bs_bs_banner

Environmental Microbiology (2014)

doi:10.1111/1462-2920.12569

Host-dependent differences in abundance, composition and host range of cyanophages from the Red Sea Naama P. Dekel-Bird, Gazalah Sabehi, Bar Mosevitzky and Debbie Lindell* Faculty of Biology, Technion – Israel Institute of Technology, Haifa 32000, Israel. Summary Cyanobacteria coexist in the oceans with a wealth of phages that infect them. While numerous studies have investigated Synechococcus phages, much less data are available for Prochlorococcus phages. Furthermore, little is known about cyanophage composition. Here, we examined the abundance and relative composition of cyanophages on six cyanobacterial hosts in samples collected during spring and summer from the Red Sea. Maximal abundances found on Synechococcus of 35 000 phages/ml are within ranges found previously, whereas the 24 000 phages/ml found on Prochlorococcus are approximately 10-fold higher than previous findings. T7-like, T4-like and ‘unknown’ phages were isolated on all hosts, including many T4-like phages on high-light adapted Prochlorococcus strains, whereas TIM5-like phages were found only on Synechococcus. Large differences in cyanophage abundance and composition were found for different hosts on the same sampling date, as well as for the same host on different dates, with few predictable patterns discerned. Host range analyses showed that T7-like and TIM5-like phages were quite host-specific, whereas the breadth of hosts for T4-like phages was related to host type: those isolated on high-light adapted Prochlorococcus were considerably more host-specific than those on low-light adapted Prochlorococcus or Synechococcus. These host-related differences likely contribute to the complexity of host–phage interactions in the oceans.

Received 27 April, 2014; accepted 15 July, 2014. *For correspondence. E-mail [email protected]; Tel. (+972) 4829 5831; Fax (+972) 4822 5153.

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd

Introduction The unicellular cyanobacteria of the genera Synechococcus and Prochlorococcus are the most abundant photosynthetic organisms in the oceans and are estimated to be responsible for 25% of global oceanic primary production (Flombaum et al., 2013). They are most predominant in tropical and subtropical waters, but differ in their abundances over coastal to open ocean transects (Partensky et al., 1999; Zwirglmaier et al., 2008) and over annual cycles in seasonally stratified waters (Olson et al., 1990; Lindell and Post, 1995; DuRand et al., 2001; Malmstrom et al., 2010). Prochlorococcus is more abundant in oligotrophic, stably stratified open waters, whereas Synechococcus is more prevalent in mesotrophic waters typical of coastal sites and conditions during subtropical spring blooms. Both Synechococcus and Prochlorococcus are made up of a number of distinct ecotypes or clades that also differ in their relative abundances under various environmental conditions (Johnson et al., 2006; Zinser et al., 2007; Zwirglmaier et al., 2008; Malmstrom et al., 2010; Post et al., 2011; Ahlgren and Rocap, 2012). High-light adapted (HL) Prochlorococcus ecotypes are abundant in the upper layers of the photic zone in oligotrophic waters, and the two main ecotypes (HLI and HLII) differ in their geographical distribution in correlation with surface temperatures. The low-light adapted (LL) Prochlorococcus ecotypes are found deeper in the water column (Johnson et al., 2006; Zinser et al., 2007; Malmstrom et al., 2010). Certain Synechococcus clades (II and III) are common in oligotrophic tropical and subtropical waters, while other clades (I and IV) are more prevalent in colder, nutrient-replete waters (Zwirglmaier et al., 2008; Tai and Palenik, 2009; Post et al., 2011; Ahlgren and Rocap, 2012). Cyanobacteria coexist in the oceans with phages (cyanophages) that infect them. These cyanophages have been estimated to lead to the mortality of a small but significant fraction of cyanobacteria daily (Proctor and Fuhrman, 1990; Suttle and Chan, 1993; Waterbury and Valois, 1993). Cyanophages are also thought to strongly impact the diversity and evolution of their hosts (Lindell et al., 2004; 2007; Avrani et al., 2011; Marston et al., 2012).

2

N. P. Dekel-Bird, G. Sabehi, B. Mosevitzky and D. Lindell

The marine cyanophages isolated so far are dsDNA tailed phages that belong to the order Caudovirales (Suttle and Chan, 1993; Waterbury and Valois, 1993; Sullivan et al., 2003; Wang and Chen, 2008). They are divided into three major families based on tail morphology: the Myoviridae, Siphoviridae and Podoviridae, which have long contractile, long non-contractile and short tails respectively (Ackermann, 2003). Within each morphotype, distinct lineages (or types) that differ in their replication and morphogenesis genes exist (Lavigne et al., 2008; 2009). The best characterized marine cyanophages are the T4-like myoviruses and the T7-like podoviruses that resemble the Escherichia coli T4 and T7 archetypes, respectively, in their morphology, genome architecture, and homologous sets of replication and morphogenesis genes (Chen and Lu, 2002; Mann et al., 2005; Sullivan et al., 2005; 2010; Pope et al., 2007; Weigele et al., 2007; Millard et al., 2009; Labrie et al., 2013). Each of these cyanophage lineages is quite diverse and can be further subdivided into a number of discrete phylogenetic clades (Marston and Sallee, 2003; Sullivan et al., 2008; Wang and Chen, 2008; Dekel-Bird et al., 2013). Recently, a novel lineage of myoviruses that infects marine Synechococcus spp. was discovered and was named the TIM5-like myoviruses (Sabehi et al., 2012). A number of siphoviruses have also been isolated. They are quite diverse and do not appear to belong to a single lineage (Sullivan et al., 2009; Huang et al., 2012; Ponsero et al., 2013). A single report suggests that non-tailed ssDNA phages may also infect marine Synechococcus (McDaniel et al., 2006). The discrete cyanophage types differ in their intrinsic host ranges. The T7-like podoviruses and the siphoviruses are considered to be host-specific, often infecting only the host of isolation (Sullivan et al., 2003; Wang and Chen, 2008; Dekel-Bird et al., 2013). The host range of S-TIM5 is also quite narrow, infecting only two Synechococcus hosts (Sabehi et al., 2012). In contrast, the T4-like myoviruses tend to have a broader host range, generally infecting more than one host (Suttle and Chan, 1993; Waterbury and Valois, 1993; Sullivan et al., 2003; Wang and Chen, 2008). Some T4-like cyanophages can even infect members of both Synechococcus and Prochlorococcus (Sullivan et al., 2003; Millard and Mann, 2006). Many studies have investigated the abundance of cyanophages on spatial and temporal scales. These studies, generally using a select few Synechococcus strains as hosts, showed that cyanophage abundances ranged from a few hundred phages up to 105 phages/ml. Their abundances decrease along coastal to open water transects and change seasonally, mirroring changes in the abundance of Synechococcus (Suttle and Chan, 1993; 1994; Waterbury and Valois, 1993; Marston and

Sallee, 2003; Sullivan et al., 2003; Wang and Chen, 2004). So far, only a single study has investigated the relative abundance of cyanophages infecting Prochlorococcus and reported low overall numbers of less than 3000 phages/ml in the Sargasso and Red Seas (Sullivan et al., 2003). The composition of cyanophages in the environment has been inferred from isolation studies over the years. T4-like myoviruses are most commonly isolated on Synechococcus hosts and are often considered to be the most abundant cyanophage type (Marston and Sallee, 2003; Sullivan et al., 2003; Wang and Chen, 2004; Millard and Mann, 2006). Furthermore, a recent quantitative polymerase chain reaction (PCR) assessment has indicated that the T4-like cyanophages are abundant in the oceans (Matteson et al., 2013). However, T7-like podoviruses, TIM5-like myoviruses and siphoviruses have also been found on Synechococcus hosts (Marston and Sallee, 2003; Wang et al., 2011; Huang et al., 2012; Sabehi et al., 2012; Dekel-Bird et al., 2013). In comparison, a predominance of T7-like podoviruses were isolated on HL Prochlorococcus strains, whereas both T4-like myoviruses and T7-like podoviruses were isolated on LL Prochlorococcus hosts (Sullivan et al., 2003). Here, we investigated the prevalence of three major cyanophage lineages, the T4-like, T7-like and TIM5-like phages, in the subtropical oligotrophic waters of the Gulf of Aqaba, Red Sea. This body of water undergoes cycles of seasonal stratification in spring-summer and deep mixing in winter (Wolf-Vecht et al., 1992; Carlson et al., 2013). Cyanobacterial abundances vary quite dramatically over this annual cycle, with Synechococcus most abundant in the spring and Prochlorococcus most abundant in the summer (Lindell and Post, 1995, Fuller et al., 2005, Penno et al., 2006). Sampling in the spring and summer of two consecutive years, we estimated cyanophage abundances using the plaque assay with Synechococcus and Prochlorococcus ecotypes commonly found in oligotrophic environments as hosts. We then used PCRs of cyanophage lineage-specific signature genes for the identification of the plaques to determine the relative composition of the different cyanophage types. While the three cyanophage types were found on each sampling date, their abundance and relative composition varied greatly with host strain on a single sampling date, as well as on a single strain on different sampling dates. Host range assessments revealed that while the T7-like and TIM5-like phages were quite host-specific, the breadth of the host range within the T4-like myoviruses was dependent on the host: cyanophages infecting HL Prochlorococcus were predominantly host-specific, whether T7-like or T4-like, while the T4-like phages infecting other hosts had considerably greater host ranges.

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology

Cyanophage abundance, composition and host ranges Results and discussion Cyanophage abundance In order to estimate the abundance of cyanophages during the spring and summer in the Gulf of Aqaba when Synechococcus and Prochlorococcus dominate, respectively (Lindell and Post, 1995, Fuller et al., 2005, Penno et al., 2006), we carried out plaque assays with six different cyanobacterial strains as hosts. The six chosen strains are representatives of types commonly found in open-water environments, including the Red Sea, and include three Synechococcus strains – two from clade II (WH8109 and CC9605) and one from clade III (WH8102) – as well as three Prochlorococcus strains – one each from clade HLI (MED4), clade HLII (MIT9215) and clade

3

LLI (NATL2A). We used two Synechococcus clade II strains to assess whether the same results are obtained for two strains from the same clade. Three water samples were investigated for each sampling date: two were from Station A, one within the surface mixed layer at 20 m and one below it at 60 m. This is an open-water site with a bottom depth of approximately 700 m. The third sample was taken from the surface off the pier of the Interuniversity Institute for Marine Sciences (IUI) above a bottom depth of approximately 4 m. Phages were found to infect each of the six cyanobacterial strains from all sites and on all sampling dates (Fig. 1). Abundances ranged from just a few hundreds of phages to tens of thousands of phages per millilitre but were generally on the order of thousands

Fig. 1. Estimates of cyanophage abundances in the Gulf of Aqaba, Red Sea. Abundances (plaque forming units per millilitre), using six cyanobacterial strains as hosts, are shown for the three sampling sites in spring and summer of 2009 and 2010: (A) 16 March 2009, (B) 19 August 2009, (C) 24 March 2010 and (D) 16 August 2010. Prochlorococcus host strains, MED4, MIT9215 and NATL2A, are shown in green above the graphs, and belong to the HLI, HLII and LLI ecotypes respectively. Synechococcus host strains, WH8102, WH8109 and CC9605, are shown in red above the graphs, and belong to clades III, II and II respectively. Abundances are the average and standard deviation of three to four plaque assays per sample. N/A, data not available due to lack of growth of Synechococcus WH8109 lawn for this sampling date (see text).

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology

4

N. P. Dekel-Bird, G. Sabehi, B. Mosevitzky and D. Lindell

per millilitre of seawater. Maximum phage numbers were found in August 2009, with approximately 35 000 phages/ml when Synechococcus WH8102 was the host and 24 000 phages/ml when Prochlorococcus MED4 or NATL2A were the hosts (Fig. 1). These maximal Synechococcus phage abundances are within the ranges reported previously for different bodies of water (Suttle and Chan, 1993; 1994; Waterbury and Valois, 1993; Marston and Sallee, 2003), but are considerably higher than those reported for Station A in the Gulf of Aqaba throughout the year of 1999 (Mühling et al., 2005; Millard and Mann, 2006). Furthermore, the maximal abundances for Prochlorococcus phages found in this study were approximately an order of magnitude higher than those found previously for September samples collected at the same sampling site in the Gulf of Aqaba as well as from the Sargasso Sea (Sullivan et al., 2003). The high degree of natural variation in cyanophage abundances (Figs 1 and 2), together with the current scarcity of data on Prochlorococcus phage abundances, may explain why higher abundances of Prochlorococcus phages have not been reported previously. It is also partly possible that higher abundances were reported in this study as phage titre assays were carried out within 24 h of sampling, thus circumventing rapid decay in infective phage titres found under common storage conditions (see Experimental procedures for analysis of decay of infective phages). One particularly noticeable finding was that large differences in cyanophage abundances were often found for the same sample when different cyanobacterial strains served as hosts, even when two strains from the same clade were used (Fig. 1). Furthermore, abundances on

Fig. 2. Estimates of cyanophage abundances at the pier sampling site during summer stratification in multiple years. Abundances are shown for surface samples collected from the pier during August 2009, 2010, 2012 and September 2012 when using (A) Prochlorococcus MED4 as host and (B) Synechococcus WH8102 as host. Abundances are the average and standard deviation of three to four plaque assays per sample. The values for August 2009 and 2010 are the same as those appearing in Fig. 1, and those for August and September 2012 are those presented as V1 in Fig. 5. See the legend of Fig. 5 for statistical analysis of the differences in phage abundances between August and September of 2012.

any particular host differed greatly in the different samples collected on a certain date (Fig. 1), as well as on different dates and years of the same season at the same site (Fig. 2). These temporal, spatial and host straindependent differences in cyanophage abundances are similar to findings from previous studies (Suttle and Chan, 1993; Waterbury and Valois, 1993; Marston and Sallee, 2003; Sullivan et al., 2003). While this study does not attempt to assess seasonal differences in abundances at the different sampling sites, a few observations are worthy of mention. First of all, combined cyanophage abundances from the three samples for all three hosts within a genus showed that significantly more cyanophages were detected on Synechococcus hosts than on Prochlorococcus hosts for both spring sampling dates as well as for the summer 2009 sampling date (P < 0.01). This is despite the fact that Prochlorococcus concentrations are higher than Synechococcus during the summer (Lindell and Post, 1995; Fuller et al., 2005; Penno et al., 2006). See the discussion below on the breadth of host ranges for an explanation of why Synechococcus phages may be more abundant than Prochlorococcus phages. Second, more phages for both Synechococcus and Prochlorococcus hosts were found in August than in March of 2009 (P < 0.05 and P < 0.005 for Prochlorococcus and Synechococcus phages, respectively), but no such differences were found for 2010 (P = 0.56 and P = 0.2 for Prochlorococcus and Synechococcus phages respectively). Third, during summer stratification, some clear trends were observed at different sites for certain hosts. For example, fewer Prochlorococcus phages were present at the site close to shore (pier) than at the openwater site (Station A), and cyanophage abundances on the LL Prochlorococcus strain were higher below the mixed surface layer (60 m) than within it (20 m). In comparison, Synechococcus phages were of similar abundance in the surface samples both at the pier and at Station A for all hosts in the summer of both years. However, fewer Synechococcus phages were detected at 60 m on two of the three hosts (WH8102 and CC9605). Therefore, while combined cyanophage abundances do not necessarily follow overall seasonal differences in the abundance of the host genera, some site- and depthrelated trends appear to follow those known for certain Synechococcus and Prochlorococcus ecotypes. Data are not available for Synechococcus WH8109 from the August 2009 sampling date due to a problem with the formation of the host lawn. Repeat plating 10 days later of the three samples for this date with all three Synechococcus hosts provided phage estimates that were at least twofold lower than the initial plating 1 day after sampling (data not shown). Therefore, we excluded these results from our analyses. This finding suggested a

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology

Cyanophage abundance, composition and host ranges rapid decay of infective phages using common storage conditions of 4°C in the dark. See Experimental procedures for a more formal assessment of phage decay, which showed a significant decline in phage abundances on both Synechococcus and Prochlorococcus hosts within 1–4 weeks of sample collection using various glass and plastic containers. Cyanophage composition Cyanophage abundances determined from the plaque assay provide estimates of the number of phages that can infect the particular host used. However, these data do not provide any information on the composition of the phage types infecting these hosts. To gain an understanding of the relative composition of the cyanophage types in our samples, we carried out PCRs targeting lineage-specific signature genes: for T4-like cyanophages, we used the portal protein gene, g20 (Sullivan et al., 2008), and for the T7-like and TIM5-like cyanophages we used their respective DNA polymerase genes (Wang and Chen, 2008; Sabehi et al., 2012; Dekel-Bird et al., 2013). See Table S1 for a list of the PCR primers and Table S2 for the PCR conditions used in this study. Approximately 60 plaques were randomly picked and analysed for each cyanobacterial host collected from each sampling site on the four sampling dates. Remarkably similar, yet not identical, relative compositions of the different cyanophage lineages were found for the same host in the three samples on a particular sampling date (Fig. 3). This was the case even on sampling dates for which large differences in the abundances of total cyanophages were found at the different sites (compare Fig. 3 with Fig. 1). This is easily seen for the Prochlorococcus MED4 and Synechococcus WH8102 hosts in August of both years. However, quite large differences in composition were found for the different hosts on a particular sampling date (Fig. 3). These do not appear to be related to the host genus. This was most obvious for March 2009, when T7-like podoviruses were most abundant on Prochlorococcus MED4 and Synechococcus CC9605, while T4-like myoviruses were mainly found on Prochlorococcus MIT9215 and Synechococcus WH8102. Furthermore, the relative composition of cyanophages changed from sample date to sample date when using the same host for the assay. For example, the phages identified on Synechococcus CC9605 were primarily T7-like podoviruses in March 2009, T4-like myoviruses in August 2009, a mix of T4-like and TIM5-like myoviruses in March 2010, and mainly TIM5-like and unknown viruses in August 2010. These data indicate that while compositions are fairly constant for a particular host on a particular date, they vary quite dramatically with host strain and with time.

5

Taking the findings from all four sampling dates together, we found that T7-like and T4-like cyanophages infect all of the cyanobacterial strains used in this study. However, T7-like podoviruses appeared less often overall and were rare on Prochlorococcus NATL2A and Synechococcus WH8102 on these sampling dates. In contrast to the cosmopolitan nature of the T7-like and T4-like cyanophages, TIM5-like phages were found only on Synechococcus WH8102 and CC9605. On some sampling dates, this phage type was quite rare, such as in March 2009, whereas on other dates they made up most of the phages infecting these two strains, such as in August 2010. Estimations of the abundance of each phage type for a particular host can be done by combining total cyanophage abundances with the proportion of each phage type identified. This indicated that the abundances of each phage type can be quite significant on different sampling dates, with maximal abundances of approximately 25 000 T4-like phages/ml on Synechococcus WH8102 in August 2009, 5000 T7-like podoviruses/ml on Synechococcus CC9605 in March 2009 and 10 000 TIM5like myoviruses/ml onSynechococcus WH8102 in August 2010. We sequenced a random sampling of the PCR fragments from different cruises for all three phage types (Table S3) in order to verify their identity and assess their diversity and phylogeny. This revealed that the T7-like phages were quite diverse and were dispersed throughout clade B, but not clade A, of the DNA polymerase tree (Fig. S1). The most distant sequences had approximately 60% nucleotide identity, although many of the 35 sequences were identical or near identical to others (Table S3). The T4-like phages were also highly diverse. Fifteen of the 16 g20 sequences evaluated were different to each other, and the most distant g20 sequences displayed about 60% nucleotide identity. Despite the limited number of phages investigated, the sequences spanned the known diversity for T4-like cyanophages, with representatives clustering in all four cyanophage g20 clades (Fig. S2). In contrast, the TIM5-like phages were highly similar to each other, with only four unique sequences out of the 67 phages investigated, all of which clustered closely with the S-TIM5 archetype phage (Fig. S3). Furthermore, the most diverse sequences were 95% identical at the nucleotide level, and all differences were synonymous and thus did not cause a change in amino acid sequence. The low degree of sequence diversity found, together with the limited number of Synechococcus hosts that this phage type seems to infect, raises the question as to whether the TIM5-like phages are an inherently low-diversity cyanophage type. Alternatively, perhaps more diverse members exist that infect a broader range of hosts, including Prochlorococcus, but were not identified

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology

6

N. P. Dekel-Bird, G. Sabehi, B. Mosevitzky and D. Lindell

Fig. 3. Cyanophage lineage composition in the Gulf of Aqaba, Red Sea. The relative composition of three cyanophage types and unknown phages are shown for the three sampling sites in the spring and summer of 2009 and 2010: (A) 16 March 2009, (B) 19 August 2009, (C) 24 March 2010 and (D) 16 August 2010. The three cyanophage lineages, T4-like, T7-like and TIM5-like phages, are shown in red, blue and green, respectively, as determined by PCR on lysates derived from randomly picked plaques from each of the six hosts (shown above the graphs and as described in Fig. 1). Lysates from plaques that failed to produce a PCR fragment for any of the three phage lineages was scored as ‘unknown’, and their relative contribution is shown in grey. N/A, data not available due to lack of growth of the Synechococcus WH8109 lawn for this sampling date.

with our current primer set. Further work is required before we can answer this question. Previous isolation studies suggested that HL adapted Prochlorococcus strains are predominantly infected by podoviruses, that LL Prochlorococcus strains are infected by podoviruses and myoviruses (Sullivan et al., 2003; 2008), and that Synechococcus strains are infected mainly by myoviruses (Waterbury and Valois, 1993; Marston and Sallee, 2003; Sullivan et al., 2003). Different from the above, we found that both T4-like myoviruses and T7-like podoviruses were commonly isolated on HL Prochlorococcus types (Fig. 3), and that at least certain Synechococcus strains were often infected by T7-like podoviruses (Fig. 3). However, the predominance of each phage type varied quite a lot on the different sampling

dates. Recently, Wang and colleagues (2011) also found that Synechococcus WH7803 can be infected by either T7-like or T4-like phages, and that the predominant phage type differs with sampling date and station in the Chesapeake Bay. Thus, it appears that considerably more data on the composition of the phage lineages infecting Synechococcus and Prochlorococcus from multiple sites and sampling expeditions are required before robust generalizations can be made. The above analyses also revealed a large proportion of unknown phages infecting each of the hosts, particularly in the summer samples (Fig. 3). In one particular sample (60 m depth in August 2009), approximately 50% of the phages infecting Prochlorococcus NATL2A did not yield PCR fragments for any of the three phage lineages

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology

Cyanophage abundance, composition and host ranges investigated (Fig. 3). This is equivalent to approximately 12 000 phages/ml and was the maximal number of unknown phages measured. These unknown phages may belong to one of the phage lineages investigated here but be sufficiently diverse to go undetected with our primer sets. For example, based on the genome sequence of the T7-like P-RSP2 cyanophage, it is unlikely that the DNA polymerase gene would be amplified with our primer set. This phage is an outlier, being significantly different from the other T7-like podoviruses isolated to date (Labrie et al., 2013, Fig. S1). Other unknown phages are likely to belong to the siphovirus family, not investigated here. It is also feasible that these phages belong to lineages not currently known to infect marine cyanobacteria, or even to novel phage types, as was found for the recently identified TIM5-like phage lineage (Sabehi et al., 2012). Either way, our findings suggest that numerous diverse cyanophage types await discovery in the oceans, and emphasizes the need to use a variety of cyanobacterial hosts for their detection and isolation. Host range We next wished to assess whether the host ranges found previously for the three cyanophage lineages hold for the phages found in this study. This was of particular interest for the T4-like myoviruses isolated on HL Prochlorococcus, which have not been investigated so far, as well as for the TIM5-like myoviruses for which only a single

7

representative has been examined. We investigated the host range of over 400 phages obtained on our six host strains for the ability to infect the other five cyanobacterial strains (see Experimental procedures). As expected, the T7-like podoviruses were very host-specific, generally infecting only the host used for initial plaque formation (Fig. 4). The TIM5-like phages also had quite a narrow host range, with all phages from this type being capable of infecting both Synechococcus WH8102 and CC9605 but not the other hosts. This host range is the same as that determined for S-TIM5, the archetype phage for this family (Sabehi et al., 2012). In comparison to the consistently narrow host range of the T7-like and TIM5-like phages, the T4-like cyanophages showed a range of specificities (Fig. 4). Many T4-like phages were specific for the host of isolation, while many others infected other hosts within the same genus. Yet other T4-like phages could infect strains from both cyanobacterial genera, as has been reported previously for this phage lineage (Sullivan et al., 2003; Millard and Mann, 2006). Intriguingly, we found that the degree of host specificity appears to be related to the host of isolation. Many of the phages isolated on the two HL Prochlorococcus strains (MED4 and MIT9215) were quite host-specific, while others also infected additional Prochlorococcus but not Synechococcus strains (Fig. 4). The majority of phages isolated on Synechococcus WH8109 and CC9605 were less host-specific, being capable of infecting other hosts

Fig. 4. Breadth of cyanophage host ranges as a function of phage lineage and the host of isolation. The relative number of phages per host with different host range categories is shown for each phage type isolated on each of the six hosts. The following are the host range categories: infect only the host of isolation (blue), infect other hosts within the same genus (red) and infect members of both genera (green). The host strain of isolation and the number of plaques investigated for each host are shown above the graph, while the phage type, as determined by PCR, is shown on the x-axis. Samples analysed were those collected in March 2009 and August 2010 from 60 m depth on each of the six hosts and were tested for the ability to infect the other five hosts. Only phages that lysed the host of isolation at the time of testing were included in the analysis.

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology

8

N. P. Dekel-Bird, G. Sabehi, B. Mosevitzky and D. Lindell

within the same genus, but rarely Prochlorococcus hosts. Lastly, the phages isolated on the LL Prochlorococcus NATL2A and Synechococcus WH8102 were the least host-specific, with many phages capable of infecting hosts from both cyanobacterial genera: 45% and 19% of the T4-like phages on NATL2A and WH8102 respectively (Fig. 4). However, none of the phages infected all six hosts. The above findings suggest that certain cyanobacterial strains are more prone to infection by broad host range phages than others, and that this is not related to host genus within the cyanobacteria. Combining our current findings with those of Sullivan and colleagues (2003), it appears that HL Prochlorococcus strains are primarily infected by rather narrow host range phages whether these are T7-like or T4-like phages. In contrast, other cyanobacterial strains, such as LL Prochlorococcus NATL2A and Synechococcus WH8102, appear to be more prone to infection by broad host range phages. Narrow host range cyanophages would need to infect highly abundant hosts to maintain their populations, as was suggested previously by Sullivan and colleagues (2003). Indeed, HLII Prochlorococcus, which had the highest number of host-specific T4-like cyanophages (Fig. 4), is more abundant than HLI and LL Prochlorococcus types as well as Synechococcus in the Gulf of Aqaba (Fuller et al., 2005). In contrast, broad host range phages may find sufficient hosts through the ability to infect multiple different host types, each of which has relatively lower abundances in the oceans. Consistent with this, the clade III Synechococcus strain WH8102 resulted in the highest number of broad host range phages out of the three Synechococcus strains and is less abundant than clade II Synechococcus (represented here by WH8109 and CC9605) in this body of water (Fuller et al., 2005; Post et al., 2011). These differences in host range confound attempts to determine overall cyanophage abundances using culture-dependent means. This is because it is not possible to simply sum up the number of T4-like cyanophages found on the different hosts due to their overlapping host ranges, nor is it possible to use a single ‘highly susceptible’ host, as numerous T4-like cyanophages are very host-specific and will not infect this host. On the other hand, accurate abundances cannot be determined for the highly host-specific T7-like cyanophages either: combining abundances found for different hosts will lead to large underestimations of their overall abundances unless all susceptible host types are used. This will be highly problematic to achieve since multiple Synechococcus and Prochlorococcus subpopulations with different viral susceptibility regions exist in the oceans (Rodriguez-Valera et al., 2009; Avrani et al., 2011). Therefore, even though our culture-dependent

results (Fig. 3) and those of others (Marston and Sallee, 2003; Sullivan et al., 2003; Millard and Mann, 2006) suggest that the T4-like myoviruses are the dominant cyanophage lineage in the oceans, this impression may be greatly skewed by the fact that many of these phages infect multiple hosts, whereas the T7-like podoviruses and TIM5-like myoviruses are much more host-specific. Indeed, fairly equal abundances of T4-like and T7-like cyanophages have been suggested recently from metagenomic analyses (Angly et al., 2006; Bench et al., 2007; Labrie et al., 2013). Furthermore, the impression that more Synechococcus than Prochlorococcus phages were present on three of the four sampling dates in this study may be partly due to the greater breadth of host range of the phages infecting the Synechococcus than the Prochlorococcus hosts used in this study. Conclusions In summary, our results show that a large degree of variability is found in cyanophage abundances and lineage composition, both for a single host at different times and across different cyanobacterial hosts in a single sample. These findings, combined with those of others (Waterbury and Valois, 1993; Marston and Sallee, 2003; Wang et al., 2011), suggest that cyanophage populations are highly dynamic, and that their community structure is also quite diverse and variable. Thus, based on currently existing data, cyanophage community structure does not appear to be stable or predictable, despite the predictable nature of the abundances and distribution patterns of their cyanobacterial hosts (Zinser et al., 2007; Zwirglmaier et al., 2008). This may partially be due to the fact that a phage’s ability to infect a host is related to non-core host genes that do not follow host phylogeny (Avrani et al., 2011). However, a dearth of currently available information for the cyanophages may simply be preventing us from seeing existing patterns at present. The complex cross-infection patterns found for cyanophages (this study, Suttle and Chan, 1993; Waterbury and Valois, 1993; Sullivan et al., 2003) that go hand-in-hand with differential degrees of susceptibility of different host types to the various phage lineages (this study, Sullivan et al., 2003) indicates a great deal of complexity for host–virus interactions in the oceans. This complexity is not only a function of the intrinsic differences in host susceptibility per se, but is also related to the co-evolutionary process between cyanobacteria and their phages that leads to continuous host diversification (Avrani et al., 2011; Marston et al., 2012). In fact, it is likely that these co-evolutionary processes have led to the differences in host susceptibility patterns observed here.

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology

Cyanophage abundance, composition and host ranges Experimental procedures Sampling Water samples were collected from Station A (29° 28′ N, 34° 55′ E) in the Gulf of Aqaba, Red Sea in 10 l Niskin bottles and transferred into amber collection bottles. Samples from the IUI pier were taken directly into amber collection bottles. For estimating phage abundances using the plaque assay, samples were filtered through 0.2 μm acrodisc syringe filters (Pall Life Sciences), and the filtrate was kept at 4–10°C in 25 ml acid-washed glass tubes (Moore et al., 2007) during transportation to the lab and prior to plating. The samples were plated within 24 h of collection.

Cyanobacterial growth Prochlorococcus strains were grown in the seawater-based Pro99 medium (Moore et al., 2007) using seawater collected from the Mediterranean Sea. Synechococcus strains were grown in an artificial seawater (ASW) medium (Wyman et al., 1985; Lindell et al., 1998). All strains were grown at 22°C under 5–15 μmol photons·m−2·s−1 white light with a 14:10 h light : dark cycle. Cyanobacterial growth was monitored by chlorophyll a fluorescence as a proxy for biomass using either a Turner Designs 10AU field fluorometer (Ex/Em: 340– 500 nm/ > 665 nm) when grown in glass tubes, or with a BioTek Synergy 2 microplate reader (Ex/Em: 440/680 nm) when grown in microtiter plates.

Plaque assays and phage lysate enrichment The filtrate of the seawater samples were serially diluted in autoclaved seawater prior to plating with exponentially growing host cells in pour plates (Brahamsha, 1996; Moore et al., 2007) with a number of changes as described in Lindell (2014). Briefly, 0.28% ultrapure low melting point agarose (Invitrogen) in Pro99 or ASW medium, with the addition of 1 mM sodium sulfite, was cooled to below 30°C, and combined with the cyanobacterial host and serially diluted seawater samples. For lawns of Prochlorococcus strains, the Alteromonas sp. EZ55 helper strain was also added (Morris et al., 2008; Lindell, 2014). Plates were then incubated under growth conditions, and plaques were counted three to four times over a 7- to 14-day period until no new plaques appeared. All samples were plated within 24 h of sample collection, except for the decay experiments which where plated at the stated times after collection (see below). Three to four replicate plaque assays were carried out per sample. Approximately 60 well-separated plaques on each host were randomly picked from the highest dilution plates in which plaques formed and were placed into 50 μl of host growth medium in wells of microtiter plates. These phage isolates were enriched by adding more host cells and medium and allowing the wells to clear. The original plaques or enriched liquid lysates were used for PCR analysis of phage type, and the enriched lysates were used for host range tests.

Host range Host range assays were carried out with the enriched lysates produced from the primary plaque isolates obtained from the

9

60 m depth samples from Station A from March 2009 and August 2010. We did not isolate the phages by triple plaque purification prior to the assays as we wished to assess a large number of phages that were representative of the phages in these samples. Prior isolation would have left us with much fewer phages to investigate due to loss during the isolation process and would have prevented the assessment of nearly as many phages. Phage lysates originating from each plaque were added to exponentially growing cultures from all six hosts. Host growth was monitored by chlorophyll a fluorescence and compared with control wells containing uninfected host cultures. Well clearings indicated lysis of the host. Only phages that maintained the capacity to infect the original host were used; approximately 25% of the plaques lost infectivity. Plaques containing a mix of phage types (15%), as determined by positive PCR amplification of multiple phage types from the one plaque, were not used for the assessment of host range. However, we cannot rule out the possibility that some wells contained a mix of known and unknown phages, the latter of which would not be detected by PCR. This left us with a total of 422 phages to use for the investigation of host range out of the 660 originally picked plaques.

Determining phage family composition by PCR PCR assays were carried out on each plaque (re-suspended in 50 μl medium) or enriched lysate for each of the three signature genes for the T4-like, T7-like and TIM5-like phage families respectively. Enriched lysates were used when no amplicon was obtained from the original plaques for either of the T4-like or T7-like genes, which have a threshold of detection of 104 phage particles per PCR reaction and were assayed first. Enriched lysates yielded PCR fragments in 25% of these cases. PCR for the TIM5-like family was carried out on the enriched lysates. The T4-like myoviruses were targeted by PCR with primers for the g20 portal protein gene, using degenerate primers CPS1.1 and CPS8.1 following Sullivan and colleagues (2008), which produce a 592 bp fragment; the T7-like podoviruses were targeted with a mix of four degenerate primers for the DNA polymerase gene as described previously (Dekel-Bird et al., 2013): DPOL_341Fd and DPOL_534Rd, together with primers DPOL_349Fd and DPOL_533Rd modified from Chen and colleagues (2009), which yield a 579 bp fragment; and the TIM5-like myoviruses were targeted with primers for the DNA polymerase genes that are nearly identical to those used previously (Sabehi et al., 2012): DPOL2_1F and DPOL2_R that yield a 378 bp fragment. While the same gene is targeted for the T7-like and TIM5-like phages, these genes are highly diverged between the phage types such that the primers are specific for each lineage. See Table S1 for a list of the PCR primers used in this study. Representative PCR fragments (generally 5–10 per phage type for each sampling date) were sequenced and compared with known cyanophage sequences to verify that the amplicons resulted in the expected gene for each family. This was the case for all amplicons except for phages isolated on the WH8109 host from the August 2010 cruise: positive fragments with the T7-like primers were obtained, but sequencing indicated that they were not T7-like DNA polymerase sequences. Therefore, the WH8109 plaques that pro-

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology

10

N. P. Dekel-Bird, G. Sabehi, B. Mosevitzky and D. Lindell

duced a fragment with these primers on this sampling date were considered ‘unknown’ phages. PCR amplifications were carried out for the T4-like g20, the T7-like DNA polymerase and the TIM5-like DNA polymerase genes, as described previously in Sullivan and colleagues (2008), Dekel-Bird and colleagues (2013), and Sabehi and colleagues (2012) respectively. Phage lysate (1 μl) was used as the template with 1.25–2 μM of forward and reverse primers in the presence of 200 μM dNTPs, 1X OptiBuffer and 1–4 units BIO-X-ACT Short DNA polymerase (Bioline), without the addition of magnesium chloride, in 25 μl reactions. See Table S2 for differences in the conditions used for the three signature genes. Thirty-five to forty cycles were carried out using the conditions outlined in Table S2 for each signature gene on a DNA Engine Peltier Thermal Cycler (Bio-Rad). An initial 5 min denaturation step at 95°C was performed prior to cycling, and a 5–10 min extension at the elongation temperature followed the cycling.

Sample variance In order to determine how representative samples are of a particular sampling period, cyanophage abundances from

Fig. 5. Variance of cyanophage abundances. Cyanophage abundances were determined for five samples collected 2 min apart (V1–V5) at the pier site in August (A, C) and September (B, D) of 2012 with Prochlorococcus MED4 (A, B) and Synechococcus WH8102 (C, D) as hosts. Variances ranged between 7% and 11% for each sampling date on each of the hosts. Abundances within a sampling cluster on the same date were not significantly different from each other: P = 0.28 for (A), P = 0.55 for (B), P = 0.68 for (C) and P = 0.07 for (D). In comparison, Prochlorococcus phage abundances were significantly higher in September than in August (P < 0.01), whereas no significant differences were found for Synechococcus phage abundances between these two sampling dates (P = 0.29). Abundances are the average and standard deviation of three plaque assays per sample.

multiple samples were compared. Five samples were collected 2 min apart at the pier site in August and September 2012, and cyanophage abundances were assayed using a single host from each genus: Synechococcus WH8102 and Prochlorococcus MED4. Highly similar abundances were found for all five samples on each of the hosts for each sampling date (Fig. 5), with a coefficient of variance of 7–11%. While significantly more Prochlorococcus phages were found in September relative to August of 2012, Synechococcus phage abundances were not significantly different on these two sampling dates. These results indicate that cyanophage abundances from a single sample are representative of abundances found over at least a short time period, and that differences between samples and sampling dates can be clearly discerned.

Decay of infective phage using different storage vessels Following the finding that the number of plaques declined when re-plated 10 days after the initial plating in August of 2009 (see Results and discussion), we carried out a more extensive assessment of the temporal decay of infective cyanophages over a 6-month period using Synechococcus WH8102 and Prochlorococcus MED4 as hosts during which time the samples were stored at 4°C in the dark in glass tubes. In all cases, we found a significant decline in cyanophage abundances within the first month (P < 0.01) (Fig. 6). In some cases, this was apparent within the first week, while in others the decline was observed between the first and fourth week. Abundances generally declined less after the first month (Fig. 6). The reduction in infective phage abundances was observed for sampling on different dates and stations, and when vastly different initial phage abundances were present (ranging from 1600 to 24 500 phages/ ml). Thus, even under common laboratory storage conditions without the damaging effects of UV light, infective cyanophage abundances declined rather quickly. These findings are in agreement with reports of a daily decay of 4–13% even in the absence of light (Suttle and Chen, 1992) and differ from reports of no significant decline in infective cyanophage over the period of a year (Sullivan et al., 2003). Differences in decay between the studies may be related to potential differences in phage composition in the different water samples, as differences in the rate of decay have been measured for different phages (Suttle and Chen, 1992). In order to determine whether different storage vessels may lead to less of a decline in cyanophage abundances, we compared storage in a variety of containers: glass tubes, coated glass tubes (Sigma-cote), and three types of plastic containers, namely sterile polypropylene 50 ml tubes (Greiner Bio-One), square polycarbonate containers (Nalgene) and high-density polyethylene rectangular amber bottles (Nalgene) using Synechococcus WH8102 as host. All containers were acid-washed except for the sterile polypropylene tubes. A similar and significant decline in phage abundances was found during 1–4 weeks for all sample containers [except the polycarbonate containers, which were significantly different only after 6 months (P = 0.0158) and for which the initial plating had a high standard deviation] (Fig. 6C). These findings indicate that storage of samples for even short periods of time is likely to underestimate phage abundances

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology

Cyanophage abundance, composition and host ranges

11

Fig. 6. Decay of infective cyanophages. (A, B) The per cent of initial cyanophages with time after sample collection when plated on Synechococcus WH8102 (A) and Prochlorococcus MED4 (B) as hosts. Samples were collected from the pier site at the surface or from Station A at 20 m depth on different sampling dates and stored at 4°C in the dark in acid-washed glass tubes. The date of sampling and site are shown on the x-axis, and the number of initial cyanophages (on day 1) is shown above the graphs in pfu (plaque forming units). (C) The number of cyanophages with time after sample collection in August 2012 when stored at 4°C in the dark in different storage vessels and plated on Synechococcus WH8102. PP, polypropylene; PC, polycarbonate; HDPE, high-density polyethylene. Abundances are the average and standard deviation of three to four plaque assays per sample. t-Tests or Mann–Whitney tests (see Experimental procedures) were used to assess differences in phage abundances between day 1 and other assay times: *P < 0.05, **P < 0.01, ***P < 0.001. Note that the value of alpha after the Bonferroni correction is 0.0167. Abundances declined significantly within the first month for all sampling dates and for all containers, except for the polycarbonate containers which had a high standard deviation for the first sample point and was significantly different only after 180 days (P = 0.0158).

by more than two to threefold. Further work is needed to determine whether freezing of samples with a cryoprotectant, such as glycerol or dimethyl sulphoxide, prevents the decay in infective cyanophages. Our findings suggest that until an appropriate storage procedure is found, it is best to assay samples for infective phage as soon as possible and at the same relative time after collection for all samples in comparative studies.

Phylogenetic analyses Multiple amino acid sequences were aligned using the CLUSTALX1.81 or MUSCLE programmes and were verified visually. Phylogenetic analysis was carried out using the Phylogeny.fr website (Dereeper et al., 2008). Amino acid sequences were used for the T7-like DNA polymerase tree (150 positions) and the T4-like g20 portal protein (95 positions) trees. The TIM5-like DNA polymerase tree was based on nucleotide sequences (318 positions) and was aligned according to the amino acid alignment. Neighbour-joining (NJ) analyses were performed with the distance programme ProtDist/FastDist + BioNJ, and the maximum likelihood (ML) analyses were performed using the PHYLML programme. Tree topologies were drawn with TreeFig. The overall topologies were the same for both NJ and ML gene trees. Bootstrap re-samplings were performed to obtain confidence estimates for inferred tree topologies, with 1000 re-samplings carried out for NJ trees and 400 for ML trees.

Statistical analyses Data were tested for normality using the Kolmogorov– Smirnov and Shapiro–Wilk tests. t-Tests were used to test the differences between the abundances of phages infecting Prochlorococcus relative to those infecting Synechococcus for each cruise, and for phages infecting a single genus on the different sampling dates, using pooled data from all stations and hosts within a genus, after being found to be normally distributed. Statistical analyses for differences between phage abundances at different sites for a single host during summer stratification were not carried out as only single samples were collected from each station on each sampling date. Thus, the differences that repeated themselves in both years are reported as trends. One-way analysis of variance was carried out to assess sample variance after being found to be normally distributed. For analysis of the decay of infective cyanophage between day 1 and other assay times, t-tests were used when the data were normally distributed, whereas Mann–Whitney tests were used when the data were not normally distributed, followed by Bonferroni corrections for three multiple testings. All tests were carried out using SPSS 15 (Rel. 15.0.1. November 2006, SPSS, Chicago).

Nucleotide sequence accession numbers The new sequences presented in the phylogenetic trees and described in Table S3 have been submitted to the GenBank

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology

12

N. P. Dekel-Bird, G. Sabehi, B. Mosevitzky and D. Lindell

database. The T7-like DNA polymerase sequences appear under the accession numbers KM113808-KM113824. The T4-like g20 (portal protein) sequences appear under the accession numbers KM113829-KM113842. The TIM5-like DNA polymerase sequences appear under the accession numbers KM113825-KM113828.

Acknowledgements We thank the Interuniversity Institute for Marine Sciences in Eilat (IUI) for use of facilities, and the National Monitoring Program for allowing us to participate in cruises. We also thank Irena Pekarsky for help with the decay experiments, Alisa Zhilin for technical assistance, Sarit Avrani for help with statistics, Daniel Schwartz and Sarit Avrani for comments on the manuscript, and Lindell lab members for discussions. This research was funded by a European Council FP6 Marie Curie Reintegration grant (#046549) and an Israel Science Foundation Individual grant (749/11) to D.L.

References Ackermann, H. (2003) Bacteriophage observations and evolution. Res Microbiol 154: 245–251. Ahlgren, N.A., and Rocap, G. (2012) Diversity and distribution of marine Synechococcus: multiple gene phylogenies for consensus classification and development of qPCR assays for sensitive measurement of clades in the ocean. Front Microbiol 3: 213. Angly, F.E., Felts, B., Breitbart, M., Salamon, P., Edwards, R.A., Carlson, C., et al. (2006) The marine viromes of four oceanic regions. PLoS Biol 4: e368. Avrani, S., Wurtzel, O., Sharon, I., Sorek, R., and Lindell, D. (2011) Genomic island variability facilitates Prochlorococcus-virus coexistence. Nature 474: 604–608. Bench, S.R., Hanson, T.E., Williamson, K.E., Ghosh, D., Radosovich, M., Wang, K., and Wommack, K.E. (2007) Metagenomic characterization of Chesapeake Bay virioplankton. Appl Environ Microbiol 73: 7629–7641. Brahamsha, B. (1996) A genetic manipulation system for oceanic cyanobacteria of the genus Synechococcus. Appl Environ Microbiol 62: 1747–1751. Carlson, D.F., Fredj, E., and Gildor, H. (2013) The annual cycle of vertical mixing and restratification in the Northern Gulf of Eilat/Aqaba (Red Sea) based on high temporal and vertical resolution observations. Deep Sea Res 84: 1– 17. Chen, F., and Lu, J. (2002) Genomic sequence and evolution of marine cyanophage P60: a new insight on lytic and lysogenic phages. Appl Environ Microbiol 68: 2589–2594. Chen, F., Wang, K., Huang, S., Cai, H., Zhao, M., Jiao, N., and Wommack, K.E. (2009) Diverse and dynamic populations of cyanobacterial podoviruses in the Chesapeake Bay unveiled through DNA polymerase gene sequences. Environ Microbiol 11: 2884–2892. Dekel-Bird, N.P., Avrani, S., Sabehi, G., Pekarsky, I., Marston, M.F., Kirzner, S., and Lindell, D. (2013) Diversity and evolutionary relationships of T7-like podoviruses infecting marine cyanobacteria. Environ Microbiol 15: 1476–1491.

Dereeper, A., Guignon, V., Blanc, G., Audic, S., Buffet, S., Chevenet, F., et al. (2008) Phylogeny.fr: robust phylogenetic analysis for the non-specialist. Nucleic Acids Res 36: W465–W469. DuRand, M.D., Olson, R.J., and Chisholm, S.W. (2001) Phytoplankton population dynamics at the Bermuda Atlantic time-series station in the Sargasso Sea. Deep Sea Research Part II 48: 1983–2003. Flombaum, P., Gallegos, J.L., Gordillo, R.A., Rincón, J., Zabala, L.L., Jiao, N., et al. (2013) Present and future global distributions of the marine Cyanobacteria Prochlorococcus and Synechococcus. PNAS 110: 9824– 9829. Fuller, N.J., West, N.J., Marie, D., Yallop, M., Rivlin, T., Post, A.F., and Scanlan, D.J. (2005) Dynamics of community structure and phosphate status of picocyanobacterial populations in the Gulf of Aqaba, Red Sea. Limnol Oceanogr 50: 363–375. Huang, S., Wang, K., Jiao, N., and Chen, F. (2012) Genome sequences of siphoviruses infecting marine Synechococcus unveil a diverse cyanophage group and extensive phage–host genetic exchanges. Environ Microbiol 14: 540–558. Johnson, Z.I., Zinser, E.R., Coe, A., McNulty, N.P., Woodward, E.M.S., and Chisholm, S.W. (2006) Niche partitioning among Prochlorococcus ecotypes along ocean-scale environmental gradients. Science 311: 1737– 1740. Labrie, S., Frois-Moniz, K., Osburne, M., Kelly, L., Roggensack, S., Sullivan, M., et al. (2013) Genomes of marine cyanopodoviruses reveal multiple origins of diversity. Environ Microbiol 15: 1356–1376. Lavigne, R., Seto, D., Mahadevan, P., Ackermann, H.-W., and Kropinski, A.M. (2008) Unifying classical and molecular taxonomic classification: analysis of the Podoviridae using BLASTP-based tools. Res Microbiol 159: 406–414. Lavigne, R., Darius, P., Summer, E.J., Seto, D., Mahadevan, P., Nilsson, A.S., et al. (2009) Classification of Myoviridae bacteriophages using protein sequence similarity. BMC Microbiol 9: 224. Lindell, D. (2014) The genus Prochlorococcus, phylum cyanobacteria. In The Prokaryotes – Other Major Lineages of Bacteria and the Archaea, 4th edn. Rosenberg, E., Thompson, E.S.F., DeLong, E.F., Lory, S., Stackebrandt, E., and Thompson, F. (eds). Berlin, Germany: SpringerVerlag. In press. Lindell, D., and Post, A.F. (1995) Ultraphytoplankton succession is triggered by deep winter mixing in the Gulf of Aqaba (Eilat), Red Sea. Limnol Oceanogr 40: 1130–1141. Lindell, D., Padan, E., and Post, A.F. (1998) Regulation of ntcA expression and nitrite uptake in the marine Synechococcus sp. strain WH 7803. J Bacteriol 180: 1878–1886. Lindell, D., Sullivan, M.B., Johnson, Z.I., Tolonen, A.C., Rohwer, F., and Chisholm, S.W. (2004) Transfer of photosynthesis genes to and from Prochlorococcus viruses. Proc Natl Acad Sci USA 101: 11013–11018. Lindell, D., Jaffe, J.D., Coleman, M.L., Futschik, M.E., Axmann, I.M., Rector, T., et al. (2007) Genome-wide expression dynamics of a marine virus and host reveal features of co-evolution. Nature 449: 83–86.

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology

Cyanophage abundance, composition and host ranges Malmstrom, R.R., Coe, A., Kettler, G.C., Martiny, A.C., Frias-Lopez, J., Zinser, E.R., and Chisholm, S.W. (2010) Temporal dynamics of Prochlorococcus ecotypes in the Atlantic and Pacific oceans. ISME J 4: 1252–1264. Mann, N.H., Clokie, M.R., Millard, A., Cook, A., Wilson, W.H., Wheatley, P.J., et al. (2005) The genome of S-PM2, a ‘photosynthetic’ T4-type bacteriophage that infects marine Synechococcus strains. J Bacteriol 187: 3188–3200. Marston, M.F., and Sallee, J.L. (2003) Genetic diversity and temporal variation in the cyanophage community infecting marine Synechococcus species in Rhode Island’s coastal waters. Appl Environ Microbiol 69: 4639–4647. Marston, M.F., Pierciey, F.J., Shepard, A., Gearin, G., Qi, J., Yandava, C., et al. (2012) Rapid diversification of coevolving marine Synechococcus and a virus. PNAS 109: 4544–4549. Matteson, A.R., Rowe, J.M., Ponsero, A.J., Pimentel, T.M., Boyd, P.W., and Wilhelm, S.W. (2013) High abundances of cyanomyoviruses in marine ecosystems demonstrate ecological relevance. FEMS Microbiol Ecol 84: 223–234. McDaniel, L.D., DelaRosa, M., and Paul, J.H. (2006) Temperate and lytic cyanophages from the Gulf of Mexico. J Mar Biolog Assoc UK 86: 517–527. Millard, A.D., and Mann, N.H. (2006) A temporal and spatial investigation of cyanophage abundance in the Gulf of Aqaba, Red Sea. J Mar Biolog Assoc UK 86: 507– 515. Millard, A.D., Zwirglmaier, K., Downey, M.J., Mann, N.H., and Scanlan, D.J. (2009) Comparative genomics of marine cyanomyoviruses reveals the widespread occurrence of Synechococcus host genes localized to a hyperplastic region: implications for mechanisms of cyanophage evolution. Environ Microbiol 11: 2370–2387. Moore, L.R., Coe, A., Zinser, E.R., Saito, M.A., Sullivan, M.B., Lindell, D., et al. (2007) Culturing the marine cyanobacterium Prochlorococcus. Limnol Oceanogr Methods 5: 353–362. Morris, J.J., Kirkegaard, R., Szul, M.J., Johnson, Z.I., and Zinser, E.R. (2008) Facilitation of robust growth of Prochlorococcus colonies and dilute liquid cultures by ‘helper’ heterotrophic bacteria. Appl Environ Microbiol 74: 4530–4534. Mühling, M., Fuller, N.J., Millard, A., Somerfield, P.J., Marie, D., Wilson, W.H., et al. (2005) Genetic diversity of marine Synechococcus and co-occurring cyanophage communities: evidence for viral control of phytoplankton. Environ Microbiol 7: 499–508. Olson, R.J., Chisholm, S.W., Zettler, E.R., Altabet, M.A., and Dusenberry, J.A. (1990) Spatial and temporal distributions of prochlorophyte picoplankton in the North Atlantic Ocean. Deep Sea Res Part A 37: 1033–1051. Partensky, F., Blanchot, J., and Vaulot, D. (1999) Differential distribution and ecology of Prochlorococcus and Synechococcus in oceanic waters: a review. Bulletin de l’Institut Oceanographie, Monaco 19: 457–475. Penno, S., Lindell, D., and Post, A.F. (2006) Diversity of Synechococcus and Prochlorococcus populations determined from DNA sequences of the N-regulatory gene. ntcA. Environ Microbiol 8: 1200–1211. Ponsero, A.J., Chen, F., Lennon, J.T., and Wilhelm, S.W. (2013) Complete genome sequence of cyanobacterial

13

siphovirus KBS2A. Genome Announcements 1: e00472– 00413. Pope, W.H., Weigele, P.R., Chang, J., Pedulla, M.L., Ford, M.E., Houtz, J.M., et al. (2007) Genome sequence, structural proteins, and capsid organization of the cyanophage Syn5: A ‘Horned’ bacteriophage of marine Synechococcus. J Mol Biol 368: 966–981. Post, A.F., Penno, S., Zandbank, K., Paytan, A., Huse, S.M., and Welch, D.M. (2011) Long term seasonal dynamics of Synechococcus population structure in the Gulf of Aqaba, Northern Red Sea. Front Microbiol 2: 131. Proctor, L.M., and Fuhrman, J.A. (1990) Viral mortality of marine bacteria and cyanobacteria. Nature 343: 60–62. Rodriguez-Valera, F., Martin-Cuadrado, A.-B., RodriguezBrito, B., Pasic, L., Thingstad, T.F., Rohwer, F., and Mira, A. (2009) Explaining microbial population genomics through phage predation. Nat Rev Microbiol 7: 828–836. Sabehi, G., Shaulov, L., Silver, D.H., Yanai, I., Harel, A., and Lindell, D. (2012) A novel lineage of myoviruses infecting cyanobacteria is widespread in the oceans. PNAS 109: 2037–2042. Sullivan, M.B., Waterbury, J.B., and Chisholm, S.W. (2003) Cyanophages infecting the oceanic cyanobacterium Prochlorococcus. Nature 424: 1047–1051. Sullivan, M.B., Coleman, M.L., Weigele, P., Rohwer, F., and Chisholm, S.W. (2005) Three Prochlorococcus cyanophage genomes: signature features and ecological interpretations. PLoS Biol 3: e144. Sullivan, M.B., Krastins, B., Hughes, J.L., Kelly, L., Chase, M., Sarracino, D., and Chisholm, S.W. (2009) The genome and structural proteome of an ocean siphovirus: a new window into the cyanobacterial ‘mobilome’. Environ Microbiol 11: 2935–2951. Sullivan, M.B., Coleman, M.L., Quinlivan, V., Rosenkrantz, J.E., DeFrancesco, A.S., Tan, G., et al. (2008) Portal protein diversity and phage ecology. Environ Microbiol 10: 2810–2823. Sullivan, M.B., Huang, K.H., Ignacio-Espinoza, J.C., Berlin, A.M., Kelly, L., Weigele, P.R., et al. (2010) Genomic analysis of oceanic cyanobacterial myoviruses compared with T4-like myoviruses from diverse hosts and environments. Environ Microbiol 12: 3035–3056. Suttle, C.A., and Chan, A.M. (1993) Marine cyanophages infecting oceanic and coastal strains of Synechococcus: abundance, morphology, cross-infectivity and growth characteristics. Mar Ecol Prog Ser 92: 99–99. Suttle, C.A., and Chan, A.M. (1994) Dynamics and distribution of cyanophages and their effect on marine Synechococcus spp. Appl Environ Microbiol 60: 3167–3174. Suttle, C.A., and Chen, F. (1992) Mechanisms and rates of decay of marine viruses in seawater. Appl Environ Microbiol 58: 3721–3729. Tai, V., and Palenik, B. (2009) Temporal variation of Synechococcus clades at a coastal Pacific Ocean monitoring site. ISME J 3: 903–915. Wang, K., and Chen, F. (2004) Genetic diversity and population dynamics of cyanophage communities in the Chesapeake Bay. Aquat Microb Ecol 34: 105–116. Wang, K., and Chen, F. (2008) Prevalence of highly hostspecific cyanophages in the estuarine environment. Environ Microbiol 10: 300–312.

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology

14

N. P. Dekel-Bird, G. Sabehi, B. Mosevitzky and D. Lindell

Wang, K., Wommack, K.E., and Chen, F. (2011) Abundance and distribution of Synechococcus spp. and cyanophages in the Chesapeake Bay. Appl Environ Microbiol 77: 7459– 7468. Waterbury, J.B., and Valois, F.W. (1993) Resistance to co-occurring phages enables marine Synechococcus communities to coexist with cyanophages abundant in seawater. Appl Environ Microbiol 59: 3393– 3399. Weigele, P.R., Pope, W.H., Pedulla, M.L., Houtz, J.M., Smith, A.L., Conway, J.F., et al. (2007) Genomic and structural analysis of Syn9, a cyanophage infecting marine Prochlorococcus and Synechococcus. Environ Microbiol 9: 1675–1695. Wolf-Vecht, A., Paldor, N., and Brenner, S. (1992) Hydrographic indications of advection/convection effects in the Gulf of Eilat. Deep Sea Res 39: 1393–1401. Wyman, M., Gregory, R., and Carr, N. (1985) Novel role for phycoerythrin in a marine cyanobacterium, Synechococcus strain DC2. Science 230: 818–820. Zinser, E.R., Johnson, Z.I., Coe, A., Karaca, E., Veneziano, D., and Chisholm, S.W. (2007) Influence of light and temperature on Prochlorococcus ecotype distributions in the Atlantic Ocean. Limnol Oceanogr 52: 2205–2220. Zwirglmaier, K., Jardillier, L., Ostrowski, M., Mazard, S., Garczarek, L., Vaulot, D., et al. (2008) Global phylogeography of marine Synechococcus and Prochlorococcus reveals a distinct partitioning of lineages among oceanic biomes. Environ Microbiol 10: 147–161.

Supporting information Additional Supporting Information may be found in the online version of this article at the publisher’s web-site: Fig. S1. Clustering patterns of T7-like cyanophages based on the DNA polymerase gene. The neighbour-joining (NJ) tree was inferred from 150 amino acids. Bootstrap values greater than 50% for NJ and maximum likelihood (ML) trees are shown (NJ/ML). Phages are listed by their name followed by the name of the host used for isolation: red and green boxes indicate Synechococcus and Prochlorococcus hosts respectively. The new sequences reported in this study are shown in blue type and were amplified from a random collection of primary phage isolates from the different cruises (see Table S3). The digits to the right of certain phages indicate

the number of sequences that were greater than 99% identical to each other. The clade and subclade designations to the right of the tree follow those in Dekel-Bird and colleagues (2013). The archetype T7 phage was used as an outgroup. Note that all the sequences from this study cluster with clade B phages and are dispersed throughout this clade. Fig. S2. Clustering patterns of T4-like cyanophages based on the portal protein (g20) gene. The neighbour-joining (NJ) tree was inferred from 95 amino acids. Bootstrap values greater than 50% for NJ and maximum likelihood (ML) trees are shown (NJ/ML). Phages are listed by their name followed by the name of the host used for isolation: red and green boxes indicate Synechococcus and Prochlorococcus hosts respectively. The new sequences reported in this study are shown in blue type and were amplified from a random collection of primary phage isolates from the different cruises (see Table S3). The ‘x2’ to the right of the 7Gmp sequence indicates that two sequences with greater than 99% identity to each other were found. The clade designations to the right of the tree follow those in Sullivan and colleagues (2008). The archetype T4 phage sequence was used as an outgroup. Note that sequences from this study are dispersed throughout the tree and are found in all four cyanophage containing clades. Fig. S3. Clustering patterns of TIM5-like cyanophages based on the DNA polymerase gene. The neighbour-joining (NJ) tree was inferred from 318 nucleotides. Bootstrap values greater than 50% for NJ and maximum likelihood (ML) trees are shown (NJ/ML). Phages are listed by their name followed by the name of the host used for isolation: red boxes indicate that only Synechococcus hosts were found. The new sequences reported in this study are shown in blue type and were amplified from a random collection of primary phage isolates from the different cruises (see Table S3). The digits to the right of certain phages indicate the number of sequences that were 100% identical to each other. ECV67914, ECV33575 and ECW18666 are environmental sequences. The mitochondrial DNA polymerase genes from Homo sapiens, Xenopus laevis and Saccharomyces cerevisiae were used as the outgroup. Note the high similarity of the sequences to the S-TIM5 archetype phage. Table S1. PCR primers used in this study. Table S2. PCR conditions used in this study. Table S3. Signature gene sequences from phages and primary isolates collected from the four cruises reported in this study.

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology