Identification of the SaPI2-Coded Inhibitor(s) of Phage 80.

To determine whether SaPI2 uses the same interference mechanisms against phage 80 as it does against phage 80α, we cloned the cpmAB and ppi loci of SaPI2 separately to the vector pCN51 (9), behind the exogenous cadmium-inducible promoter (Pcad) (10) and tested these clones for inhibition of phage 80. Although either of these clones completely blocked plaque formation by phage 80α (7), we were surprised to find that neither, even when induced, had any effect on plaque formation by phage 80 (Fig. 1A). To confirm this effect in situ, we deleted both ppi and cpmAB from SaPI2 and found that the interference was unaffected. This result showed that there must be a third interference mechanism; to identify it, we cloned each of the SaPI2 ORFs separately, using the same vector, and found that one of the ORFs, 17, caused interference. ORF17, when cloned, blocked plaque formation by phage 80 (Fig. 1A) (7). One other ORF, ORF13, was toxic to the host cells and could not be expressed adequately from a plasmid. It was found to cause interference on the basis of a deletion (see below).

• Download figure
• Open in new tab
• Download powerpoint

Fig. 1.

(A) Phage interference mediated by cloned SaPI2 genes. The indicated genes were cloned to plasmid pCN51 under a cadmium-inducible promoter (Pcad). Strain RN4220 containing the indicated plasmids was infected with phages 80α (∼150 pfu per plate) or 80 (∼250 pfu per plate) plated on phage bottom agar containing 1.0 μM CdCl₂ and incubated for 48 h at 32 °C. Plates were stained with 0.1% TTC in TSB and photographed. (B) Map of SaPI2 operon 1. Genes previously shown to cause phage interference are shown in red; hypothetical genes of unknown function are shown in blue; numbered ORFs newly determined to be involved in interference are shown in orange; and the SaPI2 small terminase subunit is shown in green.

ORF13 and -17 are located in operon 1, which includes cpmA and B, ORF16, and terS_S. ppi is upstream of the operon and is equivalent to ORF10 (Fig. 1B). No other SaPI2 gene had any detectable inhibitory effect on phage 80.

ORF17-Mediated Interference.

We began with ORF17 because it could be overexpressed. To identify the phage gene(s) targeted by ORF17, we isolated ORF17-resistant mutants as rare plaques on a strain containing the cloned and overexpressed ORF17. Thirteen of the mutant phages were sequenced. All 13 had point mutations in a single gene, 32, and all had amino acid replacements at a single site in gp32: tyrosine 67 was replaced by histidine in 11 mutants and by cysteine in two mutants (Fig. S1).

The sequence of gene 32 showed it to be late transcriptional regulator C, ltrC (11, 12), which corresponds to 80α rinA (13). LtrC and RinA are regulatory proteins that activate the transcription of the late phage operon (morphogenetic and lysis genes) (Fig. 2A). Five families of these proteins have been described, constituting a superfamily of late transcriptional regulators (Ltr) (11, 12). These are essential phage genes (14, 15) corresponding to coliphage lambda gene Q. To test phage 80 LtrC for its role in transcriptional activation, we needed to delete the gene. Because the gene is presumed to be essential for phage reproduction, this deletion would have to be done with the prophage. This deletion could not be done with phage 80, which cannot form a lysogen (13), so we turned to a close relative, phage 52A, which can form lysogens, can induce SaPI2, and whose LtrC and late gene operon are identical to those of phage 80. Using a phage 52A lysogen, we introduced an in-frame deletion in ltrC and found that the phage, as predicted, was unable to lyse the cell or to produce any infectious phage particles. We next cloned the β-lactamase reporter downstream of the late phage promoter in plasmid pCN41 in RN450 with either the WT or ltrC mutant 52A prophage. The ltrC mutant did not detectably activate the late gene promoter (Fig. 2B); following mitomycin C treatment, the culture did not undergo lysis, and no detectable phage were produced, confirming the essentiality of the gene for phage 52A and, by inference, for phage 80. The presence of SaPI2 had no effect on the β-lactamase activity in the strain with the ltrC mutant but, as predicted, had a profound effect on the activity in the strain with the WT phage (Fig. 2B). Using the WT phage, we next demonstrated inhibition of late phage transcription by the cloned SaPI2 ORF17. These results are presented in Fig. 2C and have suggested the designation “phage transcription inhibitor A” (ptiA) for the gene (ORF17). Because PtiA blocked expression of the entire late phage gene module, we predicted it would block lysis as well as plaque formation. Note that the lysis of a culture and the formation of plaques on an indicator strain are rather different tests of phage function: Plaque formation can be blocked by a modest reduction in burst size, whereas lysis can be blocked only by preventing the expression of the lysis module. Unlike the other two interference mechanisms, which blocked plaque formation but not lysis (7), PtiA, as predicted, blocked both (Fig. 3A), making it the most severe of the interference mechanisms yet identified. Also, predictably, the cloned ptiA blocked plaque formation by other phages encoding LtrC, as shown in Fig. S2.

• Download figure
• Open in new tab
• Download powerpoint

Fig. 2.

(A) Comparison of the rinA and ltrC genomic region from 80α and 80. Note that the position of ltrC (ORF32) in phage 80 corresponds to that of rinA in phage 80α. “P” represents the late gene promoter. (B) Activation of phage 52A late transcript promoter by LtrC. The late transcript promoter was cloned to plasmid pCN41 as a transcriptional fusion to β-lactamase. The resulting reporter plasmid was introduced into strains lysogenic for either the WT prophage or a derivative prophage containing a deletion of ltrC in the absence or presence of SaPI2. β-Lactamase activity was measured 2 h after induction with mitomycin C (MC) and in a parallel culture without induction. (C) Effects of cloned ptiA and ptiM on the phage late transcript promoter. ptiA and ptiM were cloned singly or together to pCN51, and their effects on LtrC activation of the late transcript promoter–β-lactamase fusion were determined. For these tests, the pti genes were induced with 1.0 µm CdCl₂, the β-lactamase fusion construct was integrated in the chromosome at the SaPI4 att site (lab plasmids collection), and LtrC was provided by superinfection with phage 80. Cultures were assayed at 2 h postinfection, and assays were performed in triplicate.

• Download figure
• Open in new tab
• Download powerpoint

Fig. 3.

(A) Modulation of PitA activity by PtiM. (Upper) Plaque assay. Phage platings were as in Fig. 1A. (Lower) Lysis assay. RN4220strains carrying pCN51 (vector) or carrying the pti clones used were grown with the inducer (1.0 μM CdCl₂), infected with phage 80 at a multiplicity of infection (MOI) of 0.2, and incubated at 32 °C at 60 rpm for 3 h. (B) Bacterial adenylate cyclase-based two-hybrid (BACTH) analysis. Spots in each row represent three independent colonies. Plasmid combinations are numbered as follows: 1, pKT25-zip + pUT18C-zip (positive control); 2, pKT25-LtrC (WT80) + pUT18-PtiA_S2; 3, pKT25-LtrC (PtiA_S2-resistant 80) + pUT18-PtiA_S2; 4, pKT25-PtiM_S2 + pUT18-PtiA_S2; 5, pKT25 + pUT18 (negative control). Blue color indicates cAMP-dependent lacZ expression following reconstitution of adenylate cyclase activity by the interaction of fusion proteins.

Mechanism of Action of PtiA.

Because all the PtiA-resistant mutations mapped in the LtrC coding sequence, it was likely that PtiA acted by direct protein binding. Accordingly, we performed a two-hybrid assay, as shown in Fig. 3B. As can be seen (Fig. 3B, row 2), there was a strong positive reaction between PtiA and the LtrC WT protein, but no reaction was detected between PtiA and a PtiA-resistant mutant of LtrC (Fig. 3B, row 3). Therefore we conclude that PtiA interferes with phage 80 growth by binding to LtrC and directly inhibiting its ability to activate late gene transcription.

It is interesting that the phage 80 proteins corresponding to the 80α targets of Ppi (TerS_P) and of CpmAB (presumably the phage head scaffolding protein) are considerably different from the corresponding 80α proteins and are unaffected by these inhibitors. In fact, phage 80 virion proteins are not formed into small capsids by any of the three prototypical SaPIs (13); moreover, 80α is indifferent to PtiA, presumably because of the insensitivity of RinA to the inhibitor (Fig. 1A). PtiA, in turn, also occurs in variant forms, and we have cloned those of SaPI1 and SaPI1bov1. The SaPI1 variant PtiA_S1 differs from PtiA_S2 by a single amino acid substitution and is equally effective against phage 80; however, the SaPI1bov1 variant PtiA_SB1 differs at 10 sites and does not inhibit phage 80 (Fig. S3). PtiA homologs are encoded by all intact SaPIs, and their sequences suggest that they fall into different groups. Perhaps PtiA variants encoded by different SaPIs inhibit LtrC variants encoded by different phages.

Modulation of PtiA Activity.

As noted above, when overexpressed (cloned to a plasmid and driven by an inducible promoter), PtiA_S2 not only blocks phage 80 plaque formation but totally prevents lysis (Fig. 3A). The logic of SaPI-phage parasitism would indicate that lysis prevention cannot be the normal effect of PtiA_S2 because it would be suicidal for the SaPI as well as for the phage. However, SaPI2 is not a suicide bomber: Although WT SaPI2 eliminates plaque formation by phage 80, the infected cultures undergo lysis. So we next attempted to find the basis for this difference. Because, when overproduced, PtiA_S2 totally blocks phage 80 reproduction, and because overproduction is expected during SaPI replication, it seemed unlikely that the difference is based simply on relative gene dosage. In fact, even in single copy, ptiA_S2 has a profound effect on phage 80 reproduction (Fig. S4). Therefore, we considered the possibility that PtiA_S2 action is modulated by a second SaPI2 gene. As seen in Fig. 1B, ORF16 is immediately upstream of ptiA, and sequence analysis suggested that the two genes probably are translationally coupled. Therefore, ORF16 was a likely candidate for the putative modulator of PtiA_S2. To test this possibility, we compared the effects of the cloned ptiA_S2 with those of a clone containing both ORFs on the activity of phage 80. As seen in Fig. 3A, the cloned ptiA_S2 totally inhibited phage 80, whereas the ORF16-ptiA_S2 clone caused only modest inhibition of the phage and did not prevent lysis. Furthermore, ORF16 also reduced the PtiA_S2-mediated inhibition of the phage late gene transcription (Fig. 2C). Because the clone of ORF16 alone had little, if any, effect on plaque formation, it is suggested that ORF16 is, in fact, the modulator of PtiA_S2. If so, ORF16 could act at the level of ptiA_S2 transcription or at the level of translation of the product, or it could form an inhibitory complex with the putative protein. Because the two genes almost certainly are cotranscribed, it seemed highly unlikely that ORF16 controlled the transcription of ptiA_S2. As noted above, modulation of gene dosage is unlikely to be the mechanism of control by ORF16. Therefore, it also seemed unlikely that ORF16 controlled the translation of ptiA_S2. Accordingly, we considered it more likely that there would be direct binding between the two proteins. We tested for this direct binding by performing a second bacterial two-hybrid assay to test for possible binding of the ORF16 product, gp16, to PtiA_S2. As shown in Fig. 3B row 4, a positive binding reaction was readily demonstrated, suggesting that gp16 modulates the activity of PtiA_S2 by a direct binding interaction. Because PtiA_S2 has two binding partners, which could be mutually exclusive, the equilibrium between these partners could account for a subtle and precise modulation system. On the basis of these results we propose that gp16 be designated “modulator of PtiA_S2,” PtiM_S2.

To confirm the effects of PtiA_S2 and PtiM_S2 in the intact SaPI, we constructed in-frame deletions of the two genes and tested them individually and together for their effects on phage 80 plaque formation. As seen in Fig. 4, deletion of ptiA_S2 resulted in small plaques; deletion of ptiM_S2 eliminated plaque formation, as expected, because of the absence of its modulatory effect on ptiA_S2. Also as expected, deletion of both caused the formation of small plaques indistinguishable from those seen with deletion of ptiA_S2 alone; however, the failure of these deletions to restore the formation of full-sized plaques is consistent with interference by ORF13, because, as noted above, none of the other SaPI2 ORFs showed interference. Accordingly, we then constructed an in-frame deletion of ORF13 and, indeed, this deletion resulted in partial restoration of plaque formation (Fig. 4); deletion of both ptiA_S2 and ORF13 restored plaques to full size (Fig. 4). More definitive results were obtained by one-step growth analysis of phage 80 in the presence of SaPI2 with and without deletions of the interference genes studied here. As shown in Fig. 5A, deletions of the various interference genes restored the growth of phage 80 to different extents, roughly paralleling the plaque formation tests.

• Download figure
• Open in new tab
• Download powerpoint

Fig. 4.

Effects of SaPI2 ptiA, -B, and -M deletions on SaPI2 interference with phage 80. Strain RN4220 containing WT SaPI2 or various SaPI2 deletion mutants was infected with phage 80 (∼250 pfu per plate), plated on phage bottom agar, and incubated for 48 h at 32 °C. Plates were stained with 0.1% TTC in TSB and photographed.

• Download figure
• Open in new tab
• Download powerpoint

Fig. 5.

(A) One-step phage growth analysis of ptiA- and ptiB-mediated interference of phage 80. Bacteria were infected with phage 80 at an MOI of 10, washed to remove unadsorbed phage, diluted to contain ∼10⁴ infective centers/mL, and then incubated for 90 min. Samples were removed at the indicated times and plated for plaque formation using RN4220 as the indicator. Three replicates were used for each strain. (B) Effect of deletion of ptiA, ptiB, ptiM, and their combinations on phage 80 LtrC activity. The late phage transcript promoter–β-lactamase reporter used for the experiments in Fig. 2 B and C was tested in RN4220 with SaPI2 and several of its deletion mutants. Strains were infected with phage 80 at an MOI of 0.2. The infected cultures were incubated at 32 °C, 60 rpm and assayed for β-lactamase at 2.5 h postinfection. Three replicates were used for each strain.

ORF13-Mediated Interference.

Because of its bacterial toxicity, ORF13 could not be cloned in Escherichia coli and could be cloned in Staphylococcus aureus only behind a promoter (Ptet) with an extremely low basal level of expression. Bacteria could grow on plates only at concentrations of the inducer anhydrotetracycline (ATc) ≤2 ng/mL; growth on an ATc gradient plate (0–20 ng/mL) is shown Fig. S5A. Plaque size was slightly larger at the borderline of inhibition, suggesting that the host cells are more sensitive to ORF13 than the phage. Thus, our usual strategy for identifying interference targets, namely the formation of rare mutant plaques by the inhibited phage, could not be used in this case. Because we already had a very useful β-lactamase fusion to the late phage promoter, it seemed worthwhile to perform the extremely simple test of measuring the effects of the ORF13 deletion on that promoter before attempting to identify the unknown phage function that was inhibited. In fact, the ORF13 deletion had a profound effect on the late phage promoter, as shown in Fig. 5B (light green bar). It relieved the inhibition by SaPI2 to about the same extent as the ptiA_S2 deletion (aqua bar) and, more strikingly, in conjunction with the ptiA_S2 deletion, it fully restored promoter activity (orange bar). On the basis of these results, we have designated ORF13 ptiB_S2. It is likely, but not certain without mutational evidence, that this is the only interference effect of PtiB_S2.

The question that then arises is whether PtiB_S2 is toxic to host cells in its native context. In the absence of SaPI or SOS induction, it would not be expressed. Because the host cells lyse after the induction of a helper phage, PtiB_S2 could not be so toxic to the host cells that it would interfere with the phage lytic cycle. Indeed, after Ptet induction in broth, the ptiB_S2-containing strain grows for six generations and then stops sharply (Fig. S5B), suggesting that the synthesis of an essential protein is inhibited and that there is an ∼64-fold excess of this protein in exponential cells, an amount that surely is more than sufficient to support the lytic cycle. Therefore, its host cell toxicity is irrelevant to phage propagation. In principle, one could test for host inhibition in the absence of any phage, because SaPI operon 1, in which the gene is located, is repressed by LexA (16). Although SOS induction relieves LexA repression, the effect is transient, and transient expression of ptiB_S2 would not provide a clear answer because cells would have six generations to recover from any toxicity. We suggest that, in any case, the possible toxicity of PtiB_S2 is of little or no importance to the phage or SaPI life cycle.

Another question that arises is whether PtiB_S2, like PtiA_S2, is modulated by a separate SaPI gene product. The simplest route to answering this question would be via host mutants resistant to PtiB_S2. Although colonies appear on inducer-containing plates at a frequency of ∼10⁻⁶, analysis of 25 such colonies revealed that all have mutations in Ptet. Screening of additional colonies is in progress. Alternatively, an approach involving further cloning and deletion studies has been initiated; however, that work is beyond the scope of this report.