75  A550 Þ=ðt  V  A590 Þ

Tris–HCl, 100 mM NaCl and 1 mM EDTA, pH 8.0. The oligonucleotide mixture was then heated on a boiling water bath for 15 min and allowed to cool slowly to room temperature. The concentration of the double-stranded DNA was estimated from its optical density at 260 nm. This DNA was used at a final concentration of 10 nM in a total volume of 2.5 ml. The titration of DNA by CII in presence of His6-HflD was performed in the assay buffer containing 20 mM Tris–HCl, 100 mM potassium glutamate, 10% glycerol and 1 mM EDTA, pH 8.0. The experiment was carried out in two ways: (a) 150 nM of His6-HflD was mixed with the DNA which was titrated with increasing concentrations of CII, or (b) CII and His6HflD were pre-incubated at equimolar concentrations (up to 900 nM) of each for 1 h at 4 °C and was then added to DNA. Measurements were carried out in a Hitachi F-4500 spectrofluorimeter with excitation and emission wavelengths set at 494 and 520 nm, respectively. Bandwidth on each side was 5 nm. Anisotropy was measured by scanning the sample for 2 min at each titration point and recording the averaged value of the anisotropy.

where t is the time of reaction (in min) and V is the volume (in ml) of the culture used in the assay [34] .

In vivo stability of CII in presence of HflD

Table 2 Effects of HflD on in vivo transcription activation by CII. E. coli strain

plasmids carried

AK525

pKP109 pKP109 pKP219 pKP219 pKP419 pKP419

& & & & & &

pKP908 pKP917 pKP908 pKP917 pKP908 pKP917

(His)6-protein expressed

b-Galactosidase activity (Miller Units)a

– HflD CII CII & HflD CIIA CIIA & HflD

0.2 (±0.3) 0.4 (±0.4) 47.9 (±6.5) 6.1 (±3.2) 52.8 (±7.8) 10.2 (±4.3)

a All values shown are the average of three independent experiments. The respective standard deviations are indicated within brackets.

0.3 ml of 1 M Na2CO3 after sufficient yellow colour had developed. The absorbances at 420 and 550 nm were then recorded. The unit of b-galactosidase activity was calculated from

In vitro transcription assays The effect of His6-HflD on CII-mediated transcription from pE was quantitated by in vitro transcription assays as described in [17]. Reactions were performed in a 20-ll reaction volume in transcription buffer (40 mM Tris–HCl, 0.1 M potassium glutamate, 1 mM DTT,1 20 mM MgCl2; pH 8.0) using the 540 bp DNA fragment [17] as the template, which contained the promoters pE (CII-dependent, would generate a 144-base run-off transcript) and poop (CIIindependent, would give a 77-base terminated transcript). The transcript from the latter promoter was used as an internal positive control. In all reactions, DNA and E. coli RNA polymerase were used at 5 and 60 nM, respectively. The experiment was carried out in two ways. In the first, 250 nM of CII and a specific concentration of His6-HflD (0, 50, 100, 250 or 500 nM) were incubated at 37 °C for 10 min before RNA polymerase and template DNA were added. This was followed by further incubation at 37 °C for 10 min. Transcription was initiated by the addition of nucleotide mix (0.1 mM each of ATP, GTP, CTP; 0.01 mM of UTP and 5 lCi of [a-32P] UTP and 1 lg of heparin). The reactions were terminated after 20 min by the addition of 5 ll of formamide stop dye. In the second, CII was mixed at a concentration of 250 nM with RNA polymerase and template DNA, and reactions were incubated at 37 °C for 20 min followed by initiation of transcription by the addition of NTP mix. After 4 min, His6HflD was added at concentrations of 0, 50, 100, 250 and 500 nM and the reactions were further incubated at 37 °C for an additional 16 min. Finally, reactions were terminated by the addition of the stop dye. Transcripts were resolved by electrophoresis in a 10% (w/ v) polyacrylamide–7 M urea gel, which was dried and autoradiographed at 20 °C. DNA binding studies DNA binding studies for CII in the presence of His6-HflD were carried out by fluorescence anisotropy measurements [35]. In this study, a 20 bp oligonucleotide containing the recognition sequence for CII (pE promoter) was used. The sense (50 -fluorescein-TCGT TGCGTTTGTTTGCACG-30 ) and anti-sense (50 -CGTGCAAACAAACGC AACGA-30 ) strands were purchased from BioTechdeskTM as HPLCpurified DNA. The strands were mixed with each other to a final concentration of 100 lM of each in a buffer containing 20 mM 1 Abbreviations used: DTT, dithiothreitol; HTH, helix-turn-helix; CRP, cyclic AMP receptor protein.

In vivo stability of His6-CII in the presence of His6-HflD was carried out following the protocol of [36]. Here, E. coli XL1 Blue or AK525 cells carrying the pair of plasmids pKP219 and pKP908 or pKP219 and pKP917 was used as the host. Cells were grown till A590 = 0.6–0.7 in Luria–Bertani medium supplemented with ampicillin (100 lg/ml) and kanamycin (50 lg/ml) at 37 °C, followed by induction with 300 lM of IPTG for 60 min and treatment with spectinomycin (100 lg/ml) to inhibit protein synthesis. 500 ll aliquots of cell culture were collected at different time intervals after the addition of spectinomycin. Cells were harvested by centrifugation at 6600g at 4 °C for 8 min and resuspended in 1 gel-loading buffer. Samples were then electrophoresed on a 15% SDS–polyacrylamide gel followed by electroblotting using Mini Trans-BlotÒ Cell (Bio-Rad, USA) onto a PVDF membrane. Proteins were visualized by immunoblotting with rabbit polyclonal anti-CII antibody. The blot pattern was subjected to densitometric analyses using the volume analysis routine of the Molecular Analyst software (Bio-Rad, USA). Possible HflD–CIIA interactions in vitro GST pull-down assays were carried out to check possible interactions between HflD and CIIA, according to Halder et al. [24]. Fifty micrograms of purified GST-HflD was immobilized on Glutathione SepharoseTM 4B beads at 4 °C for 30 min in buffer B (50 mm NaP, 100 mM NaCl and 5% Glycerol, pH 7.0). To the same beads, 50 lg of purified His6-CII or His6-CIIA was added and incubated for 30 min at 4 °C with mixing by continuous rotation. The beads were then washed thrice with buffer B. Bound GST-HflD was eluted by the elution buffer (buffer B plus 10 mM of reduced glutathione). Each fraction (loading, each wash and elution) was analysed by 15% SDS–PAGE. Proteins were then electroblotted onto a PVDF membrane and visualized by immunoblotting using polyclonal anti-CII IgG. Modelling and computation To get an idea about the possible interacting region(s) of CII and HflD, molecular docking between HflD and CII was carried out using the GRAMM docking program in a low resolution matching mode [37–39]. The molecular structures of HflD (1SDI) and CII (1XWR) from the Protein Data Bank were used. The parameters of docking are shown in Supplementary Table S1. The best model structures having the minimum energy were used to obtain

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information on the binding interaction between HflD and CII, and on intermolecular hydrogen bond formation. Results Accessibility of cysteine residues of HflD Upon incubation of HflD with DTNB, an increase in A412 was observed, which reached a saturation value of 0.14 at 45 min. This value was used to calculate the number of accessible cysteine(s) in HflD. It was found that HflD had one (0.95) surface-accessible cysteine residue. From the three-dimensional crystal structure of HflD it was observed that of the two cysteines in HflD, Cys31 was surface accessible, while Cys16 was buried. Oligomeric state of HflD The oligomeric state of HflD was examined both by glutaraldehyde crosslinking and size-exclusion chromatography. The antiHis antibody immunoblotting profile of His6-HflD crosslinked with glutaraldehyde (Fig. 1a) showed that in the presence of glutaraldehyde, a weaker band corresponding to the molecular weight of the His6-HflD dimer (48 kDa) appeared, in addition to the monomer band observed in the absence of the crosslinker. The size-exclusion chromatogram of His6-HflD (Fig. 1b) clearly showed that recombinant HflD existed mostly as a monomer with a small amount of dimer. Fig. 1c shows a plot of the Rf value against the logarithm of molecular weight obtained from gel-filtration. In this plot, the positions of the His6-HflD dimer (46.70 kDa) and monomer (24.20 kDa) are shown. Interaction between CII and HflD: effects on the conformations of CII and HflD It is known that HflD interacts with CII both in vivo and in vitro [26]. Whether this interaction causes any structural change in either of the proteins is not known. The possible structural change(s) in HflD and in CII were examined by various means, results of which are presented below. Effect of CII on cysteine accessibility of HflD HflD has two cysteines (Cys16 and Cys31), of which only one (most likely, C31) is available to react with DTNB. Does the binding of CII change the accessibility of this cysteine? DTNB titration of His6-HflD in the presence of CII was carried out to examine this. It may be noted that CII has no cysteine residue; therefore DTNB would react only with the accessible cys of HflD. In the presence of different concentrations of CII (up to 20 lM), His6-HflD (5 lM) was treated with DTNB and the number of accessible cysteines was determined for each concentration of CII present. The final concentration of DTNB was 150 lM. These results are shown in Fig. 2. It was found that with increasing concentrations of CII, the number of accessible cys in HflD decreased drastically from 1 to 0.2. Is this reduction due to a molecular crowding effect by CII as its concentration was increased? To exclude this possibility, the accessibility of cys in HflD was also measured in the presence of PEG 6000, a straight chain polymer of a simple repeating unit H(OCH2CH2)nOH that induces macromolecular crowding of solutes in aqueous solution [40]. Increasing concentrations of PEG 6000 did not affect the cysteine accessibility in HflD (Fig. 2). To examine if the effect of CII on HflD was due to a specific interaction between the proteins, E. coli CRP, a protein that is not expected to interact with CII, was chosen. The accessibility of the cysteine residues of CRP, a homodimer having two accessible cysteines per dimer [32], was measured in the presence of the same concentrations

Fig. 1. Oligomeric state of HflD. (a) Glutaraldehyde crosslinking. 13.5% SDS–PAGE showing HflD alone (lane 1) or crosslinked for 5 or 10 min (lanes 2 and 3). Anti-his antibody was used for immunoblotting. Lane 4 shows molecular weight markers (97, 66, 43, 30 and 20 kDa). Arrows indicate dimeric (Di) and monomeric (Mono) HflD. (b) Elution profile of His6-HflD. HflD peaks corresponding to dimeric (A) and monomeric (B) His6-HflD are indicated. A SuperdexÒ75 column was used. (c) A plot of Rf against the logarithm of molecular weight for the four standard proteins (Albumin, 67 kDa; Ovalbumin, 43 kDa; Chymotrypsinogen A, 25 kDa; Ribonuclease A, 13.7 kDa) obtained by size-exclusion chromatography. The linear fit is shown along with its equation. The positions of dimeric (s) and monomeric (h) His6-HflD are shown along with molecular weight standards ().

Fig. 2. Number of accessible cysteine residues in HflD. The number of modified cys residues in HflD was determined by DTNB treatment, as a function of different concentrations of kCII (h) or PEG 6000 (j). The number of accessible cysteines per monomer of E. coli CRP as a function of varying kCII concentrations is also shown (s).

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of CII as for HflD. No effect of CII on the number of accessible cys residues of CRP was observed (Fig. 2). Therefore, it is clear that the observed decrease of cysteine accessibility of HflD upon addition of CII was due to a specific interaction between the two proteins, which could be due to either of the following: (a) HflD interacted with CII through its cysteine accessible surface area or (b) as a result of the interaction with CII, structural changes occurred within HflD causing the surface exposed cysteine to be inaccessible. Quenching of tryptophan fluorescence of CII Possible structural changes within CII due to its interaction with HflD was also monitored by examining the intrinsic tryptophan fluorescence of CII in the presence of mHflD, an HflD mutant devoid of any tryptophan residue (verified by the abolition of intrinsic tryptophan fluorescence of HflD, data not shown). It was observed that the relative fluorescence intensity of CII increased with increasing concentrations of mHflD, reaching saturation at 130 nM (Fig. 3). This result indicates that binding of HflD caused a conformational change in CII, leading to an increase in the quantum yield(s) of one or more of the three tryptophan residues in CII.

Fig. 4. Stoichiometry of interaction between CII and His6-HflD from glutaraldehyde crosslinking. (a) CII (8 lM) and His6-HflD (8 lM) were crosslinked and run on a 12– 15% gradient SDS–PAGE, followed by electroblotting onto a PVDF membrane and immunoblotting by anti-CII antibody. Lane 1: CII without crosslinker; lanes 2–5: a mixture of CII and HflD, crosslinked for 2, 4, 8 and 12 min, respectively, on ice; lane 6: protein molecular weight standards (85, 48, 34, 26 and 19 kDa). The band corresponding to CII–HflD complex (70 kDa) is shown. The bands corresponding to monomer (Mono), dimer (Di), trimer (Tri) and tetramer (Tetra) of CII are indicated. (b) CII alone was subjected to crosslinking followed by immunoblotting with anti-CII-antibody. Lane 1: protein molecular weight standards (85, 48, 34, 26 kDa); lane 2: CII without crosslinker; lane 3: CII crosslinked for 10 min on ice.

Glutaraldehyde crosslinking

Effect of HflD on transcription activation by CII

To determine the stoichiometry of interaction between CII and HflD, His6-HflD and native CII (without any tag) were subjected to glutaraldehyde crosslinking (Fig. 4a). It was observed that besides the monomer and higher order oligomers of CII, another higher molecular weight protein band corresponding to a molecular weight of 70 kDa appeared. This band, which was recognized by anti-CII polyclonal antibody, did not appear when CII was crosslinked alone (Fig. 4b), indicating that it corresponded to a CII–HflD complex. The molecular weights of CII and His6-HflD are 11 and 24 kDa, respectively. Therefore, the probable molar ratio of HflD and CII in the complex could be 1:4 or 2:2. Thus, one molecule of HflD may complex with four monomers of CII giving rise to a molecular weight of 68 kDa for the complex. Alternatively, a 2:2 complex comprising two molecules of both CII and HflD may result in the higher molecular weight band (70 kDa). Since in solution CII exists predominantly as a tetramer and HflD as a monomer, the first possibility, i.e. interaction of CII tetramer with HflD monomer is more likely. This result also suggests that CII retains its native tetrameric state in the presence of HflD.

The effect of HflD on CII-mediated transcription activation from pE was examined both in vitro and in vivo. The rate of in vitro transcription activation of pE decreased with increasing concentrations of HflD, when CII was pre-incubated with His6-HflD (Fig. 5). Interestingly, the maximum inhibition (35–40%) occurred around 60– 70 nM of HflD, which was 25% of the concentration of CII (250 nM of monomers) used in the reaction. This fact is consistent with an interaction of monomeric HflD with tetrameric CII with a 1:1 stoichiometry, as indicated in the previous section. However, HflD had no effect on in vitro transcription from pE when His6-HflD was added after initiation (Fig. 5). The in vivo effect of HflD on CII-dependent transcription activation from pE was monitored by measuring the amount of b-galactosidase expressed from lacZ as a reporter gene under the control of pE, the results of which are shown in Table 2. When CII was expressed alone, the unit of b-galactosidase activity was observed to be 47.9. This activity was solely due to the action of CII on pE up-

Fig. 3. Effect of addition of mHflD on the intrinsic tryptophan fluorescence of kCII. A plot representing the change in relative fluorescence intensity [(F0  F)/F0, where F0 and F are the observed fluorescence intensity at 343 nm in the absence and presence of mHflD, respectively] as a function of the concentration of mHflD (kex = 295 nm).

Fig. 5. Effect of HflD on CII-dependent transcription activation in vitro. Amount of transcripts from pE and poop were quantified using densitometry. The ratio of the two (taken as 100% when HflD was absent) was plotted against the concentration of HflD added (up to 500 nM). Results for HflD pre-incubated with CII (250 nM) before initiation of transcription (j) or when HflD was added after 4 min of initiation of transcription by 250 nM CII (s) are shown. Each data point represents the average value from three independent experiments.

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increasing concentrations of CII, a gradual increase in anisotropy was observed (Fig. 6), indicating binding of CII to the DNA. However, the anisotropy value remained unchanged when the same experiment was repeated with an equimolar mixture of CII and His6-HflD (Fig. 6), indicating a lack of binding. Clearly, CII–HflD interaction prevented CII from binding to its cognate site on pE. When His6-HflD (150 nM) was pre-incubated with DNA prior to addition of CII, an interesting result was obtained. At lower (up to 150 nM) concentrations of CII, there was no change in anisotropy of DNA. Above 150 nM, the anisotropy increased, though the rate of increase was less compared to that for CII alone (Fig. 6). Therefore, it is likely that HflD–CII interaction is stronger than pE–CII interaction, and binding of HflD to CII inhibits the DNA binding ability of the latter. In vivo proteolysis of CII: effect of HflD Fig. 6. Binding of CII to pE from fluorescence anisotropy measurements. Fluorescence anisotropy was plotted for a 20-bp ds DNA (10 nM) containing pE and labelled by fluorescein at the 50 end of the sense strand in presence of increasing concentrations of CII alone (j) or an equimolar mixture of CII–HflD (4). Results from a similar experiment when DNA was pre-incubated with 150 nM HflD before titration with CII are also shown (s).

stream of the lacZ gene, since the activity measured in the absence of CII was nominal (0.2). Upon simultaneous overexpression of HflD with CII, the activity underwent an 87% reduction, decreasing to a value of 6.1. Thus, HflD inhibited CII-dependent transcription both in vitro and in vivo. The results of the in vitro and in vivo transcription studies reveal that HflD inhibits CII-dependent transcription activation. This may happen due to either of the following reasons: (a) upon interaction with CII, HflD may block recognition of its cognate DNA, (b) HflD– CII interaction disrupts specific CII–RNA polymerase interaction essential for transcription activation, or (c) the stimulating effect of HflD on in vivo proteolysis of CII may predominate. To distinguish among these possibilities, DNA binding and in vivo proteolysis of CII was carried out in the presence of HflD, as described below. Effect of HflD on DNA binding by CII The effect of HflD on binding of CII at pE was examined by fluorescence anisotropy measurements. When a 50 -fluorescein labelled 20 bp ds DNA containing the pE sequence was titrated with

As shown above, HflD inhibits the CII-dependent transcription activation from pE in vivo. To test whether this inhibition resulted from a stimulation of in vivo CII proteolysis by HflD, the proteolysis was carried out in the presence of His6-HflD, as shown in Fig. 7. HflD significantly accelerated (by 25%) the in vivo proteolysis of CII in wild type XL1 Blue cells (Fig. 7a), while there was no effect of HflD in mutant AK525 cells that lacked HflB (Fig. 7b). Thus, the destabilization of CII by HflD occurred only in the presence of HflB. From this result one may also conclude that the observed inhibition of CII-mediated transcription from pE by HflD (carried out in AK525 cells) was not due to the depletion of CII but due to a specific interaction between CII and HflD that resulted in a loss of DNA binding by CII. Interactions between CII and HflD by molecular modelling The three-dimensional structure of HflD (formerly known as YcfC), an all-helix protein, has been reported by D. Borek and Z. Otwinowski (PDB ID: 1QZ4 and 1SDI; unpublished). The crystal structure of CII in the free state (PDB ID: 1XWR) and as a DNAbound complex (PDB ID: 1ZS4) are also available [14,15]. We used the structure 1XWR to build a reasonable molecular model in order to unravel the possible interactions between CII and HflD by docking the two proteins using the GRAMM docking program [38,39] in a low resolution matching mode. Such a minimum-energy model of the CII–HflD complex obtained from docking is shown in

Fig. 7. In vivo proteolysis of CII. (a) Western blot (using rabbit polyclonal anti-CII antibody) of whole cell (XL1 Blue) lysates for His6-CII at various time points in the presence (s) or absence (j) of overexpressed His6-HflD after arresting the protein synthesis by spectinomycin (100 lg/ml). The bands were quantified using Molecular Analyst software (Bio-Rad, USA) and the percentage of intracellular CII remaining was plotted. (b) Similar experiments as in (a) but carried out in AK525 cells (DhflB). Each experiment was repeated four times.

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Fig. 8. Molecular models for CII–HflD interaction (PyMOL view). (a) CII–HflD complex. The four chains (A, B, C and D) of CII (cartoon) and HflD molecule (lines) are shown, along with the Cys31 of HflD. (b) CII–HflD–pE complex. When the above CII–HflD complex was docked with DNA, the HTH motifs of A and C chains (indicated by white circle) of CII were unable to interact with the TTGC repeats in DNA within successive major grooves (indicated by red arrow). Instead, only the HTH motif from the B chain of CII (indicated by white arrow) contacted DNA. In each case, minimum energy structures generated by the GRAMM docking program was used. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

Fig. 8a. The interface residues of both the proteins were identified using a web-server, ProFace [41], developed for dissecting protein– protein interfaces and deriving various physicochemical parameters. In this model, HflD was found to interact with CII through its surface containing the Cys31 residue. As a result of the interaction, Cys31 of HflD was buried. This model is consistent with the experimental results obtained in this study: (a) decrease of surface accessibility of a cysteine residue of HflD upon interaction with CII, (b) interaction of monomeric HflD with tetrameric CII and (c) retention of the tetrameric nature of CII upon interaction with HflD. From this model, potential residues that may form intermolecular hydrogen bonds were also identified (Table 3). A preliminary experiment to test the model was carried out, using a mutant in which four residues of HflD (Q28, E67, D124 and E128) were all changed to alanine. In vivo transcription by CII decreased by 10–20% in the presence of this mutant (data not shown), while for wild type HflD the decrease was 90% (see ‘‘Effect of HflD on Transcription Activation by CII” section, Table 2). In the published crystal structures of CII used for the above docking studies, 17 residues at the disordered C-termini are missing [14]. Therefore, a possibility remains that HflD may interact with CII at the C-terminal residues of the latter. Such an interaction would be missed in our molecular modelling. To explore this possibility, a C-terminal deleted version of CII (CIIA) that contains residues 1–82 was prepared. Binding of GST-HflD to this mutant was carried out using GST pull-down assays. As is evident from Fig. 9, both CII and CIIA showed comparable binding to HflD, supporting the idea that the C-terminal end of CII was not required for CII– HflD interaction. This was further vindicated by the results of in vivo transcription experiments carried out with CIIA alone or

in the presence of HflD (Table 2). The b-galactosidase activity decreased from 52.8 to 10.2 (a reduction of 80%) in the presence of HflD. However, how HflD blocks the DNA binding of CII and inhibits transcription activation at pE (both in vivo and in vitro) remains unresolved from this model. To understand this aspect, another docking experiment was carried out where the generated model structure of CII–HflD complex was docked with a 27 bp DNA containing the pE promoter sequence (coordinates of which were obtained from the crystal structure of CII–pE complex; PDB ID: 1ZS4 [15]). The result (Fig. 8b) shows that even at the best position at which the CII–HflD complex could be docked to DNA, the HTH motifs in the A and C chains of CII, which are involved in CII–DNA interactions [14,15], were unable to contact the cognate CII sites (TTGC) at the major grooves. In the CII–HflD–DNA model (Fig. 8b), CII was able to interact with DNA only through the HTH motif of chain B instead of the HTH motifs of A and C chains. Such an interaction would essentially be weak and insufficient to hold CII onto DNA tightly. Indeed, in the presence of HflD, CII–DNA binding was inhibited (Fig. 6). This would explain the observed loss in transcription activation (Fig. 5 and Table 2) in the presence of HflD. Discussion HflD was identified at the gene level, as an additional locus in E. coli, mutations to which increased the frequency of lysogenization by lambda [26]. Apart from reports that it interacts with CII and has an effect upon degradation of the latter by HflB [26], little is known about this protein. Kihara et al. [26] had suggested that

Table 3 Probable intermolecular hydrogen bond forming residues identified from the CII–HflD complex model structure (Fig. 8a). Residue of CII

Residue of HflD

Glu31 (Chain A) Arg7 (Chain B) Glu14 (Chain B) Arg45 (Chain B) Arg69 (Chain C) Arg12 (Chain D) Gln73 (Chain D)

Arg78 Thr152 Gln28 Glu67 Gln28 Asp124 Glu128

Fig. 9. In vitro interactions between HflD–CII and HflD–CIIA by GST pull-down assays. GST-HflD was immobilized on Glutathione Sepharose beads and purified His6-CII or His6-CIIA was applied and incubated to allow binding. After washing, GST-HflD was eluted from the beads and all fractions were run on 15% SDS–PAGE and visualized by immunoblotting with polyclonal anti-CII antibody. L, W1, W2, W3 and E indicate loading, wash 1, wash 2, wash 3 and elution fractions, respectively.

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HflD may exist as dimers or higher oligomers. In this work, however, we found that HflD existed predominantly as a monomer, though a small amount of dimer was also present under our experimental conditions. It was proposed that HflD sequestered CII from its target promoters (pE, pI and paQ) and recruited it to the proximity of HflB for rapid degradation [26]. Our results show that monomeric HflD interacted with tetrameric CII which retained its tetrameric nature after the interaction. The accessibility of the accessible cysteine residue of HflD (likely to be Cys31, as seen in the crystal structure of HflD) decreased upon interaction with CII, indicating that either HflD interacts with CII through its cysteine accessible surface or as a result of the interaction, a structural change is occurring which causes Cys31 to be buried. From the model structure of the HflD–CII complex, it was found that HflD interacted with CII through its Cys31-accessible surface area, explaining the decrease in the accessibility of Cys31 of HflD upon interaction with CII. This study also shows that HflD inhibited CII-dependent transcription from pE. The down regulation of the lysogenization frequency of lambda by HflD could be a result of this inhibition, rather than by the stimulation of the in vivo proteolysis of CII [26]. Both the effects may work together. We also found that HflD was unable to inhibit CII-dependent transcription once CII was bound to DNA. This result contradicts the sequestering activity of HflD proposed by Kihara et al. [26]. Further, our results suggest that HflD blocked recognition of DNA by CII due to a higher affinity of CII towards HflD than towards its cognate DNA site. A structural change in CII upon its interaction with HflD, as indicated from fluorescence quenching (see ‘‘Quenching of Tryptophan Fluorescence of CII” section), could be responsible for the loss of DNA binding by CII in the presence of HflD. As our docking results show, a CII–HflD complex would make an effective CII–DNA interaction extremely weak. This might be the possible reason for the inhibitory role of HflD on CII-mediated transcription activation, and on its effect on lambda lysogeny.

Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at doi:10.1016/j.abb.2009.10.010. References [1] [2] [3] [4] [5] [6] [7] [8] [9] [10] [11] [12] [13] [14] [15] [16] [17] [18]

[19] [20] [21] [22] [23] [24] [25] [26] [27]

Conclusions We find that HflD, a peripheral membrane protein, acts as a transcription inhibitor. This is an unusual feature for an inhibitor protein that acts through a direct interaction with an activator protein (in this case, kCII). All proteins that have an effect on the lysogenization of k (such as kCIII, HflB, HflKC and HflD) are believed to act through their effects on the stability of CII. Our results show that in addition, HflD binds to CII and prevents the binding of the latter to its cognate DNA site(s), leading to an inhibition of CII-dependent transcription. This role of HflD makes it a unique component in the k lysis–lysogeny switch. Acknowledgments This work was funded by Institutional Project 5 (Microbial Genomics) of Bose Institute. Shrihari Sonavane (Bose Institute) helped in carrying out the molecular docking experiments. Pabitra Parua was supported by a fellowship from CSIR, India.

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[28] [29] [30] [31]

[32] [33] [34] [35] [36] [37] [38] [39] [40] [41] [42]

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