Factors affecting virus dynamics and microbial hostvirus interactions in marine environments


  • Kristina D.A. Mojica,

    Corresponding author
    1. Department of Biological Oceanography, Royal Netherlands Institute for Sea Research (NIOZ), Den Burg, The Netherlands
    • Correspondence: Kristina D.A. Mojica, Department of Biological Oceanography, Royal Netherlands Institute for Sea Research (NIOZ), P.O. Box 59, 1790 AB Den Burg, Texel, The Netherlands. Tel.: +31 222 369449; fax: +31 222 319674; e-mail: Kristina.mojica@nioz.nl

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  • Corina P.D. Brussaard

    1. Department of Biological Oceanography, Royal Netherlands Institute for Sea Research (NIOZ), Den Burg, The Netherlands
    2. Department of Aquatic Microbiology, Institute for Biodiversity and Ecosystem Dynamics (IBED), University of Amsterdam, Amsterdam, The Netherlands
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Marine microorganisms constitute the largest percentage of living biomass and serve as the major driving force behind nutrient and energy cycles. While viruses only comprise a small percentage of this biomass (i.e., 5%), they dominate in numerical abundance and genetic diversity. Through host infection and mortality, viruses affect microbial population dynamics, community composition, genetic evolution, and biogeochemical cycling. However, the field of marine viral ecology is currently limited by a lack of data regarding how different environmental factors regulate virus dynamics and host–virus interactions. The goal of the present minireview was to contribute to the evolution of marine viral ecology, through the assimilation of available data regarding the manner and degree to which environmental factors affect viral decay and infectivity as well as influence latent period and production. Considering the ecological importance of viruses in the marine ecosystem and the increasing pressure from anthropogenic activity and global climate change on marine systems, a synthesis of existing information provides a timely framework for future research initiatives in viral ecology.


Since the discovery of high viral abundance in marine environments in marine environments, the ecological importance of viruses to aquatic systems has become increasingly evident. Most of these viruses infect the numerically dominant microorganisms, which constitute over 90% of the ocean's biomass and serve as the major driving force behind nutrient and energy cycles (Cotner & Biddanda, 2002; Suttle, 2007; Sorensen, 2009). Aside from driving host population dynamics and horizontal gene transfer, viruses influence microbial community structure and function through the conversion of biomass to dissolved and particulate organic matter via host cell lysis (Suttle, 2007). Viral activity thus effectively regulates biodiversity and food web efficiency. The extent and efficiency to which viruses are able to drive microbial processes can be regulated by both abiotic and biotic aspects of the environment in which they occur.

At any spatiotemporal point in the ocean, viral abundance reflects the balance between rates of removal and production through host lysis. When viral progeny are released from their hosts, they are present in the environment as free virus particles and are directly exposed to environmental factors which may reduce infectivity, degrade or remove virus particles, and adversely affect adsorption to host, thereby reducing the chance of a successful host encounter and infection (Fig. 1a). Moreover, as obligate parasites, viruses are reliant upon their host to provide not only the cellular machinery but also the necessary energy and resources required for viral replication and assembly. Consequently, factors regulating the physiology of the host, as well as its production and removal, are also important in governing virus dynamics (Fig. 1b).

Figure 1.

Schematic overview of environmental factors and processes in the marine environment that have been found thus far to affect virus dynamics and virus–host interactions. (a) A synopsis of environmental factors that can lead to the removal or inactivation of virus particles reducing the chance of a successful host encounter and infection. (b) Overview of aspects that may influence the viral pool by altering host dynamics and decreasing susceptibility to infection or by modifying characteristics of viral proliferation. Heterotrophic nanoflagellates are abbreviated as HNF.

In the face of continued anthropogenic activity (marine utilization, eutrophication, urbanization, tourism, and global climate change), it will become increasingly important to unravel how environmental factors regulate virus dynamics and virus–host interactions and thus influence the role that viruses have in the marine environment. Reviews on aquatic viruses have thus far only limitedly conversed the influence of ‘the environment’ in viral ecology. It is therefore an opportune time to synthesize the current available knowledge on factors affecting host–virus interactions in the marine pelagic environment and identify any remaining gaps. The present minireview focuses on microorganism–viruses, both in culture and in the field (with emphasis on the pelagic).


Due to the dependency of viruses on a host for replication, the actual distribution of viruses can be expected to be constrained either by their own sensitivity to an environmental factor or by that of their hosts. Viruses may be more resistant to thermal stress than their host systems, indicating that the temperature distribution of the virus–host system is set by the host. Based on the available data from culture studies, the inactivation temperatures of most marine viruses fall outside of those at which host growth can be maintained (Table 1). Interestingly, the inactivation temperature of the psychrophilic filamentous phage SW1 infecting Shewanella piezotolerans showed the largest divergence from the host's optimum growth temperature. To our knowledge, this is the only marine filamentous phage tested thus far and it would be interesting to uncover if this is a general feature of this virus morphotype (Supporting Information, Table S1). Apart from the filamentous phage, phages BVW1 and GVE1 of the hydrothermal field bacteria Bacillus and Geobacillus, respectively, were inactivated at temperatures comparable to that of their thermophilic host's optimum. The fact that non-hydrothermal field viruses have lower absolute inactivation temperatures suggests an ongoing adaption to the lower optimum growth temperature of their hosts. One may then speculate that marine viruses have retained (from evolutionary origin) the genetic blueprint for thermal stability that is neutral in the current environment and may be useful for adaptation to (future) environmental increases in temperature.

Table 1. Temperature (°C) range and optimum for host growth and the range tested and values for inactivation, successful host lysis and maximum plaque forming unit (PFU) of associated viruses. When optimum temperature of the host was not reported, the culture temperature of host employed was assumed to be optimum. Parenthesis indicates that range provided is not strain-specific but obtained from the literature
HostRangeOptimumVirusGenome TypeTestedInactivationHost lysisMax PFUReferences
  1. a

    Values are the reported natural temperature range where strains are found.

  2. b

    Highest identities based on 16S analysis.

  3. =, equal efficiency.

Phaeocystis globosa Pg-I8 to 2015Group I PgVdsDNA20 to 753515 Baudoux & Brussaard (2005) and Brussaard unpublished data (host growth)
P. globosa Pg-I8 to 2015Group II PgVdsDNA20 to 752515 Baudoux & Brussaard (2005)
M. pusilla LAC384 to 2215MpRNAV-01BdsRNA20 to 95404 to 15 Brussaard et al. (2004) and Brussaard unpublished data (host lysis)
M. pusilla LAC384 to 2215MpV-03T, 06T, 08-12T, 14T, R3-4, B4-5dsDNA4 to 45404 to 15 J. Martinez-Martinez & C.P.D. Brussaard, unpublished data
M. pusilla CCMP15458 to 2420MpV-02T, 04-05T, 07T, 13T, R1-R2, SP1dsDNA4 to 45404 to 15 J. Martinez-Martinez & C.P.D. Brussaard, unpublished data
Chaetoceros debilis Ch48(9 to 30)a15CdebDNAV18ssDNA4 to 20> 2015 Tomaru et al. (2008, 2011a)
C. lorenzianus IT-Dia51(9 to 27)a15ClorDNAVssDNA4 to 20> 2015 Tomaru et al. (2011b)
C. setoensis IT07-C11(7 to 28)a15CsetDNAVssDNA4 to 20> 2015 Tomaru et al. (2013)
C. socialis (7 to 28)a15CsfrRNAVssRNA4 to 20> 2015 Tomaru et al. (2009b)
C. tenuissimus 2 to 10(9 to 30)a15CtenRNAV01ssRNA4 to 20> 2015 Shirai et al. (2008) and Tomaru et al. (2011a)
Heterosigma akashiwo H93616(5 to 30)20HaV01dsDNA4 to 20> 2015 to 30 Tomaru et al. (2005), Nagasaki & Yamaguchi (1998) and Graneli & Turner (2007) (host growth
H. akashiwo NM96(5 to 30)20HaV01dsDNA4 to 20> 2015 to 25 Nagasaki & Yamaguchi (1998)
H. akashiwo H93616(5 to 30)20HaV08dsDNA4 to 20> 2020 to 30 Nagasaki & Yamaguchi (1998)
H. akashiwo NM96(5 to 30)20HaV08dsDNA4 to 20> 2020 to 25 Nagasaki & Yamaguchi (1998)
H. akashiwo H93616(5 to 30)20HaV53dsDNA4 to 20> 2020 Tomaru et al. (2005)
H. akashiwo H93616(5 to 30)20HaRNAVssRNA4 to 20> 2020 Tomaru et al. (2005)
Heterocapsa circularisquama HU9433-P(15 to 30)20HcV03dsDNA4 to 20> 2020 Tomaru et al. (2005) and Yamaguchi et al. (1997) (host growth)
H. circularisquama HU9433-P(15 to 30)20HcV05dsDNA4 to 20> 2020 Tomaru et al. (2005)
H. circularisquama HU9433-P(15 to 30)20HcV08dsDNA4 to 20> 2020 Tomaru et al. (2005)
H. circularisquama HU9433-P(15 to 30)20HcV10dsDNA4 to 20> 2020 Tomaru et al. (2005)
H. circularisquama HU9433-P(15 to 30)20HcRNAV34ssRNA4 to 20> 2020 Tomaru et al. (2005)
H. circularisquama HCLG-1(15 to 30)20HcRNAV109ssRNA4 to 20> 2020 Tomaru et al. (2005)
Pseudomonas putrefaciens P19X2 to 272Phage 27dsDNA−5 to 26, 5555−5 to 13−5 to 2Delisle & Levin (1972a,b)
P. putrefaciens P10 2 to 272Phage 27dsDNA  2 to 202Delisle & Levin (1972a)
P. putrefaciens P132 to 2720Phage 23dsDNA2 to 26, 55552 to 2620 to 26Delisle & Levin (1972a)
P. putrefaciens P2 2 to 272Phage 25FdsDNA2 to 26, 55552 to 26=Delisle & Levin (1972a)
Pseudoalteromonas marina KCTC 12242b(2 to 25)25φRIO-1dsDNA20 to 504010 to 2520 to 25Hardies et al. (2013)
Vibrio sp. ATCC196486 to 3018unknowndsDNA6 to 30, 50506 to 25=Johnson (1968)
Vibrio (Beneckea) natriegens ATTC 14048(4 to 40)27nt-1dsDNA5 to 605027 Zachary (1976) and Farmer & Janda (2005) (host growth)
V. natriegens ATTC 14048(4 to 40)27nt-6dsDNA5 to 603727 Zachary (1976)
V. fischeri MJ-1(5 to 30)15rp-1dsDNA23 to > 454525 Levisohn et al. (1987) and Waters & Lloyd (1985) (host growth)
Pseudomonas sp.25 to 3725 to 2806N-58PssRNA5, 45 to 504525 Hidaka & Ichida (1976)
Bacillus sp. w1345 to 8568BVW1dsDNA60 to 8070> 6060Liu et al. (2006)
Geobacillus sp. E2632345 to 8565GVE1dsDNA60 to 8070> 6060Liu et al. (2006)
Colwellia psychrerythraea 34H−18 to 1810 to 18Phage 9AdsDNA−12 to 5525−6 to 4 Wells & Deming (2006b,a) and Bowman et al. (1998)
C. demingiae ACAM 459T−10 to 1810 to 18Phage 9AdsDNA  −6 to 8 Bowman et al. (1998), Wells & Deming (2006b)
21C (C. psychrerythraea)b0 to 15421cdsDNA  0 to 5 Borriss et al. (2003)
Aeromonas sp.0 to 3312UnknowndsDNA45 to 60450 to 235 to 12Wiebe & Liston (1968)
1A (Shewanella frigidimarina LMG 19867)b0 to 2141adsDNA  0 to 14 Borriss et al. (2003)
S. piezotolerans WP20 to 2815 to 20SW1ssDNA4 to 25, 60, 70,704 to 154Wang et al. (2004, 2007)

Even though marine viruses are typically more stable to temperature than their host, it does not necessarily mean that virus–host interactions within the host's growth temperature range will lead to successful viral proliferation. For example, Pseudomonas putrefaciens (P19X) can grow well up to 27 °C, but phage-27 was unable to form plaques above 20 °C (Delisle & Levin, 1972a) Furthermore, the lower temperature stability of a marine sediment phage (0–23 °C for phage vs. 0–33 °C for the host Aeromonas sp.) was due to an apparent inability to irreversible adsorp to host cells, as phage titers only demonstrated limited reduction when exposed to 30 °C for 24 h in the presence of their host bacterium (Wiebe & Liston, 1968). However, whether the inactivation was a consequence of thermal alterations to phage structure or host receptors remains unknown.

Temperature affects the structural conformation of proteins and the elasticity of biomolecules such as proteins and membrane lipids, therefore variability in the response of different viruses to modifications in temperature will most likely arise from molecular or structural differences that regulate the sensitivity of viral lipid membranes or capsid proteins to thermal deformation or thermal fracture (Selinger et al., 1991; Evilevitch et al., 2008). Group I PgVs-infecting Phaeocystis globosa were inactivated above 35 °C, while infectivity of Group II PgVs could only be maintained below 25 °C (Baudoux & Brussaard, 2005). These viruses differ in their phylogenetic origin, genome size, and in the size and composition of capsid proteins, most likely underlying the observed variation (Table S1). (Baudoux & Brussaard, 2005; Santini et al., 2013). In contrast, the larger dsDNA viruses infecting Heterocapsa circularisquama were more sensitive to losses of infectivity at over different temperatures tested compared to the smaller ssRNA virus infecting the same species (Tomaru et al., 2004, 2005; Nagasaki et al., 2005). Although very different virus types, it would be interesting to test whether the smaller size of the putative major capsid protein of the HcV03 (591 nt) as compared to HcRNAV109 (678 nt) may explain the discrepancy in the expected viral stability (Table S1; Hickey & Singer, 2004; Tomaru et al., 2009a). Although the underlying mechanisms remain unknown, variation in temperature sensitivity provides a driving force for virus and host population dynamics, and can be expected to affect the outcome of adaptation to changing environments (Bolnick et al., 2011). It is important to note that in general, unfiltered or 0.2-µm pore-size filtered water which is commonly used for investigating the stability of viruses may include components such as extracellular enzymes that can contribute to the inactivation of viruses in a temperature-dependent manner.

Temperature can also regulate infection dynamics and can vary among viruses infecting the same host as demonstrated for Heterosigma akashiwo viruses. The dsDNA virus HaV01 only infects H. akashiwo strain H93616 between 15 and 30 °C, while the comparable virus strain HaV08 is infective between 20 and 30 °C (Nagasaki & Yamaguchi, 1998). In addition, phenotypic variability can also be dependent on the host stain being infected. Heterosigma akashiwo strain H93616 was infected by HaV01 and HaV08 up to 30 °C, whereas strain NM96 (with same growth optimum temperature) was not sensitive to infection above 25 °C (Nagasaki & Yamaguchi, 1998). Similar results have also been found in a bacterium-phage system, wherein Phage 27 could successfully form plaques between 2 and 20 °C on host P. putrefaciens P10, but was restricted to 2–13 °C on host P19X (Delisle & Levin, 1972a). However, in this case, it was not due to an inability of the virus to adsorb to host cells, as temperature (0 and 26 °C) had no effect on the absorption of Phage 27 to P19X. Such virus–host co-occurring variability in temperature sensitivity, within the optimum range host growth, will enhance the temporal intraspecies diversity index.

Temperature is a major regulatory factor for microbial growth (through the regulation of enzyme kinetics, molecular diffusion, and membrane transport) and therefore can be expected to affect viral life strategy and viral production (White et al., 1991; Wiebe et al., 1992). Indeed, prophage (φHSIC ) induction correlated to seasonal variations in temperature (from 15 to 30 °C) in a eutrophic estuary was found to be a consequence of a twofold higher growth rate of the host (Listonella pelagia) at 28 °C compared to 18 °C (Cochran & Paul, 1998; Williamson et al., 2002; Williamson & Paul, 2006). Temperature-induced difference in growth rate is also the most probable cause for the delay in the onset of viral lysis of infected H. akashiwo, that is, two- to threefold delay in lysis under suboptimal temperature conditions of host (Nagasaki & Yamaguchi, 1998). Unfortunately, the authors did not sample for virus abundance, so no conclusions can be made on alterations to adsorption, latent period, or burst size. Nevertheless, this study illustrates the importance of studying different host strain and virus strain models to accurately extrapolate to natural virus–host dynamics (e.g., red-tide bloom dynamics in the case of H. akaswhiwo). In contrast, production of a filamentous phage (SW1) in the deep-sea bacterium S. piezotolerans WP3 only occurred at temperatures below the optimum of the host, producing two- to ninefold more plaques at 4 °C compared to 10 and 15 °C (Wang et al., 2007). Moreover, SW1 was shown to have a negative effect on the swarming ability of the host at low temperatures, which may provide energy for SW1 proliferation under suboptimal growth conditions of the host and therefore may play a role in adjusting the fitness of the host cells to the cold deep-sea environments (Jian et al., 2013).


Osmotic shock assays demonstrate that viral capsids have differing permeabilities to water and salt ions, which can lead to inactivation or virus particle destruction when exposed to rapid changes in ionic strength (Cordova et al., 2003). There is also evidence that suggests phage morphology plays a role in resistance. Tailed viruses appear to be the most resistant to changes in ionic strength. In addition, membrane viruses are more sensitive compared to non-lipid containing, and of the membrane containing viruses, enveloped viruses are more sensitive than those with internal membranes (Kukkaro & Bamford, 2009). Similar to temperature, the realized niche of viruses can be constrained either by their own sensitivity to variations in ionic strength or by that of their hosts. Bacteriophages and archaeoviruses isolated from a wide range of ionic strength environments have been found to be more resistant to variations in ionic strength than their host (Table 2). Moreover, marine bacteriophages appear to have specific ionic requirements to maintain structural stability and remain infective indicating an adaptation to the marine environment. Both sodium and magnesium ions were necessary for retention of viability for bacteriophages NCMB384 and 385 infecting the marine Cytophaga sp. NCMB397 (Chen et al., 1966). Keynan et al. (1974) found sodium ion concentration was the most important for the stability of hv-1 phage of the marine luminous bacterium Beneckea harveyi (maximum stability in seawater, followed by 3% NaCl; Keynan et al., 1974). Stability, infectivity, plating efficiency, and uniformity of plaque formation, but not adsorption to host, were improved by divalent ionic such as Mg2+ and Ca2+, indicating that the requirement was related to infection of phage DNA. Thus, salt stress may affect the survival and successful infection of bacteriophages, potentially through decreases in capsid pressure and consequent reductions to DNA injection efficiency or release of phage DNA though cracks in the capsid, which has been shown for the coliphage λ. Wherein the injection of DNA into host is driven by energy stored in the DNA due to its confinement, therefore any changes in ion concentrations can interact with the DNA and change the state of stress and hence the ejection force (Cordova et al., 2003; Evilevitch et al., 2008).

Table 2. Salinity (expressed as M NaCl) ranges for host growth and the range tested and values for successful infectivity and adsorption of associated viruses. Parenthesis indicates that value was assumed based on range tested for absorbance of virus to host
HostRangeVirusIons requiredTestedInfectiveMax adsorptionReferences
  1. a

    Required for stability.

  2. b

    Required for adsorption.

  3. c

    Required for lysis of host.

  4. =, equal efficiency.

Salmonella enterica 0 to 0.75PRD1 0 to 4.50< 4.250Kukkaro & Bamford (2009)
S. enterica 0 to 1.00P22 0 to 4.500 to 4.500Kukkaro & Bamford (2009)
Pseudomonas syringae 0 to 0.25φ6 0 to 4.50< 2.750.25Kukkaro & Bamford (2009)
Pseudolateromonas sp.0 to 1.75PM2 0 to 4.50< 3.250.75Kukkaro & Bamford (2009)
Halorubrum sp.2.00 to 4.50HRTV-1 0 to 4.500 to 4.50> 4.00Kukkaro & Bamford (2009)
H. hispanica 2.25 to 4.50HHTV-1 0 to 4.500 to 4.504.00Kukkaro & Bamford (2009)
H. hispanica 2.25 to 4.50HHPV-1 0 to 4.50> 1.753.50Kukkaro & Bamford (2009)
H. hispanica 2.00 to 4.50SH1Mg2+, Na+a0 to 4.500> 4.00Kukkaro & Bamford (2009) and Porter et al. (2005)
H. californiae 2.25 to 4.50HCTV-1 0 to 4.500 to 4.503.00Kukkaro & Bamford (2009)
Salicola sp.1.00 to 3.50SCTP-1 0 to 4.500 to 4.503.50Kukkaro & Bamford (2009)
Salicola sp.1.00 to 4.50SCTP-2 0 to 4.50c0 to 4.503.00Kukkaro & Bamford (2009)
Vibrio (Beneckea) natriegens 0.06 to 0.40nt-1Na+, K+b0 to 0.160 to 0.16> 0.16Zachary (1976)
V. natriegens  nt-6=b0 to 0.16≥ 0.06=Zachary (1976)
V. harveyi 0.10 to 0.60hv-1Na+, Ca2+, Mg2+a0 to 0.60> 0.085 Keynan et al. (1974)
V. fischeri MJ-1(0 to 1.03)rp-1 0 to 1.370 to 1.370.34 to 0.68Levisohn et al. (1987)
Aeromonas sp.0.085 to 0.50unknownMg2+, Ca2+c   Wiebe & Liston (1968)
Cytophaga sp. NCMB 384Mg2+, Na+a   Chen et al. (1966)
Cytophaga sp. NCMB 385Mg2+, Na+a   Chen et al. (1966)

Colwellia psychrerythraea

34 H

0.29 to 0.96Phage 9A 0.29 to 0.740.29 to 0.74 Wells & Deming (2006b)

C. demingiae


0.29 to 0.96Phage 9A 0.40 to 0.500.40 to 0.50 Wells & Deming (2006b)

There is high variability in the effect of salt concentration on the adsorption of virus to host, even up to four orders of magnitude, suggesting different mechanisms for host binding (Zachary, 1976; Torsvik & Dundas, 1980; Kukkaro & Bamford, 2009; Wigginton et al., 2012). Marine bacteriophages appear to have maximum host cell binding at salt concentrations similar to seawater, suggesting that viruses adapt to the ionic strength of their native environment. However, this could also be due to the host, as cationic imbalance, particularly deficiencies of various cations are believed to affect the permeability and other properties of cells surface structures in certain marine bacteria, which would impede attachment or penetration mechanisms of the phage (Brown, 1964). Slowest binding kinetics were found among viruses isolated from Archaea-dominated high-salt environments (Kukkaro & Bamford, 2009). The slower adsorption kinetics of archaeoviruses compared to bacteriophages might be explained by dissimilarity of surface structures of bacteria and archaeal hosts (i.e., as Archaea have lower membrane permeability; Valentine, 2007). Alternatively, halophages may have evolved to exert minimal selective pressure on their sensitive hosts (Santos et al., 2012). Prokaryotic hosts are more sensitive to changes in ionic strength, and the physiological state of cells decreases at high-salt concentrations (Kukkaro & Bamford, 2009; Bettarel et al., 2011). Therefore, slow absorption, in combination with slow decay rate, might be selected for as a mechanism to avoid decimating host population under higher ionic stress or could in turn be tied to the low-generation times of hosts, as fast adsorption and infection dynamics could decimate host populations (Bettarel et al., 2011). Indeed, over a wide range of salinities (10–360) along the coast of Senegal, the frequency of infected prokaryotes was negatively correlated with salinity, whereas a high-percentage of lysogenic prokaryotes at the higher salinities (> 150) was found to correlate to the abundance of archaeal cells (Bettarel et al., 2011).

Although salinity has been found to trigger the marine temperate phage φHSIC to switch to a lysogenic existence when incubated at brackish salinity, it is likely that this was due to a reduction of host growth rate (Williamson & Paul, 2006). However, there is also some indication that salinity can alter viral proliferation independent of host growth. A study employing an estuarine salt marsh bacterium B. natriengens reported phage-specific effects on production with alterations in salinity (Zachary, 1976). Phage nt-1 showed longer latent periods and highly reduced burst sizes (plaque forming units) at salinities below 18, while nt-6 revealed highest phage production rates at brackish salinities (which was below the host's growth optimum). The differences between these phages exemplify the potential importance of salinity on virus–host interactions and suggest a mechanism for alterations in viral population dynamics under changing salinity, both particularly meaningful for estuarine viral ecology.


Biologically harmful ultraviolet radiation (UV, 100–400 nm) can penetrate to depths exceeding 60 m in clear oceanic waters (Booth et al., 1997; Whitehead et al., 2000) and has been found to be a principal factor contributing to the decline of viral infectivity in bacteriophages, cyanophages, and viruses infecting eukaryotic hosts, with average losses of 0.2 h−1 and rates up to 0.8 h−1 in phage isolates (Table 3). Solar radiation can directly affect free viruses by degrading proteins, altering structure, and decreasing infectivity (Suttle & Chen, 1992; Wommack et al., 1996; Wilhelm et al., 1998a; Weinbauer et al., 1999). However, viral particles appear more vulnerable to inactivation than to destruction (Wommack et al., 1996; Jacquet & Bratbak, 2003). While a strong link between UV-A and loss of infectivity of marine viruses has not been found, UV-B shows a clear correlation (Table 3; Weinbauer et al., 1997; Wilhelm et al., 1998a; Jacquet & Bratbak, 2003). The shorter wavelength (290–320 nm) can result in the modification to viral proteins and the formation of photoproducts such as cyclobutane pyrimidine dimers (CPD; Kellogg & Paul, 2002; Hotze et al., 2009; Wigginton et al., 2010). As common lethal photoproducts of UV are thymine dimers, DNA viruses (containing thymine) are generally more sensitive to damage by UV than RNA viruses (not containing thymine). Furthermore, double-stranded DNA or RNA viruses are more resistant to UV than single-stranded viruses (Lytle & Sagripanti, 2005). However, these differences have yet to be demonstrated for marine viruses. Interestingly, Kellogg & Paul (2002) found a significant negative correlation between the G+C content of marine phage DNA and the degree of DNA damage induced by solar radiation. Viruses with AT-rich genomes and thus higher potential dimer (T-T) sites had a higher potential for UV damage (Kellogg & Paul, 2002). In addition, AT-rich DNA also enhances the generation of oxygen species, which cause oxidative damage (Wei et al., 1998). Repair mechanisms can reduce the lethal effect of UV, especially for viruses possessing double-stranded DNA (Lytle & Sagripanti, 2005). The dsDNA virus PBCV of Chlorella contains a DNA repair gene giving it access to two DNA repair mechanisms, that is, photoreactivation using host-encoded gene products and a virus-encoded enzyme that initiates dark repair (Furuta et al., 1997). The combined activities of these repair systems should enhance survival and maintenance of viral activity, particularly in the relatively UV-rich surface waters.

Table 3. Average decay rates (h−1) reported for losses in infectivity of virus isolates in the dark, under full sunlight, and in the absence of UV-B
HostViral isolateInfectivitySample informationReference
DarkSunlightNo UV-BLocationTime of year
  1. a

    0.2-μm filtered seawater or artificial seawater.

  2. b

    Estimated from figures in referred paper.

  3. c

    Light transmission: UV-C (200–290 nm) 3–23%, UV-B (290–320 nm) 23–26%, UV-A (320–400 nm) 26–32%, PAR (400–700 nm) 32–55%.

  4. d

    Light transmission: UV-B (290–320 nm) 67%, UV-A (320–400 nm).

  5. Numerical superscripts link data in table to the appropriate reference.

LMG1LMG1-P4 0.681, 0.2720.181Gulf of MexicoMay1, various2Suttle & Chen (1992)1,b and Suttle & Chan (1994)2
PWH3aPWH3a-P10.00a0.351, 0.242, 0.8030.081Gulf of MexicoMay1, various2, June3Suttle & Chen (1992)1,b, Suttle & Chan (1994)2 and Wilhelm et al. (1998a)3
Photobacterium leiognathi (LB1VL)LB1VL-P1b0.00a0.521, 0.2820.151Gulf of MexicoMay1, various2Suttle & Chen (1992)1,b and Suttle & Chan (1994)2
CB 38CB 38Φ0.05a0.11a York river estuaryOctoberWommack et al. (1996)c
CB 7CB 7Φ0.04a0.06a York river estuaryOctoberWommack et al. (1996)c
H2H2/10.02a0.07 Santa Monica BayMarch-JulyNoble & Fuhrman (1997)b,d
H11H11/10.02a0.07 Santa Monica BayMarch-JulyNoble & Fuhrman (1997)b,d
H40H40/10.01a0.090.06Santa Monica BayMarch–JulyNoble & Fuhrman (1997)b,d
H85H85/10.03a0.070.04Santa Monica BayMarch–JulyNoble & Fuhrman (1997)b,d
PR1PR1/10.02a0.05 Santa Monica BayMarch–JulyNoble & Fuhrman (1997)b,d
PR2PR2/10.02a0.04 Santa Monica BayMarch–JulyNoble & Fuhrman (1997)b,d
PR3PR3/10.02a0.05 Santa Monica BayMarch–JulyNoble & Fuhrman (1997)b,d
PR4PR4/10.02a0.04 Santa Monica BayMarch–JulyNoble & Fuhrman (1997)b,d
Synechococcus S-PWM1 0.19 Gulf of MexicoVariousSuttle & Chan (1994)b
Synechococcus Natural Community Syn DC2 phages 0.19 Gulf of Mexico1 yearGarza & Suttle (1998)
Synechococcus sp.Syn DC2 isolates (S-PWM1 and S-PWM3) 0.39 Gulf of Mexico1 yearGarza & Suttle (1998)b
Micromonas pusilla MpV SP10.000.30 Gulf of MexicoMarch-AprilCottrell & Suttle (1995)

While some large dsDNA algal viruses may encode for their own DNA repair enzymes, most viruses rely on repair mechanisms of their hosts, which are achievable only after DNA has been inserted into the host cell (Furuta et al., 1997; Weinbauer et al., 1997; Shaffer et al., 1999; Orgata et al., 2011; Santini et al., 2013). In the summer, Garza & Suttle (1998) found twofold lower decay rates of natural cyanophage communities as compared to isolates, whereas in the winter, they were equal, suggesting (seasonal) selection for viruses that encoding host-mediated repair mechanisms. Alternatively, this may be explained by the rapid inactivation and removal of more sensitive cyanophages. Either way, it is important to note that UV-impact studies employing viral isolates may not represent the natural community response and that viruses in surface waters may be more infective than previously thought based on the literature decay values.

CPDs in marine viruses have been found to increase over a latitudinal gradient (i.e., 41°S to 4°N), from 250 in the south to 2000 Mb−1 DNA near the equator, consistent with longer solar days and decreased solar angle. In addition, in the Gulf of Mexico, higher rates (i.e., 0.35 h−1) for the loss of infectivity were recorded in natural cyanophage communities and cyanophage isolates (S-PWM1 and S-PWM3) during the summer and early fall when solar insolation was the highest, compared to undetectable levels in winter and spring (Garza & Suttle, 1998; Wilhelm et al., 2003). Variations in level of DNA damage in waters with similar transparency and optical properties but different mixing depths have been found (Wilhelm et al., 1998b). When the mixing depth was reduced by half, photoreactivation was prevented and resulted in increased levels of CPD beyond what could be repaired overnight, leading to accumulation of damage over time (Wilhelm et al., 1998b). Similarly, Wilhelm and coworkers found high residual CDP levels in the surface water viral community off the Pacific coast of South America during an upwelling event (Wilhelm et al., 2003). Viruses were, therefore, constricted to the surface waters leading to residence times and DNA damage levels exceeding normal daily levels. There is also substantial evidence that marine viruses may adapt to local conditions of solar radiation; making them less susceptible to the degradation. Phages isolated from the coastal waters of Santa Monica Bay (USA) were 50–75% less susceptible to decay under local solar radiation than non-native phages of the North Sea (Noble & Fuhrman, 1997). In addition, phages isolated from tropical waters have higher G+C content and higher survival rates over a range of UV radiation compared to phages isolated from temperature regions (Kellogg & Paul, 2002). Similarly, the proportion of lysogens induced by sunlight was found to be lower at oceanic than at coastal stations, which may be due to higher resistance to induction in the more transparent oligotrophic open ocean, or to induction of most UV-inducible lysogens. Natural solar radiation may thus alter the viral life cycle by inducing lysogenic phage production, but does not seem to be an important source of phage production (max. 3.5%; Wilcox & Fuhrman, 1994; Jiang & Paul, 1998; Weinbauer & Suttle, 1999, 1996).

Due to the difference in susceptibility and abilities of viruses to repair the damaging effects of UV, it is not surprising that a considerable amount of variability exists in the sensitivity of viral particles to UV radiation, which has important implications for host population dynamics and species diversity (Suttle & Chen, 1992; Wommack et al., 1996; Noble & Fuhrman, 1997; Kellogg & Paul, 2002; Jacquet & Bratbak, 2003; Lytle & Sagripanti, 2005). Five large dsDNA algal viruses showed varying sensitivities to UV-B from no effect for PoV-infecting Pyramimonas orientalis to complete inactivation for PpV-infecting Phaeocystis pouchetii (Jacquet & Bratbak, 2003). Interestingly, the same study showed that some of these algal viruses, that is, of P. pouchetii and M. pusilla, had a protective effect on surviving host cells when exposed to UV-B subsequent to infection (Jacquet & Bratbak, 2003). Although the mechanisms of UV-B stress and resistance to viral infection remain largely unclear, it demonstrates the complexity of how environmental factors interact with host–virus systems.

Photosynthetic active radiation (PAR)

Light is the essential energy source for photosynthetic organisms and most often drives synchronization of phytoplankton cell division and thus DNA synthesis and mitosis. Production of the virus-infecting P. orientalis was found to depend on the host cell cycle, with a three- to eightfold increase in progeny viruses when infection occurred at the end of cell division cycle (around the onset of the light period; Thyrhaug et al., 2002). During a mesocosm study of E. huxleyi blooms, EhV abundance increased during the first part of the day (light period), suggesting that viral production was also synchronized to host cell cycle (Jacquet et al., 2002). A diel cycle-dependent cyanophage infection has been hypothesized with maximal phage production and reinfection occurring at night to explain the sharp decline in Synechococcus abundance at the onset of darkness (Suttle, 2000). However, support for this hypothesis from field observations varies (Bettarel et al., 2002; Clokie et al., 2006). Light-dependent viral infection and proliferation which triggers infection by dawn and lysis by dusk or dark would reduce exposure of the viruses to light (UV) and allow viral replication to align with its host's reproduction cycle (Clokie & Mann, 2006). Diel patterns have even been described in virally infected bacterioplankton (Winter et al., 2004), that is, viral lysis of bacteria with high viral progeny occurring around noon or early afternoon when bacterial activity was most likely responding to photosynthetic extracellular release (in combination with increased bioavailability of dissolved organic carbon by UV radiation). In addition, the newly released phages may accumulate less DNA damage (by UV) in the afternoon (Winter et al., 2004). This concept would also explain the diel variability described for a natural microbial community of NW Mediterranean Sea (Bettarel et al., 2002). One mechanism by which viruses could synchronize infection with host cell cycle is to have light-dependent absorption. The effect of light on the adsorption of nine cyanophages to Synechococcus sp. (WH7803) was found to be either light independent (S-PWM1, S-BM3, S-MM4, S-MM1, S-MM5) or light dependent (S-BnM1, S-BP3, S-PWM3, S-PM2; Jia et al., 2010). However, the adsorption rate and dependence on light were host strain-specific. Light-dependent adsorption may be due to light-induced charge neutralization at the cell surface or by light-induced alterations to the ionic composition of the host cell surfaces, which could vary according to the host (Cseke & Farkas, 1979).

In contrast, the production of PpV-infecting P. pouchetii was cell cycle-independent which was in agreement with earlier work showing that the duration of the lytic cycle of PpV was of similar duration in darkness as in light and therefore not dependent on photophosphorylation (Bratbak et al., 1998). Similarly, the latent period of algal viruses infecting Chlorella (PBCV-1, first algal virus characterized, although not marine) and H. akashiwo (one ssRNA and two uncharacterized DNA viruses) was unaffected by darkness (Van Etten et al., 1983; Juneau et al., 2003; Lawrence & Suttle, 2004). However, the viral burst size strongly decreased (50% for PBCV and 90% for PpV), implying that light-independent processes such as exploitation of host energy via chlororespiration, ATP reserves, and/or production via respiration could provide the energy needed for viral replication and host cell lysis (Juneau et al., 2003). The degree to which darkness affects viral production can also depend upon the previous light conditions experienced by the host (Baudoux & Brussaard, 2008). Viral production in P. globosa pre-adapted to a low irradiance level (25 μmol quanta m−2 s−1) was inhibited under darkness, but resumed once light was reinitiated. Conversely, mid- and high-light (100 and 250 μmol quanta m−2 s−1) pre-adapted host cells did not show additional viral production when reintroduced to the light. The burst sizes of the low- and high-light-adapted P. globosa cells were only half of the mid-light cultures indicating PgV proliferation is sensitive to shortage of energy (low light) as well as high irradiance inhibition that likely induces reactive oxygen species formation. Conversely, light level did not affect the virus growth cycle of MpV-infecting Micromonas pusilla (Baudoux & Brussaard, 2008). Yet, prolonged darkness (48–65 h) did delay host cell lysis and consequent release of virus progeny (Brown et al., 2007; Baudoux & Brussaard, 2008). It is presumed that energy and potentially reductants derived from stored metabolic intermediates were sufficient to permit viral multiplication to proceed, but at the expense of the host DNA replication (Brown et al., 2007). Late stages of infection and lysis, however, may be too energy expensive to be overcome in the dark and thus required host photosynthetic energy. We speculate that such a response to darkness is related to cell size, with small-sized picophytoplankton having insufficient reserves to complete the virus growth cycle in the dark. Darkness is an extreme condition of light limitation; in nature, phytoplankton cells are exposed periodically to dark at night, and prolonged darkness only occurs once cells sink out of the euphotic zone. However, within the euphotic zone, light conditions are far from static and algae can experience changes in light of several orders of magnitude throughout the day depending on mixing conditions and cloud cover.

These studies show that viral production of phytoplankton species may occur at low light levels and even below the photic zone, although the extent is species-specific and dependent on the growth conditions prior to viral infection. Typically, the light dependence of viral replication is characterized by a gradually shut down of host photosynthesis, with a portion of photosynthetic capacity being maintained until the end of the lytic cycle (Waters & Chan, 1982; Suttle & Chan, 1993; Juneau et al., 2003; Brown et al., 2007; Baudoux & Brussaard, 2008). In some hosts, this dependence was investigated in more detail, and key photosynthetic complexes, such as the chloroplasts, and ratios of several key photosynthetic proteins (rubisco, PSI, PSII, and ATP synthase) were maintained during the course of infection (Juneau et al., 2003; Brown et al., 2007). The importance of light in marine virus–host systems is further exemplified by the acquisition of key photosynthetic functional genes by cyanophages infecting Prochlorococcus and Synechococcus during the course of evolution (Sullivan et al., 2006). These photosynthetic genes are expressed during viral replication and aid in maintaining host photosynthesis and ensuring the provision of energy for viral replication until the onset of lysis (Lindell et al., 2005).

To our knowledge, no studies on the potential impact of the color (wavelength) of the photosynthetically active radiation (PAR, 400–700 nm) have been reported.


Lysogeny often prevails in systems with a lower trophic status, independent of geographical location (Williamson et al., 2002; Weinbauer et al., 2003; Payet & Suttle, 2013). In the deep-sea, microorganisms, experiencing a low-nutrient flux and rapidly changing conditions, have high numbers of lysogenic hosts (Weinbauer et al., 2003; Williamson et al., 2008; Anderson et al., 2011). Hence, lysogeny seems to represent a survival strategy under conditions of low host productivity and abundance, and exemplifies the crucial role that host physiology plays in determining viral life strategy. Seasonal studies and nutrient addition experiments demonstrate that viral production can be enhanced through alterations in bacterial host metabolism either by increasing host growth rate or by prophage induction (Williamson et al., 2002; Motegi & Nagata, 2007; Payet & Suttle, 2013). Likewise, P-limitation of cyanobacterium Synechococcus sp. induces lysogens, while P-addition stimulated the production of temperate cyanophage from natural P-depleted Synechococcus sp. (Wilson et al., 1996, 1998). Viral-induced lysis of host resulting from lytic infection may then act as a switch for lytic infection of prophages of the community in uninfected host which can utilize the cellular compounds released from lyzed cells. These findings illustrate the highly dynamic and responsive nature of viral life strategies to environmental factors. As the impact on host population dynamics, food web functioning, and biogeochemical cycling is very different for lysogenic or lytic viral infection, there is need for more detailed studies on this topic.

In addition to altering virus life strategies, environmental conditions that affect host physiology can also regulate the characteristics of a lytic viral infection. The latent period of marine bacteriophages typically corresponds closely with host generation time (Proctor et al., 1993; Middelboe, 2000). Similarly, viral production is often found to have a negative relationship to host growth phase, that is, lowest for host cells in stationary phase in comparison with exponentially growing cultures (Moebus, 1996; Middelboe, 2000). Interestingly, no distinct trend has been found between burst size (mostly determined by whole-cell TEM analysis) and bacterial production across different systems (Parada et al., 2006), which may be due to high bacterial host diversity under these natural conditions or selection for lysogeny under non-favorable conditions. However, in some algal host–virus model systems, burst size has been linked to host growth phase (Van Etten et al., 1983; Bratbak et al., 1998). Experiments with Chlorella virus PBCV-1 revealed that this was not due to differences in adsorption but rather enhanced viral replication in actively growing cells (Van Etten et al., 1983). Shirai et al. (2008) found that while host growth phase had no effect on the burst size of the ssRNA virus CtenRNAV01 infecting the diatom Chaeotoceros tenuissimus, viral lysis occurred earlier in the stationary-phase culture. Interestingly, Nagasaki and Yamaguchi (1998) showed that the harmful algal bloom-forming H. akashiwo was sensitive to infection by both dsDNA viruses HaV01 and HaV08 when growing exponentially, but became resistant to HaV01 when in stationary phase. While the underlying mechanism is unknown, the results indicate that the functional status of the host cell is an important determinant of virus–host interactions. Despite the potential importance of host growth as a driving factor of virus–host dynamics and subsequent organic matter cycling, there is surprisingly little attention focused on this area of research. A large proportion of the seas and oceans are oligotrophic, and phytoplankton growth is often limited by inorganic nutrient (P, N, Si, Fe) availability, increasing the potential importance of host growth as a regulatory factor of virus–host interactions. Only two studies have investigated the consequence of N-depletion in host cells on viral production and demonstrate either no effect or reduced virus yield (Bratbak et al., 1993, 1998). Alternatively, the few studies which have focused on the effect of P-depletion on algal host–virus interactions consistent show reductions in the production of viruses, that is, on average 70% for EhV-infecting E. huxleyi, 30% for PpV-infecting P. pouchetii after correction for growth phase differences, and 80% for MpV-infecting M. pusilla (Bratbak et al., 1993, 1998; Jacquet et al., 2002; Maat et al., 2014). Furthermore, the length of the latent period of M. pusilla virus MpV-08T was positively correlated to the degree of P-limitation (Maat et al., 2014). Although in theory, viral production can be depressed in P-limited cultures due to insufficient intracellular P for the production of nucleic acid-rich (and thus P-rich) viruses, it may also be caused by reduced energy availability (Clasen & Elser, 2007; Maat et al., 2014).

Wikner et al. (1993) have shown that bacterial host nucleic acids provide a major source of nucleotides for marine bacteriophages and suggest a mechanism by which marine phages limit their sensitivity to P-limitation which may be common in some open ocean areas (Paytan & McLaughlin, 2007). Interestingly, Prochlorococcus cyanophage genomes may contain the putative ribonucleotide reductase (RNR) domain, which could function as extra nucleotide-scavenging genes in P-limited environments (Sullivan et al., 2005). The highest fraction of cyanophage genomes containing host-like P-assimilation genes originate from low-P source waters (Sullivan et al., 2010; Anderson et al., 2011; Kelly et al., 2013). Moreover, host-derived pho-regulon genes, which regulate phosphate uptake and metabolism under low-phosphate conditions, are found specifically in marine phages (40% of marine vs. 4% of non-marine phage genomes (Goldsmith et al., 2011). Recently, Zeng and Chisholm (2012) showed enhanced transcription of the Prochlorococcus cyanophage-encoded alkaline phosphatase gene (phoA) and the high-affinity phosphate-binding protein gene (pstS), both of which have host orthologs, in phages infecting P-starved hosts. Such adaptations suggest that manipulation of host-PO4 uptake may be an important adaptation strategy for viral proliferation in many marine ecosystems (Monier et al., 2011). Moreover, phage genes are controlled by the host's PhoR/PhoB system, nicely illustrating the regulation of lytic phage genes by nutrient limitation of host.

Inorganic particles

Turbidity not only affects light penetration in sea but may also passively adsorb viruses. In natural waters, viruses possess a net negative surface charge due primarily to the ionization of carboxyl groups present on the external surfaces of viral capsid proteins (Wait & Sobsey, 1983). Low molecular weight peptides and amino acids have a natural binding affinity for clay minerals, with the amount being absorbed and bound dependent on the type of clay and the type of cation saturating the clay (Dashman & Stotzky, 1984). The addition of functional groups, such as amino or carboxyl groups, enhances absorption, suggesting that these molecules play an important role in the absorption kinetics (Dashman & Stotzky, 1984). Many studies have demonstrated the capacity of enteric viruses to adsorb and bind to sediment and clay particles in the marine system (Kapuscinski & Mitchell, 1980; Kimura et al., 2008). These reveal that association with particles can enhance survival and persistence of viruses by providing protection against UV radiation (most likely due to shading) and chemical pollutants relative to free viruses in seawater (Vettori et al., 2000; Templeton et al., 2005; Kimura et al., 2008). In addition, the ability of marine viruses to irreversibly or reversibly bind to clay particles depends on virus and clay type and can be affected by environmental factors such as temperature, mixing, changes in ionic strength, organic matter type, size and concentration that either enhance adsorption or induce desorption of viruses from some particles (Kapuscinski & Mitchell, 1980; Lipson & Stotzky, 1984). Hewson & Fuhrman (2003) reported that between 20% and 90% of natural marine viruses can be absorbed by mineralogically uncharacterized suspended sediments, dependent on sediment concentration, size, and source. The marine (cold-active) heterotrophic bacteriophage-9A failed to be inactivated by incubation with different ecologically relevant clay types; however, this could have been due to the high concentration of organics in the medium used, as organic matter can inhibit phage adsorption to clays, presumably by outcompeting phage for binding site (Lipson & Stotzky, 1984; Wells & Deming, 2006a). Alternatively, a marine bacteriophage under simple media conditions has been shown to serve as nuclei for iron adsorption and precipitation, presumably via the same binding mechanism as clays, that is, carboxyl and amino functional group reactive sites enabling iron atoms to penetrate and bind to the protein capsid (Daughney et al., 2004). Anthropogenic pollution has also led to the introduction of non-native particles such as black carbon to the marine environment, which have been shown to also adsorb viruses (Cattaneo et al., 2010). As very few studies have focused on the effect of inorganic particles on native marine viruses, additional research is needed to determine their ecological role, either as protective or removal agents, for viruses in marine systems.

Organic particles

In the marine environment, phytoplankton and bacteria generate large amounts of extracellular polysaccharides (EPS). In addition, intracellular substances released during viral lysis, or sloppy feeding also contribute to the organic matter pool. One important type of EPS is transparent exoploymer particles (TEP). TEP originate from colloidal DOM precursors, which may themselves bind to viruses and lead to inactivation, which might explain the discrepancy in inactivation rates in the < 0.2 μm fraction of seawater (Mitchell & Jannasch, 1969; Suttle & Chen, 1992; Noble & Fuhrman, 1997; Passow, 2002; Finiguerra et al., 2011). TEP form a chemically and heterogeneous group of particles, and their chemical composition and physical properties are dependent on the species releasing them and the prevailing environmental conditions. TEP aggregation is non-selective implying that all categories of particles present in the water become incorporated into aggregates (including viruses). In addition, TEP are primarily composed of negatively charged polysaccharides which would have a high affinity to viruses.

TEP are abundant in all marine waters (1–8000 mL−1 for the TEP > 5 μm and 3000–40 000 mL−1 for the > 2 μm fraction) and are often in the same size range as phytoplankton dramatically increasing the potential collusion frequency of particles. Suttle and Chen (1992) estimated that viruses can adsorb to microaggregates at a rate of 0.41 day−1 (averaged over the upper 10 m of the water column), which rivaled the loss due to solar radiation (0.38 day−1), indicating that oceanwide virus association with TEP could be a significant mechanism leading to the inactivation or removal of viruses from the pelagic system. Weinbauer et al. (2009) reviewed that viral abundance on suspended matter ranges between 105 and 1011 viruses cm−3 of aggregate, although it remains largely unclear how many viruses are truly attached and how many occur in the pore water of the TEP matrix and aggregates. In P. globosa mesocosms, TEP production resulting from colony disintegration and viral lysis adsorbed large quantities of the viral progeny (10–80%, also depending on whether N or P was limiting algal growth; Brussaard et al., 2005b). This could provide a mechanism by which viruses can be rapidly removed from the pelagic system after the collapse of a bloom. Conversely, reversible virus association may prolong survival and infectivity (Weinbauer et al., 2009). In addition, TEP colonized by microorganisms may continue to produce viral progeny, as viral production measurements of TEP have been shown to rival those of surrounding seawater (Proctor & Fuhrman, 1991; Wommack & Colwell, 2000; Weinbauer et al., 2009). Viral-induced cell lysis of the prokaryotic hosts releases organic matter that can further stimulate aggregation. While bacterial exoenzymes (such as aminopeptidase) and dissolved extracellular proteases and nucleases (through cell lysis) may degrade viral capsid proteins and inactivate the viruses (Simon et al., 2002; Bongiorni et al., 2007) in a similar manner to that found in seawater (although not always, Finiguerra et al., 2011) and sediment (Cliver & Herrmann, 1972; Noble & Fuhrman, 1997; Corinaldesi et al., 2010) and thereby contribute to the disintegration of TEP aggregates. The magnitude to which viruses are associated with different type, quality, size, and age of aggregates thus represents the sum of passive adsorption and the active production by the microbial community living on the aggregates (Weinbauer et al., 2009).


Removal of viruses by heterotrophic nanoflagellate (HNF) grazing seems to play only a minor role in the removal of viruses (0.1% of virus community h−1; Suttle & Chen, 1992). Similarly, Gonzalez et al. (1993) demonstrated that fluorescently labeled viruses were ingested and digested by cultured and natural HNFs at clearance rates of about 4% of those for bacterial prey, with rates depending on abundance and species of grazer and virus grazed. In contrast, Hadas et al. (2006) showed a removal of viruses by a coral reef sponge at an average efficiency of 23%, which may affect the virus-to-host ratios in the surrounding waters (depending on the removal rate of bacteria by the sponge; Hadas et al., 2006). Enteric viruses have been found to accumulate in filter-feeding shellfish (oysters, clams, and mussels), revealing the potential of these organisms to dilute ambient virus concentrations (Rao et al., 1986; Enriquez et al., 1992; Faust et al., 2009). In addition to the direct removal of virus particles, organic particles present in seawater can also be grazed (Passow, 2002). As these particles may have adsorbed viruses, the rate of viral removal by grazing might actually be underestimated.

Virus-specific selective grazing has the potential (when in high enough rates) to influence the specific virus–host dynamics and affect biodiversity. This effect can be further influenced by selective grazing on the virally infected host. Grazing of infected host cells will also alter the contact rate between virus and uninfected host cell by reducing the number of progeny viruses released from the remaining infected host cells (Ruardij et al., 2005). Preferential grazing of infected cells has been observed for E. huxleyi (Evans & Wilson, 2008), but was unconfirmed using lower, more ecologically relevant algal abundances (J. Martínez Martínez & C.P.D. Brussaard unpublished data). Preferential grazing has also been hypothesized as a response to the inhibited release of starlike structures from infected P. globosa cells (Sheik et al., 2012). These rigid chitinous filaments are thought to provide a protective benefit against grazers (Zingone et al., 1999; Dutz & Koski, 2006). In addition to increasing the susceptibility to grazers, this process directly reduces the availability of newly released PgVs by the release of hydrated flocculants, that is, the intracellular precursors of the starlike structures which exist in a fluid state within vesicles in the cell (Chretinennot-Dinet et al., 1997), which passively adsorb a high percentage of the viral progeny (c. 68%; Sheik et al., 2012). Through the use of NanoSIMS technology and single-cell investigations, it has been revealed that viral infection of P. globosa results in a leakage or excretion of 13C-labeled compounds prior to lysis which elicited an immediate response by the microbial community (Sheik et al., 2012). Leakage of intracellular material prior to lysis would provide a chemical trail which could be followed by chemotaxic grazers, thereby providing a mechanism by which preferential grazing of infected cells could occur.

Grazing may also result in the release of viral antagonists. Upon grazing of E. huxleyi cells by Oxyrrhis marina, dimethyl sulfide (DMS) and acrylic acid were released which diminished the viral titers of EhV (Evans et al., 2006, 2007). While viral lysis of E. huxleyi also led to the production of DMS and acrylic acid, the rate was reduced, which has been postulated to serve as a counter-strategy of the virus to protect the progeny infectivity (Evans et al., 2006). The same mechanism might explain the earlier results by Thyrhaug et al. (2003) who demonstrated that the viral lysate of E. huxleyi contained inhibitory compounds that delayed cell lysis. The finding that the DMS concentrations differ between E. huxleyi strains (because of diverse intracellular dimethylsulfoniopropionate (DMSP) concentrations and DMSP lyase activities; Steinke et al., 1998), clearly illustrates how quickly multiple ecologically relevant factors complicate natural virus–host dynamics and promote coexistence of host and virus. Moreover, uninfected E. huxleyi cells subjected to viral glycosphingolipids, normally produced by infected E. huxleyi to induce the release of EhV progeny, promptly executed programmed cell death (Vardi et al., 2009). It has been suggested that during blooms of E. huxleyi, production of viral glycosphingolipids may act as a strategy to limit viral propagation through clonal host populations.

Host morphology

Grazing on heterotrophic prokaryotes can also lead to alterations in bacterial phenotypes (Pernthaler, 2005). Filamentation, and the formation of microcolonies or biofilms, reduces the likelihood that a specific prokaryotic host cell encounters a phage due to partial shading (Abedon, 2012). On the other hand, if successfully infected, progeny viruses might have easy access to host in such an arrangement, decreasing encounter time (Abedon, 2012). However, the extent to which these processes influence viral encounter rate and host survival in the marine environment remains unknown, as detailed studies using marine phage-bacteria model systems are largely missing. Such information is essential as many marine bacteria are found in filaments or attached to particles (Tang et al., 2012).

It has been argued that not all host cell morphotypes of E huxleyi are found equally sensitive to viral infection, that is, exposure of E. huxleyi diploid cells to EhV would promote transition to the more infection resistant haploid phase, thereby ensuring that that genes of dominant diploid clones are passed on to the next generation in a virus-free environment (Frada et al., 2008). However, during a natural bloom of E. huxleyi, both the diploid coccolith-bearing C-cells and the haploid scale-bearing S-cells were virally infected (Brussaard et al., 1996). Another prymnesiophyte, Phaeocystis, also has a polymorphic life cycle phase which consists of solitary cells and cells embedded in a colony matrix. In this case, the haploid flagellated single cells are readily infected, whereas the colonial stage protects against viral infection (Brussaard et al., 2005a; Jacobsen et al., 2007; Rousseau et al., 2007). Model evidence shows that the probability of a virus coming in contact with an individual colonial cell decreased with the size of the colony (Murray & Jackson, 1992; Ruardij et al., 2005). P, globosa colony formation requires sufficient light for excess carbon fixation necessary to form the colonial matrix. Under reduced light conditions (20 μmol quanta m−2 s−1), only exponential growth of the flagellated single-cell morphotype was maintained and consequently viruses were able to control host abundance at low abundance and prevent bloom formation (Brussaard et al., 2005a, 2007).


The present minireview summarizes our current understanding of how environmental factors can influence virus dynamics and regulate virus–microorganism host interactions. Marine viruses affect microbial host population abundance, community structure, and biogeochemical cycling in the ocean. Identifying environment factors which regulate these processes is therefore essential to our understanding of global geochemical cycling and ecosystem functioning. This review illustrates a variety of environmental factors which can influence viruses at all stages of their life cycle (Fig. 2). Moreover, it highlights the fact that we are currently restricted by the availability of information regarding the effect that different environmental factors have on marine viruses and by the scarcity of reported rates. Factors which have been studied in more detail, for example, UV radiation, provide useful insights into how viruses have multiple strategies by which they can adapt to their environment emphasizing the need for more detailed studies. We therefore encourage further research aimed at unraveling the role that the environment plays in regulating virus dynamics and virus–host interactions and recommend using both prokaryotic (both bacterial and archaeal) and eukaryotic virus model systems from a variety of locations and depths. We would also like to stress the importance of reporting the physicochemical and biological characteristics during field studies which is crucial for optimal interpretation. Moreover, standardization of approaches is warranted to allow comparison between different studies.

Figure 2.

Conceptual diagram illustrating the influence of environmental factors on the different stages of virus life cycle. Horizontal bars indicate the amount of information known about the effect of the specific environmental variable on life cycle stage.

To fully understand the ecological relevance of environmental factors to viral ecology, the mechanisms behind losses of infectivity and absorption under natural conditions need to be elucidated. It may therefore be beneficial to consider extreme environments, where adaptations maybe more apparent due to their necessity for survival under such conditions, for example, the stability of viral proteins in and around deep-sea hydrothermal vents, resistance to extreme pH values near black smokes, and viral G+C content adaptation to UV in the surface vs. the deep ocean. In addition, some aspects which have not been addressed here, due to the scarcity of data, may have devastating consequences for marine ecosystems and should be considered in future endeavors. For example, lysogens have been found to be very sensitive to anthropogenic pollution. Pesticides, PCBs, trichloroethylene, PAHs, fuel oil, and sunscreen, have all been reported to cause substantial prophage induction, even at low concentrations (Cochran et al., 1998; Paul et al., 1999; Danovaro et al., 2003). In addition, organic UV filters originating from sunscreen triggered lytic infection in prophages of symbiotic zooxanthellae resulting in the release of large amounts of coral mucous and complete bleaching within a few days of exposure (Danovaro et al., 2008). There is also very little known about the stability of marine viruses to changes in pH, particularly for eukaryotic viruses. Although the few studies including pH sensitivity demonstrate that virus inactivation only occurred at relatively low pH values < 7, pH could be an important environmental factor in some marine systems, as pH can vary both on local and seasonal scales (Hidaka & Ichida, 1976; Borsheim, 1993; Brussaard et al., 2004; Hofmann et al., 2011; Traving et al., 2013).

Finally, it is important to recognize that changing environmental conditions are most often comprised of multiple factors, thus shifting of one factor that influences viral infectivity, production, or decay may change the sensitivity of the viruses to other factors. Therefore, it would be valuable to investigate interactions between different environmental stimuli. Especially considering that changing environmental conditions are most often comprised of multiple factors, for example, alterations in sea surface temperature may be accompanied by changes in salinity, as well as UV exposure and nutrient limitation due to alterations in stratification. The lack of mechanistic understanding strongly restrains insight and predictive capacity of how, for example, global warming-induced climate change (affecting multiple environmental variables) will influence viral production, activity, and decay. Realizing the important ecological role viruses have for biodiversity and element fluxes, we would advocate for additional focus on this particular topic.

In summary, at this moment in time, it is difficult to identify general patterns on how environmental factors regulate virus dynamics and virus–host interactions. To provide a broader overview which would permit viral ecologists to identify ecological functional patterns, virus–host systems need to be investigated in more detail, across different types of environment and/or factors. While the current review is far from exhaustive, it provides a useful framework for identifying gaps in our understanding of (1) model host/virus systems (Table S1) and (2) field-based testing which will likely lead to exciting new discoveries in marine viral ecology.


We thank Douwe Maat for his useful discussions during the initial stages of the present minireview. This work is part of the STRATIPHYT project which was supported by the division for Earth and Life Sciences Foundation (ALW), with financial aid from the Netherlands Organization for Scientific Research (NWO).