Lysogenic Lytic Cycle Compare Contrast Essays

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Bacteriophages (phages), which are natural viral predators of bacteria, multiply by infecting specific host bacteria. Although there is an additional type of phage-host relationship called “steady-state infection,” which is exemplified by filamentous phages (1), phage genome replication generally occurs via two different developmental paths: the lytic cycle and the lysogenic cycle. In contrast to the lytic cycle, which results in immediate bursting of the host bacteria and the release of bacteriophage progeny, the lysogenic cycle involves the maintenance of the phage genome as a part of the host genome for several generations, typically by integrating into host chromosomes or, more rarely, by replicating as low-copy-number phage plasmids (2–4). The expression of genes necessary for progeny production and host cell lysis is tightly repressed by a phage regulatory system, but some physiological changes in the host induced by UV light irradiation or other DNA-damaging agents activate the lytic cycle by disabling the phage repressor. Phages fall into two categories: virulent phages that replicate strictly by the lytic cycle and temperate phages that can enter both the lytic cycle and the lysogenic cycle.

The lytic switch following lysogenic development has been well studied in the temperate phage lambda. In the lambda lysogenic phase, phage CI repressors form dimers and bind to specific operators to prevent expression of lambda early genes and subsequent late genes (5, 6). Upon host DNA damage, the activated host RecA protein induces CI proteolysis in a manner similar to the inactivation of the host SOS response regulator LexA (7–9). CI proteolysis leads to the expression of early and late genes, resulting in lytic development. This mechanism illustrates how lambda and other similar phages exploit the host cell SOS response to escape quickly from a potentially damaged host using the RecA-dependent cleavable repressor. Alternatively, some phages in the families Sipho- and Myoviridae utilize the LexA-regulated antirepressors instead of the cleavable repressor to associate their lytic switch to the host SOS response (10–12).

Here, in an effort to identify the factor(s) that causes a phenotypic difference between two very similar podoviral Salmonella phages, SPC32H and SPC32N, we found a novel Podoviridae phage lytic switch antirepressor. We observed that a single nucleotide change in the LexA-binding site, which overlaps with the promoter of the phage antirepressor gene, causes constitutive expression of the antirepressor Ant and consequent inhibition of phage repressor function in SPC32N. As a result, SPC32N could not establish lysogeny as clear plaque mutants. A LexA-dependent lytic switch involving an antirepressor, rather than repressor proteolysis, has been found previously in only sipho- and myoviral phages (10–12), and the podoviral SPC32H/N Ant protein had no significant homology to these known antirepressors. A database search identified many proteins with homology to Ant, suggesting the extensive use of antirepressor-mediated lytic induction among temperate phages in the order Caudovirales.

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Bacterial strains, plasmids, and growth conditions.The bacterial strains and plasmids used in this study are listed in Tables 1 and 2, respectively. All Salmonella mutants were derived from the prophage-cured Salmonella enterica serovar Typhimurium strain LT2 [referred to as LT2(c)] and its ΔLT2gtrABC1 (SR5003) derivative to exclude the effect of prophages and spontaneous phage resistance via O-antigen glucosylation, respectively (13, 14). Standard cloning procedures were used to construct the recombinant plasmids. Bacteria were grown aerobically at 37°C in LB medium supplemented with the following chemicals, as needed: ampicillin (Ap), 50 μg ml−1; kanamycin (Km), 50 μg ml−1; chloramphenicol (Cm), 25 μg ml−1; 5-bromo-4-chloro-3-indolyl-β-d-galactoside (X-Gal), 40 μg ml−1; l-arabinose, 0.2% (final concentration); isopropyl-β-d-thiogalactopyranoside (IPTG), 1,000 μM (final concentration); and mitomycin C (MMC), 1 μg ml−1 (final concentration). For the disc diffusion assay, 6-mm-diameter filter paper discs were soaked with 10 μl of arabinose, antibiotics, or MMC at the indicated concentrations, placed on the surface of the bacterium-inoculated solidified soft top agar (LB supplemented with 0.4% [wt/vol] agar and X-Gal, if necessary), and incubated at 37°C for 8 h.

Table 1

Bacterial strains and bacteriophages used in this study

Table 2

Plasmids used in this study

Bacteriophage.The bacteriophages used in this study are listed in Table 1. All phage mutants were derived from the temperate phage SPC32H, which was previously isolated from chicken fecal samples obtained from a traditional marketplace in South Korea (15). Routine phage spotting and double-agar overlay assays were conducted to determine the efficiency of plating (EOP) in specific bacteria (14, 15). For the morphological analysis, the phage stocks were negatively stained with 2% uranyl acetate (pH 4.0) as previously described (15) and were examined by transmission electron microscopy (LEO 912AB TEM; Carl Zeiss, Jena, Germany) at 120-kV accelerating voltage. The images were scanned with a Proscane 1,024 × 1,024-pixel charge-coupled device camera.

Bacteriophage genome sequencing and analysis.Phage nucleic acids that were extracted by the phenol-chloroform extraction method with protease K/SDS treatment (16) were pyrosequenced using the GS FLX Titanium system by Macrogen, Seoul, South Korea. The quality-filtered reads were assembled using the GS de novo assembler (v. 2.60), and the open reading frames (ORFs) that encode proteins of more than 35 amino acids in size were predicted using the software programs GeneMarkS (17), Glimmer 3.02 (18), and FgenesB (Softberry, Inc., Mount Kisco, NY, USA). The predicted ORFs were annotated based on the results of BLASTP (19), InterProScan (20), and NCBI Conserved Domain Database (21) analysis. tRNAscan-SE (22) and BPROM (Softberry, Inc.) were used to predict the tRNA sequences and the putative promoter/transcription factor-binding sites, respectively. Genomic comparison at the DNA level was visualized using the program Easyfig (23).

Construction of the Salmonella and phage mutants.The lambda red recombination method was used for in-frame gene deletion (24). To construct the noncleavable LexA protein, a point mutation in lexA {resulting in a G85D mutation in the amino acid sequence [lexA(G85D)]} was generated by lambda red recombination and double homologous recombination-based counterselection, as previously described (14), using the suicide vector pDS132 (25). The SPC32H lysogen [ΔLT2gtrABC1 (32H); SR5100] was isolated by sequential streaking of SPC32H-resistant clones from a lawn of phage-treated ΔLT2gtrABC1 and was verified by PCR amplification of the phage attachment (attR) site. The transcriptional recET::lacZ fusion was constructed using pCE70, as previously described (26, 27). Human influenza virus hemagglutinin (HA) epitope tagging of the specific gene(s) was also accomplished by lambda red recombination using oligonucleotides containing the HA tag sequence.

Phage mutants were induced from the SPC32H lysogen after the gene manipulations described above, with some modifications. Briefly, to generate SPC32H m1, a truncated tailspike gene (tsp::Kmr), which was constructed by lambda red recombination in the SPC32H lysogen, was replaced with the m1-containing tsp gene by double homologous recombination-based counterselection. The presence of the m1 mutation in the induced phage was confirmed by DNA sequencing. Similar methods were used to construct SPC32H m2 and SPC32H m12. The oligonucleotides used in this study are listed in Table 3.

Table 3

Oligonucleotides used in this study

Bioluminescence reporter assay.The 197-bp fragment upstream of the ant gene in SPC32H (designated Pant_H) or SPC32N (designated Pant_N) was PCR amplified and cloned into pBBRlux (28), resulting in the transcriptional fusion of the operon luxCDABE to the putative ant gene promoter. S. Typhimurium strains harboring this reporter plasmid were cultured in 200 μl of fresh LB broth supplemented with appropriate antibiotics in a 96-well plate. The cellular bioluminescence of the culture and the absorbance at 600 nm (A600) were measured periodically using an Infinite 200 Pro plate reader (Tecan, Männedorf, Switzerland), and the results were expressed in arbitrary relative light units (RLU). To trigger the SOS responses, MMC was added to the culture after a 3-h incubation. The three independent assays with triple technical replications were performed.

Western blot analysis.At the mid-exponential phase, culture of the HA-tagged-gene(s)-containing Salmonella was treated by MMC, and portions of the culture were sampled at the indicated time points. Bacterial cells were harvested by centrifugation and were lysed with the B-Per reagent (Thermo Scientific, Illinois, USA). Soluble proteins (10 μg) from cell lysates were separated by 15% SDS-PAGE and electrotransferred to the polyvinylidene difluoride (PVDF) membrane. HA-tagged proteins and DnaK were detected with anti-HA and anti-DnaK antibodies, respectively. The chemiluminescence signals were developed using the West-Zol plus Western blot detection system (iNtRON Biotechnology, Gyeonggi-do, South Korea) after the goat anti-mouse IgG-horseradish peroxidase (HRP) (Santa Cruz Biotechnology, CA, USA) treatment, and then X-ray film was exposed to chemiluminescent light to detect the signals.

Bacterial two-hybrid assay.Protein-protein interaction was determined by the recovery of adenylate cyclase (CyaA) activity through heterodimerization of fusion proteins in the Escherichia coli BTH101 reporter strain (cyaA mutant) (29). The reporter strain harboring the fusion plasmid pair (e.g., pKT25-rep and pUT18c-ant) was streaked on LB agar supplemented with Km, Ap. and X-Gal or subjected to the β-galactosidase assay (30) to quantitatively measure the interaction.

Purification of proteins, rTEV protease treatment, and analytical size exclusion chromatography.Cultures of E. coli BL21(DE3) harboring pHIS-LexA, -Rep, or -Ant (optical density at 600 nm [OD600] = ∼0.15) were treated by 100 μM IPTG and incubated at 25°C for an additional 4 h. Cells were harvested by centrifugation and lysed in lysis buffer (20 mM Tris [pH 8.0], 500 mM NaCl, and 20 mM imidazole) by sonication on ice. Centrifuged (16,000 × g, 4°C, for 30 min) and filtered (0.22-μm filter; Millipore, Ireland) cell lysate was subjected to nickel-nitrilotriacetic acid (Ni-NTA) affinity chromatography (Qiagen, California, USA) according to the manufacturer's protocol with elution buffer (lysis buffer with 250 mM imidazole). The eluted protein was concentrated using a Vivaspin 20 instrument (3,000-molecular-weight cutoff [MWCO] polyethersulfone [PES]; Sartorius, Goettingen, Germany), and the buffer was changed (20 mM [Tris pH 8.0], 500 mM NaCl, and 50% glycerol) using a PD MidiTrap G-25 column (GE Healthcare, Buckinghamshire, United Kingdom). To remove the His6 tag from the purified proteins, recombinant tobacco etch virus (rTEV) protease (1:5 ratios in concentration) was treated for 6 h at 4°C in a cleavage buffer (10 mM Tris [pH 8.0], 150 mM NaCl, 0.5 mM EDTA, and 100 mM dithiothreitol [DTT]). For analytical size exclusion chromatography, a Superdex 200 10/300 GL column (GE Healthcare) was used. The column was equilibrated with a buffer consisting of 500 mM NaCl and 20 mM Tris [pH 8.0], and then purified proteins (500 μl of 0.8 μg μl−1) were loaded on to the column at a flow rate of 0.5 ml min−1.

Electrophoretic mobility shift assay (EMSA).The purified PCR fragments of the ant gene promoter region (APR) was γ-32P labeled using T4 polynucleotide kinase (TaKaRa, Japan). The labeled DNA (approximately 4 nM) was incubated with various concentrations of LexA for 30 min at 37°C in 20 μl of reaction mixture containing 1× binding buffer (10 mM HEPES [pH 8.0], 10 mM Tris [pH 8.0], 50 mM KCl, 1 mM EDTA, 1 mM dithiothreitol, and 5% glycerol) and 1.1 μg of poly(dI-dC). For determination of Rep binding, various amounts of Rep were incubated for 15 min at 20°C with the 4 nM labeled DNA in the 20-μl reaction mixture. When appropriate, Rep was preincubated with various concentrations of Ant for 30 min at 20°C prior to incubation with labeled DNA. The samples were resolved by 6% native PAGE in 0.5× TBE buffer (45 mM Tris-borate [pH 8.3] and 1 mM EDTA). The gels were vacuum dried, and the radioactivity was analyzed using a BAS2500 system (Fujifilm, Tokyo, Japan).

Nucleotide sequence accession numbers.The genome sequences of SPC32H and SPC32N are available at GenBank under accession numbers KC911856 and KC911857, respectively.

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Phenotypic and genomic characterization of the two related S. Typhimurium phages SPC32H and SPC32N.Previously, we isolated nine phages specific for S. Typhimurium from chicken fecal samples (15). Two of these phages, which originated from the same sample collection, exhibited distinct plaque morphologies on a lawn of S. Typhimurium: one phage (SPC32H) formed turbid plaques surrounded by a halo, but the other phage (SPC32N) formed clear plaques without a halo (Fig. 1A and B). Transmission electron microscopy (TEM) analysis revealed that both phages belonged to the family Podoviridae, since they had an isometric head (∼62.3 nm in diameter) and a short noncontractile tail (∼15.4 nm in length) with tail shaft and tail spikes (Fig. 1C and D). These two phages infected identical repertories of Salmonella strains using the O antigen (O-Ag) of Salmonella as the host receptor (data not shown).

Fig 1

Two similar S. Typhimurium-specific Podoviridae phages, SPC32H and SPC32N, produce morphologically distinct plaques. (A and B) Plaque morphology of SPC32H (A) or SPC32N (B). Dilutions (10 μl) of each phage stock were spotted onto a lawn of the S. Typhimurium LT2(c) ΔLT2gtrABC1 strain (SR5003). (C and D) TEM image of SPC32H (C) or SPC32N (D). Inset at the bottom left of each panel shows the enlarged virion morphology with a black scale bar (50 nm). The white arrow and arrowheads indicate the tail shaft and tail spikes, respectively.

Sequencing of SPC32H and SPC32N revealed that both phages contain 38,689 bp of double-stranded DNA with an identical G+C content of 50.16% and 51 predicted open reading frames (ORFs) with one Arg-tRNA. About half of the ORFs (24 ORFs) were annotated as hypothetical proteins, whereas the other annotated proteins were classified into the following modules: DNA packaging, virion structure morphogenesis, lysogenic conversion, host lysis, and DNA replication/recombination (Fig. 2A; see also Table S1 in the supplemental material). The predicted proteins included a phage integrase as well as a putative repressor, indicating that both phages might be temperate phages. BLASTP searches revealed that the SPC32H and SPC32N genomes closely resembled those of Salmonella phage ε15 and other ε15-like phages (31, 32). Indeed, whole-genome comparisons made at the DNA level revealed a significant degree of synteny between the genomes of SPC32H, ε15, and the ε15-like phage phiV10 (Fig. 2A). In particular, 34 out of 51 SPC32H gene products, including a small/large terminase, a head-to-tail joining protein, a putative major coat protein, a putative holin/endolysin, an integrase, a repressor, and a putative DNA replication protein, were highly similar (50 to ∼100% identity at the amino acid level) to those of ε15 (see Table S1). Genes for the putative SPC32H tail structure module (e.g., SPC32H_016, 017, and 018) had higher similarity to those of phiV10 than ε15 (Fig. 2A). Since these phages infect different hosts (i.e., S. Typhimurium for SPC32H, Salmonella enterica serovar Anatum for ε15, and E. coli O157:H7 for phiV10), differences were observed in the genes encoding tailspike proteins and the flanked lysogenic conversion module (which converts O-Ag to prevent superinfection). Taken together, these results suggest that SPC32H and SPC32N should be assigned to the class of ε15-like phages.

Fig 2

There are two single nucleotide differences between the genomes of ε15-like phage SPC32H and SPC32N. (A) DNA alignment of the genomes of phage ε15 (NC_004775.1), SPC32H, and phiV10 (NC_007804.2) using Easyfig. High sequence similarity between the genomes is indicated by the gray regions. SPC32H ORFs are indicated by numbered or annotated arrows. Phage functional modules are indicated under the arrows. ant, antirepressor; tsp, tailspike; oac, o-acetyltransferase; hol, holin; end, endolysin; int, integrase; rep, repressor. Note that the SPC32N genome is identical to that of SPC32H with the exception of two single nucleotide differences (see panel B). (B) Schematic representation of the location of the two single nucleotide differences, m1 and m2. The partial SPC32H genome sequence surrounding the two single nucleotide differences is shown. m1 (located within the tsp gene) and m2 (located in the intergenic region between SPC32H_020 and tsp) are indicated in bold, uppercase letters. The predicted −10 and −35 sites of the putative promoter for SPC32H_020 gene are boxed. The putative LexA-binding site (SOS box) and the putative repressor-binding site are underlined and doubly underlined, respectively. (C) Consensus sequence of the LexA-binding site from E. coli (8, 33, 34) and the putative LexA-binding sites from phage SPC32H and SPC32N. m2 is indicated with a gray background. Note that the LexA-binding site sequences for SPC32H and SPC32N shown here are reverse complements of the sequence shown in panel A.

A single nucleotide change is responsible for the phenotypic difference between the two phages.Interestingly, a comparison of the full genome sequences of SPC32H and SPC32N revealed only two nucleotide differences. One nucleotide difference, designated m1, is located within the tsp gene (SPC32H_021), which encodes a phage tailspike, and the other nucleotide difference, m2, is located in the intergenic region between a gene (SPC32H_020) encoding a hypothetical protein and the tsp gene (Fig. 2B). To verify whether m1, m2, or both single nucleotide differences were responsible for the differences between SPC32H and SPC32N, we mutated the SPC32H sequence to match that of SPC32N. As shown in Fig. 3A, we observed no significant changes in the turbidity of the lysis zone when the SPC32H m1 sequence was changed to that of SPC32N. However, the turbidity decreased dramatically when the SPC32H m2 sequence was replaced by that of SPC32N. We therefore investigated in detail how the single nucleotide difference at the m2 locus leads to this phenotypic difference.

Fig 3

Introducing the m2 sequence from SPC32N induces SPC32H to enter the lytic cycle. (A) High-titer phage stocks (>107 PFU ml−1; 10 μl) of SPC32H, SPC32N, and three mutant phages derived from SPC32H were spotted onto a lawn of S. Typhimurium LT2(c) ΔLT2gtrABC1. (B) SPC32H can lysogenize host Salmonella, whereas SPC32N cannot. Various template samples were PCR amplified with an attR-specific primer pair. M, DNA marker 1 Kb+ (Invitrogen); i, inner part of the lysis zone; e, edge of the lysis zone; gDNA, genomic DNA; ΔLT2gtrABC1(32H), SPC32H lysogen (SR5100). (C) DNA isolated from the lysis zones shown in panel A was PCR amplified using primers specific for the attR site to determine the lysogenization of each phage. Lanes 1 to 5 correspond to each lysis zone shown in panel A. Note that the introduction of m2 resulted in a disappearance of the lysogen-specific attR band (lanes 4 and 5).

Supplementation with the repressor induces the lysogenic development of the lytic cycle-biased phage SPC32N.Because lysogen formation is normally associated with plaque morphology, we investigated the ability of SPC32H and SPC32N to lysogenize. The SPC32H and ε15 integrases have 93% identity at the amino acid level, and both phages contain the highly conserved common core regions and arm-type binding sequences that are required for phage genome integration (31). This suggests that both phages may integrate their genome into the same attachment site, near the end of the Salmonella guaA gene. Therefore, to detect lysogenization by SPC32H or SPC32N, we PCR amplified the right end of the phage genome attachment site (attR site) using a primer pair that specifically anneals to the upstream region of the phage integrase gene (int) and within the guaA gene. The specific attR band was amplified from DNA isolated from the SPC32H lysis zone, whereas no band was detected using DNA isolated from the SPC32N lysis zone (Fig. 3B). The specific attR band was amplified from both colony and genomic DNA from the putative SPC32H lysogen [ΔLT2gtrABC1 (32H)] but not from DNA isolated from the parental Salmonella strain, SPC32H, or SPC32N (Fig. 3B). Furthermore, the SPC32H lysogen spontaneously produced phages that formed halo plaques during prolonged incubation (data not shown). These results clearly demonstrate that Salmonella can be lysogenized by SPC32H but not by SPC32N. The specific attR band was amplified from DNA isolated from the lysis zone of SPC32H m1 but not SPC32H m2 or m12 (Fig. 3C), confirming that m2 is the reason for phenotypic differences between SPC32H and SPC32N.

Because the phage repressor plays a critical role in the maintenance of the lysogenic state by repressing the expression of lytic genes and both phages have a putative repressor gene (rep), we assessed the deficiency of repression in SPC32N. When SPC32H and SPC32N were spotted onto lawns of a Salmonella strain overexpressing repLT2gtrABC1 + prep), the EOP of both phages was significantly reduced (<10−5 for SPC32H and <10−2 for SPC32N), and both strains exhibited a more turbid lysis zone than the control (Fig. 4A). Moreover, the specific attR band was amplified from DNA isolated from the lysis zone of both phages (data not shown), suggesting that supplementation with the repressor can promote lysogenic development in SPC32N. These results suggest that SPC32N is defective in maintaining lysogeny, most likely due to an insufficient amount of the active repressor.

Fig 4

The novel antirepressor, encoded by SPC32H_020 (ant), induces the lytic development of SPC32H. (A) Supplementation with the putative repressor leads to the lysogenic development of the lytic cycle-biased phage SPC32N, while supplementation with the putative antirepressor results in the lytic development of SPC32H. Salmonella strains transformed with a control plasmid (pBAD24), a putative repressor-overexpressing plasmid (prep), or an SPC32H_020-overexpressing plasmid (pant) were infected with serially diluted (10-fold) stocks of SPC32H or SPC32N. l-Arabinose (0.2%, final concentration) was added to induce SPC32H_020 expression from pant. (B) The expression of the SPC32H_020 protein promotes the switch from lysogenic to lytic development. The SPC32H lysogen [ΔLT2gtrABC1 (32H); SR5100] and nonlysogen (ΔLT2gtrABC1; SR5003) strains were transformed with pant or a control plasmid (pBAD24), and the resulting strains were subjected to a disc diffusion assay with 10 μl of 15% l-arabinose. pant* indicates the plasmid encoding a frame-shifted ant gene. Arabinose-induced bacterial lysis was observed only in the SPC32H lysogen harboring pant.

A novel antirepressor encoded by SPC32H_020 governs the lytic switch.The m2 mutation is located 24 bp upstream of the start codon of the hypothetical protein SPC32H_020, suggesting that m2 may cause the observed phenotypic differences by affecting the expression of this protein. SPC32H_020 is a small, 86-amino-acid protein with no known conserved domain or motif. A BLASTP search identified 40 hypothetical proteins with more than 64% identity with SPC32H_020 but did not identify any protein with a known function. The proteins exhibiting high homology to SPC32H_020 were from members of the Enterobacteriaceae, such as E. coli, Salmonella spp., Klebsiella spp., Citrobacter spp., and Cronobacter spp., and from ε15-like phages, including ε15, TL-2011b, phiV10, SPN1S, and SPN9TCW (see Table S2 in the supplemental material). Interestingly, some larger proteins (>218 amino acids) with a relatively low identity (<42%) were annotated as putative antirepressors, suggesting the possibility of an antirepressor role for SPC32H_020.

To determine whether SPC32H_020 functions as an antirepressor, we measured the prophage induction efficiency from a Salmonella strain harboring a SPC32H mutant which lacks the gene SPC32H_020LT2gtrABC1 (32H Δant) strain]. Compared with results for the WT phage lysogen, the spontaneous induction rate of the mutant phage lysogen was significantly lower (ca. 6 × 10−6-fold lower than that of the WT phage) (Table 4), indicating the critical role of SPC32H_020 in normal prophage induction. Furthermore, MMC treatment did not notably enhance induction of the mutant phage (1.11-fold increase) but did cause a 78.65-fold increase in induction of the WT phage (Table 4), suggesting a potential network between the SPC32H_020 gene and the host SOS response. Because the EOPs in Salmonella of the WT and mutant phage were similar (1.8 × 107 and 4.0 × 107 PFU ml−1, respectively), these results suggest that SPC32H_020 might act as an antirepressor. The function of the SPC32H_020 gene was further tested by a phage spotting assay using Salmonella harboring a plasmid overexpressing SPC32H_020 from a pBAD promoter (pant). As expected, both SPC32H and SPC32N generated clear lysis zones/plaques (Fig. 4A), and the lysogen-specific attR band was not PCR amplified from DNA isolated from the SPC32H lysis zone (data not shown). To verify the function of SPC32H_020 in lytic switching and prophage induction, an arabinose disc diffusion assay was conducted using Salmonella (ΔLT2gtrABC1) and the SPC32H Salmonella lysogen [ΔLT2gtrABC1 (32H)], both harboring pant. The SPC32H lysogen carrying pant underwent lysis in the presence of 15% arabinose, but no lysis was observed in the absence of pant (Fig. 4B). When the arabinose-inducible plasmid contained a frame-shifted ant gene (ant*), which was generated by inserting an additional adenine directly downstream from the SPC32H_020 start codon, the arabinose treatment did not induce lysis (Fig. 4B), suggesting that lytic switching is induced by the SPC32H_020 protein rather than the RNA. Taken together, our results strongly suggest that SPC32H_020 encodes a novel antirepressor protein that plays a significant role in the switch from the lysogenic cycle to the lytic cycle. We have annotated the SPC32H_020 gene as ant (antirepressor) and its gene product as Ant.

Table 4

Comparison of prophage induction efficiencies

The m2 sequence in the SOS box causes constitutive expression of the antirepressor by SPC32N.The results described above indicate that the m2 mutation may allow the overexpression of ant in SPC32N. Using the BPROM software program, the −10 and −35 sites of the putative ant promoter and one LexA-binding site (SOS box), which overlaps the predicted −10 site, were predicted in the upstream region of the ant gene (Fig. 2B). Intriguingly, m2 is located in the consensus LexA-binding site sequence (8, 33, 34) (Fig. 2C). LexA is a transcriptional repressor that represses various SOS regulons, including LexA itself and the RecA protein, via binding to the SOS box. DNA damage induces the formation of activated RecA nucleoprotein filaments that promote autocleavage of LexA and consequent derepression of SOS regulons (8). Therefore, we hypothesized that the ant gene is an SOS regulon controlled by LexA and that m2 in the consensus LexA-binding site sequence might prevent the LexA-mediated repression of the ant gene in SPC32N.

To test this hypothesis, we first examined the promoter activity of the ant gene in both SPC32H and SPC32N via a bioluminescence reporter assay using luxCDABE. In contrast to the low number of RLU (relative light units) detected using the ant promoter from SPC32H (Pant_H), the promoter from SPC32N (Pant_N) exhibited approximately 2-log higher values (Fig. 5A and B). Treatment with MMC significantly increased the RLU produced from a clone harboring pPant_H::lux but not from a clone harboring pPant_N::lux (Fig. 5A and B), suggesting that Pant_H expression was activated by DNA damage, whereas Pant_N was expressed constitutively and independent of DNA damage. To elucidate whether these responses were associated with LexA, we constructed Salmonella mutants without the lexA gene or expressing a noncleavable form of LexA [lexA(G85D)] and measured the bioluminescence from the reporter plasmid pPant_H::lux. Both mutants were constructed in a ΔsulA background to suppress the lethality of the lexA deletion (35). This sulA deletion did not affect reporter gene expression (data not shown). In the absence of lexA, Pant_H activity was comparable to that observed in the lexA+ background in the presence of MMC, and the Pant_H activity was not affected by MMC treatment (Fig. 5B, ΔlexA). In addition, the lexA+ phenotype was partially rescued by in trans complementation of lexA (data not shown). In contrast, replacing LexA with the noncleavable form of LexA prevented promoter activation by MMC [Fig. 5B, lexA(G85D)], indicating that DNA damage activates the ant promoter through LexA proteolysis. The results were similar regardless of the presence of the SPC32H prophage [Fig. 5B, lexA+ versus lexA+(32H)], suggesting that no other factors, including the SPC32H repressor, are involved in ant gene regulation, despite the presence of a repressor-binding site immediately upstream of the ant promoter (Fig. 2B; also see below).

Fig 5

The ant promoter of SPC32H is activated by DNA damage via LexA proteolysis, whereas the SPC32N ant promoter is constitutively active, due to the inability of LexA to bind to the m2-containing consensus LexA-binding site. The RLU (relative light units) were calculated by dividing the measured bioluminescence by the A600 value. The mean and SD for three independent assays are shown on a log scale on the y axis (A and B). (A) Time course observation of ant promoter activity in the presence or absence of DNA damage. Salmonella strains harboring the bioluminescence reporter plasmid pPant_H::lux (luxCDABE fused to the putative ant promoter of SPC32H) or pPant_N::lux (luxCDABE fused to the putative ant promoter of SPC32N) were incubated at 37°C, and the bioluminescence, as well as the A600 of the culture, was measured every half-hour. The vertical arrows indicate MMC treatment (1 μg ml−1, final concentration; 3 h after incubation). (B) ant promoter activities of the various Salmonella strains at an A600 of ∼0.6, harboring the bioluminescence plasmid. MMC (1 μg ml−1, final concentration) was added after 3 h of incubation. lexA+, ΔLT2gtrABC1, SR5003; lexA+(32H), ΔLT2gtrABC1(32H), SR5100; ΔlexA, ΔLT2gtrABC1 ΔsulA ΔlexA, SR5158; lexA(G85D), ΔLT2gtrABC1 ΔsulA lexA(G85D), SR5176. ∗∗∗, P < 0.001. (C) LexA specifically binds to the putative ant gene promoter region of SPC32H but not to that of SPC32N, which contains m2. The γ-32P-labeled DNA fragment of the ant gene promoter region from SPC32H (APRH*) or from SPC32N (APRN*) was incubated with the indicated amounts of purified Salmonella LexA and was subjected to an electrophoretic mobility shift assay (EMSA). Corresponding unlabeled DNA fragments (APRH and APRN) were used for the competition analysis. The position of the unbound fragments (F) and fragments retarded by LexA binding (B) are indicated.

We next performed an electrophoretic mobility shift assay (EMSA) to show the binding of LexA to the SOS box within Pant_H or Pant_N. When the radiolabeled DNA fragment APRH* (ant gene promoter region from SPC32H) was incubated with an increasing amount of purified Salmonella LexA, a specific mobility shift was observed, and the APRH* fragment was released by the addition of the unlabeled competing cold probe APRH (Fig. 5C, lanes 1 to 8). In contrast, the unlabeled cold probe APRN (ant gene promoter region from SPC32N) was unable to compete with APRH* for LexA, and the APRN* fragment was not shifted in the presence of LexA (Fig. 5C, lanes 9 to14), confirming that LexA cannot repress ant expression in SPC32N due to an inability to bind to the SOS box containing m2.

Based on these results, we investigated the overall cascade of SPC32H induction using Salmonella strains lysogenized by a derivative of SPC32H containing lacZ transcriptionally fused to the putative recET genes. Because the phage recE and recT gene products, a 5′ → 3′ exonuclease and a single-strand DNA binding/annealing protein, respectively, promote homologous recombination to mediate the integration/excision of phage genome to/from the host chromosome (36–38), the expression of recET (and its orthologous genes) can be used as a reporter for prophage induction. As shown in Fig. 6, treatment with MMC but not other antibiotics activated the recET::lacZ fusion in the lexA+ background but not in the lexA(G85D) background, indicating that the DNA damage generated by MMC induces SPC32H induction dependent on LexA proteolysis. The expression of Ant clearly resulted in lysogen-specific lysis (Fig. 4B), supporting the idea that derepression of the ant gene via MMC-induced LexA proteolysis leads to phage induction. Taken together, these results demonstrate that the ant gene of SPC32H is negatively regulated by LexA and that m2 in the SOS box causes the dramatic phenotype differences between SPC32H and SPC32N by influencing ant expression.

Fig 6

DNA damage-induced LexA proteolysis followed by SPC32H ant expression induces the switch to lytic development. The lacZ gene, transcriptionally fused to the putative recET genes, was introduced into the SPC32H lysogens harboring an intact (lexA+) or noncleavable [lexA(G85D)] LexA, and the resulting strains were subjected to a disc diffusion assay with the following solutions: MMC, 0.5 mg ml−1 mitomycin C; Cm, 2.5 mg ml−1 chloramphenicol; Ap, 10 mg ml−1 ampicillin; D.W., distilled water. Note that the blue zone appears to surround the MMC disc in the lexA+ background only.

The antirepressor Ant interacts directly with the cognate repressor Rep.The putative repressor from SPC32H (designated Rep), encoded by SPC32H_041, is a 198-amino-acid protein that contains a helix-turn-helix motif. The RecA-mediated autocleavage site (Ala-Gly or Cys-Gly), a highly conserved site in cleavable repressors such as lambda CI (39), is not present in SPC32H Rep, strongly supporting the notion that SPC32H prophage induction involves the inhibition of Rep through means other than autocleavage assisted by RecA nucleofilaments. The immunodetection of HA epitope-tagged Rep demonstrated that the expression level of Rep remained virtually constant (i.e., was not cleaved) throughout a 1-h treatment with MMC (Fig. 7A). The lytic switch was activated by MMC in this experiment, as shown by the fact that HA-tagged Ant was expressed and accumulated after treatment with MMC (Fig. 7A, lower panel). The HA-tagged versions of the Rep and Ant proteins are fully functional (data not shown).

Fig 7

DNA damage induces Ant accumulation but not Rep cleavage, and the consequent binding of Ant to Rep inhibits the binding of Rep to specific operators. (A) Salmonella strains lysogenized by SPC32H expressing HA-tagged Rep (upper panel; ΔLT2gtrABC1 [32H rep-HA], SR5192) or both HA-tagged Rep and HA-tagged Ant (lower panel; ΔLT2gtrABC1 [32H rep-HA ant-HA], SR5197) were exposed to MMC for 1 or 2 h, respectively. The MMC-treated bacterial cultures were sampled at the indicated time points and subjected to the Western blotting to immunodetect the HA-tagged proteins. DnaK was used as an internal control. (B) Bacterial two-hybrid assays revealed the direct binding of Ant to Rep. The β-galactosidase activity of E. coli BTH101 reporter strains harboring the indicated plasmid pairs were measured. The activities are presented in Miller units. B, a backbone plasmid. (C) EMSA with purified Rep and Ant demonstrates the Ant-mediated inhibition of Rep binding to its operators. Mixtures of APRH* and the indicated amounts of Rep were incubated at 20°C for 15 min in 1× binding buffer supplemented with 1.1 μg of poly(dI-dC) and then electrophoresed on a 6% native acrylamide slab gel for EMSA. For competition analysis, unlabeled APRH fragments were added as cold probes to the mixture. When appropriate, Rep was preincubated with the indicated amounts of Ant at 20°C for 30 min and further incubated with APRH* as described above. The positions of the unbound fragments (F) and fragments retarded by Rep binding (B1 and B2) are indicated.

To explore the possible interaction between Rep and Ant, we performed a bacterial two-hybrid assay based on the restoration of β-galactosidase activity in E. coli cyaA mutant strain BTH101 (29). The reporter strain E. coli BTH101 expressing the combination of hybrid proteins (i.e., T25-Rep/T18-Ant or T25-Ant/T18-Rep) produced blue colonies on X-Gal plates (data not shown) and exhibited a significantly higher (approximately 10- to 50-fold) level of β-galactosidase activity than the negative control (i.e., E. coli BTH101 expressing the unfused T18 and T25 peptides) (Fig. 7B), indicating a heterodimerization of the hybrid proteins via interaction between Rep and Ant. Strong Rep-Rep and Ant-Ant interactions were also observed, implying the possibility of multimerization by each protein. Indeed, the results of analytical size exclusion chromatography demonstrated that Rep and Ant were able to dimerize and tetramerize, respectively (data not shown). Taken together, these results suggest that Rep and Ant can interact with themselves and each other.

EMSA using purified Rep and Ant revealed that Ant inhibits Rep target site binding. The high homology (97% identity) between SPC32H Rep and the ε15 repressor suggests that Rep may recognize the same DNA sequence (5′-ATTACCnnnnGGTAAT −3′) as the ε15 repressor. Radiolabeled APRH*, which includes the putative repressor-binding site as well as the SOS box, was also used in this assay. Two DNA-protein complex bands with different mobilities appeared when purified Rep was incubated with APRH* (Fig. 7C, lanes 1 to 4), suggesting that APR may have two Rep-binding sites with different affinities for Rep. A competition assay using a nonlabeled cold probe demonstrated the specificity of Rep-binding for APR (Fig. 7C. lanes 5 and 6). Notably, preincubation of Rep with purified Ant prevents the mobility shift of the APRH* fragment in an Ant concentration-dependent manner (Fig. 7C, lanes 7 to 9), suggesting that the specific interaction between Ant and its cognate repressor Rep interferes with Rep binding to its target DNA. Note that the protein concentrations indicated were calculated based on the assumption that the Rep and Ant stocks consisted entirely of active dimers and tetramers, respectively. The APRH* fragment clearly did not exhibit a mobility shift when incubated with Ant alone (Fig. 7C, lane 10), excluding the possibility that Ant inhibits Rep activity by competing for the Rep-binding site with Rep.

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The goal of this study was to determine the cause for the phenotypic differences between two highly similar podoviral ε15-like phages, SPC32H and SPC32N. We detected two nucleotide differences between the two phage genomes, but only one, located within a noncoding region, was responsible for the phenotypic differences. This nucleotide polymorphism, m2, was located within a consensus LexA-binding site sequence that overlaps the −10 site of the promoter for SPC32H_020, which encodes a hypothetical protein (Fig. 2). This sequence difference prevents LexA from binding to its binding site, allowing the constitutive expression of a small hypothetical protein (Fig. 5), which we have identified as a novel antirepressor of the family Podoviridae. This antirepressor inhibits the binding of its cognate phage repressor to regulatory regions (Fig. 7C), resulting in a switch of the phage life cycle from lysogenic to lytic.

To date, at least two categories of lytic switch antirepression systems have been identified in temperate phages. The first system is represented by the Cro protein of several lambdoid phages, such as phage lambda, HK022, and HK97. In this system, the binding of the Cro protein to target operator sites prevents expression of the cI gene, which encodes the phage repressor CI (40). In contrast, the second system controls repressor activity at the protein level. For example, the antirepressor Tum from myoviral coliphage 186 binds directly to the phage repressor CI, preventing CI from binding to its operator sites (12). Notably, the latter system has been found in only a few temperate phages, including siphoviral coliphage N15 (11) and the siphoviral prophages Gifsy-1 and Gifsy-3 identified in S. Typhimurium strain 14028 (10). In the present study, we have elucidated the mechanism and regulation of an antirepressor, which belongs to the second category of lytic switch antirepression systems and is the first example of this type of system in the Podoviridae family of the order Caudovirales. A notable common feature of this second system type is the LexA-regulated initiation of antirepressor expression. Although the phage P22 also produces an antirepressor that inactivates the c2 repressor and prevents RecA-dependent c2 proteolysis (41, 42), it is unknown whether LexA regulates the expression of the antirepressor. However, the presence of a consensus LexA-binding sequence 38 bp upstream of the start codon of the antirepressor protein suggests that LexA may be involved in the regulation of the P22 antirepressor and consequently that the P22 antirepressor may be a member of the second category of antirepression systems.

Linking the host SOS response to the lytic switch is a fundamental strategy used by prophages to escape from damaged host cells. Compared to the RecA-dependent cleavable repressor system, such as the lambda CI (7, 40), the antirepression system appears to be more advantageous to the prophages. If host bacteria are able to repair DNA damage before prophage induction and survive (43), it would be more beneficial for the prophages to remain in the host cell. If lysis occurred, the induced phages would need to reestablish the prophage state in new host bacterial cells to stably maintain their genome as a part of a host genome. This superfluous step could easily be prevented by expressing the antirepressor in a LexA cleavage-dependent manner. Since antirepressor levels are reduced by the replenished LexA pool, lysogenic development could resume because inactivated, rather than degraded, repressors can be restored to function by dissociating from the antirepressor. Although we did not demonstrate the reversible binding of Ant and the recovery of Rep activity after Ant dissociation in the present study, Rep levels were stably maintained without degradation during MMC treatment (Fig. 6), suggesting that recycled Rep could be used in the resumed SPC32H lysogenic development. Indeed, the antirepressor Tum/repressor CI pair from coliphage 186 exhibits reversible Tum binding and the recovery of CI activity after dissociation from Tum (12). We are currently attempting to elucidate this issue by investigating the structure of the Rep-Ant complex as well as the individual proteins.

Considering the advantages of rapid resumption of the regulatory circuit, it is possible that this type of repressor/antirepressor system is widespread among the temperate phages. Remarkably, several homologues of SPC32H Ant (38 to 100% amino acid similarities) were identified in other Podoviridae phages and various bacteria in the family Enterobacteriaceae (see Table S2 in the supplemental material), most likely as a gene product of unknown function of prophages. Moreover, the phage antirepressors Tum (from Myoviridae coliphage 186), AntC (from Siphoviridae coliphage N15), GfoA (from Siphoviridae phage Gifsy-1), and Ant (from Podoviridae phage SPC32H) are distinct from each other at the amino acid sequence level (Fig. 8), suggesting that diverse repressor/antirepressor pairs are present in the order Caudovirales to allow for more prudent control of lytic/lysogenic switching. Therefore, as suggested by Mardanov and Ravin, the cleavable repressor system may not be the exclusive mechanism for lytic/lysogenic regulation in temperate phages (11). As recently illustrated by Lemire et al., antirepressors can mediate cross talk between prophages in polylysogenic strains (10). Thus, further studies regarding the trans activity of diverse phage antirepressors, including SPC32H Ant, would provide insight into the coordinated behavior of temperate phage subversion of their bacterial prey.

Fig 8

Amino acid alignment of the phage antirepressors. The amino acid sequences of Tum (from coliphage 186), AntC (from coliphage N15), GfoA (from Gifsy-1) and Ant (from SPC32H) were aligned using ClustalW2. There are no noticeable consensus residues, demonstrating the diversity of phage antirepressors in the order Caudovirales.

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This work was supported by a National Research Foundation of Korea (NRF) grant funded by the Ministry of Education, Science and Technology (no. 20090078983).


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Bacteriophages or phages are the most abundant organisms in the biosphere and they are a ubiquitous feature of prokaryotic existence. A bacteriophage is a virus which infects a bacterium. Archaea are also infected by viruses, whether these should be referred to as ‘phages’ is debatable, but they are included as such in the scope this article. Phages have been of interest to scientists as tools to understand fundamental molecular biology, as vectors of horizontal gene transfer and drivers of bacterial evolution, as sources of diagnostic and genetic tools and as novel therapeutic agents. Unraveling the biology of phages and their relationship with their hosts is key to understanding microbial systems and their exploitation. In this article we describe the roles of phages in different host systems and show how modeling, microscopy, isolation, genomic and metagenomic based approaches have come together to provide unparalleled insights into these small but vital constituents of the microbial world.

Key words: bacteriophage, ecology, cyanophages, archaeal viruses, animal microbiome


We live in a microbial driven world that only exists because Bacteria and Archaea tempered the previously hostile environment on early Earth to create atmospheric conditions that allow eukaryotic life forms to flourish. Bacterial and archaeal encoded enzymes catalyze all the major processes involved in global biogeochemical cycling, playing key roles in the carbon and nitrogen cycles, and producing approximately half of the oxygen in the Earth's atmosphere.1 In macro-organisms (animals) prokaryotic cells generally outnumber eukaryotic cells, where they assist in important aspects of survival such as nutrition and defense. So what roles are phages playing in this microbial mix? Once ignored, it is now becoming increasingly accepted that phages play key roles in the biology of microbes, which themselves impact environments at large.24 Many previous excellent reviews have highlighted the importance of bacteriophages in specific environments for example.57 In this article we present three case studies to illustrate how an appreciation of the roles of the viruses is pertinent to understanding microbial physiology, population dynamics and evolution. We show how our microbial driven world is tempered by bacteriophages. To contextualize the case studies we summarize the history of phage research and give an introduction to the biology of bacteriophages. We review their distribution and describe how they are enumerated and characterized. Finally we discuss the ways in which phages may influence their host's evolution and population dynamics.

Brief history of bacteriophage discovery and research.

Bacteriophages were first discovered in 1915 by William Twort, and in 1917 by Felix d'Herelle realized that they had the potential to kill bacteria. After a pre-antibiotic era heyday they were then essentially disregarded as significant therapeutic agents in the West, primarily due to the comparative ease by which antibiotics could be administered. Research and the practice of using bacteriophages did continue in some countries such as Georgia (as part of the former USSR), where they were, and continue to be routinely isolated and used to treat a large number of diseases.8 Bacteriophage research then focused on a number of model phages which primarily infected E. coli. These studies provided the back-bone of modern molecular biology, for example phages were used to identify the basis of genetic material, and that 3 nucleotides code for an amino acid.9 They also allowed the identification of restriction enzymes.9 For several decades, only a handful of phages were studied in great detail. The recent renaissance seen in phage biology has been triggered due to a growing awareness of the number of phages in all bacterial dominated environments (as revealed by epiflourescent and electron microscopy, and from molecular studies), and indeed in the genomes of bacteria following whole genome sequencing projects. This checkered history has resulted in a patchy knowledge of phage biology but with enough observations for scientists to realize that phages are dictating many aspects of Bacterial/Archaeal biology. These observations have invigorated an invigorated interested in bacteriophages, and are part of the stimulation for this journal Bacteriophage, in which this article is written to illustrate the roles that bacteriophages play in the natural world.

Phage life cycles.

In order to appreciate the roles of phages in nature, an understanding of their possible interactions with their hosts is necessary. Phages have various possible life cycles which, along with interaction with their physical environment, dictate their role in bacterial/archaeal biology. The lytic life cycle is where phages infect and rapidly kill their infected host cells, thereby shaping bacterial population dynamics and occasionally assisting in their long term evolution via generalized transduction.24 The lysogenic life cycle in contrast, is where phages instead of directly killing their hosts, integrate into their host genome, or exist as plasmids within their host cell.10 This lysogenic life cycle can be stable for thousands of generations and the bacteriophage may alter the phenotype of the bacterium by expressing genes that are not expressed in the usual course of infection in a process known as lysogenic conversion. A well known example of this is the gene associated with Vibrio chlolerae which encodes the toxins that cause cholera symptoms.11 Phages may also have a psuedolysogenic component to their life cycle. This is a controversial concept, and has many different definitions within phage biology.2 We define it here as the situation that occurs when a phage has entered a bacterial cell and doesn't integrate in a stable fashion, but will stay in this ‘mode’ until conditions occur which trigger them to enter into the lytic or lysogenic life cycle.2,12 We illustrate how pseudolysogeny is difficult to study, yet may be important in markedly different systems. Finally there is the chronic infection lifestyle found in some archaeal viruses, in filamentous phages (rod shaped single stranded DNA phages), and in plasmaviruses which infect Mycoplasma. In this life cycle phages are slowly shed from the cell over a long time period without obvious cell death.

Phage abundance and diversity.

Having considered the possible phage life cycles, it is logical to review where phages are found, and how they are enumerated and characterized. Specific examples of phages and their characterization will then be given in the case studies. The first approaches that led to the realization of phage abundance were based on epifluorescent microscopy following DNA staining which suggested that in sea water there are around 10 phages in existence for each bacterial/archaeal cell.7,13,14 Similar figures have been shown for freshwater environments, but for other more complex environments the situation is less clear and virus numbers may either be higher or lower than that of their bacterial/archaeal hosts.15

As phages have an obligate requirement for a host, their abundance and distribution is likely to be based on that of their host organisms. Therefore to make sense of viral abundance, one must establish where the majority of their hosts exist. Although we are often focused on bacterial pathogens, most of the Earth's Bacteria and Archaea are found in the open ocean, the soil and in ocean sediments, and terrestrial sub-surfaces where there are an estimated 1.2 × 1029, 2.6 × 1029, 3.5 × 1030 and 0.25–2.5 × 1030 cells respectively.16 Bacteria and Archaea are often associated with humans and animals which provide many niche environments within them, often where these micro-organisms have become an essential symbiont. Despite animal abundance on the Earth the total number of prokaryotes associated with them is several orders of magnitude less than for the major land and ocean environments. For example, in humans the majority of prokaryotes are found in the colon, so multiplying the total human population of 6.8 × 109 by the number of prokaryotes per gram of human colonic matter (3.2 × 1011), by the average amount of colonic material per human of 220 grams, gives an estimated total of 4.8 × 1023 prokaryotes (based on ref. 14, and on the UN 2009 current estimate of the human population size). Although not as numerically significant, bacteria are of essential importance when associated with humans, particularly either in a disease, or a food producing context when bacteria are associated with causing disease, or where we are reliant on bacteria, for example for cheese production, which can fall prey to bacteriophage attack. Therefore, in terms of human impact, an appreciation of the roles of bacteriophages which infect these bacteria is of paramount importance.

Traditional approaches to the quantification and characterization of bacteriophages.

There is no single method that can be used to establish how many phages in an individual sample can infect a specific host however traditional and molecular approaches can be usefully combined to build up a picture of the viral community. The number of phages which infect all hosts can be determined using epifluorescent microscopy, or flow cytometry17,18 and the morphological diversity of phages using transmission electron microscopy (TEM).19 Currently the number of phages which infect specific hosts can only be determined from isolation approaches.20 For isolation studies, suitable hosts can either be isolated specifically from the environment of interest, or a model permissive host can be used. Clearly these approaches only identify phages that infect the specific strains being used as a host and so it is difficult to establish what proportion of phages present are being isolated. Phages may be present which infect the species being used in isolation, but may not infect the model strain if the strain lacks the appropriate phage receptors, has a restriction system, if abortive infection occurs, or if it has a CRISPR (clustered regularly interspaced short palidromic repeats) defense system.2123 Furthermore, phages isolated are amenable to propagation, and not necessarily representative of the most abundant phages in natural populations. It is hoped that future work based on single cell sequencing will provide additional data on the noncultureable viruses that are important in natural populations.

Molecular approaches to the quantification and characteriszation of bacteriophages.

There is no universal marker for phages in the same way the 16S rRNA gene can be used to reliably place the phylogenetic affinity of all bacteria. This is because there no genes that are suitably conserved within all phages, or even for example present within one taxonomic group such as the bacterial virus order Caudovirales.24 However, there are several examples of smaller taxonomic group specific markers, which are extremely useful for assessing phage diversity and abundance. For example, researchers commonly target genes, which encode structural proteins as phylogenic markers. One gene which has been widely used is the gene which encodes the portal protein which is located at the top of the neck of the phage and through which DNA passes en route down the tail sheath.2528 The same primer sets have been used to investigate these sequences in T4-type phages which are known to infect a wide range of bacterial hosts.29 As well as providing estimates of diversity, molecular markers can also offer new ways of quantifying bacteriophage abundance which are free from the isolation based complications discussed in the paragraph above. For example molecular markers based on the Q gene and a gene encoding for a shiga toxin, revealed a far greater abundance of shiga toxin phages present in the soil than was observed using standard isolation based approaches.30

Other molecular approaches to assessing bacteriophage diversity are where markers based on restriction fragment length polymorphisms (RFLP), or on denaturing gradient gel eletrophoresis (DGGE) are used to assess the diversity of a bacteriophage genome, or of a particular gene respectively. Examples of how these techniques have been effectively used are given in the cyanophage case study, and in the animal phage case studies.


The newest way of assessing phage diversity and indirectly abundance is using viral metagenomics. This is where the total viral component from a particular environment is collected and sequenced. This approach has been made possible due to the progress in sequencing technology, and the reduction in cost which has made it relatively affordable. Protocols vary according to the sample in which the phages are present but bacteria are always removed, and often where the total amount of viral DNA is low, enrichment steps are carried out to amplify the total viral community DNA so there is enough to sequence.3133 Metagenomics can be used to identify phages or phage genes of environmental significance, such as those that are highly abundant or specific to particular niches.3436 This allows data to be collected on the dominant viral genomes present in a specific location, without having to culture their hosts and isolate phages, and provides a great starting point for understanding the roles that bacteriophages may be playing. It can also provide information on phages that are not amenable to propagation, or that do not have hosts in culture. It is estimated that 95% of bacteria cannot be cultivated under laboratory conditions, so consequently the phages which infect them cannot currently be isolated either.37,38 Metagenomics can also potentially provide abundance information based on the amount of coverage of particular phages/gene sets present in sample sets. Clearly there may be amplification, or sequencing bias but over, or under representation of particular genes can yield useful information about phage biology.

The main current drawback to metagenomic studies is that because viral genome diversity is so high, a large proportion of predicted genes are ‘unknown’ or ‘hypothetical’, and therefore currently much of the information collected by this approach is not immediately useful. This situation will improve as additional genomes from isolated bacteriophages are sequenced and annotated, and as bioinformatic tools based on structural protein homologies are developed to assist amino acid or nucleotide sequence comparisons. Finally, it often is difficult to test hypotheses made from sequence data. Although genes of interest from metagenomes can be cloned, expressed and biochemically characterized, their relevance to specific phages can only be established if cultured phages with those genes are in existence.

The big picture; phage abundance and diversity studies.

While each enumeration/identification technique contributes one piece of information to our understanding of phage abundance and diversity, few studies attempt all methods in combination. Fewer still collect the necessary metadata needed to establish phage number and hosts identity. In other words most studies either count total phage number, or identify a sub-set of phages which are associated with one host bacterial strain. However, each approach has merit, and the body of research based on them is gradually improving our understanding of the phage world. It is really an exciting time to be a phage biologist because very few environments have been well characterized and those that have, have revealed endless surprises in terms of gene content. Therefore, it is likely that significant surprises and unexpected findings will abound as new systems are studied.

An example of the synergy of different approches can be seen in a recent study which compared cyanophage genomes present in the large scale metagenomes present in the ocean derived CAMERA (Community Cyberinfrastructure for Advanced Microbial Ecology Research & Analysis) data set to those found in culture. Rather pleasingly, the data in the metagenomes reflects the gene content and diversity of the cyanophages that are in culture.39 Some studies have used multiple methods to characterize viral assemblages; for example a recent study used EM, flow cytometry and metagenomics to characterize the viral community associated with Antarctic lakes throughout an annual cycle. These combined approaches revealed many interesting features of the virus communities in Arctic systems such as the shift from a ssDNA virus dominated community in spring when lakes are generally iced over, to a dsDNA dominated community in the summer.40

Bacteriophage biogeography and persistance.

Recent evidence suggests that prokaryotes may exhibit biogeography i.e. be endemic to particular environments which goes against the idea that “everything is everywhere.”41,42 This idea leads to the possibility that phages can also show biogeography. Studies have shown that some phage have a global distribution while others may be endemic to particular environments. A recent review on this subject reports that a 2009 meeting of the Scientific Committee on Ocean Research Viral ecology Working Group concluded that this question remained unanswered.43 In this review we consider this notion more under our case studies.

Generally phages are pretty stable if the environment is not hostile. They are broken down in UV light, and can be damaged by abrasion, or exposure to chemicals, but researchers have been known to keep phages in their fridges for over 40 years with no reduction in titre (Ackerman HW, personal communication). Unpublished work by Suttle, and by Clokie has demonstrated that cyanophages can be isolated from sediments that are several decades old (Clokie MRJ, unpublished). Finally some bacteriophages seem remarkably unstable in the laboratory and Clokie et al. have observed that both Clostridium difficile bacteriophages, and those which infect Streptococcus pneumoniae reduce in titre on a weekly basis regardless of the buffer/media they are stored in (Clokie MRJ et al., unpublished).

Impact of phages on host populations.

Having defined the possible life cycles and ascertained what is known about phage diversity and abundance, it is pertinent to review what is known about how bacteriophages impact their host populations. Several approaches have been used to determine the impact phages have on their host's populations. Experimental evidence from chemostats and observations of phages/hosts in open systems have shown that for some bacterial species, populations of phages and hosts oscillate with time.44,45 The relationship between phages and their hosts has been modeled, and in a simple environment if there is no cost to host resistance the same oscillation in populations occurs.46 However, if there is a cost to phage resistance, then bacteriophages have been theoretically and experimentally shown to drive host diversification.4749 Diversification of bacteria may occur in the phage receptor region, which may be related to nutrient uptake, or it may occur possibly on a faster timescale on CRISPR elements that can quickly evolve to provide a host defense system.50 An alternative dynamic between hosts and phages exists where phages have a temperate life cycle whereby phages may contribute to the success of their host bacteria by encoding useful genes; examples of this dynamic have been hypothesized following many bacterial genome sequencing projects, and through recent metagenomic studies.35,36

To illustrate the major roles that phages play in microbial ecology, physiology and evolution we describe in detail case studies from three contrasting systems of phages in their natural settings. The first case study describes marine cyanophages because they have been the most extensively studied phages in the marine environment, and, at a broader level, they probably constitute the group of phages that has been studied from the greatest number of perspectives and therefore significant data exists to begin to unravel their roles in cyanobacterial biology. They have been isolated from around the world, have been the attention of 10s of genome sequencing projects, and much metagenomic sequencing effort, their impact on host dynamics has been studied in natural systems and expression work and modeling have begun to establish the biological significance of particular features of their genomes. The second case study reviews our knowledge of the roles that phages play in animal bodies. Despite this environment being closer to home, the complex nature of the environments, and a surprising paucity of research effort mean that it is less well understood. We predominantly focus on phages which infect gut E. coli (coliphages) as they are abundant in animal bodies and enough significant data exist on them for an understanding of their relevance to be postulated. Finally the we discuss the ‘phages’ which infect the Archaeal domain as an example of how little is known about this intriguing group of organisms. Within each case study we briefly summarize the history of research in the field and describe how the phages have been isolated and characterized. We describe the impact of molecular and metagenomic studies on the fields with a view in all three systems to ascertaining the impact of phage on their bacterial/archaeal hosts, how they shape and control their host populations. In all systems, we also highlight the areas where conflicting data, or lack of it means that the role of phages has yet to be established. We hope that the concepts brought out in this review will form an appropriate framework which will be helpful when considering novel, or less studied groups of phages, or when considering the exploitation of phages.

Case Study: Marine Cyanophages

Cyanophages: Discovery and distribution.

Although several heterotrophic phages have been studied, this review focuses only on the cyanophages for which most data is available for. Their study in the marine environment began in the early 1980's when phages infecting both unicellular and filamentous cyanobacteria were observed in the Black Sea.51 In the early 1990s, research into began in earnest with the isolation and characterization of phages infecting marine Synechococcus.5254 Ten years later cyanophages that infect the closely related Prochlorococcus were also isolated.55 These two genera of cyanobacteria are the predominant primary producers of the nutrient poor (oligotrophic) areas of the ocean which cover around 70% of the surface of Earth. Remarkably, these two genera Synechococcus and Prochlorococcus account for up to 50 % of primary production the world's oceans, Prochlorococcus is generally found in the warmer oceanic waters between the 40° latitude north and 40° latitude south whereas Synechococcus is much hardier and is found on either side of those latitudes.56,57 Therefore the study of the viruses which infect these organisms has significant ecological interest in terms of global carbon cycling.

Isolation and characterization.

The early work on cyanophages concentrated on determining their abundance in the environment using isolation based approaches, and then characterising the isolated phages using well or plate assays to determine host range and burst size.53,58 These studies revealed that cyanophages are widespread in the environment, at concentrations as high as 1 × 106 pfu ml−l.58 They all had relatively long latent periods of several hours and their burst sizes range from ∼20 for the cyanophage S-PM2 59 to ∼250 for cyanophage S-BBP1.53 TEM analysis has revealed a large diversity of cyanophage morphotypes, with phages observed from the Myoviridae, Podoviriade and Siphoviridae.5254 Most cyanophages that have been isolated to date however belong to the Myoviridae, and are morphologically similar to T4-like phages.60

Molecular approaches to cyanophage study.

An important milestone in the study of cyanophages was the development of molecular tools to study their distribution and diversity. The discovery by Fuller et al. that cyanophages have a gene that is homologous to g20 in T4, was surprising at the time, and allowed the development of a set of PCR primers to specifically amplify this gene from cyanophages.61 Several studies using this initial primer set, or improvements on it, to target g20 from a wide range of geographical locations, and ocean habitats.27,6266 A similar approach was taken to target the DNA polymerase gene of cyanophages from the Podoviridae family.67 Both of these studies have revealed how widespread cyanophages are within the oceans, and have revealed a clear lack of cyanophage biogeography based on either gene.

Surprising synteny and homology between cyanophages and enteric virues.

The work of Hambly et al. extended the work above by sequencing the genes adjacent to g20 and to show that the cyanophage S-PM2 shares a conserved module of genes with the model E. coli phage T4.68 The subsequent sequencing of cyanomyovirus genomes has revealed that there is an even greater amount of synteny and homology between them and T4.6972 Although this may seem like a diversion from the main thread of examining bacteriophage roles in nature, it suggests that bacteriophages do share a commonality in terms of the way in which they function in many bacterial groups. Figure 1 shows a dendrogram describing the relationship between cyanophages and other myoviruses. This is based on the presence or absence of specific genes. Although it clearly demonstrates that cyanophages are more closely related to each other than to other T4-like phages, they do share some genes with the T4-like phages KVP40, RB49 and T4. Podoviruses which infect cyanobacteria are also related to enteric phages with the genomes of cyanophages P60 and P-SPP7 being T7-like in terms of architecture and gene content.70,73 However, this appears not to be the same in the siphoviruses. For example the genome of the siphovirus P-SS7 that infects Prochlorococcus is very distinct from other lambdoid like phages.74 This is the only Prochlorococcus siphovirus reported so far, others may of course be different and share similarity with other known siphoviruses.

Cyanophage genomes.

The availability of sequenced genomes has allowed detailed genome comparison studies of cyanophages. Twenty-one marine cyanophages have been sequenced to date; fifteen myoviruses, five podoviruses and a siphovirus.7580 This number is expected to rise rapidly in the next few years; for example one initiative being funded by the Gordon and Betty Moore Foundation is sequencing dozens more cyanophage genomes and viromes (all viruses in a habitat). This is likely to provide major insights into the biology of these organisms, and to highlight further the roles phages are playing in host biology.

Impact of cyanophages on the physiology of their hosts during infection.

Clues to how cyanophages influence the biology of their hosts have been obtained from gene expression studies of infected model systems. Research has shown that while cyanomyoviruses share many genes with T4-like phages, they do not have the same expression patterns. T4 has a beautifully choreographed expression pattern with genes transcribed with early, middle and late profiles. Cyanophages lack a “middle” mode of transcription pattern, and consistent with this they lack the middle promoter activational genes MotA and coactivator AsiA which control the middle expression genes in T4.81,82 The cyanopodoviruses also encode and express a number of genes that are homologues to host genes.81,83 The Prochlorococcus cyanophage P-SSP7 has been demonstrated to increase the expression of a number of host-encoded genes during infection. This observation is thought to result from both the host stress response to infection, and to activation by phage encoded factors.

‘Host’ encoded phage genes illustrate the intertwined relationship between phage and host.

A number of genes have been found in cyanophages that are homologous to genes found in their cyanobacterial hosts. These genes are often referred to as “host genes” or more recently AMGs (auxiliary metabolic genes).84 Among the most interesting AMGs identified in cyanophages are the genes involved in photosynthesis which has led to some cyanophages being referred to as “photosynthetic phages”. S-PM2 was the first cyanophage discovered to carry the essential photosynthesis genes psbA and psbD which encode the D1 and D2 proteins respectively.76,85 The D1 and D2 proteins form a heterodimer at the core of photosystem II. An unavoidable consequence of oxygenic photosynthesis is the release of reactive oxygen species, that can damage the PSII complex, specifically the D1 polypeptide.86 As a consequence of this, a repair mechanism has evolved in all oxygenic phototrophs, which removes and replaces the damaged D1 protein.86 It is postulated that the expression of phage encoded D1 protein ensures a source of energy for phage replication, by maintaining photosynthesis after host protein synthesis has ceased to be expressed.87 This is supported by the fact that both phage encoded psbA transcripts81,83,88 and the corresponding D1 polypeptides are increased during the infection cycle. Furthermore recent modeling studies using in silico deletions of psbA from P-SSP7 predicts that under highlight conditions there is an increase in burst size for phages carrying psbA.89,90

Genome sequencing, PCR screening and comparative genome hybridization have revealed that psbA and psbD are widely distributed in cyanophage isolates and the environment at large.9195 Although psbA and psbD are widespread, they are not found in all cyanophages, and only psbA is thought to be part of the “core-genome” of cyanomyoviruses.71,78 The siphovirus P-SS2 lacks both psbA and psbD; at present it is not known if this is common feature of siphoviruses.74

Cyanophage genomes contain a many other ‘bacterial’ genes that might maintain or alter host physiology.39 Many of these genes are thought to have been acquired as a response to the harsh environment that cyanophages inhabit; the high-light oligotrophic open ocean. Other genes linked to high light conditions include the high-light inducible (hli) genes,69,88 a PTOX-encoding gene postulated to provide an alternative mechanism to reduce photo-damage,71,72,78 electron transport protein encoding genes71,72 and genes encoding proteins required for synthesis of the light-harvesting phycobilisomes complex.96 Additionally several genes that encode enzymes found in the pentose-phosphate pathway have been found to be common in cyanophages and are thought to allow optimization of necessary NADH and ATP for phage replication.71,78 A phosphate transport encoding gene found only in cyanophages isolated from phosphorous limited waters78 provides further evidence that the environment shapes cyanophage genomes to have several features that may benefit them.

Metagenomic studies have extended observations of ‘bacterial’ genes in phage genomes, and they have shown the vast extent to which the cyanobacteria and phages have evolved together.39,97

Life cycles and marine cyanophages.

P-SS2 is also the first marine cyanophage that has been isolated and had its genome sequenced that is thought to be temperate.74 At present the overwhelming majority of cyanophages that have been isolated are obligately lytic.60 This is somewhat surprising as there are numerous reports of lysogeny occurring in the environment in cyanobacteria.98100 However, despite the sequencing of over 20 Synechococcus and Prochlorococcus genomes, no complete prophages have been identified.101,102 Interestingly, McDaniel et al have reported the isolation of a putative temperate phage induced from Synechococcus, however it has a ssDNA genome which is unlike all other cyanophage isolates.99 The bioinformatic analysis of the P-SS2 genome sequence suggests that it capable of intergration into a host genome, although this has not been experimentally demonstrated.74

Temperate cyanophages are either rare, or just inherently so unstable to work with that we are just not able to isolate them well yet, and therefore we do not appreciate their roles in cyanobacteria. The lack of isolated cyanobacterial lysogens means that nothing is known on how cyanophages may change host physiology via lysogenic conversion. Consistent with no isolated cyanobacterial lysogens, no prophages have been observed in cyanobacterial genomes as there are for heterotrophic bacteria.102,103

The psuedolysogenic life cycle may be ecologically relevant in cyanophages. It has been shown that when cyanobacteria are grown in phosphate deplete media (as they would often be growing in the open oceans), then cyanophages enter the cells but do not enter the lytic cycle.12 Although the genetic nature of this interaction has not been characterized, the observation is consistent with Abedon's definition of pseudolysogeny being a carrier state where ‘lytic’ phages can remain inside bacterial cells until conditions are suitable to enter most probably the lytic infection cycle.104 Unpublished data has shown that in this psuedolysogenic state AMG's are expressed at a higher level than structural phage genes (Clokie MRJ et al., unpublished).

Influence of cyanophages on cyanobacterial populations.

A study over an annual cycle in the Red Sea showed that cyanophages co-vary in abundance and genetic diversity with that of Synechococcus, which is consistent with the hypothesis that cyanophages are an important factor in controlling cyanobacterial secondary ecological succession.63 The diversity of phages was estimated using DGGE following DNA extraction from the viral and host fraction of the water samples. Similar studies have indicated that cyanophages are involved in structuring the population dynamics of their host community, and driving biogeochemical cycling.105 Also consistent these observations is a laboratory based study that has showed that when Synechococcus develop resistance to cyanophage infection, they appear to have reduced growth rates.106 Therefore in a natural setting, infection from phages is a necessary ‘risk’ to achieve suitable growth rates to outcompete the ‘phage resistant’ strains. The ‘cost of resistance’ may also manifest itself in other forms. A recent study found that a spontaneous phage-resistance mutant of Synechococcus was more susceptible to grazing by heterotrophic nanoflagellates.107 Evidence of cyanophages shaping the populations of hosts has also been observed at a genetic level, with evidence of intragenic recombination of psbA genes between phages and hosts108,109 and the horizontal transfer of hli genes to and from cyanophages and Prochlorococcus.110

Cyanophage spatial and temporal dynamics.

Cyanophage abundance and diversity has been studied at a number of scales from short diel cycles111 to annual cycles.54,58,62,112 Generally cyanophage numbers are at their highest when their hosts are most abundant, which coincides with the warmer summer months.54,62,112 As one might expect, cyanophage abundance also varies through the water column, with cyanophage numbers generally found to decrease with depth.112 The diversity of cyanophages has also been observed to change over temporal and spatial scales with particular genotypes observed only at certain periods of the year or particular depths.62,63,66

Cyanophage future perspectives and summary.

Although we are beginning to understand the importance of cyanophage interactions with their hosts, there are many unknowns to be unraveled. For example no cyanophage tail fibers, or indeed their corresponding receptors have been experimentally demonstrated. A further example of a molecular mechanism used by cyanophages to manipulate and control their hosts was demonstrated with the discovery of the expression of an antisense RNA by the cyanophages S-PM2.113 While this is the first antisense RNA found in a lytic phage, it is unlikely to be the only one that exists. Antisense RNAs form part of large group of RNAs commonly referred to as non coding RNA that are becoming increasingly found in bacterial genomes, where they act as regulators, adjusting physiology in response to changes in the environment.114 Therefore, it is likely the antisense RNA in the cyanophage S-PM2 (and other viruses) is also important as part of a response to environmental change.

In summary, estimates of cyanophage infection rates are high with an estimated 50% of all cyanobacteria being infected at any one time.115 Whether or not they have a defined biogeography is uncertain. Cyanophages were initially thought to be important in terms of diverting the flow of carbon that is fixed by cyanobacteria into the microbial loop, i.e. by infection and the subsequent release of carbon by cell lysis. However, the discovery that cyanophages carry and express photosynthetic genes, suggests that they may also be directly responsible for a significant proportion of carbon fixation in the oceans.116 Most known cyanophages are essentially lytic and they have been shown to drive host diversity over monthly timescales. Cultured cyanophages appear to be consistent with metagenomic data,39 and at least 50% of the genes within their genomes are unique. The development of a genetic system will allow the significance of the ‘host’ encoded genes to be understood, and also allow the function of novel genes to be elucidated.

Case Study 2: The Animal Environment

Animal-associated phages: Discovery and distribution.

The animal or human organism is a complex microcosm of multiple interconnected ecological systems, many of which are densely populated by microorganisms. An indication of the extent of the microbial biomass can be seen from the fact that it contributes up to 54% of the total weight of human feces.117 Significant numbers of individual species of bacteria are found in the intestines, the oral cavity, the vagina, the respiratory tract and the skin where the number of bacteria present frequently overwhelmingly exceeds the expected threshold levels required for efficient phage multiplication. Where this is the case, a high impact of phage infection on the dynamics of the bacterial populations can be expected.118

Although Felix D'Herelle first noticed that bacterial viruses are a normal part of the microbiota of healthy animals and humans,119 our understanding of the role of bacteriophages in shaping and maintaining human symbiotic micro-flora is scarce and fragmentary. The available literature on this subject was extensively analyzed in a recent review.120 Here we review the key roles that phages play in the ecology of the symbiotic micro-flora found in animals.

Isolation and characterization.

The high abundance of phage-like particles in the intestinal microbial systems was first demonstrated by electron microscopy observations in the 1960s120123 these particles represented a vast majority of all viruslike particles (VLP) observed in these kind of samples collected from healthy humans or animals. Despite the early start of the studied of animal and human non-cultured intestinal viral communities, the precise measurements of VLPs concentrations in intestinal contents or feces of any animal were never published. Based on the yield of the total virus (phage) DNA reported in recent metagenomic studies124,35 the concentration of VLPs in feces on humans can be estimated as 1010 ml−1 and in horses up to 1011 ml−1.

Molecular approaches to animal phage sudies.

The diversity of non-cultured intestinal phages was initially characterized by their morphology observed by TEM. This was followed by studies based on the purification of viral communities and analysis of their nucleic acids, initially by pulse-field gel electrophoresis separation to determine genome size, and more recently by metagenomic approaches (reviewed in ref. 120; see also ref. 35). It appears that the vast majority of the intestinal virus-like particles are related to the tailed phages (except for RNA viruses that are mostly plant viruses ingested with food). The estimation of phage diversity based on these data is in the order of hundreds to thousands of distinct genotypes present in one sample of rumen content or feces. In a recent study, as many as 69 morphologically distinguishable bacteriophage types were detected in one specimen of horse feces in over total 200 particles which were examined (Fig. 2).126 These results agree well with the data of the metagenomic analyses on horse feces which have shown that even the most abundant phage type only constitutes between 5–10% of the total population.

Figure 2

Morphology of selected virus like - particles in non-cultured viral community of horse feces. M - myoviruses, S - siphoviruses, P - podoviruses. Note the high prevalence of the siphoviruses with very long tails. The morphotype S18 was the most prevalent...

Animal phage spatial and temporal dynamics.

Although many bacteriophages and phage-host systems have been isolated and characterized from animal-related sources using culture based approaches, the vast majority of these studies were designed to use phages as biological indicators of water fecal contamination, or for other practical applications. The presence and variation of the phage populations in time and in space have only been analyzed in a limited number of studies.

A recently published metagenomic study of viromes of the feces obtained from monozygotic twins and their mothers indicated that their viral communities are stable in time, but differ markedly between individuals within one family, despite the fact that the individual bacterial communities of the close relatives were quite similar.35 This is consistent with metagenomic bacterial data.125 Consistent with human intestinal data, the composition of the non-cultured viral community in rumenal or intestinal microbial systems also seems to vary significantly between subjects. One major difference however is that unlike the situation in the human intestinal virome, the rumenal virome changes significantly with time (reviewed in ref. 120). In conclusion, unlike our appreciation of bacteria and their niches, there are not sufficient data to attribute some types of phages to be “normally” associated with particular ecological processes, or with particular animal species.

Interaction between phages and hosts in their physical environment.

The environments created by the animal-host physiology, and by the microbial activity in different densely populated niches appears to profoundly influence the mode of interaction between bacterial and phage populations. The intestinal coliphage ecology is one of the best studied examples of this complex interaction and the impact of the phages on host populations, mechanisms of phage-host mutual regulation and adaptation have been shown to vary considerably in different animal species. For example, the environmental conditions found in the mouse gut are not favorable for the replication of E. coli phages for some reason, and no studies have shown that mice excrete any natural coliphages.127,128 Furthermore, the resident mouse E. coli populations are almost completely resistant in vivo to externally administrated cocktails of T4 related bacteriophages that kill up to 100% of the same strains in vitro. In contrast to the mouse gut, recent work by Golomidova et al. has confirmed a previously published observation that in a natural situation, healthy horses frequently do excrete coliphages. This work also showed that coliphage populations exhibit significant temporal variation, with up to 4 orders of magnitude difference in phage abundance during 15 days of monitoring.129 In terms of coliphages in other animals, a low fecal coliphage prevalence has been reported in dogs.130

The role that phages play in the ecology of bacteria, also differs between specific ecosystems within different body sites in the same animal species. The failure to obtain phage isolates from the vaginal and oral cavity in humans suggests that the phage impact in these systems is less than their impact in other environments, such as colonic ecosystems of the same species (see here below and the refs. in ref. 120).131133 However direct electron microscopy observations of some samples obtained from dental plaque material showed that high numbers of VLPs were observed.134 Therefore the problem of the phage activity in the oral cavity has to be further addressed, and may be a reflection of our inability to isolate these viruses. A further complication to our understanding of phages associated with animals is the observation that despite relatively high concentrations (about 109–1010 ml−1) of phage particles in the rumenal contents of ruminants, the viruses appear to be unable to control the density of the host populations (reviewed in ref. 120). This observation could be due to the physical and chemical impact of these environments, for example the tannic acid and other chemical compounds in the rumen of sheep and cattle appears to significantly inhibit bacteriophage infection of E. coli.135,136

Generally, it seems that the phage ecology in animal-associated microbial systems should be considered as a tripartite interplay between the bacteriophage, the host bacterium and the environment within the macroorganism. In this play, the macroorganism influences both the bacteria and the phages. A direct influence on phages can be seen from their destruction by digestive enzymes and macrophages, their translocation by blood and their transmission between organisms that is facilitated, or restricted due to specific behavior. Finally, the efficacy of phage infection can be markedly influenced by compounds secreted by the macro-organism such as bile salts (reviewed in ref. 120). This interplay is genuinely three way, and there is evidence that the macro-organism is directly, and indirectly influenced by both phages and microbes. An example of phage influence on the macro-organism can be seen from recent data which suggests that phage particles may interact directly with immune competent cells, exerting immuno-modulatory activity.120,137 It has also been suggested that there may be a direct involvement of phage particles in pathogenesis of inflammatory bowel disease.138

Life cycles and animal bacteriophages.

The identification of a number of coliphages isolated on laboratory E. coli strains and on the field isolates of horse fecal coliform bacteria from the same samples, suggests that the vast majority of the naturally occurring coliphages of the horse gut are lytic. This observation contradicts the observations that Furuse and colleagues made on human associated coliphages. These authors found that in feces of healthy subjects, the coliphages are present at fairly low titers, and are mostly temperate.139 They also demonstrated that both abundance and predominant life-cycle observed, changed when patients were ill either with internal or leukemic diseases. In patients, in contrast to healthy subjects the coliphage numbers were considerably higher and of a substantial fraction of them were virulent phages.139 In several patients, phage titers were shown to increase with the severity of the clinical symptoms. Recent metagenomic data has also suggested that the majority of all phages present the human gut are temperate, and many are involved in the processes associated with anaerobic gut microbiota.35

Impact of animal phages on host bacterial populations.

Like the situation described with cyanobacteria, resistance to bacteriophages which infect the chicken gut bacterium Campylobacter jejuni comes at a price. In the chicken gut, phages do seem to exert substantial selective pressure on the population composition of their bacterial hosts, selecting for phage resistance. The cost however is a reduced ability for the bacteria to colonize the gut.140

Phages shaping bacterial populations dynamics.

Unlike the studies discussed for cyanophages, the impact of naturally occurring phages on bacterial population dynamics has not been quantitatively measured in any animal-associated habitats. However, in some cases there is indirect evidence of the phage pressure. Recently, the intra-species diversity of coliform bacteria in horse gut has been shown to be remarkably high, with over 1000 strains distinguishable by high-resolution PCR fingerprinting, present in a single sample of feces (Fig. 3).129,141 A molecular analysis an TEM based analysis of the equine intestinal coliform-coliphage community is consistent with the predictions from of mathematical and experimental modeling of phage-host communities in which the co-evolution of both components is allowed.48,142 Field observations indicated that the phages isolated using the indigenous E. coli strains, were able to lyse only 2–8% of the E. coli strains occurring in the same sample (Tarasyan KK, Letarov AV, unpublished data). Again consistent with this is the observation that each indigenous bacterial strain could only be infected by 1, or less frequently 2 bacteriophage genotypes present in the same sample suggesting that there is competition between the viruses for host cells.

Figure 3

PCR-fingerprinting of 18 random field isolates of coliform bacteria obtained from the single sample of horse feces. Taken from Isaeva et al., 2010.

There is still much to be learned about phage host dynamics in gut systems, and although there is a high overall density of bacteria for example in the equine intestinal system, most coliphages appear to only infect a small subset of host E. coli strains. The concentration of certain phages ‘types’ has been shown to fall below 102 PFU (Plaque forming units) g−1, and the concentration of their corresponding host cells below 104 CFU (colony forming units) g−1. Under this scenario one would expect that the population of a given phage strain would not be stable and therefore would be eliminated completely, however this is clearly not the case. A further complication is that some bacterial cells may have mechanisms which inhibit bacteriophage intracellular development, causing the extinction of phage lineages. However despite all these theoretical problems posed to bacteriophages, the long term maintenance of some coliphage strains (traced by repeated isolation and genomic DNA RFLP analysis) was observed in the intestine of the horse for more than 2 years (Letarov AV et al., unpublished data).

Possible implications of psuedolysogenic infections.

The mechanisms that allow phages to avoid extinction are still not clear; however, a possible explanation may be due to phages from field isolates being able to form psuedolysogenic associations with their hosts. The normally lytic phages may have formed quasi-stable relationships with their hosts. Putative psuedolysogenic infection in E. coli can be observed by growing E. coli cells derived from phage plaques. When material from phage plaques is streaked on to fresh Petri-dishes a large proportion of the cells are resistant to the phage whose plaque they were isolated from. This resistance and can be maintained for several passages. In addition to displaying resistance, to the bacteria are able to produce phages active against the parental bacteria strains for between 5–15 passages. These observations lead to the possibility that some phage populations in the horse gut are maintained due to psuedolysogenic micro-colonies or biofilm patches on the surfaces of the food particles and mucosa within the gut environment.

An alternative (or complementary) explanation of the longtime persistence of bacteriophage populations that appear to be severely limited by availability of the hosts was suggested by Kunisaki and Tanji.143 These authors discovered that some E. coli strains have become phage resistance in anaerobic continuous culture, are resistant because they the bacteriophages used can no longer absorb to them. However approximately 1–2% of the cells grown from these colonies were phenotypic revertants which could be infected by the phage. Interestingly, these cells were not true genetic revertants as authors were unable to isolate phage sensitive derivatives despite screening several hundred sub-clones of the resistant strain. If this observation is confirmed in future experiments, and explanation for it established, it could become a significant milestone in our understanding of bacteriophage ecology in the environments with high density of microbial life. Finally, it is worth mentioning the nascent phage phenomenon whereby phages have broader host ranges immediately after they are released from a bacterial cell than they do after several hours, or longer, have passed.144

Summary and future perspectives in phages associated with animals and humans.

To summarize our knowledge of phage ecology in the human and animal microflora, the “rules of the game” of the interplay between viruses and the hosts here appear to be complicated in animal environments because a significant role of the mutual interactions of phage and host within the macroorganism. This game has been going on for hundreds of millions of years and understanding the “rules” is essential if we are to successfully understand the role of bacteriophage in disease, or to exploit them. For example, these natural patterns will inform us as to how we should expect the dynamics of bacteria/phages to stabilize in phage therapy, or in other technologies which use phages to control bacterial populations.

Case Study 3: Archaeal Viruses

Are archaeal viruses phages?

The final case study discussed in this paper is that of phages in extreme environments, and this necessitates a brief consideration of what we actually mean by a bacteriophage. More than 30 years ago the seminal work of Carl Woese demonstrated that cell based life on this planet was not best described by a dualistic partition into the prokaryotes and eukaryotes.145 Rather, as now taught by entry level microbiology courses and introductory microbiology texts, living diversity is better described by a division into three domains; the Archaea, Bacteria and Eukarya, each with an associated virus population.146,147 There is still a tendency by many to pool viruses associated with the domains Bacteria and Archaea and think of them as bacteriophages. This tendency is deeply and institutionally rooted e.g. by the International Committee on the Taxonomy of Viruses.148 A visit to the NCBI genomes page provides links to all completely sequenced genomes which can be searched in a variety of ways e.g. within the three domains of cellular life. However, it is not possible to search for viruses infecting cells specifically of the domain Bacteria or specifically of the domain Archaea; these are linked together under ‘Phages’ and with this in mind they are considered in the final section of this paper.

Archaeal virus discovery and distribution.

The domain Archaea was proposed about 35 years ago and the study of viruses associated with it has used classical bacteriophage techniques. Thus far most studies have depended upon lawn/plaque approaches.149 Since this pioneering work it has become apparent that the Archaea are not just niche players. They may be almost as common as the Bacteria in oceans, soils and subterranean environments and also show enormous diversity.4143 They also occupy and can predominate in so-called extreme environments, such as hot-springs, salt and soda lakes.

Likewise, there is every reason to assume that viruses of the Archaea are as numerous and diverse as those of the bacteria. Reflecting our greater knowledge of the domain Bacteria, almost 6000 bacteriophage have been described,150 compared with only 50 or so viruses which infect Archaea.150,151 In terms of sequenced ‘bacteriophages’, the NCBI genomes page listed 580 ‘phages’ on June 16, 2010 of which just 32 were viruses of the Archaea, and many of these are sequences of closely related viruses. If we have seen the ‘tip of the iceberg’ with respect to bacteriophages, then we are just beginning to scratch the ‘tip of the iceberg’ with viruses of the Archaea. Therefore, our knowledge of their roles in nature is also fragmentary, far more so than for the cyanophages and phages associated with animals. Obviously if we are to begin to understand biogeochemical cycling and environmental ecology then a much more complete genetic census of archaeal viruses in the environment and their effects upon host is essential.

Isolation and characterization of Archaeal viruses.

To summarize our current knowledge about these viruses we have to first describe the main divisions within the Archaea. They are divided into two kingdoms, the Crenarchaeota which mainly contains hyperthermophiles, and the Euryarchaeota mainly consists of halophiles and methanogens. Many viruses which infect members of the two kingdoms have been identified.5 Most viruses have double stranded DNA genomes varying in size from 10 kb to over 100 kb.152 No viruses of the Archaea with an RNA genome have yet been identified, but it would seem likely that they exist, just as they do in the domains Bacteria and Eukarya.

Around 30 viruses infecting the Crenarchaeota have been described and they are notable not only for generally being isolated from hyperthermophilic hosts growing at temperatures > 80°C but also for an array of unusual morphologies not observed in viruses of the Eukarya or Bacteria.152156 These include spindle shaped virions of the Fuselloviridae e.g. SSV1, infecting members of the genera Sulfolobus and possibly Acidianus, which have small circular genomes from ∼15–24 kb. The rod shaped Rudiviridae, SIRV1 and SIRV2 with 35 kb genomes, infect Sulfolobus species.157 SIFV, an enveloped flexible filamentous virus with a 41 kb genome infecting Sulfolobus is a member of the Lipothrixviridae.158 These latter two families have recently been grouped into a new order the Ligamenvirales. The droplet shaped virions of the Guttaviridae, Sulfolobus neozealandicus, SNDV, all have circular genomes of around 20 kb.159 The Ampullaviridae, are exemplified by the Acidianus bottle shaped virion ABV, which has a linear 24 kb genome.160 The Acidianus two tailed virion of the Bicaudaviridae, Acidianus ATV, has a genome of 62 Kb.161 This virus is notable for exiting the cell as a lemon shaped fusiform particle which then develops long tails at each pointed end at temperatures above 75°C. This unprecedented extracellular morphological development is entirely independent of the host cell.162 Spherical viruses ‘the Globuloviridae’ also exist, for example the Pyrobaculum spherical virus PSV.163

Euryarchaeota predominate in high salt environments, morphological studies again show a diversity of morphotypes with the head and tail variety being in the minority. About 20 viruses have been studied from these environments, infecting members of the genera Halobacteriales, morphologically most resemble head and tail viruses.164 These are most similar to bacteriophages which infect Bacteria and solely on this morphological basis are classified as Caudovirales including the myoviruses and siphoviruses. An example of these viruses is the temperate myovirus ϕCh1 which has a 58.5 kb linear genome, and infects the haloalkaliphilic host Natrialba magadii. The lytic viruses HF1 and the closely related HF2, have linear genomes of 75.9 kb and 77.7 kb and infect the haloarchaea Haloferax lucentense and Halorubrum coriense, respectively. BJ1 with a 43 kb genome infects Halorubrum kocuri.42,165 A lytic icosahedral virus SH1 having a linear genome of 31 kb infects Haloarcula hispanica.166 The Salterprovirus include His1 and 2 have linear genomes of 14.5 and 16 kb respectively and infect Har. hispanic.167 The only archaeal virus thus far identified not having a dsDNA genome is the Halorubrum pleomorphic virus 1 which has a ssDNA genome 7048 nucleotides in size.152 The Euryarchaeotal methanogens also have an identified virus, psi M1 with a linear dsDNA 30 kb genome isolated from Methanothermobacter marburgensis.168

Molecular characterization of Archaeal viruses.

Genomic analysis of archaeal viruses generally demonstrates very low identity to any other virus sequences.42

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