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Specific hydrophobic residues in the α4 helix of λCII are crucial for maintaining its tetrameric structure and directing the lysogenic choice

Pabitra K. Parua, Ajit B. Datta, Pradeep Parrack

  • Department of Biochemistry, Bose Institute, P-1/12, C.I.T. Scheme VIIM, Kolkata 700 054, India
  • Correspondence
    Pradeep Parrack
    pradeep{at}bic.boseinst.ernet.in

    • Journal of General Virology 2010; 91(1):306–312 · https://doi.org/10.1099/vir.0.015040-0

    View at publisher PubMed

    Abstract

    The CII protein of the temperate bacteriophage λ is the decision-making factor that determines the viral lytic/lysogenic choice. It is a homotetrameric transcription activator that recognizes and binds specific direct repeat sequences TTGCN6TTGC in the λ genome. The quaternary structure of CII is held by a four-helix bundle. It is known that the tetrameric organization of CII is necessary for its activity, but the molecular mechanism behind this requirement is not known. By specific site-directed mutagenesis of hydrophobic residues in the α4 helix of CII that constitutes the four-helix bundle, we found that residues leu70, val74 and leu78 were crucial for maintaining the tetrameric structure of the protein. When any of these residues was substituted by a polar one, CII lost its activity and failed to promote lysogeny. This loss of activity was accompanied by the inability of CII to form tetramers, to bind DNA or to activate transcription.

    • ↵†Present address: Department of Biophysics and Biophysical Chemistry, Johns Hopkins University School of Medicine, Baltimore, MD 21205, USA.

    • Three supplementary figures and two supplementary tables are available with the online version of this paper.

    INTRODUCTION

    The temperate bacteriophage λ can switch between its different developmental stages, lytic or lysogenic, in response to various environmental signals, including those that influence the growth of its bacterial host, Escherichia coli (Wegrzyn & Wegrzyn, 2005; Court et al., 2007). Lambda responds to diverse growth conditions through the phage protein CII, whose concentration is the deciding factor for its developmental choice (Hoyt et al., 1982; Rattray et al., 1984; Banuett et al., 1986; Echols, 1986). Elevated levels of CII promote lysogeny, while depletion of CII owing to its rapid degradation by the host protease HflB (also known as FtsH) drives λ toward the lytic pathway (Shotland et al., 1997, 2000).

    CII is a transcription activator that acts upon the phage promoters pE, pI and paQ, leading to products that help establish the lysogenic pathway (Kaiser, 1957; Simatake & Rosenberg, 1981; Ho et al., 1983; Ho & Rosenberg, 1985; Friedman, 1992). A homotetramer of 97-residue subunits (Ho et al., 1982), CII is an all-helix protein with each monomer consisting of four alpha helices α1–α4 (Datta et al., 2005a; Jain et al., 2005). The α4 helix (residues 64–77) from each of the four monomers in tetrameric CII contributes to the four-helix bundle structure (Datta et al., 2005a), while about 15 residues from the C-terminal end are disordered (Datta et al., 2003, 2005b). This disordered C-terminal tail is the target for the proteolysis of CII by HflB and makes CII unstable, a property essential for its function (Herman et al., 1998; Kobiler et al., 2002). Tetrameric organization of CII is also essential for its activity. It has been reported that various point mutants of CII that exhibit cII− phenotypes fail to form tetramers (Ho et al., 1988). Moreover, CIIB, a truncated form of CII (residues 4–69) lacking a major portion of the α4 region does not form tetramers, is inactive and is unable to bind operator DNA (Datta et al., 2005b). Therefore, the α4 helix is absolutely required for maintaining the 3D structure of CII and for its effective functioning.

    Consequently, CII exists predominantly as a tetramer (Ho et al., 1982). The 3D structure of CII suggests that it is roughly a ‘dimer of dimers’ (Datta et al., 2005a; Jain et al., 2005). While at low concentrations monomeric CII has been reported (Ho et al., 1982), CII is not found to exist as dimers. Interestingly, the C-terminal truncated protein CIIB that lacks the α4 helix does not tetramerize and instead, forms dimers (Datta et al., 2005b). Nevertheless, the specific residues responsible for the tetramerization of CII are not known, though there are some clues from its 3D structure (Datta et al., 2005a; Jain et al., 2005). A helical wheel plot of the residues forming the α4 helix of CII is shown in Fig. 1⇓. As indicated, residues A71, L78, M67, V74 and L70 form the core of the four-helix bundle that holds the tetrameric structure of CII. Among these, L70, V74 and L78 from each of the four CII subunits line up to form zipper-like structures between different subunits (Supplementary Fig. S1, available in JGV Online). We therefore attempted to disrupt the hydrophobic zipper-like core of tetrameric CII by specifically substituting each of these three residues by acidic residues, alone or together. The resulting mutant proteins were expressed, purified and studied for their ability to (i) promote lysogeny, (ii) activate transcription from pE, (iii) bind DNA and (iv) form tetramers.

    Figure image not available in archive
    Fig. 1.

    Helical wheel representation for the α4 helix (residues 64–78) of λCII. Hydrophobic residues are indicated by grey circles and polar residues are indicated by open circles. Residues in the core region of the four-helix bundle that holds the tetramer are marked by asterisks.

    RESULTS

    Complementation by CII mutants

    The CII defective mutant phage λcII68 (Kaiser, 1957) was used to check the functional activity of intracellular CII (wild-type or any of the six mutants) expressed from a plasmid. Upon infection with λcII68, a turbid plaque is observed only if the CII protein expressed from the plasmid is functional. Each of the mutants produced clear plaques, unlike native CII (Table 1⇓). Thus, none of them could complement a cII− phenotype, indicating that for each, the mutation had abolished its activity.

    Table 1.

    Effect of CII and its mutants on λ plaque morphology

    –, No plasmid/protein. E. coli strain BL21 (DE3) was used.

    Oligomeric status of the CII mutants

    Native CII exists as a homotetramer, with each subunit having a molecular mass of 11 kDa (Ho et al., 1982). The present mutations were aimed at disrupting this tetrameric organization of the protein. Glutaraldehyde cross-linking was carried out to check the oligomeric nature of each of the mutant proteins (Fig. 2a⇓). Upon immunoblotting the proteins with anti-CII polyclonal antibody after 15 % SDS-PAGE, dimers, trimers and tetramers were observed for native CII, as the individual subunits of CII were cross-linked in various combinations. Conversely, none of the mutants tested showed a cross-linked species larger than a dimer, indicating that all the mutants existed mostly as dimers. The oligomeric status of the proteins was also tested by analytical gel filtration chromatography (Fig. 2b⇓). While CII eluted as an approximately 53 kDa species (corresponding to a tetramer), the mutants corresponded to molecular masses of ∼27–30 kDa, indicating that they moved as dimers. Thus, it is clear that the three double mutants (CII7074, CII7078 and CII7478) as well as the single mutants (CII70, CII74 and CII78) existed as dimers, and did not form tetrameric assemblies, unlike native CII.

    Figure image not available in archive
    Fig. 2.

    Oligomeric status of native and mutant CII. (a) Western blots of the glutaraldehyde-cross-linked proteins. Lanes: 1, CII; 2, CII70; 3, CII78; 4, CII7074; 5, CII7078; 6, CII7478; 7, protein molecular mass markers (85, 48, 34, 26 and 19 kDa; Sigma); 8, CII74. Bands corresponding to the monomers (Mono), dimers (Di), trimers (Tri) and tetramers (Tetra) of CII are indicated. (b) Size exclusion chromatography. A plot of Rf against log molecular weight (MW) for the four standard proteins (albumin, 67 kDa; ovalbumin, 43 kDa; chymotrypsinogen A, 25 kDa; ribonucelase A, 13.7 kDa). The linear fit is shown along with its equation. The positions of the protein molecular mass standards (○), native CII (•), CII70 (▴), CII78 (▪), CII7074 (□), CII7078 (◊) and CII7478 (△) are shown.

    Circular dichroism (CD) spectroscopy

    It is apparent from the CD spectra of native CII and its mutants (Fig. 3⇓) that the mutant proteins had a higher molar ellipticity (θ), indicating a higher α-helical content in the mutants (68–69 %) compared with that in the native protein (61 %), as calculated from the individual CD spectra using the neural network algorithm-based CDNN program (Bohm et al., 1992). The general nature of all the spectra were similar, suggesting that the mutants maintained their secondary structure despite having lost their tetrameric organization.

    Figure image not available in archive
    Fig. 3.

    CD spectra of CII and its mutants in the far-UV region; 6 μM of each protein was used. The curve shows the plot of molar ellipticity (θ) against wavelength (nm) for native CII (•), CII70 (□) and CII7478 (△).

    Transcription activation of pE by CII mutants

    The α4 mutations caused CII to be dimeric. In vitro and in vivo transcription assays were carried out to determine whether these mutants still have the ability to activate transcription from the pE promoter. A 144 base run-off transcript corresponding to transcription from pE was produced only in the case of native CII (Fig. 4⇓), while the CII independent transcript poop was produced in all cases. Even 1 μM of any of the mutants could not activate transcription, though a much lower concentration of native CII (15 nM) is sufficient for activation (Datta et al., 2005b).

    Figure image not available in archive
    Fig. 4.

    In vitro transcription from pE in the presence of CII and its mutants. Transcripts in the presence of native CII (lane 1) or the mutants CII70 (2), CII78 (3), CII7074 (4), CII7078 (5), CII7478 (6) and CII74 (7) are shown. The expected 144 base run-off transcript from pE and the 77 base terminated transcript from poop (used as an internal control) are indicated by arrows.

    In vivo transcription experiments were carried out by measuring the β-galactosidase activity (Table 2⇓). Unlike native CII, none of the mutants tested (CII70, CII74, CII78, CII7074, CII7078 or CII7478) showed much activity.

    Table 2.

    In vivo transcription activation by native CII and mutants

    E. coli strain BL21 (DE3) was used.

    DNA binding

    CII binds and activates transcription at the pE promoter as a tetramer (Datta et al., 2005b). It has been reported that the tryptic fragment CIIB (4–69 residues; 7.5 kDa) is dimeric and is unable to bind or activate transcription at pE (Datta et al., 2005b). In the present study, we found that even the full-length α4 mutants of CII existed as dimers and were unable to activate transcription from pE either in vivo or in vitro. This loss of activation could be due to a loss in DNA binding by the mutant or at a subsequent step. DNA binding properties of the mutants were examined by fluorescence anisotropy measurements (Fig. 5⇓) using a fluorescently labelled DNA fragment containing the pE sequence and tagless proteins. For native CII, the anisotropy increased with increasing concentrations, reaching saturation at around 400 nM CII, indicating that CII could bind pE under the experimental conditions. However, no such increment in the fluorescence anisotropy was observed for any of the CII mutants tested, implying that they failed to bind to the cognate CII binding site pE.

    Figure image not available in archive
    Fig. 5.

    Binding of CII and its mutants to DNA. 5′-Fluorescein-labelled DNA (10 nM) containing the pE promoter sequence was titrated with increasing concentrations of native CII (▪), CII7074 (△), CII7078 (○), CII7478 (□) or CII78 (▽). Fluorescence anisotropy, A, is plotted against the concentration of the proteins.

    DISCUSSION

    Most prokaryotic transcription factors bind to their cognate DNA sequences via a helix–turn–helix (HTH) motif (Pabo & Sauer, 1984; Brennan & Matthews, 1989). This binding causes dimeric proteins to recognize inverted repeat sequences in different strands of DNA. The λCII protein represents a different class of transcription activators that recognize direct repeat sequences, and is the first, perhaps the sole, representative of this class of proteins. In each of the promoters it activates, CII binds the sequence TTGCN6TTGC (Ho et al., 1983). Such a direct repeat binding necessitates that CII binds to sites on the same strand of DNA, a fact borne out by the crystal structure of the CII–DNA complex (Jain et al., 2005).

    The repeating TTGC tetrad of CII flanks the −35 sequence of the promoters it activates (Ho et al., 1983), making CII a class II activator (Hawley & McClure, 1983). In most promoters that depend on class II activators, the σ factor of RNA polymerase (RNAP) and the activator protein bind to the same face of the DNA helix, while the C-terminal domain of the α subunit of RNAP (αCTD) is displaced upstream of the bound activator (Kedzierska et al., 2004). CII is different from other activators in this respect; the αCTD, bound near −41, enables interaction between its 261 determinant with the σ subunit of RNAP while CII is bound to the opposite face of DNA (Ho et al., 1983; Kedzierska et al., 2004). Activation of pE by CII possibly depends on both αCTD–CII and σ–CII contacts (Marr et al., 2004; Jain et al., 2005).

    So why is CII a tetramer? Other tetrameric proteins that bind DNA (such as the lac repressor) bind to two different sites with each dimer binding to one inverted repeat sequence through its HTH motif (Lewis et al., 1996; Brenowitz et al., 1991). Tetrameric CII, on the other hand, binds to DNA using only two of its four HTH motifs (Datta et al., 2005a; Jain et al., 2005). Formation of a tetrameric structure (by a four-helix bundle involving the α4 helix from each subunit) is probably required to maintain the correct conformation of CII so that it may bind to the two TTGC sequences that are 10 bp apart on the same DNA strand (Ho et al., 1983). Indeed, a slight distortion of DNA is required for proper positioning of the two HTH motifs from the A and C subunits of CII for optimum binding to the two TTGC sites (Datta et al., 2005a; Jain et al., 2005). The helical axis of DNA interacting with CII bends by about 2 °, which may facilitate specific binding of RNAP to DNA (Jain et al., 2005). Truncated CII lacking the α4 helix (CIIB) is dimeric and cannot activate transcription, since it fails to bind DNA (Datta et al., 2005b). It is also known that the tetrameric organization of native CII is essential for its activity (Ho et al., 1988).

    Here, we have presented evidence that critical residues in the α4 helix of CII that constitute the four-helix bundle are necessary for its structure and function. These residues are conserved in several viral transcription activators, including putative ones (Supplementary Fig. S2, available in JGV Online). Among these, P22 C1, a close analogue of λCII that shares ∼50 % sequence homology with the latter, also exists as a homotetramer (Ho et al., 1992). Even point mutations at these positions disrupt the tetrameric organization of CII, which can only form dimers, through large dimeric interfaces (Datta et al., 2005a). As a result, CII is rendered inactive, and cannot support lysogeny. The secondary structure of the protein appears to be generally undisturbed by these mutations, apart from a slight increase in the helical content that may arise from an altered packing of the subunits into dimeric assemblies. We have also ascertained that the inability to form tetramers by CII hinders its ability to bind to DNA, which is apparently the reason behind the protein losing its functional attributes. This would explain why tetramerization-defective mutants of CII were non-functional (Ho et al., 1988). Thus, specific hydrophobic residues at positions 70, 74 and 78 are crucial for maintaining the 3D and quaternary structure of CII, and are essential for cognate DNA binding and transcription activation by the protein.

    METHODS

    Bacterial strains and plasmids.

    E. coli XL1 Blue strain was used for cloning and harvesting plasmids and E. coli BL21 (DE3) strain was used for expression of proteins and other in vivo experiments. Details of the plasmids and primers (MWG Biotech) used are given in Supplementary Tables S1 and S2 (available in JGV Online). λcII68 was a gift from S. Adhya (NCI, NIH, Bethesda)

    Mutation and cloning of cII.

    Mutations within the cII gene were carried out to replace L70 with Glu, V74 with Asp and L78 with Asp separately or in different combinations. The substituted amino acid was chosen so that a polar residue replaced a hydrophobic one, while the helical structure of the α4 helix was retained. Site-directed mutagenesis was carried out by PCR using the overlapping extension method (Higuchi et al., 1988). For each set of mutations, the pAB305 plasmid vector (Datta et al., 2001) was used as the template and a pair of mutation primers (complementary to each other and containing the desired mutation), as well as reverse and forward end primers (complementary to the terminal sequence of the cII gene) were used. In the first step, two separate PCRs were done: one involving the forward end primer CIIF and a reverse mutation primer (e.g. P70R) and the other involving the reverse end primer CIIR and the corresponding forward mutation primer complementary to the former (i.e. P70F). The two PCR products thus obtained were mixed (in a 1 : 1 molar ratio) and used as the template in the second PCR, where overlap of the correct templates and amplification with the forward and reverse primers (CIIF and CIIR) ultimately produced the full-length cII gene containing the mutation(s) at the desired site(s). These mutated cII genes were cloned into pET28a vector at the NdeI and BamHI sites. Primer sequences are given in Supplementary Table S2.

    Expression and purification of proteins.

    His6-tagged native and mutant CII were overexpressed and purified by using the following method. E. coli BL21(DE3) cells harbouring the plasmids carrying native or mutated cII genes were grown in 500 ml Luria–Bertani (LB) medium supplemented with ampicillin (100 μg ml−1) or kanamycin (50 μg ml−1) at 37 °C. The cultures were induced with 500 μM IPTG at A590 ∼0.6–0.7, at 37 °C for 3 h. The cells were collected by centrifugation at 4 °C at 7000 r.p.m. (Sorvall RC5B+, SA600) for 7 min. After washing with 0.9 % NaCl, cells were lysed by sonication in lysis buffer I [20 mM Tris/HCl, 0.5 M NaCl, 5 mM MgCl2, 10 mM imidazole, 10 % (v/v) glycerol, 0.5 mM PMSF, pH 8.0]. The supernatant was collected by centrifugation at 4 °C for 20 min at 12 000 r.p.m. and loaded on to a Ni2+–NTA affinity column previously equilibrated in the same buffer, followed by washing with buffer II (20 mM Tris/HCl, 0.5 M NaCl, 50 mM imidazole, pH 8.0) and elution in buffer III (20 mM Tris/HCl, 0.3 M NaCl, 500 mM imidazole, pH 8.0). Eluted samples were analysed in a 15 % SDS-polyacrylamide gel. These purified His6-tagged proteins were used in all experiments other than DNA binding studies.

    For DNA binding experiments, native CII (without any tag) was obtained by overexpressing the recombinant plasmid pAB305 in BL21 (DE3) E. coli cells and purification by two steps of ion-exchange chromatography as described previously (Datta et al., 2001). Tagless mutant proteins were obtained by removal of the N-terminal His6-tag followed by chromatography using a Superdex 75 column (Amersham/GE Healthcare Biosciences) as described below. Between 5 and 8 mg of the His6 mutant CII was digested with 1.5 U thrombin at 22 °C for 16–20 h. The reaction mixture was loaded on a Superdex 75 column, pre-equilibrated with 20 mM Tris/HCl (pH 8.0), 300 mM NaCl and 1 mM EDTA. Chromatography was carried out at 25 °C with a flow rate of 1.0 ml min−1. The eluted fractions were run on 15 % SDS-PAGE and the fractions containing only a single band corresponding to the desired size were pooled together and concentrated with an Amicon Ultra spin column with a cut-off of 5 kDa (Millipore).

    Glutaraldehyde cross-linking.

    Glutaraldehyde cross-linking experiments were carried out to identify the oligomeric state of the CII mutants. The cross-linking reactions were done essentially as described by Kobiler et al. (2007), with some modifications. Briefly, 0.05 % of glutaraldehyde was added to 10 μM of each protein solution in buffer P (20 mM Tris/HCl, 300 mM NaCl, 1 mM EDTA, pH 8.0) and the reaction mixtures were incubated for 1 min at room temperature. Reactions were immediately terminated by the addition of 200 mM glycine followed by the addition of 1× SDS-PAGE gel-loading buffer and boiling. The samples were separated by electrophoresis on 15 % SDS-PAGE, transferred to a PVDF membrane and immunoblotted with anti-CII polyclonal antibody. The blot was subjected to densitometric analysis using the volume analysis routine of the molecular analyst software (Bio-Rad).

    Size-exclusion chromatography.

    For size-exclusion chromatography, an ÄKTA FPLC system (Amersham Biosciences) was used with a Superdex 75 HR column (Datta et al., 2003) pre-equilibrated with buffer P. Between 250 and 500 μg protein was injected at a time. The flow-rate was maintained at 0.5 ml min−1. The Rf value was calculated from the equation,

    Figure image not available in archive
    where Ve=elution volume, V0=void volume and Vt=total volume of the column.

    In vitro transcription.

    For all CII-dependent transcription activation studies, a double-stranded (ds)DNA fragment containing the pE promoter was used as the template. It was prepared as described by Datta, 2003</xref>. The 540 bp DNA (transcription template, IVTT) was generated by PCR using the pAB801 plasmid (pBluescript SK− plasmid carrying 651 bp BglII DNA fragment of the λ genome) as the template and FPivt (5′-AAAGCCCTTCCCGAGTAAC-3′) and RPivt (5′-TCTGCCACATTACGCTCC-3′) as the forward and the reverse primers, respectively. The PCR-amplified DNA fragment contained the promoters pE (CII-dependent, would generate a 144-base run-off transcript) and poop (CII-independent, would give a 77-base terminated transcript acting as an internal positive control), as shown in Supplementary Fig. S3(a) (available in JGV Online).

    In vitro transcription reactions were performed essentially in the same manner as described by Datta et al. (2005b), in a 20 μl reaction volume in transcription buffer (40 mM Tris/HCl, 0.1 M potassium glutamate, 1 mM DTT, 20 mM MgCl2, pH 8.0) using 5 nM IVTT and 60 nM sigma-saturated E. coli RNA polymerase (Epicentre). CII (or its mutants) was added at a concentration of 1 μM. Reactions were incubated at 37 °C for 20 min and transcription was initiated by the addition of the nucleotide mix [0.1 mM each of ATP, GTP, CTP, 0.01 mM of UTP (all ribonucleotides were from Amersham Biosciences) and 5 μCi of [α-32P]UTP (BRIT) and 1 μg heparin]. After 20 min, the reactions were terminated by the addition of 5 μl formamide stop dye (90 % formamide, 20 mM EDTA, 0.05 % bromophenol blue, 0.05 % xylene–cyanol) followed by electrophoresis in a 10 % (w/v) polyacrylamide–7 M urea gel and autoradiography at −20 °C.

    Preparation of construct for in vivoβ-galactosidase assays.

    The pE promoter was amplified by PCR using IVTT as the template, and PEF and PER (Supplementary Table S2) as the forward and reverse primers respectively (Supplementary Fig. S3a). The resulting 298 bp DNA fragment was then cloned into the pSD5B vector (Jain et al., 1997) at the XbaI site, upstream of the lacZ gene, in the correct orientation as checked by control PCRs using the appropriate intermediate primer 5′-TCGCTATTACGCCAGCTG-3′ (200 bp downstream from the start codon of the lacZ gene in pKP109; see Supplementary Fig. S3b) and PEF. The mutant cII genes were subcloned into pET15b at the NdeI and BamHI sites, to change the antibiotic selection marker from kanamycin to ampicillin. The resulting plasmids were derivatives of pAB412 (Halder et al., 2007) which carries the wild-type cII gene (Supplementary Fig. S3c).

    In vivo transcription (β-galactosidase) assays.

    β-Galactosidase assays were carried out according to the method described by Miller (1972). E. coli BL21 (DE3) cells were cotransformed with plasmid pAB412, or its derivatives carrying mutated cII (pKP401 to pKP406; Supplementary Table S1), and pKP109 (bearing the lacZ gene under promoter pE), and were used as the host. Both of these plasmids were compatible with each other with respect to their replicons p15A and colE1 (Supplementary Fig. S3b, c). BL21 (DE3) cells carrying pKP109 and pET15b (i.e. lacking cII) were used as the negative control. The cells were allowed to grow in 5 ml LB medium (supplemented with 100 μg ampicillin ml−1 and 50 μg kanamycin ml−1) at 37 °C, followed by induction with 100 μM IPTG at A590 ∼0.6–0.7 for 10 min. Following this, they were immediately transferred onto ice and the absorbance at 590 nm was recorded. Aliquots (0.1 ml) of these cultures were added separately to 0.9 ml Z-buffer (60 mM Na2HPO4, 40 mM NaH2PO4, 10 mM KCl, 1 mM MgSO4, 50 mM β-mercaptoethanol, pH adjusted to 7.0). The cells were lysed by the addition of 10 μl 1 % SDS, 20 μl 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 ONPG (4 mg ml−1; dissolved in Z buffer) at 28 °C. Reactions were stopped by the addition of 0.3 ml 1 M Na2CO3 after sufficient yellow colour had developed due to the cleavage of the glycosidic bond of ONPG by β-galactosidase, leading to the production of ortho-nitrophenyl ions. The absorbance at 420 and 550 nm were recorded. The β-galactosidase activity (Miller units) was calculated from the formula,

    Figure image not available in archive
    where t=time of reaction (min) and V=volume (ml) of the culture used in the assay (Miller, 1972).

    DNA binding studies.

    A 20 bp dsDNA fragment containing the recognition sequence for CII (pE) was used to check the DNA-binding ability of the proteins, using fluorescence anisotropy (LeTilly & Royer, 1993). The sense and anti-sense strands (complementary to each other, see Supplementary Table S2 for sequences) were obtained from BioTechdesk as HPLC-purified DNA, with the sense strand 5′-labelled with fluorescein. The strands were annealed in a buffer containing 20 mM Tris/HCl, 100 mM NaCl and 1 mM EDTA, pH 8.0, mixed with each other to a final concentration of 100 μM of each. The oligonucleotide mixture was then heated in a boiling water bath for 15 min and allowed to cool slowly to room temperature. The concentration of the dsDNA was estimated from its absorbance at 260 nm.

    Fluorescence anisotropy was measured in a FluoroMax-3 spectrofluorometer (HORIBA/Jobin Yvon). The fluorescein-labelled double stranded oligonucleotide (10 nM) was taken in a total volume of 2.5 ml. The titration was performed in assay buffer containing 20 mM Tris/HCl, 100 mM potassium glutamate and 5 % glycerol, pH 8.0. The proteins were diluted to the desired concentration in the same assay buffer. Increasing concentrations of protein (native or mutant CII, up to 900 nM) were added. After each addition, the DNA–protein mixture was incubated for 2 min at room temperature before taking the reading. Fluorescence at 520 nm was measured, with λex=495 nm and bandwidths of 5 nm on each side. All the proteins used in this study were without the His6-tag. Anisotropy was measured six times for each protein concentration and the mean anisotropy values were found.

    CD spectroscopy.

    CD measurements for CII (or its mutants) were carried out in buffer P, both in the presence or absence of 0.2 M guanidium chloride at room temperature using a JASCO J600 spectropolarimeter, as described previously (Datta et al., 2003). The far-UV CD spectrum (250–200 nm) was recorded with 6 μM protein solution.

    Complementation assays.

    The in vivo activity of His6-tagged native and mutant CII was tested by complementation assays following the method described by Halder et al. (2007), as described below. E. coli BL21 (DE3) cells containing pAB412 (carrying the native cII gene) or any of the plasmids pKP301 to pKP306 (carrying one of the mutated cII genes, see Supplementary Table S1) were used as hosts. These host cells were subjected to infection by phage λcII68 (cII−) to test whether the expressed CII mutants were functional. The bacterial cultures were grown in TBM (1 % tryptone, 0.5 % NaCl, 0.2 % maltose and 0.2 % MgCl2) in the presence of 100 μg ampicillin ml−1 (for pAB412) or 50 μg kanamycin ml−1 (for plasmids carrying mutated cII) at 37 °C. Protein expression was induced by 50 μM IPTG at 37 °C for 30 min at A590 ∼0.8–0.9. An aliquot of culture containing 2×107 cells (expressing either native CII or one of the CII mutants) was mixed with 100 phage particles which were adsorbed at room temperature for 20 min. Soft agar (3 ml; TBM plus 0.5 % agar) was added and the mixture was immediately poured onto an LB agar plate. The plate was incubated at 32 °C for 16 h. Complementation by native or mutant CII was judged by examining the morphology of the resulting plaques (turbid or clear).

    Acknowledgments

    The authors would like to thank Dr S. Adhya (NIH, USA) for λcII68, Professor G. Basu (Bose Institute) for useful suggestions and Professor P. Chakrabarti (Bose Institute) for his comments on the manuscript. P. P. was supported by a research fellowship from CSIR, India.

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