The first known virus isolates from Antarctic sea ice have complex infection patterns

The first known virus isolates from Antarctic sea ice have complex infection patterns Abstract Viruses are recognized as important actors in ocean ecology and biogeochemical cycles, but many details are not yet understood. We participated in a winter expedition to the Weddell Sea, Antarctica, to isolate viruses and to measure virus-like particle abundance (flow cytometry) in sea ice. We isolated 59 bacterial strains and the first four Antarctic sea-ice viruses known (PANV1, PANV2, OANV1 and OANV2), which grow in bacterial hosts belonging to the typical sea-ice genera Paraglaciecola and Octadecabacter. The viruses were specific for bacteria at the strain level, although OANV1 was able to infect strains from two different classes. Both PANV1 and PANV2 infected 11/15 isolated Paraglaciecola strains that had almost identical 16S rRNA gene sequences, but the plating efficiencies differed among the strains, whereas OANV1 infected 3/7 Octadecabacter and 1/15 Paraglaciecola strains and OANV2 1/7 Octadecabacter strains. All the phages were cold-active and able to infect their original host at 0°C and 4°C, but not at higher temperatures. The results showed that virus–host interactions can be very complex and that the viral community can also be dynamic in the winter-sea ice. virus-host interactions, virus isolation, sea ice microbiology, VLP in sea ice INTRODUCTION Almost 10% of the world's ocean is covered by sea ice at least once per year, which makes it one of the largest biomes on Earth (Dieckmann and Hellmer 2010). Although being a cold and harsh environment, sea ice is full of life. Specialized organisms live inside brine channels and pockets that are formed during freezing conditions, when salts and nutrients from the seawater become concentrated between the ice crystals (Thomas and Dieckmann 2002). Brine remains a liquid inside the ice, due to its high salinity, and this makes it a suitable habitat for the sea-ice microbial community, which comprises protists, bacteria, archaea and their viruses (Maranger, Bird and Juniper 1994; Mock and Thomas 2005; Arrigo, Mock and Lizotte 2010; Deming and Collins 2017). Here, we use the term bacteria instead of prokaryotes, even if in some cases archaea may also be involved. Microbes affect the biogeochemical properties of the sea ice, gas exchange between the ocean and atmosphere, and provide food for ice-associated animals, e.g. krill (Arrigo and Thomas 2004). Viruses are the most abundant biological entities and are presumed to play important roles in the biogeochemical cycles of the oceans (Fuhrman 1999; Suttle 2007). The most commonly found viruses, bacteriophages (phages) i.e. viruses infecting bacteria, are possibly the main cause of bacterial mortality (Weinbauer 2004; Suttle 2007). Since viruses can multiply only within their host cells, their activity is dependent on the abundance and activity of their hosts (Maranger, Bird and Juniper 1994; Marchant et al. 2000). Due to their host specificity, they are crucial to the control of bacterial community composition and activity (Proctor and Fuhrman 1990; Wommack and Colwell 2000; Suttle 2005; 2007). However, most studies of marine environments are conducted in the water column, whereas knowledge of viruses and their functions in sea ice are still very limited. The numbers of virus-like particles (VLPs) measured, range between 105 and 108 ml−1 in bulk Arctic and Antarctic sea ice from spring to autumn. The lowest values have been observed during the winter in Antarctic bulk ice (Paterson and Laybourn-Parry 2012), whereas the highest numbers of VLPs occur during freezing or spring algal mass growth (Maranger, Bird and Juniper 1994; Collins and Deming 2011). Moreover, VLPs may also contain particles other than viruses, e.g. gene transfer agents or membrane vesicles (Forterre et al. 2013; Soler et al. 2015). VLPs are positively correlated with bacterial abundance, activity and chlorophyll-a (chl-a) concentrations (Maranger, Bird and Juniper 1994; Gowing et al. 2002, 2004). In aquatic environments, a typical virus-to-bacteria ratio (VBR; more precisely VLP-to-prokaryotic cell ratio) is 10:1 (Maranger and Bird 1995). Bacterial and viral density dictates their contact rate, which is one of the key controls in virus–host interactions. The semi-enclosed environment of brine channels may increase this contact rate, especially during winter, when the brine channels are narrower and the brine even more concentrated (Wells and Deming 2006a). Sea ice may, therefore, be a place where virus–host interactions can be enhanced, compared with the open ocean. To understand these effects, virus–host systems need to be isolated to examine their interactions in detail. To the best of our knowledge, only three virus–host systems to date have been isolated from Arctic and seven from Baltic Sea ice (Borriss et al. 2003; Luhtanen et al. 2014, Yu et al. 2015), but none from Antarctic sea ice. The viruses isolated represented different phage morphologies. The Arctic virus isolates Shewanella phage 1a and Colwellia phage 21c are icosahedral viruses with either a contractile or noncontractile tail, resembling double-stranded DNA (dsDNA) phages of the order Caudovirales (Borriss et al. 2003; 2007). The filamentous nonlytic phage f327 from the Arctic infects a Pseudoalteromonas strain and is reminiscent of viruses in the family Inoviridae (Yu et al. 2015). In addition, seven tailed icosahedral dsDNA phages infecting strains from either Flavobacterium or Shewanella were isolated from Baltic Sea ice (Luhtanen et al. 2014; Senčilo et al. 2015). All virulent sea-ice phage isolates have a narrow host range, are cold-active (capable of infection and production at ≤4°C; Wells and Deming (2006b)), and produce plaques (clear zones on bacterial lawns used to determine the number of infectious viruses) only at the lower end of their host bacterial temperature growth range (Borriss et al. 2003; Luhtanen et al. 2014). In addition, a cold-active siphophage 9A was isolated from Arctic nepheloid layer seawater (Wells and Deming 2006b) for Colwellia psychrerythraea strain 34H, isolated originally from Arctic shelf sediments (Huston, Krieger-Brockett and Deming 2000). It was reminiscent of the isolates from sea ice, because it also has a narrow host range, is cold-active, and has a more restricted growth temperature range than the host bacteria. Here, we report the isolation of the first cultivable phage–host systems and VLP abundance from the winter-sea ice in the Weddell Sea, Antarctica. Studying phage–host systems gives us valuable information on sea-ice microbial communities and their potential roles in the sea-ice ecosystem. MATERIALS AND METHODS Sea-ice sampling and ice properties Pack ice samples were collected from the Weddell Sea (Antarctica) during the austral winter as part of the Antarctic Winter Ecosystem Climate Study (AWECS) aboard R/V Polarstern in June–August 2013 (leg ANT-XXIX/6). Sampling was performed either with a metal basket (pancake ice at stations 486, 488 and 489) or using a motorized, trace-metal-clean (electropolished steel) CRREL-type ice-coring auger (Lannuzel et al.2006), 14 cm in diameter. For this study, full-depth ice cores (one or two) were taken from 10 locations (ice-stations 486, 488, 489, 493, 496, 500, 503, 506, 515 and 517; Table 1; Fig. S1, Supporting Information), as described in Tison et al. (2017). Bulk ice was used, because some of the microbes may have been attached to the brine channel walls and could have been lost if only the liquid brine was sampled (Meiners, Krembs and Gradinger 2008). The ice cores were cut into one, three, five or seven layers, depending on the ice thickness, and crushed gently with a hammer inside a polyethylene plastic bag. When two cores were taken, the corresponding layers of the nearby ice cores were pooled (Table 1). The VLP abundances were measured from these layers. The bacterial abundances (Table 1), bacterial production (measured as thymidine incorporation) and bacterial community composition analyses from these ice cores have been published elsewhere (Eronen-Rasimus et al. 2017). For the isolation work, we used the ice samples from first-year ice-station 500 and early second-year ice-station 515a (Tison et al. 2017). The surface parts of the 515a core were removed to minimize contamination, and the core was cut into 12 layers (∼12–14 cm each; Table 2), using an electric-band saw sterilized with 70% ethanol. The 12 layers were used separately to isolate the bacterial strains. The bacterial community composition of ice core 515a has also been published under the name 515 T (0–56 cm), M (56–126 cm) and B (126–166 cm; Eronen-Rasimus et al. 2017). For virus isolation, bulk ice from station 500 and core 515a was used. The ice samples were left to melt in sterilized containers at 4°C overnight, after that the remaining ice was melted in a water bath with continuous shaking (Rintala et al. 2014). After melting, the samples were immediately transferred back to 4°C. Table 1. VLP and bacterial abundances and VBRs together with ice temperature and chlorophyll-a concentrations in the different layers of ice cores from all the AWECS sampling stations.         Ice depth (cm) from air-ice interface            Station  Date  Latitude  Longitude  Core I  Core II  Ice temperature °Cb  VLP x 105ml-1 in bulk iced  Bacteria x 105ml-1 in bulk icec,d  VBR Viruses/Bacteria  Total chl-a (µg l-1 bulk ice)b,d  486a  6/17/2013  −61.526  −0.086  0–6  –  −5.0  1.90  0.56  3.4  0.41  488a  6/18/2013  −62.928  −0.006  0–15  –  −10.9  4.90  0.56  8.8  0.29          15–20  –  −7.4  2.60  0.81  3.2  0.28          20–35  –  −4.1  6.80  1.20  5.7  1.39  489a  6/19/2013  −63.901  −0.031  0–15  –  −8.4  6.50  1.20  5.4  1.48          15–22  –  −5.8  4.90  1.30  3.8  3.89          22–37  –  −3.4  5.80  0.85  6.8  2.36  493  6/21/2013  −66.44  0.122  0–15  0–15  −8.7  13.00  3.40  3.8  9.56          15–46  15–38  −5.1  9.30  3.80  2.4  10.43          46–61  38–53  −2.7  9.30  1.80  5.2  16.59  496  6/24/2013  −67.466  −0.021  0–15  0–15  −5.3  4.70  1.30  3.6  1.81          15–45  15–57  −4.1  3.90  2.70  1.4  7.49          45–60  57–72  −2.6  5.90  2.50  2.4  15.24  500  7/3/2013  −67.949  −6.658  0–15  0–15  −2.4  16.00  2.60  6.2  3.83          15–35  15–35  −2.7  12.00  3.60  3.3  7.73          35–55  35–52  −2.6  46.00  4.00  11.5  10.34          55–75  52–72  −2.4  20.00  3.40  5.9  7.08          75–90  72–87  −2.2  9.70  2.70  3.6  9.31  503  7/8/2013  −67.187  −13.224  0–15  0–15  −7.3  8.70  0.65  13.4  0.83          15–25  15–25  −6.0  7.50  0.79  9.5  0.86          25–37  25–36  −5.0  9.20  0.81  11.4  1.05          37–47  36–46  −3.4  10.00  0.80  12.5  0.99          47–62  46–61  −2.2  6.70  0.56  12.0  0.70  506  7/11/2013  −67.19  −23.042  0–15  0–15  −6.5  3.90  0.51  7.6  0.31          15–34  15–30  −4.9  5.50  0.80  6.9  0.69          34–49  30–45  −3.1  ND  ND  ND  0.82  515a  7/26/2013  −63.456  −51.308  0–15  –  −7.6  6.90  3.60  1.9  0.78          15–45  –  −6.1  9.30  6.00  1.6  2.08          45–75  –  −6.4  21.00  7.30  2.9  30.73          75–104  –  −4.8  40.00  24.00  1.7  61.54          104–134  –  −3.9  49.00  42.00  1.2  62.57          134–164  –  −3.8  10.00  9.50  1.1  10.95          164–179  –  −2.3  8.90  12.00  0.7  11.66  517  7/30/2013  −63.509  −51.112  0–15  0–15  −3.7  5.70  0.60  9.5  0.74          15–30  15–30  −2.6  4.60  0.87  5.3  0.55          30–43  30–43  −2.2  2.20  0.71  3.1  0.22          43–58  43–58  −2.0  2.40  1.30  1.8  0.31          58–73  58–73  −1.8  5.80  1.40  4.1  0.60          Ice depth (cm) from air-ice interface            Station  Date  Latitude  Longitude  Core I  Core II  Ice temperature °Cb  VLP x 105ml-1 in bulk iced  Bacteria x 105ml-1 in bulk icec,d  VBR Viruses/Bacteria  Total chl-a (µg l-1 bulk ice)b,d  486a  6/17/2013  −61.526  −0.086  0–6  –  −5.0  1.90  0.56  3.4  0.41  488a  6/18/2013  −62.928  −0.006  0–15  –  −10.9  4.90  0.56  8.8  0.29          15–20  –  −7.4  2.60  0.81  3.2  0.28          20–35  –  −4.1  6.80  1.20  5.7  1.39  489a  6/19/2013  −63.901  −0.031  0–15  –  −8.4  6.50  1.20  5.4  1.48          15–22  –  −5.8  4.90  1.30  3.8  3.89          22–37  –  −3.4  5.80  0.85  6.8  2.36  493  6/21/2013  −66.44  0.122  0–15  0–15  −8.7  13.00  3.40  3.8  9.56          15–46  15–38  −5.1  9.30  3.80  2.4  10.43          46–61  38–53  −2.7  9.30  1.80  5.2  16.59  496  6/24/2013  −67.466  −0.021  0–15  0–15  −5.3  4.70  1.30  3.6  1.81          15–45  15–57  −4.1  3.90  2.70  1.4  7.49          45–60  57–72  −2.6  5.90  2.50  2.4  15.24  500  7/3/2013  −67.949  −6.658  0–15  0–15  −2.4  16.00  2.60  6.2  3.83          15–35  15–35  −2.7  12.00  3.60  3.3  7.73          35–55  35–52  −2.6  46.00  4.00  11.5  10.34          55–75  52–72  −2.4  20.00  3.40  5.9  7.08          75–90  72–87  −2.2  9.70  2.70  3.6  9.31  503  7/8/2013  −67.187  −13.224  0–15  0–15  −7.3  8.70  0.65  13.4  0.83          15–25  15–25  −6.0  7.50  0.79  9.5  0.86          25–37  25–36  −5.0  9.20  0.81  11.4  1.05          37–47  36–46  −3.4  10.00  0.80  12.5  0.99          47–62  46–61  −2.2  6.70  0.56  12.0  0.70  506  7/11/2013  −67.19  −23.042  0–15  0–15  −6.5  3.90  0.51  7.6  0.31          15–34  15–30  −4.9  5.50  0.80  6.9  0.69          34–49  30–45  −3.1  ND  ND  ND  0.82  515a  7/26/2013  −63.456  −51.308  0–15  –  −7.6  6.90  3.60  1.9  0.78          15–45  –  −6.1  9.30  6.00  1.6  2.08          45–75  –  −6.4  21.00  7.30  2.9  30.73          75–104  –  −4.8  40.00  24.00  1.7  61.54          104–134  –  −3.9  49.00  42.00  1.2  62.57          134–164  –  −3.8  10.00  9.50  1.1  10.95          164–179  –  −2.3  8.90  12.00  0.7  11.66  517  7/30/2013  −63.509  −51.112  0–15  0–15  −3.7  5.70  0.60  9.5  0.74          15–30  15–30  −2.6  4.60  0.87  5.3  0.55          30–43  30–43  −2.2  2.20  0.71  3.1  0.22          43–58  43–58  −2.0  2.40  1.30  1.8  0.31          58–73  58–73  −1.8  5.80  1.40  4.1  0.60  a Only one ice core sampled. b Ice temperatures and chl-a values from Tison et al. (2017). c Bacterial abundances are also published in Eronen-Rasimus et al. (2017). d Analysed from combined layers of the sibling cores I and II, when two cores were collected. VLP = virus-like particle, AWECS = Antarctic Winter Ecosystem Climate Study, chl-a = chlorophyll a, ND = not determined. View Large Table 2. Bacterial strains isolated in this study from the ice core 515a. Isolation depth from air–ice interface (cm)  Isolation mediaa  Bacterial strain  Closest match genusb  Identity % at genus level  Accession number  Used for virus isolation  0–14  Z  IceBac 363  Halomonas  100.0  KY194856  ✓  14–28  Z  IceBac 364  Polaribacter  99.7  KY194850  ✓  14–28  Z  IceBac 365  Glaciecola  99.6  KY194821  ✓  14–28  Z  IceBac 367  Paraglaciecola  99.9  KY194799  ✓  14–28  Z  IceBac 368  Paraglaciecola  99.9  KY194805  ✓  14–28  Z  IceBac 369  Marinobacter  99.3  KY194841    14–28  Z  IceBac 370  Marinobacter  99.3  KY194842  ✓  14–28  Z  IceBac 371  Glaciecola  99.6  KY194822    14–28  Z  IceBac 372  Paraglaciecola (H)  99.9  KY194800  ✓  14–28  MOX  IceBac 373  Polaribacter  100.0  KY194848  ✓  14–28  MOX  IceBac 377  Glaciecola  99.6  KY194823  ✓  28–42  Z  IceBac 378  Glaciecola  99.6  KY194828  ✓  28–42  Z  IceBac 379  Glaciecola  99.6  KY194824  ✓  28–42  Z  IceBac 380  Glaciecola  99.6  KY194825    28–42  Z  IceBac 381  Glaciecola  99.6  KY194829  ✓  28–42  Z  IceBac 382  Glaciecola  99.6  KY194831  ✓  28–42  Z  IceBac 383  Paraglaciecola  99.9  KY194801  ✓  28–42  MOX  IceBac 384  Polaribacter  100.0  KY194845    42–56  Z  IceBac 385  Polaribacter  99.7  KY194844    42–56  Z  IceBac 386  Glaciecola  99.6  KY194815  ✓  42–56  Z  IceBac 387  Glaciecola  99.5  KY194816  ✓  42–56  Z  IceBac 388  Glaciecola  99.6  KY194826  ✓  42–56  Z  IceBac 389  Glaciecola  99.6  KY194817  ✓  42–56  Z  IceBac 390  Glaciecola  99.6  KY194830    42–56  Z  IceBac 391  Marinobacter  99.3  KY194843    42–56  Z  IceBac 392  Paracoccus  99.9  KY194857  ✓  42–56  MOX  IceBac 394  Glaciecola  99.6  KY194832  ✓  56–70  Z  IceBac 396  Glaciecola  99.6  KY194827    56–70  Z  IceBac 397  Glaciecola  99.6  KY194818    56–70  Z  IceBac 398  Glaciecola  99.5  KY194820    56–70  Z  IceBac 399  Glaciecola  99.4  KY194819    56–70  Z  IceBac 400  Polaribacter  99.7  KY194849  ✓  56–70  Z  IceBac 401  Paraglaciecola  100.0  KY194802  ✓  56–70  Z  IceBac 402  Paraglaciecola  99.9  KY194806  ✓  70–84  C  IceBac 403  Colwellia  99.4  KY194851  ✓  70–84  Z  IceBac 404  Polaribacter  99.7  KY194846  ✓  70–84  Z  IceBac 405  Octadecabacter  100.0  KY194834  ✓  98–112  Z  IceBac 408  Polaribacter  99.7  KY194847  ✓  98–112  Z  IceBac 409  Glaciecola  99.6  KY194833  ✓  112–126  C  IceBac 410  Paraglaciecola  100.0  KY194813  ✓  112–126  Z  IceBac 411  Paraglaciecola  100.0  KY194808  ✓  112–126  Z  IceBac 412  Paraglaciecola  100.0  KY194803  ✓  112–126  Z  IceBac 413  Octadecabacter  100.0  KY194835  ✓  112–126  Z  IceBac 414  Paraglaciecola  100.0  KY194804  ✓  126–140  C  IceBac 415  Pseudoalteromonas  100.0  KY194855  ✓  126–140  Z  IceBac 416  Paraglaciecola  99.9  KY194807  ✓  126–140  Z  IceBac 417  Paraglaciecola  100.0  KY194809  ✓  126–140  Z  IceBac 418  Octadecabacter  100.0  KY194836  ✓  126–140  Z  IceBac 419  Octadecabacter (H)  100.0  KY194837  ✓  140–154  Z  IceBac 420  Paraglaciecola  100.0  KY194810  ✓  140–154  Z  IceBac 421  Pseudoalteromonas  100.0  KY194853  ✓  140–154  Z  IceBac 422  Pseudoalteromonas  100.0  KY194854  ✓  140–154  Z  IceBac 423  Colwellia  99.4  KY194852  ✓  140–154  Z  IceBac 424  Octadecabacter  100.0  KY194838  ✓  140–154  MOX  IceBac 426  Paraglaciecola  100.0  KY194814  ✓  154–166  Z  IceBac 428  Paraglaciecola  99.9  KY194811  ✓  154–166  Z  IceBac 430  Octadecabacter (H)  100.0  KY194839  ✓  154–166  Z  IceBac 431  Octadecabacter  100.0  KY194840  ✓  154–166  MOX  IceBac 433  Paraglaciecola  100.0  KY194812  ✓  Isolation depth from air–ice interface (cm)  Isolation mediaa  Bacterial strain  Closest match genusb  Identity % at genus level  Accession number  Used for virus isolation  0–14  Z  IceBac 363  Halomonas  100.0  KY194856  ✓  14–28  Z  IceBac 364  Polaribacter  99.7  KY194850  ✓  14–28  Z  IceBac 365  Glaciecola  99.6  KY194821  ✓  14–28  Z  IceBac 367  Paraglaciecola  99.9  KY194799  ✓  14–28  Z  IceBac 368  Paraglaciecola  99.9  KY194805  ✓  14–28  Z  IceBac 369  Marinobacter  99.3  KY194841    14–28  Z  IceBac 370  Marinobacter  99.3  KY194842  ✓  14–28  Z  IceBac 371  Glaciecola  99.6  KY194822    14–28  Z  IceBac 372  Paraglaciecola (H)  99.9  KY194800  ✓  14–28  MOX  IceBac 373  Polaribacter  100.0  KY194848  ✓  14–28  MOX  IceBac 377  Glaciecola  99.6  KY194823  ✓  28–42  Z  IceBac 378  Glaciecola  99.6  KY194828  ✓  28–42  Z  IceBac 379  Glaciecola  99.6  KY194824  ✓  28–42  Z  IceBac 380  Glaciecola  99.6  KY194825    28–42  Z  IceBac 381  Glaciecola  99.6  KY194829  ✓  28–42  Z  IceBac 382  Glaciecola  99.6  KY194831  ✓  28–42  Z  IceBac 383  Paraglaciecola  99.9  KY194801  ✓  28–42  MOX  IceBac 384  Polaribacter  100.0  KY194845    42–56  Z  IceBac 385  Polaribacter  99.7  KY194844    42–56  Z  IceBac 386  Glaciecola  99.6  KY194815  ✓  42–56  Z  IceBac 387  Glaciecola  99.5  KY194816  ✓  42–56  Z  IceBac 388  Glaciecola  99.6  KY194826  ✓  42–56  Z  IceBac 389  Glaciecola  99.6  KY194817  ✓  42–56  Z  IceBac 390  Glaciecola  99.6  KY194830    42–56  Z  IceBac 391  Marinobacter  99.3  KY194843    42–56  Z  IceBac 392  Paracoccus  99.9  KY194857  ✓  42–56  MOX  IceBac 394  Glaciecola  99.6  KY194832  ✓  56–70  Z  IceBac 396  Glaciecola  99.6  KY194827    56–70  Z  IceBac 397  Glaciecola  99.6  KY194818    56–70  Z  IceBac 398  Glaciecola  99.5  KY194820    56–70  Z  IceBac 399  Glaciecola  99.4  KY194819    56–70  Z  IceBac 400  Polaribacter  99.7  KY194849  ✓  56–70  Z  IceBac 401  Paraglaciecola  100.0  KY194802  ✓  56–70  Z  IceBac 402  Paraglaciecola  99.9  KY194806  ✓  70–84  C  IceBac 403  Colwellia  99.4  KY194851  ✓  70–84  Z  IceBac 404  Polaribacter  99.7  KY194846  ✓  70–84  Z  IceBac 405  Octadecabacter  100.0  KY194834  ✓  98–112  Z  IceBac 408  Polaribacter  99.7  KY194847  ✓  98–112  Z  IceBac 409  Glaciecola  99.6  KY194833  ✓  112–126  C  IceBac 410  Paraglaciecola  100.0  KY194813  ✓  112–126  Z  IceBac 411  Paraglaciecola  100.0  KY194808  ✓  112–126  Z  IceBac 412  Paraglaciecola  100.0  KY194803  ✓  112–126  Z  IceBac 413  Octadecabacter  100.0  KY194835  ✓  112–126  Z  IceBac 414  Paraglaciecola  100.0  KY194804  ✓  126–140  C  IceBac 415  Pseudoalteromonas  100.0  KY194855  ✓  126–140  Z  IceBac 416  Paraglaciecola  99.9  KY194807  ✓  126–140  Z  IceBac 417  Paraglaciecola  100.0  KY194809  ✓  126–140  Z  IceBac 418  Octadecabacter  100.0  KY194836  ✓  126–140  Z  IceBac 419  Octadecabacter (H)  100.0  KY194837  ✓  140–154  Z  IceBac 420  Paraglaciecola  100.0  KY194810  ✓  140–154  Z  IceBac 421  Pseudoalteromonas  100.0  KY194853  ✓  140–154  Z  IceBac 422  Pseudoalteromonas  100.0  KY194854  ✓  140–154  Z  IceBac 423  Colwellia  99.4  KY194852  ✓  140–154  Z  IceBac 424  Octadecabacter  100.0  KY194838  ✓  140–154  MOX  IceBac 426  Paraglaciecola  100.0  KY194814  ✓  154–166  Z  IceBac 428  Paraglaciecola  99.9  KY194811  ✓  154–166  Z  IceBac 430  Octadecabacter (H)  100.0  KY194839  ✓  154–166  Z  IceBac 431  Octadecabacter  100.0  KY194840  ✓  154–166  MOX  IceBac 433  Paraglaciecola  100.0  KY194812  ✓  a Z = Zobell media; MOX = MOX media; C = concentrated Zobell media. b Original isolation hosts of the viruses are marked by (H). View Large The pancake ice at stations 486 and 488 was 6 cm and 35 cm thick, the first-year ice at stations 489–506 and 517 was 37–90 cm, and the early second-year ice at station 515 was 179 cm (Table 1, Tison et al.2017). The ice temperature varied between −1.8°C and −10.9°C, with a median temperature of −3.8°C and ice chl-a median concentration of 2.4 µg l−1(Tison et al.2017). The ice at station 500 was dominated by frazil ice and at station 515 by mixed columnar and frazil ice (Tison et al.2017). The brine salinity varied from 122 to 40 practical salinity units (Tison et al.2017). Isolation of the bacterial strains Isolation of the bacterial strains was started immediately after the 12 layers of the ice core 515a samples were melted. The strains were isolated by plating 100 μl of the melted sample on (I) ZoBell plates (1000 ml Southern Ocean water, 5 g peptone, 1 g yeast extract, 15 g agar; Helmke and Weyland 1995; Middelboe et al.2003), (II) concentrated ZoBell plates (the Southern Ocean water was concentrated by boiling to half of the initial volume, otherwise similar to ZoBell) and (III) Modified Oxford (MOX) agar plates (750 ml seawater, 250 ml ultrapure water, 1 g KNO4, 0.2 g yeast extract, 10 mg FePO4, 2 g HEPES, 12 g agar). The plates were incubated at 4°C and transported from R/V Polarstern to the home laboratory with temperature-controlled courier transportation at −2.6°C to + 6.1°C (World Courier, AmerisourceBergen, Stamford, CT, USA). After 1 month of incubation in the dark at 4°C, the various colony morphotypes were picked and colony-purified at three consecutive times. Various strains were isolated from all layers, except from depths of 84–98 cm. Colony purification and cultivation were done on modified ZoBell medium, Reef Crystal (RC) medium: 33 g RCs, Aquarium Systems Inc. Sarrebourg, France, 1000 ml ultrapure water, 5 g peptone and 1 g yeast extract. The agar concentration for the plates was 1.5% (w/v). The strains were grown aerobically in RC medium at 4°C for 7 days and stored at −80°C, supplemented with 15% (v/v) glycerol. Identification of the bacterial strains The colony-purified strains were identified by 16S rRNA gene sequencing. The genomic DNA was isolated with an UltraClean Microbial DNA Isolation Kit (MO BIO Laboratories Inc., Carlsbad, CA, USA). The partial 16S rRNA genes were amplified with PCR, using primers F27 (Sait et al. 2003) and R1406 (Lane et al. 1985) or pA and pGr (Edwards et al. 1989). The Sanger sequencing was performed at the DNA Sequencing and Genomics Unit, Institute of Biotechnology (University of Helsinki), using primers pDr, pE and pFr (Edwards et al. 1989). The taxonomic identification of the strains was done with SILVA Incremental Aligner (SINA) Alignment Service (version 1.2.11, 10.6.2016, Pruesse, Peplies and Glockner 2012). The partial 16S rRNA gene sequences of the isolated bacterial strains are deposited in the NCBI GenBank database under accession numbers KY194799–KY194857 (Table 2). Phylogenetic analysis of the 16S rRNA sequences For the phylogenetic tree (Fig. 1), alignment was performed with SINA Aliqnment Service (version 1.2.11, minimum identity: 0.8, Pruesse, Peplies and Glockner 2012). Reference sequences were selected, based on SINA sequence matching, and for the nontype-strain sequence matches, their type-strain representatives were also added (EZBioCloud Database (Yoon et al.2017)). After the alignment, all sequences were truncated according to the sequence length, and the bootstrapped (1000) maximum-likelihood tree was constructed, using RAxML (version 8.2.0; Stamatakis 2014), with the GTRGAMMA evolution model. The tree was visualized with the Interactive Tree Of Life (iTOL) online tool (Leturnic and Bork 2007). Figure 1. View largeDownload slide Bootstrapped (1000) phylogenetic maximum-likelihood tree of the 16S rRNA gene sequences of the bacterial strains isolated from Antarctic sea ice. The bootstrap values (>50%) are shown with black circles. The strains are colored at the genus level (see the color key), based on their classification with SILVA Incremental Aligner (SINA) (version 1.2.11, minimum identity: 0.8, Pruesse et al. 2012). The sensitivities of the bacterial strains to isolated phages are shown as efficiency of plating (EOP; on right in gray scale). For the original host (marked by H), the EOP was set to a value of 1. The strains used as references or that do not form a lawn on solid growth media were not tested for EOP (marked with white). The scale bar indicates nucleotide substitutions per position. Figure 1. View largeDownload slide Bootstrapped (1000) phylogenetic maximum-likelihood tree of the 16S rRNA gene sequences of the bacterial strains isolated from Antarctic sea ice. The bootstrap values (>50%) are shown with black circles. The strains are colored at the genus level (see the color key), based on their classification with SILVA Incremental Aligner (SINA) (version 1.2.11, minimum identity: 0.8, Pruesse et al. 2012). The sensitivities of the bacterial strains to isolated phages are shown as efficiency of plating (EOP; on right in gray scale). For the original host (marked by H), the EOP was set to a value of 1. The strains used as references or that do not form a lawn on solid growth media were not tested for EOP (marked with white). The scale bar indicates nucleotide substitutions per position. Abundance of VLPs by flow cytometry The abundances of the VLPs were analyzed with flow cytometry from all the ice layers originating from the 10 stations (Table 1). Sample handling and measurements were done according to (Brussaard et al.2010), except that paraformaldehyde [1% (v/v) in phosphate-buffered saline, Alfa Aesar GmbH & Co KG, Karlsruhe, Germany] was used as a fixative. The samples were stained with SYBR Green I (Sigma-Aldrich Inc., Saint Louis, MO, USA) at room temperature. To measure the background, virus-sized particles were removed from the control samples (pooled from the samples originating from the various layers and stations) by ultrafiltration (Amicon Ultra-15 concentrators, MWCO 100 000 Da; Merck Millipore, Billerica, MA, USA; 4000 g, 4 min, 4°C), and the controls were processed in the same way as the actual samples. Earlier isolated and purified 1/4, 1/32, 1/40, 1/41, 1/44, 3/49 phage particles (Luhtanen et al.2014) were used as controls to define the virus population, and Fluoresbrite 0.5-μm microspheres (Polysciences Inc., Warrington, PA, USA) were used as a size standard. Enumeration of the diluted samples (dilution factor 10; molecular biology grade TE buffer, AppliChem GmbH, Darmstadt, Germany) was carried out, using a CyFlow Cube 8 (Partec GmbH, Münster, Germany) flow cytometer. A sample of 25 μl (defined by the flow cytometry electrodes) was analyzed, using a flow rate of 12 μl min−1. The data were acquired on a dot plot displaying green fluorescence (488 nm) versus side scatter signal, both on a logarithmic scale. The detection trigger was depicted as a green fluorescence. The data were analyzed, using FCS Express 4 software (De Novo Software, Glendale, CA, USA; Fig. S2, Supplementary material), and the numbers were corrected for dilution. The VBR was defined, using published bacterial abundances (Eronen-Rasimus et al.2017). Isolation of viruses For virus isolations, the melted layers of the station 500 core were pooled, as were the layers of core 515a. The melted ice was filtered through a 0.22-μm Durapore Membrane polyvinylidene difluoride filter (EMD Millipore Corporation, Billerica, MA, USA) to remove cellular organisms. The filtrates were concentrated 50 times, using Amicon Ultra-15 concentrators (MWCO 100 000 Da) and centrifugation (2000 g, 5 min, 4°C). The concentrates were stored in 15% (v/v) glycerol at −80°C and transported in liquid nitrogen to the home laboratory. All isolated and colony-purified bacterial strains (Table 2, Fig. 1) that were able to form a bacterial lawn were used to isolate viruses. Virus isolation and further cultivation were done with a plaque assay. For virus isolation, 10 µl and 100 μl of the concentrated sample with 200 μl of dense host bacterial suspension (grown for 7 days in RC medium at 4°C) and 3 ml of melted RC top-layer agar [0.4% (w/v) agar; 43°C] were mixed and poured on RC plates. After 1–3 weeks of incubation at 4°C, the plaques were individually picked and plaque-purified three consecutive times with the plaque assay. For the plaque assay, 100 μl of suitable virus dilution, 200 μl of host liquid culture, and 3 ml of RC top-layer agar were mixed and poured on RC plates, which were incubated for 5–7 days at 4°C. Production and purification of viruses The virus lysates were prepared with the plaque assay as described above, using semiconfluent plates. The top-layer agar was collected and mixed with 2 ml of RC broth per plate. The suspension was incubated for 1 h at 4°C with shaking, after which the cell debris and agar were removed (centrifugation: 10 000 g, 30 min, 4°C). Virus precipitation from the lysates was optimized with various ammonium sulfate concentrations [50%, 60%, 70% and 80% (w/v)], using either saturated solution or ammonium sulfate powder (Boulanger and Puvion 1973; Burgess 2009). Ammonium sulfate was mixed with or dissolved in the lysate for 1 h at 4°C with shaking. The precipitated viruses were collected (centrifugation: 14 000 g, 60 min), washed with SM buffer (50 mM Tris-HCl pH 7.5, 100 mM NaCl, 8 mM MgSO4; Borriss et al.2003) with or without 0.01% (w/v) gelatin and dissolved in SM buffer on ice. The virus aggregates were removed (centrifugation 9300 g, 10 min, 4°C), and the viruses in the supernatant were subjected to rate-zonal centrifugation (153 208 g, 30–70 min, 10°C), using 10–30% (w/v) linear sucrose gradients in SM buffer (Anderson et al.1966; Lawrence and Steward 2010). The gradients were fractionated (12 fractions), and the infectivity (plaque assay), absorbance (260 nm), and the protein, nucleic acid and lipid contents of the fractions were determined (see below). The virus particles in the light-scattering zones were collected with differential centrifugation (104 087 g, 3 h, 10°C), and the particles were dissolved in SM buffer overnight on ice. Virus particle analyses The absorbance values (260 nm) of the 12 sucrose gradient fractions were measured with an Eppendorf BioPhotometer D30 (Eppendorf AG, Hamburg, Germany), using 30% (w/v) sucrose in SM buffer as a blank sample. The viral structural proteins were separated with SDS-PAGE (16% (w/v) acrylamide; Olkkonen and Bamford 1989). The SM buffer used in the protein analysis samples did not contain gelatin. The protein concentrations were determined with a Coomassie Blue-based method (Bradford 1976), using bovine serum albumin as a standard. For the SDS-PAGE, when appropriate, the samples were concentrated with 10% (v/v) trichloroacetic acid precipitation (30 min, on ice). The precipitate was collected (centrifugation: 16 200 g, 30 min, 4°C). The resolving gels were stained with Brilliant Blue R (Sigma-Aldrich) for proteins and, when appropriate, stained with Sudan Black B (Sigma-Aldrich) for lipids and the stacking gels with ethidium bromide for nucleic acids. For the lipid-staining control, purified PRD1 particles (Bamford and Bamford 1991) were used as a control. To test the sensitivity of the viruses to Triton X-100, viruses were incubated in 0.1% (v/v) and 0.01% Triton X-100 (in SM buffer) for 3 h and 24 h at 4°C. SM buffer was used as a control. The infectivity of the viruses tested was determined with a plaque assay. The sensitivity of the host organisms to Triton X-100 was analyzed similarly, except that the number of colony-forming units was determined by plating. For TEM, the purified virus particles were negatively stained for 20 s, using 2% (w/v) uranyl acetate (pH 7), 3% (w/v) uranyl acetate (pH 4.5), or 1% potassium phosphotungstate (pH 7). A JEOL JEM-1400 TEM (Electron Microscopy Unit, Institute of Biotechnology, University of Helsinki) was used with 80-kV tension for detailed investigation of the viruses. Temperature ranges of host growth and virus infection The growth of the original three host strains at different temperatures was determined by plating 100 μl of diluted bacterial suspension on RC plates, which were incubated at 0, 4, 10, 15, or 20°C for 60 days. The infection ability of the bacteriophages at different temperatures was tested with a plaque assay. The host suspensions used for the plaque assay were incubated at 4°C. The plates were incubated at 0, 4, 10, or 15°C for 20 days. Virus-bacteria interactions All the bacterial isolates that were able to grow as a lawn on a plate were tested for their sensitivity to the isolated viruses (Table 2, Fig. 1). Ten microliters of undiluted and 100-times diluted virus lysates were spotted on the host strain lawns. RC medium was used as a negative control and the original virus-host pair as a positive control. The plates were incubated at 4°C for 7–14 days. The growth temperature was >0°C to prevent freezing, but still at the range of cold-active viruses. All positive results were verified with a plaque assay, using suitable dilutions of the virus. The efficiency of plating (EOP) was calculated according to the plaque count obtained with the target strain, compared with that obtained with the isolation host strain. RESULTS Sea-ice bacterial isolates We isolated 59 bacterial strains from core 515a (Table 2). The majority of the isolates (∼59%) were classified as members of the genera Glaciecola or Paraglaciecola (Fig. 1). Only one strain, IceBac 363, was isolated from the coldest top layer and belonged to the genus Halomonas. The remaining strains were identified as members of the genera Octadecabacter (seven strains), Polaribacter (seven strains), Marinobacter (three strains), Pseudoalteromonas (three strains), Colwellia (two strains), or Paracoccus (one strain). All isolated bacterial strains belonging to the same genus had identical or nearly identical (identity 99.3–100%) partial 16S rRNA gene sequences (at minimum 1290 base pairs, bp). VLP abundance and VBRs in sea ice Using flow cytometry, we determined that VLPs occurred throughout the ice cores in varying numbers. The mean VLP abundance of all the ice cores sampled was 10.9 × 105 ml−1 (range 1.9 × 105–49.0 × 105 ml−1) in the bulk ice, with the highest numbers at stations 500 and 515 (6.9 × 105–49.0 × 105 ml−1; Table 1). The mean VBR of all the ice cores sampled was 5.3 (0.7–13.4; Table 1). The highest VBR values were at station 503 (9.5–13.4; Table 1) and the lowest at station 515 (0.7–2.9; Table 1). Sea-ice bacteriophage isolates Forty-eight out of the 59 bacterial strains were able to form a bacterial lawn on a plate and were consequently used to screen phages (Table 2). Four phages were obtained, cultivated and purified in the laboratory (Table 3). The phages were named after the isolation host genus, area of isolation, and the initials of notable persons in this study (Krupovic et al. 2016). The names and their abbreviations are Paraglaciecola Antarctic GD virus 1 (PANV1), Paraglaciecola Antarctic JLT virus 2 (PANV2), Octadecabacter Antarctic BD virus 1 (OANV1) and Octadecabacter Antarctic DB virus 2 (OANV2). Phages PANV1 and PANV2 originated from station 500 and were isolated for the same host (IceBac 372), which was classified as Paraglaciecola psychrophila (similarity 99.9%). PANV1 produced clear plaques 3–4 mm in diameter, whereas the PANV2 plaques of 3–5 mm in diameter had a clear center surrounded by a turbid halo. Phages OANV1 and OANV2 were isolated from core 515a for two different bacterial hosts, IceBac 419 and IceBac 430, respectively, both identified as Octadecabacter antarcticus. Both phages produced clear plaques, but the diameters were different (3–6 mm for OANV1, 6–8 mm for OANV2). The optimized phage lysate titers varied from ∼6 × 109 to ∼5 × 1011 plaque-forming units (pfu) ml−1, depending on the phage (Table 3). The infectivity of the lysates was retained for several months when stored at 4°C. All three host strains originated from different layers in the ice core (Table 2). Table 3. Phages isolated in this study. Phage  Sampling station  Isolation host  Genus of the host (closest match)  Lysate titer (pfu/ml)  Capsid head diameter (nm)a  Tail length (nm)b  Morphotype  Paraglaciecola Antarctic GD virus 1 (PANV1)  500  IceBac 372  Paraglaciecola  1.5 × 1010  71 ± 7 (n = 20)  58 ± 22 (n = 10)  myovirus  Paraglaciecola Antarctic JLT virus 2 (PANV2)  500  IceBac 372  Paraglaciecola  5.2 × 1011  52 ± 8 (n = 29)  89 ± 30 (n = 10)  siphovirus  Octadecabacter Antarctic BD virus 1 (OANV1)  515  IceBac 419  Octadecabacter  1.2 × 1010  50 ± 8 (n = 20)  83 ± 10 (n = 10)  siphovirus  Octadecabacter Antarctic DB virus 2 (OANV2)  515  IceBac 430  Octadecabacter  5.8 × 109  53 ± 7 (n = 20)  –  podovirus  Phage  Sampling station  Isolation host  Genus of the host (closest match)  Lysate titer (pfu/ml)  Capsid head diameter (nm)a  Tail length (nm)b  Morphotype  Paraglaciecola Antarctic GD virus 1 (PANV1)  500  IceBac 372  Paraglaciecola  1.5 × 1010  71 ± 7 (n = 20)  58 ± 22 (n = 10)  myovirus  Paraglaciecola Antarctic JLT virus 2 (PANV2)  500  IceBac 372  Paraglaciecola  5.2 × 1011  52 ± 8 (n = 29)  89 ± 30 (n = 10)  siphovirus  Octadecabacter Antarctic BD virus 1 (OANV1)  515  IceBac 419  Octadecabacter  1.2 × 1010  50 ± 8 (n = 20)  83 ± 10 (n = 10)  siphovirus  Octadecabacter Antarctic DB virus 2 (OANV2)  515  IceBac 430  Octadecabacter  5.8 × 109  53 ± 7 (n = 20)  –  podovirus  a average diameter. b average length. pfu = plaque-forming unit. View Large Purification and characterization of the phages To characterize the phages, virus purification methods were optimized, based on ammonium sulfate precipitation and rate-zonal ultracentrifugation, following the recovery and purity of the infectious viruses at each step. Using ammonium sulfate precipitation, 25–54% of the infectious viruses were recovered, depending on the virus (Table 4). Both the ammonium sulfate powder and the saturated solution resulted in similar yields. PANV1 and PANV2 were precipitated with 50% ammonium sulfate, whereas OANV1 and OANV2 needed 80%. The precipitated particles were further purified with rate-zonal ultracentrifugation, and significant amounts of various noninfectious protein impurity species were detected at the top of the sucrose gradient. For all viruses, a single visible infectious light-scattering zone was detected in the middle of the sucrose gradient (Fig. 2). This zone contained several proteins of different sizes that were unique for each virus. PANV1 and OANV2 had one major protein type in sizes of ∼55 and ∼35 kDa, respectively, while PANV2 and OANV1 had two major protein types (∼40 and ∼12 kDa in PANV2; ∼35 and ∼15 kDa in OANV1). A peak in the absorbance was detected in the same light-scattering zone, as were the nucleic acids when visible. Lipids could not be detected from the gels after Sudan Black staining (not shown). In addition, treatment of phages with the nonionic detergent Triton X-100 did not affect their infectivity, suggesting that the virus particles did not contain a lipid component. Specific infectivities (∼2–9 × 1012 pfu mg−1 protein) calculated for the purified phages showed that all virus samples were highly infectious after biochemical purification (Table 4). After the final concentration step with differential centrifugation, the recoveries of infectious viruses varied from ∼10% to 20% (Table 4). Figure 2. View largeDownload slide Purification of the phages by rate-zonal centrifugation in sucrose. (A) bacteriophage PANV1, (B) bacteriophage PANV2, (C) bacteriophage OANV1, (D) bacteriophage OANV2. (A–D) Top: position of the light-scattering zone (gray) in the sucrose gradient tubes. Middle: Absorbance (closed squares) and infectivity (open circles) of the 12 sucrose gradient fractions in which the top fraction is marked as 1. Bottom: Protein content of the 12 gradient fractions analyzed with SDS-PAGE and Coomassie Blue staining. The protein patterns of the final biochemically purified and concentrated phages are shown on the right. St = molecular mass marker; PRD1 = purified phage PRD1 used as a control. The dashed line marks the position of the upper edge of the light-scattering virus zone. pfu = plaque-forming unit. Figure 2. View largeDownload slide Purification of the phages by rate-zonal centrifugation in sucrose. (A) bacteriophage PANV1, (B) bacteriophage PANV2, (C) bacteriophage OANV1, (D) bacteriophage OANV2. (A–D) Top: position of the light-scattering zone (gray) in the sucrose gradient tubes. Middle: Absorbance (closed squares) and infectivity (open circles) of the 12 sucrose gradient fractions in which the top fraction is marked as 1. Bottom: Protein content of the 12 gradient fractions analyzed with SDS-PAGE and Coomassie Blue staining. The protein patterns of the final biochemically purified and concentrated phages are shown on the right. St = molecular mass marker; PRD1 = purified phage PRD1 used as a control. The dashed line marks the position of the upper edge of the light-scattering virus zone. pfu = plaque-forming unit. Table 4. Recovery of infectious phages during biochemical purification after ammonium sulfate precipitation and rate zonal centrifugation in sucrose combined with concentration step by differential centrifugation.   Total pfusa  Recovery of infectivity %  Specific infectivity pfu/mg protein  PANV1    Virus lysate  7.5 × 10 12  100.0      50% ammonium sulfate precipitate  4.1 × 10 12  54.7      Concentrated virusb  1.5 × 10 12  20.0  1.8 × 10 12  PANV2    Virus lysate  3.2 × 10 14  100.0      50% ammonium sulfate precipitate  9.6 × 10 13  30.0      Concentrated virusb  6.2 × 10 13  19.4  8.9 × 10 12  OANV1    Virus lysate  6.0 × 10 12  100.0      80% ammonium sulfate precipitate  1.5 × 10 12  25.0      Concentrated virusb  5.7 × 10 11  9.5  3.8 × 10 12  OANV2    Virus lysate  3.3 × 10 12  100.0      80% ammonium sulfate precipitate  1.2 × 10 12  35.7      Concentrated virusb  4.4 × 10 11  13.6  6.9 × 10 12    Total pfusa  Recovery of infectivity %  Specific infectivity pfu/mg protein  PANV1    Virus lysate  7.5 × 10 12  100.0      50% ammonium sulfate precipitate  4.1 × 10 12  54.7      Concentrated virusb  1.5 × 10 12  20.0  1.8 × 10 12  PANV2    Virus lysate  3.2 × 10 14  100.0      50% ammonium sulfate precipitate  9.6 × 10 13  30.0      Concentrated virusb  6.2 × 10 13  19.4  8.9 × 10 12  OANV1    Virus lysate  6.0 × 10 12  100.0      80% ammonium sulfate precipitate  1.5 × 10 12  25.0      Concentrated virusb  5.7 × 10 11  9.5  3.8 × 10 12  OANV2    Virus lysate  3.3 × 10 12  100.0      80% ammonium sulfate precipitate  1.2 × 10 12  35.7      Concentrated virusb  4.4 × 10 11  13.6  6.9 × 10 12  a calculated per a liter of original lysate. b after rate zonal centrifugation in sucrose and concentration by differential centrifugation. pfu = plaque-forming unit. View Large Transmission electron microscopy (TEM) of the purified particles showed that phage PANV1 (Fig. 3a) had a rigid, contractile tail typical of myoviruses. Its average tail length was ∼58 nm and head diameter ∼71 nm. PANV2 (Fig. 3b) infecting the same host had an ∼89-nm noncontractile tail characteristic of the siphoviruses and an ∼52-nm head diameter. OANV1 (Fig. 3c) also had a typical siphovirus tail, with an average length of ∼83 nm and a head ∼50 nm in diameter, whereas OANV2 (Fig. 3d) seemed to have a very short tail typical of podoviruses and a head ∼53 nm in diameter. Figure 3. View largeDownload slide Transmission electron micrographs of the purified and negatively stained phages. (A) PANV1, (B) PANV2, (C) OANV1 and (D) OANV2. Figure 3. View largeDownload slide Transmission electron micrographs of the purified and negatively stained phages. (A) PANV1, (B) PANV2, (C) OANV1 and (D) OANV2. Temperature range tests for host growth and phage infection All the bacterial host strains (IceBac 372, IceBac 419 and IceBac 430) were able to form colonies at the temperatures from 0°C to 15°C, but not at 20°C (Table 5), and were therefore classified as psychrophiles (Morita 1975). The effect of temperature on phage infection (plaque formation) was tested at temperatures supporting the growth of the hosts. All the phages were able to infect their original host only at 0°C and 4°C, but not at higher temperatures (Table 5). PANV1 and PANV2 produced plaques at 0°C and 4°C in 6 days, but OANV1 and OANV2 produced plaques at 0°C in 14 days and at 4°C in 6 days. Table 5. Temperature-dependent growth of the bacterial host strains and the phages.   0°  4°  10°  15°  20°  Host bacterial growtha    IceBac 372  +  +  +  (+)  −  IceBac 419  +  +  +  +  −  IceBac 430  +  +  +  +  −  Infectivity of the phagesa  PANV1  +  +  −  −  ND  PANV2  +  +  −  −  ND  OANV1  +  +  −  −  ND  OANV2  +  +  −  −  ND    0°  4°  10°  15°  20°  Host bacterial growtha    IceBac 372  +  +  +  (+)  −  IceBac 419  +  +  +  +  −  IceBac 430  +  +  +  +  −  Infectivity of the phagesa  PANV1  +  +  −  −  ND  PANV2  +  +  −  −  ND  OANV1  +  +  −  −  ND  OANV2  +  +  −  −  ND  a + = producing colonies/plaques; ( + ) = retarded growth; − = no colonies or plaques produced; ND = not determined. View Large Phage–bacteria interactions The sensitivity of all the 48 isolated bacterial strains (that were able to grow as a bacterial lawn) to the isolated phages was tested at 4°C (Table 2; Fig. 1). In all, 17 strains (13 Paraglaciecola strains and 4 Octadecabacter strains) were sensitive to at least one of the phages (Fig. 1). Of 16 Paraglaciecola isolates, IceBac 372 was sensitive to three phages: PANV1, PANV2 and OANV1. Ten other Paraglaciecola strains were sensitive to both PANV1 and PANV2, but with different plating efficiencies (EOPs), two strains were sensitive to either PANV1 or PANV2, but with low EOP, and three strains could not be infected. All seven Octadecabacter strains had 100% identical 16S rRNA gene sequences (within 1289 bp, Fig. 1). However, only four out of seven Octadecabacter strains were sensitive to either OANV1 or OANV2 and showed different EOPs. Both the PANV1 and PANV2 phages were able to infect 12 different Paraglaciecola strains with different EOPs. However, each was able to infect only a single strain (IceBac 417 or IceBac 420, respectively) that the other could not (Fig. 1). In addition to its original host strain (IceBac 419, Octadecabacter), OANV1 was able to infect two other Octadecabacter strains, but with lower EOP. It also produced plaques with high EOP in the strain IceBac 372 ( Paraglaciecola), which was the isolation host for PANV1 and PANV2 (Fig. 1). Consequently, OANV1 was able to infect strains representing two classes: Gammaproteobacteria (Paraglaciecola) and Alphaproteobacteria (Octadecabacter). In contrast, phage OANV2 was able to infect only IceBac 430 (Octadecabacter; Fig. 1). DISCUSSION We isolated and purified four Antarctic sea-ice phages (PANV1, PANV2, OANV1 and OANV2) that could be maintained and cultivated under laboratory conditions. They were cold-active (capable of infection and production at ≤4°C; Wells and Deming 2006b), infecting bacterial strains belonging to the typical sea-ice bacterial genera Paraglaciecola or Octadecabacter. The viruses were specific for host recognition at the strain level, even though OANV1 was able to infect bacterial strains from two different classes. The highest VLP abundances were in the samples where bacteria were most abundant and active (Eronen-Rasimus et al. 2017). Isolation of sea-ice bacteria and phages We isolated 59 bacterial strains belonging to nine different genera from Antarctic winter-sea ice. The ice was melted with the direct-melting method, which has been shown to result in viable bacteria counts similar to those obtained by melting with seawater addition (Helmke and Weyland 1995), even if it may cause osmotic stress due to the rapid salinity changes. The isolated strains belonged to the following genera: Colwellia, Glaciecola, Halomonas, Marinobacter, Octadecabacter, Paracoccus, Paraglaciecola (formerly Glaciecola; Shivaji and Reddy 2014) , Polaribacter, and Pseudoalteromonas, all of which are common members in the sea-ice bacterial community (Bowman et al. 1997; Brinkmeyer et al. 2003; Deming and Collins 2017). The majority of these genera were also abundant in isolation ice core 515a, based on bacterial community composition analysis (see results in Eronen-Rasimus et al. 2017). In addition, four unique phages were isolated from the sea ice with the direct-melting and plaque assay methods, even though the viruses were exposed to a 43°C temperature for a short time during the plaque assay. This has also been successfully used previously (Borriss et al. 2003), and at least some of the cold-adapted phages can apparently tolerate high temperatures for a short time. Wide temperature tolerance can be beneficial to phage survival in natural environments throughout the various seasons. Presumably, virus reproduction is most effective when the number of susceptible hosts is high and active (Thingstad and Lignell 1997). The hosts of the bacteriophages OANV1 and OANV2 belonged to the genus Octadecabacter, which was abundant in phage isolation core 515a (Eronen-Rasimus et al. 2017). The genus Paraglaciecola (host of phages PANV1 and PANV2) could not be detected separately from the genus Glaciecola in the community analysis, likely due to the short sequence length used, but Glaciecola was present in both the 500 and 515a cores (Eronen-Rasimus et al. 2017). Our results together with those of Eronen-Rasimus et al. ( 2017) support the notion that bacteriophages of these predominant bacteria may be abundant in the viral community, resulting in increased opportunities of isolating them. Phage-host interactions We tested the sensitivity of 48 isolated bacterial strains, representing nine different genera, to our phages. PANV1 and PANV2 were able to infect several closely related Paraglaciecola strains with different EOPs, but not all the strains (Fig. 1). This may have resulted from of an arms race in which the bacterial strains evolved to inhibit phage infection, leading to diversification of bacterial strains (Thingstad et al.2014). Consequently, the phages needed to evolve to be able to survive. Since PANV1 and PANV2 have different infection patterns, this may have resulted in two different phage-host coevolution lineages. Phage OANV1 was able to infect bacterial strains from two different classes: Alphaproteobacteria (Octadecabacter) and Gammaproteobacteria (Paraglaciecola, Fig. 1). Still, it was able to infect only three of the Octadecabacter strains with identical 16S rRNA gene sequences. These phages appeared to be strain-specific in their host recognition, even though they were able to infect bacteria across classes. Virus-host interactions are commonly simplified in ecological modeling even though they can be very complex (Middelboe 2000; Holmfeldt et al.2007; Louhi et al.2016). Our results also show that the interactions can be highly complicated, while the roles of generalist and specialist viruses in the microbial communities are not clear. Even though a virus may be capable of infecting several strains of the same genus or even strains belonging to different classes, it does not necessarily mean that the virus is a generalist. The development of generalist and specialist viruses is still not totally understood, even though it has been investigated and various theories presented, most of which are based on phage-host coevolution (Flores et al.2011; Beckett and Williams 2013; Weitz et al.2013; Koskella and Brockhurst 2014). The coevolution of a phage and its host can be explained by the Killing-the-Winner model (Thingstad and Lignell 1997) with the addition of cost of resistance (Våge, Storesund and Thingstad 2013). The defense strategy of the bacteria can display a trade-off that lowers their growth rates. Competition strategists choose not to defend, which makes them easier targets for viruses. In this study all the isolated phages were strain specific, even though PANV1, PANV2 and OANV2 were able to infect several strains. The specificity of viruses primarily arises from the interaction between the receptor molecule on the cell surface and the receptor binding protein of the virus, but further molecular research is needed to understand the concept of virus specificity. Sea-ice phage isolates have very narrow host ranges (Borriss et al.2003; Luhtanen et al.2014), but based on the genomic data, cold-active phages may have broader host ranges than mesophiles (Colangelo-Lillis and Deming 2013). Myoviruses are often considered to have broader host ranges than siphoviruses and podoviruses (Suttle 2005). However, in this study, myovirus PANV1 and siphovirus PANV2 showed similar host ranges and were able to infect only closely related hosts, whereas siphovirus OANV1 was able to infect strains from two different classes. When phage host ranges are experimentally studied, the number of cultivable bacterial isolates limits the tests, and consequently the results cannot reveal the complete host range spectrum in the environment. In addition, the tests were performed only at 4°C, and consequently virus-host interactions adapted to lower temperatures were not detected. However, our results indicate that with the strain specificity observed, the phages may be able to control the bacterial community composition, as proposed earlier based on observation in the environment (Maranger, Bird and Juniper 1994), theory (Thingstad et al.2014), and experimentation (Middelboe et al.2001). Since viruses need their hosts to replicate and produce progeny, their activity is directed to the active part of the bacterial community. Purification and characterization of phages The purification process was optimized for all phages separately. Purification analysis revealed that a significant amount of impurities and host-derived complexes were separated, allowing us to obtain a light-scattering zone comprising infectious, highly purified viruses (Fig. 2). The individual protein patterns of the isolated phages (Fig. 2) and the specific infectivities calculated (Table 4) showed that each isolated phage was different and that the purification of the virus particles was successful. Efficient purification made it possible to study individual phages in more detail. Detailed TEM observations verified that all the phages isolated were icosahedral tailed phages (Fig. 3). Sudan Black staining of the sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) gel and Triton-X treatment of the virus particles indicated that the phages do not have a structural lipid component, which is in accordance with the virus morphologies observed. The virus capsid diameters (∼50–71 nm; Table 3) are similar to those in the most abundant VLP size groups (50–70 nm or <110 nm) reported in Arctic and Antarctic sea ice (Maranger, Bird and Juniper 1994; Gowing et al.2004). In three cases, the morphology of the virus tails was reliably identified, suggesting that PANV1 is a myovirus, whereas PANV2 and OANV1 are siphoviruses (Fig. 3) resembling the dsDNA bacteriophages belonging to the order Caudovirales. However, the tail of bacteriophage OANV2 was considerably difficult to detect. We propose that OANV2 is a podophage with a short noncontractile tail. Icosahedral tailed viruses from the order Caudovirales were previously the most often isolated virus types from sea ice (Borriss et al.2003; Luhtanen et al.2014), although a filamentous virus from the order Inoviridae has also been isolated (Yu et al.2015). Temperature The temperatures used here for isolation, cultivation, and tests (Table 5) for both bacteria and phages were warmer than the temperatures in the sea-ice brine (−1.8°C down to −10.9°C; Table 1; Tison et al.2017), due to methodological limitations. It is still evident that the bacteria and phages were cold-adapted, since the bacterial strains were able to grow at 0°C, but not at 20°C (Morita 1975), and the phages were capable of infecting and producing progeny at ≤4°C (Wells and Deming 2006b; Table 5). In addition, psychrophilic bacteria from sea ice can be active even at −20°C (Junge, Eicken and Deming 2004), and cold-active phages can be productive at temperatures from 8°C to −6°C (Wells and Deming 2006b) or even at −12°C (Wells and Deming 2006c). The isolated phages retained their infectivities at cold temperatures (several months at 4°C and −80°C when supplemented with 15% glycerol), consistent with the previous cold-active virus study (Wells and Deming 2006c). Viruses have adapted to cope with a wide range of temperatures from the extreme heat in hot springs (Zablocki et al.2017) to the coldness of sea ice or permafrost (Borriss et al.2003; Wells and Deming 2006b; Luhtanen et al.2014). The major responsibility for this adaptation comes from virion stability, since virions have defined the temperature ranges at which they function (Jaenicke 1991; Bischof and He 2005). Proteins adapted to low temperatures are able to function at sea-ice temperatures, but at temperatures above their optimal range the cold-active proteins are unstable (Reed et al.2013). Still, these virus isolates tolerated the temperature of the warm top-layer agar (∼43°C) used for the plaque assay and remained infectious. Based on our results, the sea-ice virus particles can probably remain infectious in the sea water during the ice-free periods and resume activity when the sea-ice microbial community is reformed. Virus plaques were formed only at temperatures about 10 degrees lower than their host could tolerate, indicating that temperature controlled the infections, as shown in previous studies on sea-ice phage-host isolates (Borriss et al.2003; Luhtanen et al.2014) and other cold-adapted phage-host systems (Wells and Deming 2006b). Temperature may affect the host, the virus and/or their interaction. The virus-receptor molecules in the host cell may only have been induced at cold temperatures, as reported previously in Yersinia enterocolitica infections (Leon-Velarde et al.2016), indicating that the receptors may be associated with the host's cryoprotection mechanisms. Temperature also affects the adsorption of Listeria phages by adsorption inhibition and unidentified post-adsorption mechanisms (Tokman et al.2016). The structure of the phage-receptor molecules could also have changed with rising temperatures, which can inhibit the infection, or the bacterial resistance mechanisms (Labrie, Samson and Moineau 2010) could have been activated at higher temperatures. The phages isolated in this study are most probably specialized for sea-ice conditions, because they were cold-active and infected bacterial strains belonging to common sea-ice genera. The temperature inside the brine channels can vary widely, depending on the air temperature and depth of the insulating snow cover on top of the ice. Therefore, it is easy to understand why adaptations to different temperatures are needed in sea-ice communities. The isolated phages can reproduce even at seawater temperatures (4°C), implying that they would be able to survive in the water column. However, the sea-ice community is typically different from the seawater community (Bowman et al.1997; Boetius et al.2015; Eronen-Rasimus et al.2015), and the contact rates of viruses and their hosts are presumably lower in the water than in the semi-enclosed brine channels. Warming global temperatures may temporarily increase the activity of the microbial community, but also broaden the brine channels flushing the community from the ice. The ice period may also become shortened, leaving less time for the community to develop. If climate change succeeds in destroying the sea-ice habitat, we may lose a significant amount of microbial diversity on our planet. Abundance of VLPs and VBRs in Antarctic winter-sea ice The VLP abundance in Antarctic winter-sea ice ranged from ∼105 to 106 ml−1 in bulk ice (Table 1). The highest abundances were measured at stations 500 and 515 (6.9 × 105–49.0 × 105 ml−1 in bulk ice). The lower range of our dataset is comparable to the values measured earlier from Antarctic winter-sea ice (∼105 ml−1 of bulk ice; Paterson and Laybourn-Parry 2012), whereas the highest abundances were similar to those in Arctic spring blooms and Antarctic late autumn and summer sea ice (∼106–108 ml−1 of bulk ice; Maranger, Bird and Juniper 1994; Gowing et al.2002, 2004, respectively). The high VLP concentrations at stations 500 and 515 may be explained by the high bacterial abundance (Eronen-Rasimus et al.2017; Table 1) and bacterial production (measured as thymidine incorporation; Eronen-Rasimus et al.2017) observed, which were positively correlated with the high chl-a concentrations (up to 113.2 mg l−1 in bulk ice; Eronen-Rasimus et al.2017; Tison et al.2017). Positive correlation of chl-a with bacterial and VLP abundances was reported in Antarctic sea ice during spring and summer ice-algal blooms (Maranger, Bird and Juniper 1994; Gowing et al.2004). Typically, the platelet layer that forms at the bottom of the ice is the most productive layer in the sea-ice environment (Arrigo, Dieckmann and Gosselin 1995). During the winter, autotrophic production is reduced, due to the low light levels. In this study, the high chl-a concentrations observed in the middle part of the ice core at station 500 (Table 1; Tison et al.2017) were likely caused by ice rafting and flooding, trapping the autumnal bottom ice biomass between two ice floes and supplying nutrients to the uppermost ice layers (Tison et al.2017). At station 500, the algal biomass (high chl-a concentration) may have been preserved from autumn algal growth, whereas at second-year ice station 515, biomass was likely preserved from the previous spring (Tison et al.2017). Our results suggest that if the chl-a concentrations and consequent bacterial abundance and activity are high, viruses may be abundant and likely active in winter-sea ice. The high VLPs also indicate that the viral winter-sea-ice community was surprisingly dynamic, considering the season. The VBR range of 0.7–13.4 (mean 5.3) corresponds to those measured previously in Antarctic winter-sea ice (1–20.8; Paterson and Laybourn-Parry 2012). The highest VBRs (Table 1) were found at first-year ice-station 503 with low bacterial abundance and activity (Table 1; see bacterial production in Eronen-Rasimus et al.2017), while the lowest VBRs were detected at young second-year ice-station 515 (Table 1) with the highest bacterial production and abundance (Eronen-Rasimus et al.2017; Table 1). The high VBR in the low-activity community may have resulted from induction of lysogenic viruses during freezing and preservation of the virus particles in sea-ice brine, similarly to young ice in the Arctic (Collins and Deming 2011). The decreasing VBR together with increasing VLP and bacterial abundances, and bacterial activity were detected during the algal spring bloom (Maranger, Bird and Juniper 1994). Similar nonlinear variation was observed during examination of microbial cell and virus abundance estimates in 25 distinct marine surveys (Wigington et al.2016). The low VBR in the active community may have resulted from change in the bacterial community composition, so that the number of bacteria resistant to the phages may have increased. Alternatively, the host bacterial activity may have been decreased, which could have lowered the viral production, possibly because the phages may have lysogenized, i.e. become prophages (Maranger, Bird and Juniper 1994). The low VBR in the high-microbial density community was also explained by the Piggyback-the-Winner model, in which viruses favor temperateness at high host densities (Knowles et al.2016). In conclusion, four phage-host systems were isolated from both first- and second-year winter-sea ice from the Weddell Sea, Antarctica. The phages seemed to be bacterial strain-specific, but some were able to infect several related bacterial strains and one from second-year ice even across classes. The phages were able to retain their infectivity for lengthy periods under cold conditions and infected their host bacteria only in the hosts’ lower growth temperature ranges, suggesting that they are cold-active. The VLP counts suggest that the viral community may also be dynamic in winter-sea ice if their hosts are active. Overall, the virus-host interactions can be very complex. SUPPLEMENTARY DATA Supplementary data are available at FEMSEC online. ACKNOWLEDGEMENTS The authors acknowledge the Electron Microscopy Unit in the Institute of Biotechnology, HiLIFE, University of Helsinki and the Finnish Environment Institute, Marine Research Centre and Finnish Marine Research Infrastructure (FINMARI) for providing the laboratory infrastructure. We also thank the Finnish Antarctic Research Program FINNARP (especially Mika Kalakoski and Eivor Lahtinen) for logistic and financial support with cargo and travel expenses, and the Alfred Wegener Institute for Polar and Marine Research, leading researcher Peter Lemke, the captain, crew, and the other participants in the AWECS expedition. We thank Sari Korhonen for skilled technical assistance in virus production and purification, Christiane Uhlig for providing the MOX medium plates, and Harri Kuosa and Daniel Delille for useful comments on the manuscript. FUNDING This work was supported by the Walter and Andrée de Nottbeck Foundation (AML, EER, JMR), Onni Talas Foundation (AML), the Academy Professor (Academy of Finland) funding grants 283072 and 255342 (DHB) and the Belgian Science Policy (Bigsouth project, SD/CA/05). BD is a research associate at the F.R.S-FNRS. 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The first known virus isolates from Antarctic sea ice have complex infection patterns

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Abstract

Abstract Viruses are recognized as important actors in ocean ecology and biogeochemical cycles, but many details are not yet understood. We participated in a winter expedition to the Weddell Sea, Antarctica, to isolate viruses and to measure virus-like particle abundance (flow cytometry) in sea ice. We isolated 59 bacterial strains and the first four Antarctic sea-ice viruses known (PANV1, PANV2, OANV1 and OANV2), which grow in bacterial hosts belonging to the typical sea-ice genera Paraglaciecola and Octadecabacter. The viruses were specific for bacteria at the strain level, although OANV1 was able to infect strains from two different classes. Both PANV1 and PANV2 infected 11/15 isolated Paraglaciecola strains that had almost identical 16S rRNA gene sequences, but the plating efficiencies differed among the strains, whereas OANV1 infected 3/7 Octadecabacter and 1/15 Paraglaciecola strains and OANV2 1/7 Octadecabacter strains. All the phages were cold-active and able to infect their original host at 0°C and 4°C, but not at higher temperatures. The results showed that virus–host interactions can be very complex and that the viral community can also be dynamic in the winter-sea ice. virus-host interactions, virus isolation, sea ice microbiology, VLP in sea ice INTRODUCTION Almost 10% of the world's ocean is covered by sea ice at least once per year, which makes it one of the largest biomes on Earth (Dieckmann and Hellmer 2010). Although being a cold and harsh environment, sea ice is full of life. Specialized organisms live inside brine channels and pockets that are formed during freezing conditions, when salts and nutrients from the seawater become concentrated between the ice crystals (Thomas and Dieckmann 2002). Brine remains a liquid inside the ice, due to its high salinity, and this makes it a suitable habitat for the sea-ice microbial community, which comprises protists, bacteria, archaea and their viruses (Maranger, Bird and Juniper 1994; Mock and Thomas 2005; Arrigo, Mock and Lizotte 2010; Deming and Collins 2017). Here, we use the term bacteria instead of prokaryotes, even if in some cases archaea may also be involved. Microbes affect the biogeochemical properties of the sea ice, gas exchange between the ocean and atmosphere, and provide food for ice-associated animals, e.g. krill (Arrigo and Thomas 2004). Viruses are the most abundant biological entities and are presumed to play important roles in the biogeochemical cycles of the oceans (Fuhrman 1999; Suttle 2007). The most commonly found viruses, bacteriophages (phages) i.e. viruses infecting bacteria, are possibly the main cause of bacterial mortality (Weinbauer 2004; Suttle 2007). Since viruses can multiply only within their host cells, their activity is dependent on the abundance and activity of their hosts (Maranger, Bird and Juniper 1994; Marchant et al. 2000). Due to their host specificity, they are crucial to the control of bacterial community composition and activity (Proctor and Fuhrman 1990; Wommack and Colwell 2000; Suttle 2005; 2007). However, most studies of marine environments are conducted in the water column, whereas knowledge of viruses and their functions in sea ice are still very limited. The numbers of virus-like particles (VLPs) measured, range between 105 and 108 ml−1 in bulk Arctic and Antarctic sea ice from spring to autumn. The lowest values have been observed during the winter in Antarctic bulk ice (Paterson and Laybourn-Parry 2012), whereas the highest numbers of VLPs occur during freezing or spring algal mass growth (Maranger, Bird and Juniper 1994; Collins and Deming 2011). Moreover, VLPs may also contain particles other than viruses, e.g. gene transfer agents or membrane vesicles (Forterre et al. 2013; Soler et al. 2015). VLPs are positively correlated with bacterial abundance, activity and chlorophyll-a (chl-a) concentrations (Maranger, Bird and Juniper 1994; Gowing et al. 2002, 2004). In aquatic environments, a typical virus-to-bacteria ratio (VBR; more precisely VLP-to-prokaryotic cell ratio) is 10:1 (Maranger and Bird 1995). Bacterial and viral density dictates their contact rate, which is one of the key controls in virus–host interactions. The semi-enclosed environment of brine channels may increase this contact rate, especially during winter, when the brine channels are narrower and the brine even more concentrated (Wells and Deming 2006a). Sea ice may, therefore, be a place where virus–host interactions can be enhanced, compared with the open ocean. To understand these effects, virus–host systems need to be isolated to examine their interactions in detail. To the best of our knowledge, only three virus–host systems to date have been isolated from Arctic and seven from Baltic Sea ice (Borriss et al. 2003; Luhtanen et al. 2014, Yu et al. 2015), but none from Antarctic sea ice. The viruses isolated represented different phage morphologies. The Arctic virus isolates Shewanella phage 1a and Colwellia phage 21c are icosahedral viruses with either a contractile or noncontractile tail, resembling double-stranded DNA (dsDNA) phages of the order Caudovirales (Borriss et al. 2003; 2007). The filamentous nonlytic phage f327 from the Arctic infects a Pseudoalteromonas strain and is reminiscent of viruses in the family Inoviridae (Yu et al. 2015). In addition, seven tailed icosahedral dsDNA phages infecting strains from either Flavobacterium or Shewanella were isolated from Baltic Sea ice (Luhtanen et al. 2014; Senčilo et al. 2015). All virulent sea-ice phage isolates have a narrow host range, are cold-active (capable of infection and production at ≤4°C; Wells and Deming (2006b)), and produce plaques (clear zones on bacterial lawns used to determine the number of infectious viruses) only at the lower end of their host bacterial temperature growth range (Borriss et al. 2003; Luhtanen et al. 2014). In addition, a cold-active siphophage 9A was isolated from Arctic nepheloid layer seawater (Wells and Deming 2006b) for Colwellia psychrerythraea strain 34H, isolated originally from Arctic shelf sediments (Huston, Krieger-Brockett and Deming 2000). It was reminiscent of the isolates from sea ice, because it also has a narrow host range, is cold-active, and has a more restricted growth temperature range than the host bacteria. Here, we report the isolation of the first cultivable phage–host systems and VLP abundance from the winter-sea ice in the Weddell Sea, Antarctica. Studying phage–host systems gives us valuable information on sea-ice microbial communities and their potential roles in the sea-ice ecosystem. MATERIALS AND METHODS Sea-ice sampling and ice properties Pack ice samples were collected from the Weddell Sea (Antarctica) during the austral winter as part of the Antarctic Winter Ecosystem Climate Study (AWECS) aboard R/V Polarstern in June–August 2013 (leg ANT-XXIX/6). Sampling was performed either with a metal basket (pancake ice at stations 486, 488 and 489) or using a motorized, trace-metal-clean (electropolished steel) CRREL-type ice-coring auger (Lannuzel et al.2006), 14 cm in diameter. For this study, full-depth ice cores (one or two) were taken from 10 locations (ice-stations 486, 488, 489, 493, 496, 500, 503, 506, 515 and 517; Table 1; Fig. S1, Supporting Information), as described in Tison et al. (2017). Bulk ice was used, because some of the microbes may have been attached to the brine channel walls and could have been lost if only the liquid brine was sampled (Meiners, Krembs and Gradinger 2008). The ice cores were cut into one, three, five or seven layers, depending on the ice thickness, and crushed gently with a hammer inside a polyethylene plastic bag. When two cores were taken, the corresponding layers of the nearby ice cores were pooled (Table 1). The VLP abundances were measured from these layers. The bacterial abundances (Table 1), bacterial production (measured as thymidine incorporation) and bacterial community composition analyses from these ice cores have been published elsewhere (Eronen-Rasimus et al. 2017). For the isolation work, we used the ice samples from first-year ice-station 500 and early second-year ice-station 515a (Tison et al. 2017). The surface parts of the 515a core were removed to minimize contamination, and the core was cut into 12 layers (∼12–14 cm each; Table 2), using an electric-band saw sterilized with 70% ethanol. The 12 layers were used separately to isolate the bacterial strains. The bacterial community composition of ice core 515a has also been published under the name 515 T (0–56 cm), M (56–126 cm) and B (126–166 cm; Eronen-Rasimus et al. 2017). For virus isolation, bulk ice from station 500 and core 515a was used. The ice samples were left to melt in sterilized containers at 4°C overnight, after that the remaining ice was melted in a water bath with continuous shaking (Rintala et al. 2014). After melting, the samples were immediately transferred back to 4°C. Table 1. VLP and bacterial abundances and VBRs together with ice temperature and chlorophyll-a concentrations in the different layers of ice cores from all the AWECS sampling stations.         Ice depth (cm) from air-ice interface            Station  Date  Latitude  Longitude  Core I  Core II  Ice temperature °Cb  VLP x 105ml-1 in bulk iced  Bacteria x 105ml-1 in bulk icec,d  VBR Viruses/Bacteria  Total chl-a (µg l-1 bulk ice)b,d  486a  6/17/2013  −61.526  −0.086  0–6  –  −5.0  1.90  0.56  3.4  0.41  488a  6/18/2013  −62.928  −0.006  0–15  –  −10.9  4.90  0.56  8.8  0.29          15–20  –  −7.4  2.60  0.81  3.2  0.28          20–35  –  −4.1  6.80  1.20  5.7  1.39  489a  6/19/2013  −63.901  −0.031  0–15  –  −8.4  6.50  1.20  5.4  1.48          15–22  –  −5.8  4.90  1.30  3.8  3.89          22–37  –  −3.4  5.80  0.85  6.8  2.36  493  6/21/2013  −66.44  0.122  0–15  0–15  −8.7  13.00  3.40  3.8  9.56          15–46  15–38  −5.1  9.30  3.80  2.4  10.43          46–61  38–53  −2.7  9.30  1.80  5.2  16.59  496  6/24/2013  −67.466  −0.021  0–15  0–15  −5.3  4.70  1.30  3.6  1.81          15–45  15–57  −4.1  3.90  2.70  1.4  7.49          45–60  57–72  −2.6  5.90  2.50  2.4  15.24  500  7/3/2013  −67.949  −6.658  0–15  0–15  −2.4  16.00  2.60  6.2  3.83          15–35  15–35  −2.7  12.00  3.60  3.3  7.73          35–55  35–52  −2.6  46.00  4.00  11.5  10.34          55–75  52–72  −2.4  20.00  3.40  5.9  7.08          75–90  72–87  −2.2  9.70  2.70  3.6  9.31  503  7/8/2013  −67.187  −13.224  0–15  0–15  −7.3  8.70  0.65  13.4  0.83          15–25  15–25  −6.0  7.50  0.79  9.5  0.86          25–37  25–36  −5.0  9.20  0.81  11.4  1.05          37–47  36–46  −3.4  10.00  0.80  12.5  0.99          47–62  46–61  −2.2  6.70  0.56  12.0  0.70  506  7/11/2013  −67.19  −23.042  0–15  0–15  −6.5  3.90  0.51  7.6  0.31          15–34  15–30  −4.9  5.50  0.80  6.9  0.69          34–49  30–45  −3.1  ND  ND  ND  0.82  515a  7/26/2013  −63.456  −51.308  0–15  –  −7.6  6.90  3.60  1.9  0.78          15–45  –  −6.1  9.30  6.00  1.6  2.08          45–75  –  −6.4  21.00  7.30  2.9  30.73          75–104  –  −4.8  40.00  24.00  1.7  61.54          104–134  –  −3.9  49.00  42.00  1.2  62.57          134–164  –  −3.8  10.00  9.50  1.1  10.95          164–179  –  −2.3  8.90  12.00  0.7  11.66  517  7/30/2013  −63.509  −51.112  0–15  0–15  −3.7  5.70  0.60  9.5  0.74          15–30  15–30  −2.6  4.60  0.87  5.3  0.55          30–43  30–43  −2.2  2.20  0.71  3.1  0.22          43–58  43–58  −2.0  2.40  1.30  1.8  0.31          58–73  58–73  −1.8  5.80  1.40  4.1  0.60          Ice depth (cm) from air-ice interface            Station  Date  Latitude  Longitude  Core I  Core II  Ice temperature °Cb  VLP x 105ml-1 in bulk iced  Bacteria x 105ml-1 in bulk icec,d  VBR Viruses/Bacteria  Total chl-a (µg l-1 bulk ice)b,d  486a  6/17/2013  −61.526  −0.086  0–6  –  −5.0  1.90  0.56  3.4  0.41  488a  6/18/2013  −62.928  −0.006  0–15  –  −10.9  4.90  0.56  8.8  0.29          15–20  –  −7.4  2.60  0.81  3.2  0.28          20–35  –  −4.1  6.80  1.20  5.7  1.39  489a  6/19/2013  −63.901  −0.031  0–15  –  −8.4  6.50  1.20  5.4  1.48          15–22  –  −5.8  4.90  1.30  3.8  3.89          22–37  –  −3.4  5.80  0.85  6.8  2.36  493  6/21/2013  −66.44  0.122  0–15  0–15  −8.7  13.00  3.40  3.8  9.56          15–46  15–38  −5.1  9.30  3.80  2.4  10.43          46–61  38–53  −2.7  9.30  1.80  5.2  16.59  496  6/24/2013  −67.466  −0.021  0–15  0–15  −5.3  4.70  1.30  3.6  1.81          15–45  15–57  −4.1  3.90  2.70  1.4  7.49          45–60  57–72  −2.6  5.90  2.50  2.4  15.24  500  7/3/2013  −67.949  −6.658  0–15  0–15  −2.4  16.00  2.60  6.2  3.83          15–35  15–35  −2.7  12.00  3.60  3.3  7.73          35–55  35–52  −2.6  46.00  4.00  11.5  10.34          55–75  52–72  −2.4  20.00  3.40  5.9  7.08          75–90  72–87  −2.2  9.70  2.70  3.6  9.31  503  7/8/2013  −67.187  −13.224  0–15  0–15  −7.3  8.70  0.65  13.4  0.83          15–25  15–25  −6.0  7.50  0.79  9.5  0.86          25–37  25–36  −5.0  9.20  0.81  11.4  1.05          37–47  36–46  −3.4  10.00  0.80  12.5  0.99          47–62  46–61  −2.2  6.70  0.56  12.0  0.70  506  7/11/2013  −67.19  −23.042  0–15  0–15  −6.5  3.90  0.51  7.6  0.31          15–34  15–30  −4.9  5.50  0.80  6.9  0.69          34–49  30–45  −3.1  ND  ND  ND  0.82  515a  7/26/2013  −63.456  −51.308  0–15  –  −7.6  6.90  3.60  1.9  0.78          15–45  –  −6.1  9.30  6.00  1.6  2.08          45–75  –  −6.4  21.00  7.30  2.9  30.73          75–104  –  −4.8  40.00  24.00  1.7  61.54          104–134  –  −3.9  49.00  42.00  1.2  62.57          134–164  –  −3.8  10.00  9.50  1.1  10.95          164–179  –  −2.3  8.90  12.00  0.7  11.66  517  7/30/2013  −63.509  −51.112  0–15  0–15  −3.7  5.70  0.60  9.5  0.74          15–30  15–30  −2.6  4.60  0.87  5.3  0.55          30–43  30–43  −2.2  2.20  0.71  3.1  0.22          43–58  43–58  −2.0  2.40  1.30  1.8  0.31          58–73  58–73  −1.8  5.80  1.40  4.1  0.60  a Only one ice core sampled. b Ice temperatures and chl-a values from Tison et al. (2017). c Bacterial abundances are also published in Eronen-Rasimus et al. (2017). d Analysed from combined layers of the sibling cores I and II, when two cores were collected. VLP = virus-like particle, AWECS = Antarctic Winter Ecosystem Climate Study, chl-a = chlorophyll a, ND = not determined. View Large Table 2. Bacterial strains isolated in this study from the ice core 515a. Isolation depth from air–ice interface (cm)  Isolation mediaa  Bacterial strain  Closest match genusb  Identity % at genus level  Accession number  Used for virus isolation  0–14  Z  IceBac 363  Halomonas  100.0  KY194856  ✓  14–28  Z  IceBac 364  Polaribacter  99.7  KY194850  ✓  14–28  Z  IceBac 365  Glaciecola  99.6  KY194821  ✓  14–28  Z  IceBac 367  Paraglaciecola  99.9  KY194799  ✓  14–28  Z  IceBac 368  Paraglaciecola  99.9  KY194805  ✓  14–28  Z  IceBac 369  Marinobacter  99.3  KY194841    14–28  Z  IceBac 370  Marinobacter  99.3  KY194842  ✓  14–28  Z  IceBac 371  Glaciecola  99.6  KY194822    14–28  Z  IceBac 372  Paraglaciecola (H)  99.9  KY194800  ✓  14–28  MOX  IceBac 373  Polaribacter  100.0  KY194848  ✓  14–28  MOX  IceBac 377  Glaciecola  99.6  KY194823  ✓  28–42  Z  IceBac 378  Glaciecola  99.6  KY194828  ✓  28–42  Z  IceBac 379  Glaciecola  99.6  KY194824  ✓  28–42  Z  IceBac 380  Glaciecola  99.6  KY194825    28–42  Z  IceBac 381  Glaciecola  99.6  KY194829  ✓  28–42  Z  IceBac 382  Glaciecola  99.6  KY194831  ✓  28–42  Z  IceBac 383  Paraglaciecola  99.9  KY194801  ✓  28–42  MOX  IceBac 384  Polaribacter  100.0  KY194845    42–56  Z  IceBac 385  Polaribacter  99.7  KY194844    42–56  Z  IceBac 386  Glaciecola  99.6  KY194815  ✓  42–56  Z  IceBac 387  Glaciecola  99.5  KY194816  ✓  42–56  Z  IceBac 388  Glaciecola  99.6  KY194826  ✓  42–56  Z  IceBac 389  Glaciecola  99.6  KY194817  ✓  42–56  Z  IceBac 390  Glaciecola  99.6  KY194830    42–56  Z  IceBac 391  Marinobacter  99.3  KY194843    42–56  Z  IceBac 392  Paracoccus  99.9  KY194857  ✓  42–56  MOX  IceBac 394  Glaciecola  99.6  KY194832  ✓  56–70  Z  IceBac 396  Glaciecola  99.6  KY194827    56–70  Z  IceBac 397  Glaciecola  99.6  KY194818    56–70  Z  IceBac 398  Glaciecola  99.5  KY194820    56–70  Z  IceBac 399  Glaciecola  99.4  KY194819    56–70  Z  IceBac 400  Polaribacter  99.7  KY194849  ✓  56–70  Z  IceBac 401  Paraglaciecola  100.0  KY194802  ✓  56–70  Z  IceBac 402  Paraglaciecola  99.9  KY194806  ✓  70–84  C  IceBac 403  Colwellia  99.4  KY194851  ✓  70–84  Z  IceBac 404  Polaribacter  99.7  KY194846  ✓  70–84  Z  IceBac 405  Octadecabacter  100.0  KY194834  ✓  98–112  Z  IceBac 408  Polaribacter  99.7  KY194847  ✓  98–112  Z  IceBac 409  Glaciecola  99.6  KY194833  ✓  112–126  C  IceBac 410  Paraglaciecola  100.0  KY194813  ✓  112–126  Z  IceBac 411  Paraglaciecola  100.0  KY194808  ✓  112–126  Z  IceBac 412  Paraglaciecola  100.0  KY194803  ✓  112–126  Z  IceBac 413  Octadecabacter  100.0  KY194835  ✓  112–126  Z  IceBac 414  Paraglaciecola  100.0  KY194804  ✓  126–140  C  IceBac 415  Pseudoalteromonas  100.0  KY194855  ✓  126–140  Z  IceBac 416  Paraglaciecola  99.9  KY194807  ✓  126–140  Z  IceBac 417  Paraglaciecola  100.0  KY194809  ✓  126–140  Z  IceBac 418  Octadecabacter  100.0  KY194836  ✓  126–140  Z  IceBac 419  Octadecabacter (H)  100.0  KY194837  ✓  140–154  Z  IceBac 420  Paraglaciecola  100.0  KY194810  ✓  140–154  Z  IceBac 421  Pseudoalteromonas  100.0  KY194853  ✓  140–154  Z  IceBac 422  Pseudoalteromonas  100.0  KY194854  ✓  140–154  Z  IceBac 423  Colwellia  99.4  KY194852  ✓  140–154  Z  IceBac 424  Octadecabacter  100.0  KY194838  ✓  140–154  MOX  IceBac 426  Paraglaciecola  100.0  KY194814  ✓  154–166  Z  IceBac 428  Paraglaciecola  99.9  KY194811  ✓  154–166  Z  IceBac 430  Octadecabacter (H)  100.0  KY194839  ✓  154–166  Z  IceBac 431  Octadecabacter  100.0  KY194840  ✓  154–166  MOX  IceBac 433  Paraglaciecola  100.0  KY194812  ✓  Isolation depth from air–ice interface (cm)  Isolation mediaa  Bacterial strain  Closest match genusb  Identity % at genus level  Accession number  Used for virus isolation  0–14  Z  IceBac 363  Halomonas  100.0  KY194856  ✓  14–28  Z  IceBac 364  Polaribacter  99.7  KY194850  ✓  14–28  Z  IceBac 365  Glaciecola  99.6  KY194821  ✓  14–28  Z  IceBac 367  Paraglaciecola  99.9  KY194799  ✓  14–28  Z  IceBac 368  Paraglaciecola  99.9  KY194805  ✓  14–28  Z  IceBac 369  Marinobacter  99.3  KY194841    14–28  Z  IceBac 370  Marinobacter  99.3  KY194842  ✓  14–28  Z  IceBac 371  Glaciecola  99.6  KY194822    14–28  Z  IceBac 372  Paraglaciecola (H)  99.9  KY194800  ✓  14–28  MOX  IceBac 373  Polaribacter  100.0  KY194848  ✓  14–28  MOX  IceBac 377  Glaciecola  99.6  KY194823  ✓  28–42  Z  IceBac 378  Glaciecola  99.6  KY194828  ✓  28–42  Z  IceBac 379  Glaciecola  99.6  KY194824  ✓  28–42  Z  IceBac 380  Glaciecola  99.6  KY194825    28–42  Z  IceBac 381  Glaciecola  99.6  KY194829  ✓  28–42  Z  IceBac 382  Glaciecola  99.6  KY194831  ✓  28–42  Z  IceBac 383  Paraglaciecola  99.9  KY194801  ✓  28–42  MOX  IceBac 384  Polaribacter  100.0  KY194845    42–56  Z  IceBac 385  Polaribacter  99.7  KY194844    42–56  Z  IceBac 386  Glaciecola  99.6  KY194815  ✓  42–56  Z  IceBac 387  Glaciecola  99.5  KY194816  ✓  42–56  Z  IceBac 388  Glaciecola  99.6  KY194826  ✓  42–56  Z  IceBac 389  Glaciecola  99.6  KY194817  ✓  42–56  Z  IceBac 390  Glaciecola  99.6  KY194830    42–56  Z  IceBac 391  Marinobacter  99.3  KY194843    42–56  Z  IceBac 392  Paracoccus  99.9  KY194857  ✓  42–56  MOX  IceBac 394  Glaciecola  99.6  KY194832  ✓  56–70  Z  IceBac 396  Glaciecola  99.6  KY194827    56–70  Z  IceBac 397  Glaciecola  99.6  KY194818    56–70  Z  IceBac 398  Glaciecola  99.5  KY194820    56–70  Z  IceBac 399  Glaciecola  99.4  KY194819    56–70  Z  IceBac 400  Polaribacter  99.7  KY194849  ✓  56–70  Z  IceBac 401  Paraglaciecola  100.0  KY194802  ✓  56–70  Z  IceBac 402  Paraglaciecola  99.9  KY194806  ✓  70–84  C  IceBac 403  Colwellia  99.4  KY194851  ✓  70–84  Z  IceBac 404  Polaribacter  99.7  KY194846  ✓  70–84  Z  IceBac 405  Octadecabacter  100.0  KY194834  ✓  98–112  Z  IceBac 408  Polaribacter  99.7  KY194847  ✓  98–112  Z  IceBac 409  Glaciecola  99.6  KY194833  ✓  112–126  C  IceBac 410  Paraglaciecola  100.0  KY194813  ✓  112–126  Z  IceBac 411  Paraglaciecola  100.0  KY194808  ✓  112–126  Z  IceBac 412  Paraglaciecola  100.0  KY194803  ✓  112–126  Z  IceBac 413  Octadecabacter  100.0  KY194835  ✓  112–126  Z  IceBac 414  Paraglaciecola  100.0  KY194804  ✓  126–140  C  IceBac 415  Pseudoalteromonas  100.0  KY194855  ✓  126–140  Z  IceBac 416  Paraglaciecola  99.9  KY194807  ✓  126–140  Z  IceBac 417  Paraglaciecola  100.0  KY194809  ✓  126–140  Z  IceBac 418  Octadecabacter  100.0  KY194836  ✓  126–140  Z  IceBac 419  Octadecabacter (H)  100.0  KY194837  ✓  140–154  Z  IceBac 420  Paraglaciecola  100.0  KY194810  ✓  140–154  Z  IceBac 421  Pseudoalteromonas  100.0  KY194853  ✓  140–154  Z  IceBac 422  Pseudoalteromonas  100.0  KY194854  ✓  140–154  Z  IceBac 423  Colwellia  99.4  KY194852  ✓  140–154  Z  IceBac 424  Octadecabacter  100.0  KY194838  ✓  140–154  MOX  IceBac 426  Paraglaciecola  100.0  KY194814  ✓  154–166  Z  IceBac 428  Paraglaciecola  99.9  KY194811  ✓  154–166  Z  IceBac 430  Octadecabacter (H)  100.0  KY194839  ✓  154–166  Z  IceBac 431  Octadecabacter  100.0  KY194840  ✓  154–166  MOX  IceBac 433  Paraglaciecola  100.0  KY194812  ✓  a Z = Zobell media; MOX = MOX media; C = concentrated Zobell media. b Original isolation hosts of the viruses are marked by (H). View Large The pancake ice at stations 486 and 488 was 6 cm and 35 cm thick, the first-year ice at stations 489–506 and 517 was 37–90 cm, and the early second-year ice at station 515 was 179 cm (Table 1, Tison et al.2017). The ice temperature varied between −1.8°C and −10.9°C, with a median temperature of −3.8°C and ice chl-a median concentration of 2.4 µg l−1(Tison et al.2017). The ice at station 500 was dominated by frazil ice and at station 515 by mixed columnar and frazil ice (Tison et al.2017). The brine salinity varied from 122 to 40 practical salinity units (Tison et al.2017). Isolation of the bacterial strains Isolation of the bacterial strains was started immediately after the 12 layers of the ice core 515a samples were melted. The strains were isolated by plating 100 μl of the melted sample on (I) ZoBell plates (1000 ml Southern Ocean water, 5 g peptone, 1 g yeast extract, 15 g agar; Helmke and Weyland 1995; Middelboe et al.2003), (II) concentrated ZoBell plates (the Southern Ocean water was concentrated by boiling to half of the initial volume, otherwise similar to ZoBell) and (III) Modified Oxford (MOX) agar plates (750 ml seawater, 250 ml ultrapure water, 1 g KNO4, 0.2 g yeast extract, 10 mg FePO4, 2 g HEPES, 12 g agar). The plates were incubated at 4°C and transported from R/V Polarstern to the home laboratory with temperature-controlled courier transportation at −2.6°C to + 6.1°C (World Courier, AmerisourceBergen, Stamford, CT, USA). After 1 month of incubation in the dark at 4°C, the various colony morphotypes were picked and colony-purified at three consecutive times. Various strains were isolated from all layers, except from depths of 84–98 cm. Colony purification and cultivation were done on modified ZoBell medium, Reef Crystal (RC) medium: 33 g RCs, Aquarium Systems Inc. Sarrebourg, France, 1000 ml ultrapure water, 5 g peptone and 1 g yeast extract. The agar concentration for the plates was 1.5% (w/v). The strains were grown aerobically in RC medium at 4°C for 7 days and stored at −80°C, supplemented with 15% (v/v) glycerol. Identification of the bacterial strains The colony-purified strains were identified by 16S rRNA gene sequencing. The genomic DNA was isolated with an UltraClean Microbial DNA Isolation Kit (MO BIO Laboratories Inc., Carlsbad, CA, USA). The partial 16S rRNA genes were amplified with PCR, using primers F27 (Sait et al. 2003) and R1406 (Lane et al. 1985) or pA and pGr (Edwards et al. 1989). The Sanger sequencing was performed at the DNA Sequencing and Genomics Unit, Institute of Biotechnology (University of Helsinki), using primers pDr, pE and pFr (Edwards et al. 1989). The taxonomic identification of the strains was done with SILVA Incremental Aligner (SINA) Alignment Service (version 1.2.11, 10.6.2016, Pruesse, Peplies and Glockner 2012). The partial 16S rRNA gene sequences of the isolated bacterial strains are deposited in the NCBI GenBank database under accession numbers KY194799–KY194857 (Table 2). Phylogenetic analysis of the 16S rRNA sequences For the phylogenetic tree (Fig. 1), alignment was performed with SINA Aliqnment Service (version 1.2.11, minimum identity: 0.8, Pruesse, Peplies and Glockner 2012). Reference sequences were selected, based on SINA sequence matching, and for the nontype-strain sequence matches, their type-strain representatives were also added (EZBioCloud Database (Yoon et al.2017)). After the alignment, all sequences were truncated according to the sequence length, and the bootstrapped (1000) maximum-likelihood tree was constructed, using RAxML (version 8.2.0; Stamatakis 2014), with the GTRGAMMA evolution model. The tree was visualized with the Interactive Tree Of Life (iTOL) online tool (Leturnic and Bork 2007). Figure 1. View largeDownload slide Bootstrapped (1000) phylogenetic maximum-likelihood tree of the 16S rRNA gene sequences of the bacterial strains isolated from Antarctic sea ice. The bootstrap values (>50%) are shown with black circles. The strains are colored at the genus level (see the color key), based on their classification with SILVA Incremental Aligner (SINA) (version 1.2.11, minimum identity: 0.8, Pruesse et al. 2012). The sensitivities of the bacterial strains to isolated phages are shown as efficiency of plating (EOP; on right in gray scale). For the original host (marked by H), the EOP was set to a value of 1. The strains used as references or that do not form a lawn on solid growth media were not tested for EOP (marked with white). The scale bar indicates nucleotide substitutions per position. Figure 1. View largeDownload slide Bootstrapped (1000) phylogenetic maximum-likelihood tree of the 16S rRNA gene sequences of the bacterial strains isolated from Antarctic sea ice. The bootstrap values (>50%) are shown with black circles. The strains are colored at the genus level (see the color key), based on their classification with SILVA Incremental Aligner (SINA) (version 1.2.11, minimum identity: 0.8, Pruesse et al. 2012). The sensitivities of the bacterial strains to isolated phages are shown as efficiency of plating (EOP; on right in gray scale). For the original host (marked by H), the EOP was set to a value of 1. The strains used as references or that do not form a lawn on solid growth media were not tested for EOP (marked with white). The scale bar indicates nucleotide substitutions per position. Abundance of VLPs by flow cytometry The abundances of the VLPs were analyzed with flow cytometry from all the ice layers originating from the 10 stations (Table 1). Sample handling and measurements were done according to (Brussaard et al.2010), except that paraformaldehyde [1% (v/v) in phosphate-buffered saline, Alfa Aesar GmbH & Co KG, Karlsruhe, Germany] was used as a fixative. The samples were stained with SYBR Green I (Sigma-Aldrich Inc., Saint Louis, MO, USA) at room temperature. To measure the background, virus-sized particles were removed from the control samples (pooled from the samples originating from the various layers and stations) by ultrafiltration (Amicon Ultra-15 concentrators, MWCO 100 000 Da; Merck Millipore, Billerica, MA, USA; 4000 g, 4 min, 4°C), and the controls were processed in the same way as the actual samples. Earlier isolated and purified 1/4, 1/32, 1/40, 1/41, 1/44, 3/49 phage particles (Luhtanen et al.2014) were used as controls to define the virus population, and Fluoresbrite 0.5-μm microspheres (Polysciences Inc., Warrington, PA, USA) were used as a size standard. Enumeration of the diluted samples (dilution factor 10; molecular biology grade TE buffer, AppliChem GmbH, Darmstadt, Germany) was carried out, using a CyFlow Cube 8 (Partec GmbH, Münster, Germany) flow cytometer. A sample of 25 μl (defined by the flow cytometry electrodes) was analyzed, using a flow rate of 12 μl min−1. The data were acquired on a dot plot displaying green fluorescence (488 nm) versus side scatter signal, both on a logarithmic scale. The detection trigger was depicted as a green fluorescence. The data were analyzed, using FCS Express 4 software (De Novo Software, Glendale, CA, USA; Fig. S2, Supplementary material), and the numbers were corrected for dilution. The VBR was defined, using published bacterial abundances (Eronen-Rasimus et al.2017). Isolation of viruses For virus isolations, the melted layers of the station 500 core were pooled, as were the layers of core 515a. The melted ice was filtered through a 0.22-μm Durapore Membrane polyvinylidene difluoride filter (EMD Millipore Corporation, Billerica, MA, USA) to remove cellular organisms. The filtrates were concentrated 50 times, using Amicon Ultra-15 concentrators (MWCO 100 000 Da) and centrifugation (2000 g, 5 min, 4°C). The concentrates were stored in 15% (v/v) glycerol at −80°C and transported in liquid nitrogen to the home laboratory. All isolated and colony-purified bacterial strains (Table 2, Fig. 1) that were able to form a bacterial lawn were used to isolate viruses. Virus isolation and further cultivation were done with a plaque assay. For virus isolation, 10 µl and 100 μl of the concentrated sample with 200 μl of dense host bacterial suspension (grown for 7 days in RC medium at 4°C) and 3 ml of melted RC top-layer agar [0.4% (w/v) agar; 43°C] were mixed and poured on RC plates. After 1–3 weeks of incubation at 4°C, the plaques were individually picked and plaque-purified three consecutive times with the plaque assay. For the plaque assay, 100 μl of suitable virus dilution, 200 μl of host liquid culture, and 3 ml of RC top-layer agar were mixed and poured on RC plates, which were incubated for 5–7 days at 4°C. Production and purification of viruses The virus lysates were prepared with the plaque assay as described above, using semiconfluent plates. The top-layer agar was collected and mixed with 2 ml of RC broth per plate. The suspension was incubated for 1 h at 4°C with shaking, after which the cell debris and agar were removed (centrifugation: 10 000 g, 30 min, 4°C). Virus precipitation from the lysates was optimized with various ammonium sulfate concentrations [50%, 60%, 70% and 80% (w/v)], using either saturated solution or ammonium sulfate powder (Boulanger and Puvion 1973; Burgess 2009). Ammonium sulfate was mixed with or dissolved in the lysate for 1 h at 4°C with shaking. The precipitated viruses were collected (centrifugation: 14 000 g, 60 min), washed with SM buffer (50 mM Tris-HCl pH 7.5, 100 mM NaCl, 8 mM MgSO4; Borriss et al.2003) with or without 0.01% (w/v) gelatin and dissolved in SM buffer on ice. The virus aggregates were removed (centrifugation 9300 g, 10 min, 4°C), and the viruses in the supernatant were subjected to rate-zonal centrifugation (153 208 g, 30–70 min, 10°C), using 10–30% (w/v) linear sucrose gradients in SM buffer (Anderson et al.1966; Lawrence and Steward 2010). The gradients were fractionated (12 fractions), and the infectivity (plaque assay), absorbance (260 nm), and the protein, nucleic acid and lipid contents of the fractions were determined (see below). The virus particles in the light-scattering zones were collected with differential centrifugation (104 087 g, 3 h, 10°C), and the particles were dissolved in SM buffer overnight on ice. Virus particle analyses The absorbance values (260 nm) of the 12 sucrose gradient fractions were measured with an Eppendorf BioPhotometer D30 (Eppendorf AG, Hamburg, Germany), using 30% (w/v) sucrose in SM buffer as a blank sample. The viral structural proteins were separated with SDS-PAGE (16% (w/v) acrylamide; Olkkonen and Bamford 1989). The SM buffer used in the protein analysis samples did not contain gelatin. The protein concentrations were determined with a Coomassie Blue-based method (Bradford 1976), using bovine serum albumin as a standard. For the SDS-PAGE, when appropriate, the samples were concentrated with 10% (v/v) trichloroacetic acid precipitation (30 min, on ice). The precipitate was collected (centrifugation: 16 200 g, 30 min, 4°C). The resolving gels were stained with Brilliant Blue R (Sigma-Aldrich) for proteins and, when appropriate, stained with Sudan Black B (Sigma-Aldrich) for lipids and the stacking gels with ethidium bromide for nucleic acids. For the lipid-staining control, purified PRD1 particles (Bamford and Bamford 1991) were used as a control. To test the sensitivity of the viruses to Triton X-100, viruses were incubated in 0.1% (v/v) and 0.01% Triton X-100 (in SM buffer) for 3 h and 24 h at 4°C. SM buffer was used as a control. The infectivity of the viruses tested was determined with a plaque assay. The sensitivity of the host organisms to Triton X-100 was analyzed similarly, except that the number of colony-forming units was determined by plating. For TEM, the purified virus particles were negatively stained for 20 s, using 2% (w/v) uranyl acetate (pH 7), 3% (w/v) uranyl acetate (pH 4.5), or 1% potassium phosphotungstate (pH 7). A JEOL JEM-1400 TEM (Electron Microscopy Unit, Institute of Biotechnology, University of Helsinki) was used with 80-kV tension for detailed investigation of the viruses. Temperature ranges of host growth and virus infection The growth of the original three host strains at different temperatures was determined by plating 100 μl of diluted bacterial suspension on RC plates, which were incubated at 0, 4, 10, 15, or 20°C for 60 days. The infection ability of the bacteriophages at different temperatures was tested with a plaque assay. The host suspensions used for the plaque assay were incubated at 4°C. The plates were incubated at 0, 4, 10, or 15°C for 20 days. Virus-bacteria interactions All the bacterial isolates that were able to grow as a lawn on a plate were tested for their sensitivity to the isolated viruses (Table 2, Fig. 1). Ten microliters of undiluted and 100-times diluted virus lysates were spotted on the host strain lawns. RC medium was used as a negative control and the original virus-host pair as a positive control. The plates were incubated at 4°C for 7–14 days. The growth temperature was >0°C to prevent freezing, but still at the range of cold-active viruses. All positive results were verified with a plaque assay, using suitable dilutions of the virus. The efficiency of plating (EOP) was calculated according to the plaque count obtained with the target strain, compared with that obtained with the isolation host strain. RESULTS Sea-ice bacterial isolates We isolated 59 bacterial strains from core 515a (Table 2). The majority of the isolates (∼59%) were classified as members of the genera Glaciecola or Paraglaciecola (Fig. 1). Only one strain, IceBac 363, was isolated from the coldest top layer and belonged to the genus Halomonas. The remaining strains were identified as members of the genera Octadecabacter (seven strains), Polaribacter (seven strains), Marinobacter (three strains), Pseudoalteromonas (three strains), Colwellia (two strains), or Paracoccus (one strain). All isolated bacterial strains belonging to the same genus had identical or nearly identical (identity 99.3–100%) partial 16S rRNA gene sequences (at minimum 1290 base pairs, bp). VLP abundance and VBRs in sea ice Using flow cytometry, we determined that VLPs occurred throughout the ice cores in varying numbers. The mean VLP abundance of all the ice cores sampled was 10.9 × 105 ml−1 (range 1.9 × 105–49.0 × 105 ml−1) in the bulk ice, with the highest numbers at stations 500 and 515 (6.9 × 105–49.0 × 105 ml−1; Table 1). The mean VBR of all the ice cores sampled was 5.3 (0.7–13.4; Table 1). The highest VBR values were at station 503 (9.5–13.4; Table 1) and the lowest at station 515 (0.7–2.9; Table 1). Sea-ice bacteriophage isolates Forty-eight out of the 59 bacterial strains were able to form a bacterial lawn on a plate and were consequently used to screen phages (Table 2). Four phages were obtained, cultivated and purified in the laboratory (Table 3). The phages were named after the isolation host genus, area of isolation, and the initials of notable persons in this study (Krupovic et al. 2016). The names and their abbreviations are Paraglaciecola Antarctic GD virus 1 (PANV1), Paraglaciecola Antarctic JLT virus 2 (PANV2), Octadecabacter Antarctic BD virus 1 (OANV1) and Octadecabacter Antarctic DB virus 2 (OANV2). Phages PANV1 and PANV2 originated from station 500 and were isolated for the same host (IceBac 372), which was classified as Paraglaciecola psychrophila (similarity 99.9%). PANV1 produced clear plaques 3–4 mm in diameter, whereas the PANV2 plaques of 3–5 mm in diameter had a clear center surrounded by a turbid halo. Phages OANV1 and OANV2 were isolated from core 515a for two different bacterial hosts, IceBac 419 and IceBac 430, respectively, both identified as Octadecabacter antarcticus. Both phages produced clear plaques, but the diameters were different (3–6 mm for OANV1, 6–8 mm for OANV2). The optimized phage lysate titers varied from ∼6 × 109 to ∼5 × 1011 plaque-forming units (pfu) ml−1, depending on the phage (Table 3). The infectivity of the lysates was retained for several months when stored at 4°C. All three host strains originated from different layers in the ice core (Table 2). Table 3. Phages isolated in this study. Phage  Sampling station  Isolation host  Genus of the host (closest match)  Lysate titer (pfu/ml)  Capsid head diameter (nm)a  Tail length (nm)b  Morphotype  Paraglaciecola Antarctic GD virus 1 (PANV1)  500  IceBac 372  Paraglaciecola  1.5 × 1010  71 ± 7 (n = 20)  58 ± 22 (n = 10)  myovirus  Paraglaciecola Antarctic JLT virus 2 (PANV2)  500  IceBac 372  Paraglaciecola  5.2 × 1011  52 ± 8 (n = 29)  89 ± 30 (n = 10)  siphovirus  Octadecabacter Antarctic BD virus 1 (OANV1)  515  IceBac 419  Octadecabacter  1.2 × 1010  50 ± 8 (n = 20)  83 ± 10 (n = 10)  siphovirus  Octadecabacter Antarctic DB virus 2 (OANV2)  515  IceBac 430  Octadecabacter  5.8 × 109  53 ± 7 (n = 20)  –  podovirus  Phage  Sampling station  Isolation host  Genus of the host (closest match)  Lysate titer (pfu/ml)  Capsid head diameter (nm)a  Tail length (nm)b  Morphotype  Paraglaciecola Antarctic GD virus 1 (PANV1)  500  IceBac 372  Paraglaciecola  1.5 × 1010  71 ± 7 (n = 20)  58 ± 22 (n = 10)  myovirus  Paraglaciecola Antarctic JLT virus 2 (PANV2)  500  IceBac 372  Paraglaciecola  5.2 × 1011  52 ± 8 (n = 29)  89 ± 30 (n = 10)  siphovirus  Octadecabacter Antarctic BD virus 1 (OANV1)  515  IceBac 419  Octadecabacter  1.2 × 1010  50 ± 8 (n = 20)  83 ± 10 (n = 10)  siphovirus  Octadecabacter Antarctic DB virus 2 (OANV2)  515  IceBac 430  Octadecabacter  5.8 × 109  53 ± 7 (n = 20)  –  podovirus  a average diameter. b average length. pfu = plaque-forming unit. View Large Purification and characterization of the phages To characterize the phages, virus purification methods were optimized, based on ammonium sulfate precipitation and rate-zonal ultracentrifugation, following the recovery and purity of the infectious viruses at each step. Using ammonium sulfate precipitation, 25–54% of the infectious viruses were recovered, depending on the virus (Table 4). Both the ammonium sulfate powder and the saturated solution resulted in similar yields. PANV1 and PANV2 were precipitated with 50% ammonium sulfate, whereas OANV1 and OANV2 needed 80%. The precipitated particles were further purified with rate-zonal ultracentrifugation, and significant amounts of various noninfectious protein impurity species were detected at the top of the sucrose gradient. For all viruses, a single visible infectious light-scattering zone was detected in the middle of the sucrose gradient (Fig. 2). This zone contained several proteins of different sizes that were unique for each virus. PANV1 and OANV2 had one major protein type in sizes of ∼55 and ∼35 kDa, respectively, while PANV2 and OANV1 had two major protein types (∼40 and ∼12 kDa in PANV2; ∼35 and ∼15 kDa in OANV1). A peak in the absorbance was detected in the same light-scattering zone, as were the nucleic acids when visible. Lipids could not be detected from the gels after Sudan Black staining (not shown). In addition, treatment of phages with the nonionic detergent Triton X-100 did not affect their infectivity, suggesting that the virus particles did not contain a lipid component. Specific infectivities (∼2–9 × 1012 pfu mg−1 protein) calculated for the purified phages showed that all virus samples were highly infectious after biochemical purification (Table 4). After the final concentration step with differential centrifugation, the recoveries of infectious viruses varied from ∼10% to 20% (Table 4). Figure 2. View largeDownload slide Purification of the phages by rate-zonal centrifugation in sucrose. (A) bacteriophage PANV1, (B) bacteriophage PANV2, (C) bacteriophage OANV1, (D) bacteriophage OANV2. (A–D) Top: position of the light-scattering zone (gray) in the sucrose gradient tubes. Middle: Absorbance (closed squares) and infectivity (open circles) of the 12 sucrose gradient fractions in which the top fraction is marked as 1. Bottom: Protein content of the 12 gradient fractions analyzed with SDS-PAGE and Coomassie Blue staining. The protein patterns of the final biochemically purified and concentrated phages are shown on the right. St = molecular mass marker; PRD1 = purified phage PRD1 used as a control. The dashed line marks the position of the upper edge of the light-scattering virus zone. pfu = plaque-forming unit. Figure 2. View largeDownload slide Purification of the phages by rate-zonal centrifugation in sucrose. (A) bacteriophage PANV1, (B) bacteriophage PANV2, (C) bacteriophage OANV1, (D) bacteriophage OANV2. (A–D) Top: position of the light-scattering zone (gray) in the sucrose gradient tubes. Middle: Absorbance (closed squares) and infectivity (open circles) of the 12 sucrose gradient fractions in which the top fraction is marked as 1. Bottom: Protein content of the 12 gradient fractions analyzed with SDS-PAGE and Coomassie Blue staining. The protein patterns of the final biochemically purified and concentrated phages are shown on the right. St = molecular mass marker; PRD1 = purified phage PRD1 used as a control. The dashed line marks the position of the upper edge of the light-scattering virus zone. pfu = plaque-forming unit. Table 4. Recovery of infectious phages during biochemical purification after ammonium sulfate precipitation and rate zonal centrifugation in sucrose combined with concentration step by differential centrifugation.   Total pfusa  Recovery of infectivity %  Specific infectivity pfu/mg protein  PANV1    Virus lysate  7.5 × 10 12  100.0      50% ammonium sulfate precipitate  4.1 × 10 12  54.7      Concentrated virusb  1.5 × 10 12  20.0  1.8 × 10 12  PANV2    Virus lysate  3.2 × 10 14  100.0      50% ammonium sulfate precipitate  9.6 × 10 13  30.0      Concentrated virusb  6.2 × 10 13  19.4  8.9 × 10 12  OANV1    Virus lysate  6.0 × 10 12  100.0      80% ammonium sulfate precipitate  1.5 × 10 12  25.0      Concentrated virusb  5.7 × 10 11  9.5  3.8 × 10 12  OANV2    Virus lysate  3.3 × 10 12  100.0      80% ammonium sulfate precipitate  1.2 × 10 12  35.7      Concentrated virusb  4.4 × 10 11  13.6  6.9 × 10 12    Total pfusa  Recovery of infectivity %  Specific infectivity pfu/mg protein  PANV1    Virus lysate  7.5 × 10 12  100.0      50% ammonium sulfate precipitate  4.1 × 10 12  54.7      Concentrated virusb  1.5 × 10 12  20.0  1.8 × 10 12  PANV2    Virus lysate  3.2 × 10 14  100.0      50% ammonium sulfate precipitate  9.6 × 10 13  30.0      Concentrated virusb  6.2 × 10 13  19.4  8.9 × 10 12  OANV1    Virus lysate  6.0 × 10 12  100.0      80% ammonium sulfate precipitate  1.5 × 10 12  25.0      Concentrated virusb  5.7 × 10 11  9.5  3.8 × 10 12  OANV2    Virus lysate  3.3 × 10 12  100.0      80% ammonium sulfate precipitate  1.2 × 10 12  35.7      Concentrated virusb  4.4 × 10 11  13.6  6.9 × 10 12  a calculated per a liter of original lysate. b after rate zonal centrifugation in sucrose and concentration by differential centrifugation. pfu = plaque-forming unit. View Large Transmission electron microscopy (TEM) of the purified particles showed that phage PANV1 (Fig. 3a) had a rigid, contractile tail typical of myoviruses. Its average tail length was ∼58 nm and head diameter ∼71 nm. PANV2 (Fig. 3b) infecting the same host had an ∼89-nm noncontractile tail characteristic of the siphoviruses and an ∼52-nm head diameter. OANV1 (Fig. 3c) also had a typical siphovirus tail, with an average length of ∼83 nm and a head ∼50 nm in diameter, whereas OANV2 (Fig. 3d) seemed to have a very short tail typical of podoviruses and a head ∼53 nm in diameter. Figure 3. View largeDownload slide Transmission electron micrographs of the purified and negatively stained phages. (A) PANV1, (B) PANV2, (C) OANV1 and (D) OANV2. Figure 3. View largeDownload slide Transmission electron micrographs of the purified and negatively stained phages. (A) PANV1, (B) PANV2, (C) OANV1 and (D) OANV2. Temperature range tests for host growth and phage infection All the bacterial host strains (IceBac 372, IceBac 419 and IceBac 430) were able to form colonies at the temperatures from 0°C to 15°C, but not at 20°C (Table 5), and were therefore classified as psychrophiles (Morita 1975). The effect of temperature on phage infection (plaque formation) was tested at temperatures supporting the growth of the hosts. All the phages were able to infect their original host only at 0°C and 4°C, but not at higher temperatures (Table 5). PANV1 and PANV2 produced plaques at 0°C and 4°C in 6 days, but OANV1 and OANV2 produced plaques at 0°C in 14 days and at 4°C in 6 days. Table 5. Temperature-dependent growth of the bacterial host strains and the phages.   0°  4°  10°  15°  20°  Host bacterial growtha    IceBac 372  +  +  +  (+)  −  IceBac 419  +  +  +  +  −  IceBac 430  +  +  +  +  −  Infectivity of the phagesa  PANV1  +  +  −  −  ND  PANV2  +  +  −  −  ND  OANV1  +  +  −  −  ND  OANV2  +  +  −  −  ND    0°  4°  10°  15°  20°  Host bacterial growtha    IceBac 372  +  +  +  (+)  −  IceBac 419  +  +  +  +  −  IceBac 430  +  +  +  +  −  Infectivity of the phagesa  PANV1  +  +  −  −  ND  PANV2  +  +  −  −  ND  OANV1  +  +  −  −  ND  OANV2  +  +  −  −  ND  a + = producing colonies/plaques; ( + ) = retarded growth; − = no colonies or plaques produced; ND = not determined. View Large Phage–bacteria interactions The sensitivity of all the 48 isolated bacterial strains (that were able to grow as a bacterial lawn) to the isolated phages was tested at 4°C (Table 2; Fig. 1). In all, 17 strains (13 Paraglaciecola strains and 4 Octadecabacter strains) were sensitive to at least one of the phages (Fig. 1). Of 16 Paraglaciecola isolates, IceBac 372 was sensitive to three phages: PANV1, PANV2 and OANV1. Ten other Paraglaciecola strains were sensitive to both PANV1 and PANV2, but with different plating efficiencies (EOPs), two strains were sensitive to either PANV1 or PANV2, but with low EOP, and three strains could not be infected. All seven Octadecabacter strains had 100% identical 16S rRNA gene sequences (within 1289 bp, Fig. 1). However, only four out of seven Octadecabacter strains were sensitive to either OANV1 or OANV2 and showed different EOPs. Both the PANV1 and PANV2 phages were able to infect 12 different Paraglaciecola strains with different EOPs. However, each was able to infect only a single strain (IceBac 417 or IceBac 420, respectively) that the other could not (Fig. 1). In addition to its original host strain (IceBac 419, Octadecabacter), OANV1 was able to infect two other Octadecabacter strains, but with lower EOP. It also produced plaques with high EOP in the strain IceBac 372 ( Paraglaciecola), which was the isolation host for PANV1 and PANV2 (Fig. 1). Consequently, OANV1 was able to infect strains representing two classes: Gammaproteobacteria (Paraglaciecola) and Alphaproteobacteria (Octadecabacter). In contrast, phage OANV2 was able to infect only IceBac 430 (Octadecabacter; Fig. 1). DISCUSSION We isolated and purified four Antarctic sea-ice phages (PANV1, PANV2, OANV1 and OANV2) that could be maintained and cultivated under laboratory conditions. They were cold-active (capable of infection and production at ≤4°C; Wells and Deming 2006b), infecting bacterial strains belonging to the typical sea-ice bacterial genera Paraglaciecola or Octadecabacter. The viruses were specific for host recognition at the strain level, even though OANV1 was able to infect bacterial strains from two different classes. The highest VLP abundances were in the samples where bacteria were most abundant and active (Eronen-Rasimus et al. 2017). Isolation of sea-ice bacteria and phages We isolated 59 bacterial strains belonging to nine different genera from Antarctic winter-sea ice. The ice was melted with the direct-melting method, which has been shown to result in viable bacteria counts similar to those obtained by melting with seawater addition (Helmke and Weyland 1995), even if it may cause osmotic stress due to the rapid salinity changes. The isolated strains belonged to the following genera: Colwellia, Glaciecola, Halomonas, Marinobacter, Octadecabacter, Paracoccus, Paraglaciecola (formerly Glaciecola; Shivaji and Reddy 2014) , Polaribacter, and Pseudoalteromonas, all of which are common members in the sea-ice bacterial community (Bowman et al. 1997; Brinkmeyer et al. 2003; Deming and Collins 2017). The majority of these genera were also abundant in isolation ice core 515a, based on bacterial community composition analysis (see results in Eronen-Rasimus et al. 2017). In addition, four unique phages were isolated from the sea ice with the direct-melting and plaque assay methods, even though the viruses were exposed to a 43°C temperature for a short time during the plaque assay. This has also been successfully used previously (Borriss et al. 2003), and at least some of the cold-adapted phages can apparently tolerate high temperatures for a short time. Wide temperature tolerance can be beneficial to phage survival in natural environments throughout the various seasons. Presumably, virus reproduction is most effective when the number of susceptible hosts is high and active (Thingstad and Lignell 1997). The hosts of the bacteriophages OANV1 and OANV2 belonged to the genus Octadecabacter, which was abundant in phage isolation core 515a (Eronen-Rasimus et al. 2017). The genus Paraglaciecola (host of phages PANV1 and PANV2) could not be detected separately from the genus Glaciecola in the community analysis, likely due to the short sequence length used, but Glaciecola was present in both the 500 and 515a cores (Eronen-Rasimus et al. 2017). Our results together with those of Eronen-Rasimus et al. ( 2017) support the notion that bacteriophages of these predominant bacteria may be abundant in the viral community, resulting in increased opportunities of isolating them. Phage-host interactions We tested the sensitivity of 48 isolated bacterial strains, representing nine different genera, to our phages. PANV1 and PANV2 were able to infect several closely related Paraglaciecola strains with different EOPs, but not all the strains (Fig. 1). This may have resulted from of an arms race in which the bacterial strains evolved to inhibit phage infection, leading to diversification of bacterial strains (Thingstad et al.2014). Consequently, the phages needed to evolve to be able to survive. Since PANV1 and PANV2 have different infection patterns, this may have resulted in two different phage-host coevolution lineages. Phage OANV1 was able to infect bacterial strains from two different classes: Alphaproteobacteria (Octadecabacter) and Gammaproteobacteria (Paraglaciecola, Fig. 1). Still, it was able to infect only three of the Octadecabacter strains with identical 16S rRNA gene sequences. These phages appeared to be strain-specific in their host recognition, even though they were able to infect bacteria across classes. Virus-host interactions are commonly simplified in ecological modeling even though they can be very complex (Middelboe 2000; Holmfeldt et al.2007; Louhi et al.2016). Our results also show that the interactions can be highly complicated, while the roles of generalist and specialist viruses in the microbial communities are not clear. Even though a virus may be capable of infecting several strains of the same genus or even strains belonging to different classes, it does not necessarily mean that the virus is a generalist. The development of generalist and specialist viruses is still not totally understood, even though it has been investigated and various theories presented, most of which are based on phage-host coevolution (Flores et al.2011; Beckett and Williams 2013; Weitz et al.2013; Koskella and Brockhurst 2014). The coevolution of a phage and its host can be explained by the Killing-the-Winner model (Thingstad and Lignell 1997) with the addition of cost of resistance (Våge, Storesund and Thingstad 2013). The defense strategy of the bacteria can display a trade-off that lowers their growth rates. Competition strategists choose not to defend, which makes them easier targets for viruses. In this study all the isolated phages were strain specific, even though PANV1, PANV2 and OANV2 were able to infect several strains. The specificity of viruses primarily arises from the interaction between the receptor molecule on the cell surface and the receptor binding protein of the virus, but further molecular research is needed to understand the concept of virus specificity. Sea-ice phage isolates have very narrow host ranges (Borriss et al.2003; Luhtanen et al.2014), but based on the genomic data, cold-active phages may have broader host ranges than mesophiles (Colangelo-Lillis and Deming 2013). Myoviruses are often considered to have broader host ranges than siphoviruses and podoviruses (Suttle 2005). However, in this study, myovirus PANV1 and siphovirus PANV2 showed similar host ranges and were able to infect only closely related hosts, whereas siphovirus OANV1 was able to infect strains from two different classes. When phage host ranges are experimentally studied, the number of cultivable bacterial isolates limits the tests, and consequently the results cannot reveal the complete host range spectrum in the environment. In addition, the tests were performed only at 4°C, and consequently virus-host interactions adapted to lower temperatures were not detected. However, our results indicate that with the strain specificity observed, the phages may be able to control the bacterial community composition, as proposed earlier based on observation in the environment (Maranger, Bird and Juniper 1994), theory (Thingstad et al.2014), and experimentation (Middelboe et al.2001). Since viruses need their hosts to replicate and produce progeny, their activity is directed to the active part of the bacterial community. Purification and characterization of phages The purification process was optimized for all phages separately. Purification analysis revealed that a significant amount of impurities and host-derived complexes were separated, allowing us to obtain a light-scattering zone comprising infectious, highly purified viruses (Fig. 2). The individual protein patterns of the isolated phages (Fig. 2) and the specific infectivities calculated (Table 4) showed that each isolated phage was different and that the purification of the virus particles was successful. Efficient purification made it possible to study individual phages in more detail. Detailed TEM observations verified that all the phages isolated were icosahedral tailed phages (Fig. 3). Sudan Black staining of the sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) gel and Triton-X treatment of the virus particles indicated that the phages do not have a structural lipid component, which is in accordance with the virus morphologies observed. The virus capsid diameters (∼50–71 nm; Table 3) are similar to those in the most abundant VLP size groups (50–70 nm or <110 nm) reported in Arctic and Antarctic sea ice (Maranger, Bird and Juniper 1994; Gowing et al.2004). In three cases, the morphology of the virus tails was reliably identified, suggesting that PANV1 is a myovirus, whereas PANV2 and OANV1 are siphoviruses (Fig. 3) resembling the dsDNA bacteriophages belonging to the order Caudovirales. However, the tail of bacteriophage OANV2 was considerably difficult to detect. We propose that OANV2 is a podophage with a short noncontractile tail. Icosahedral tailed viruses from the order Caudovirales were previously the most often isolated virus types from sea ice (Borriss et al.2003; Luhtanen et al.2014), although a filamentous virus from the order Inoviridae has also been isolated (Yu et al.2015). Temperature The temperatures used here for isolation, cultivation, and tests (Table 5) for both bacteria and phages were warmer than the temperatures in the sea-ice brine (−1.8°C down to −10.9°C; Table 1; Tison et al.2017), due to methodological limitations. It is still evident that the bacteria and phages were cold-adapted, since the bacterial strains were able to grow at 0°C, but not at 20°C (Morita 1975), and the phages were capable of infecting and producing progeny at ≤4°C (Wells and Deming 2006b; Table 5). In addition, psychrophilic bacteria from sea ice can be active even at −20°C (Junge, Eicken and Deming 2004), and cold-active phages can be productive at temperatures from 8°C to −6°C (Wells and Deming 2006b) or even at −12°C (Wells and Deming 2006c). The isolated phages retained their infectivities at cold temperatures (several months at 4°C and −80°C when supplemented with 15% glycerol), consistent with the previous cold-active virus study (Wells and Deming 2006c). Viruses have adapted to cope with a wide range of temperatures from the extreme heat in hot springs (Zablocki et al.2017) to the coldness of sea ice or permafrost (Borriss et al.2003; Wells and Deming 2006b; Luhtanen et al.2014). The major responsibility for this adaptation comes from virion stability, since virions have defined the temperature ranges at which they function (Jaenicke 1991; Bischof and He 2005). Proteins adapted to low temperatures are able to function at sea-ice temperatures, but at temperatures above their optimal range the cold-active proteins are unstable (Reed et al.2013). Still, these virus isolates tolerated the temperature of the warm top-layer agar (∼43°C) used for the plaque assay and remained infectious. Based on our results, the sea-ice virus particles can probably remain infectious in the sea water during the ice-free periods and resume activity when the sea-ice microbial community is reformed. Virus plaques were formed only at temperatures about 10 degrees lower than their host could tolerate, indicating that temperature controlled the infections, as shown in previous studies on sea-ice phage-host isolates (Borriss et al.2003; Luhtanen et al.2014) and other cold-adapted phage-host systems (Wells and Deming 2006b). Temperature may affect the host, the virus and/or their interaction. The virus-receptor molecules in the host cell may only have been induced at cold temperatures, as reported previously in Yersinia enterocolitica infections (Leon-Velarde et al.2016), indicating that the receptors may be associated with the host's cryoprotection mechanisms. Temperature also affects the adsorption of Listeria phages by adsorption inhibition and unidentified post-adsorption mechanisms (Tokman et al.2016). The structure of the phage-receptor molecules could also have changed with rising temperatures, which can inhibit the infection, or the bacterial resistance mechanisms (Labrie, Samson and Moineau 2010) could have been activated at higher temperatures. The phages isolated in this study are most probably specialized for sea-ice conditions, because they were cold-active and infected bacterial strains belonging to common sea-ice genera. The temperature inside the brine channels can vary widely, depending on the air temperature and depth of the insulating snow cover on top of the ice. Therefore, it is easy to understand why adaptations to different temperatures are needed in sea-ice communities. The isolated phages can reproduce even at seawater temperatures (4°C), implying that they would be able to survive in the water column. However, the sea-ice community is typically different from the seawater community (Bowman et al.1997; Boetius et al.2015; Eronen-Rasimus et al.2015), and the contact rates of viruses and their hosts are presumably lower in the water than in the semi-enclosed brine channels. Warming global temperatures may temporarily increase the activity of the microbial community, but also broaden the brine channels flushing the community from the ice. The ice period may also become shortened, leaving less time for the community to develop. If climate change succeeds in destroying the sea-ice habitat, we may lose a significant amount of microbial diversity on our planet. Abundance of VLPs and VBRs in Antarctic winter-sea ice The VLP abundance in Antarctic winter-sea ice ranged from ∼105 to 106 ml−1 in bulk ice (Table 1). The highest abundances were measured at stations 500 and 515 (6.9 × 105–49.0 × 105 ml−1 in bulk ice). The lower range of our dataset is comparable to the values measured earlier from Antarctic winter-sea ice (∼105 ml−1 of bulk ice; Paterson and Laybourn-Parry 2012), whereas the highest abundances were similar to those in Arctic spring blooms and Antarctic late autumn and summer sea ice (∼106–108 ml−1 of bulk ice; Maranger, Bird and Juniper 1994; Gowing et al.2002, 2004, respectively). The high VLP concentrations at stations 500 and 515 may be explained by the high bacterial abundance (Eronen-Rasimus et al.2017; Table 1) and bacterial production (measured as thymidine incorporation; Eronen-Rasimus et al.2017) observed, which were positively correlated with the high chl-a concentrations (up to 113.2 mg l−1 in bulk ice; Eronen-Rasimus et al.2017; Tison et al.2017). Positive correlation of chl-a with bacterial and VLP abundances was reported in Antarctic sea ice during spring and summer ice-algal blooms (Maranger, Bird and Juniper 1994; Gowing et al.2004). Typically, the platelet layer that forms at the bottom of the ice is the most productive layer in the sea-ice environment (Arrigo, Dieckmann and Gosselin 1995). During the winter, autotrophic production is reduced, due to the low light levels. In this study, the high chl-a concentrations observed in the middle part of the ice core at station 500 (Table 1; Tison et al.2017) were likely caused by ice rafting and flooding, trapping the autumnal bottom ice biomass between two ice floes and supplying nutrients to the uppermost ice layers (Tison et al.2017). At station 500, the algal biomass (high chl-a concentration) may have been preserved from autumn algal growth, whereas at second-year ice station 515, biomass was likely preserved from the previous spring (Tison et al.2017). Our results suggest that if the chl-a concentrations and consequent bacterial abundance and activity are high, viruses may be abundant and likely active in winter-sea ice. The high VLPs also indicate that the viral winter-sea-ice community was surprisingly dynamic, considering the season. The VBR range of 0.7–13.4 (mean 5.3) corresponds to those measured previously in Antarctic winter-sea ice (1–20.8; Paterson and Laybourn-Parry 2012). The highest VBRs (Table 1) were found at first-year ice-station 503 with low bacterial abundance and activity (Table 1; see bacterial production in Eronen-Rasimus et al.2017), while the lowest VBRs were detected at young second-year ice-station 515 (Table 1) with the highest bacterial production and abundance (Eronen-Rasimus et al.2017; Table 1). The high VBR in the low-activity community may have resulted from induction of lysogenic viruses during freezing and preservation of the virus particles in sea-ice brine, similarly to young ice in the Arctic (Collins and Deming 2011). The decreasing VBR together with increasing VLP and bacterial abundances, and bacterial activity were detected during the algal spring bloom (Maranger, Bird and Juniper 1994). Similar nonlinear variation was observed during examination of microbial cell and virus abundance estimates in 25 distinct marine surveys (Wigington et al.2016). The low VBR in the active community may have resulted from change in the bacterial community composition, so that the number of bacteria resistant to the phages may have increased. Alternatively, the host bacterial activity may have been decreased, which could have lowered the viral production, possibly because the phages may have lysogenized, i.e. become prophages (Maranger, Bird and Juniper 1994). The low VBR in the high-microbial density community was also explained by the Piggyback-the-Winner model, in which viruses favor temperateness at high host densities (Knowles et al.2016). In conclusion, four phage-host systems were isolated from both first- and second-year winter-sea ice from the Weddell Sea, Antarctica. The phages seemed to be bacterial strain-specific, but some were able to infect several related bacterial strains and one from second-year ice even across classes. The phages were able to retain their infectivity for lengthy periods under cold conditions and infected their host bacteria only in the hosts’ lower growth temperature ranges, suggesting that they are cold-active. The VLP counts suggest that the viral community may also be dynamic in winter-sea ice if their hosts are active. Overall, the virus-host interactions can be very complex. SUPPLEMENTARY DATA Supplementary data are available at FEMSEC online. ACKNOWLEDGEMENTS The authors acknowledge the Electron Microscopy Unit in the Institute of Biotechnology, HiLIFE, University of Helsinki and the Finnish Environment Institute, Marine Research Centre and Finnish Marine Research Infrastructure (FINMARI) for providing the laboratory infrastructure. We also thank the Finnish Antarctic Research Program FINNARP (especially Mika Kalakoski and Eivor Lahtinen) for logistic and financial support with cargo and travel expenses, and the Alfred Wegener Institute for Polar and Marine Research, leading researcher Peter Lemke, the captain, crew, and the other participants in the AWECS expedition. We thank Sari Korhonen for skilled technical assistance in virus production and purification, Christiane Uhlig for providing the MOX medium plates, and Harri Kuosa and Daniel Delille for useful comments on the manuscript. FUNDING This work was supported by the Walter and Andrée de Nottbeck Foundation (AML, EER, JMR), Onni Talas Foundation (AML), the Academy Professor (Academy of Finland) funding grants 283072 and 255342 (DHB) and the Belgian Science Policy (Bigsouth project, SD/CA/05). BD is a research associate at the F.R.S-FNRS. 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