Browse Subject Areas

Click through the PLOS taxonomy to find articles in your field.

For more information about PLOS Subject Areas, click here.

  • Loading metrics

Major Effect of Hydrogen Peroxide on Bacterioplankton Metabolism in the Northeast Atlantic

  • Federico Baltar ,

    Affiliation Centre for Ecology and Evolution in Microbial Model Systems (EEMiS), Linnaeus University, Kalmar, Sweden

  • Thomas Reinthaler,

    Affiliation Department of Marine Biology, University of Vienna, Vienna, Austria

  • Gerhard J. Herndl,

    Affiliations Department of Marine Biology, University of Vienna, Vienna, Austria, Department of Biological Oceanography, Royal Netherlands Institute for Sea Research (NIOZ), Den Burg, The Netherlands

  • Jarone Pinhassi

    Affiliation Centre for Ecology and Evolution in Microbial Model Systems (EEMiS), Linnaeus University, Kalmar, Sweden

Major Effect of Hydrogen Peroxide on Bacterioplankton Metabolism in the Northeast Atlantic

  • Federico Baltar, 
  • Thomas Reinthaler, 
  • Gerhard J. Herndl, 
  • Jarone Pinhassi


Reactive oxygen species such as hydrogen peroxide have the potential to alter metabolic rates of marine prokaryotes, ultimately impacting the cycling and bioavailability of nutrients and carbon. We studied the influence of H2O2 on prokaryotic heterotrophic production (PHP) and extracellular enzymatic activities (i.e., β-glucosidase [BGase], leucine aminopeptidase [LAPase] and alkaline phosphatase [APase]) in the subtropical Atlantic. With increasing concentrations of H2O2 in the range of 100–1000 nM, LAPase, APase and BGase were reduced by up to 11, 23 and 62%, respectively, in the different water layers. Incubation experiments with subsurface waters revealed a strong inhibition of all measured enzymatic activities upon H2O2 amendments in the range of 10–500 nM after 24 h. H2O2 additions also reduced prokaryotic heterotrophic production by 36–100% compared to the rapid increases in production rates occurring in the unamended controls. Our results indicate that oxidative stress caused by H2O2 affects prokaryotic growth and hydrolysis of specific components of the organic matter pool. Thus, we suggest that oxidative stress may have important consequences on marine carbon and energy fluxes.


Reactive oxygen species such as hydrogen peroxide (H2O2) are known to commonly cause oxidative stress in marine organisms (see [1] for review), and therefore, might have the potential to alter metabolic rates of marine prokaryotes, ultimately impacting the cycling and bioavailability of nutrients and organic carbon. H2O2 commonly occurs in marine and freshwater environments where it can be found in concentrations as high as 10 µM [2], although in open oceans it usually varies between <10–440 nM [3], with concentrations up to 1450 nM in Antarctic waters during autumn and winter periods [4]. Oceanic H2O2 concentrations generally decrease with depth; with concentrations up to 15 nM and 6 nM at 100 m depth and in the bathypelagic zone, respectively [5]. In the subtropical North Atlantic (our study region), H2O2 concentrations show a surface maximum of 75–220 nM, followed by a sharp decline at the shallow thermocline of ∼40 m depth [6]. In deeper waters in this region H2O2 concentrations are relatively stable, varying between 5–10 nM [6] which is in agreement with concentrations of up to 6 nM found at 150–5500 m in the subtropical North Pacific [5].

H2O2 formation is traditionally attributed to photo-oxidation of chromophoric dissolved organic matter by molecular oxygen [7], [8]. However, there is growing evidence that biological production could also be an important source of H2O2. Dark production of H2O2 has been found in unfiltered water of the coastal Mediterranean, the Red Sea and the Baltic Sea [9] and in surface waters of the Sargasso Sea [10]. Moreover, overnight production of H2O2 has been detected during in situ studies in the Atlantic [11], [12] and the Pacific [13]. Yuan and Shiller [13] suggested that biological H2O2 production might control the in situ H2O2 concentrations measured in the Northwest Pacific. The detection of H2O2 concentrations up to 6 nM at 5000 m depth was interpreted as an indication of its biological origin at depth since the short half-life of H2O2 makes it highly unlikely that photochemically produced H2O2 reaches this depth [5].

Recently, in situ H2O2 concentrations were found to be close to a steady state between dark production and decay in samples from depths of ≥10 m, suggesting that the H2O2 at those depths could be maintained primarily by particle-associated dark production and not by abiotic photochemical processes [14]. These authors suggested that extracellular H2O2 could be a common byproduct of biological processes (as is intracellular H2O2), or a product of a specific biochemical process, indicating that H2O2 production rates might vary with environmental factors and species composition. Interestingly, it was recently hypothesized that abiotic processes such as photoxidation are able to produce peroxides (i.e., hydroperoxides associated with organic matter) that might inhibit bacterial activity by singlet oxygen transfer from phytodetritus to particle-attached bacteria [15]. The oxidative stress provoked by such particle-associated production of reactive oxygen species could be particularly relevant in microhabitats and in the deep ocean due to the preferential particle-related life strategy of dark ocean prokaryotes [16], [17], [18], [19].

H2O2 can readily diffuse across cytoplasmic membranes, influencing biological processes and damaging cellular constituents [20], [21], [22]. The damage includes formation of hydroxyl radicals inside the cell that might inhibit collagen gelation, modify amino acid residues of proteins and react with cell components, thereby producing organic peroxides which can attack DNA [23], [24], [25], [26]. Lipid peroxides are formed when hydroxyl radicals interact with lipids, affecting the functioning of cell membranes, membrane-bound enzymes and other macromolecules [27]. Superoxide radicals and peroxides may also bind to DNA and alter or break the double helix structure [28].

Concentrations of ∼100 nM H2O2 influenced bacterioplankton by strongly reducing their abundance in microcosm experiments with water from the Gulf of Mexico [29], and causing oxidative stress in bacteria in Mediterranean waters [30]. In lake waters, the same H2O2 concentration (100 nM H2O2) caused a 40% reduction of the prokaryotic heterotrophic production in Lac Cromwell [22], and H2O2 concentrations of 2000–3000 nM effectively inhibited prokaryotic production in Lake Fiolen [31]. Moreover, H2O2 is an effective chemical algicide limiting cyanobacterial and green algal growth [32], [33]. If H2O2 influences bacterioplankton, then H2O2 could indirectly affect the physico-chemical characteristics of the environment (e.g., inorganic and organic nutrient quality and quantity, CO2 fixation and respiration) concomitantly affecting microbial community composition.

Despite the ubiquitous presence of reactive oxygen species such as H2O2 in the ocean and the crucial role played by marine prokaryotes in the marine biogeochemical cycles, the effect of the oxidative stress caused by reactive oxygen species on the metabolic rates of marine bacterioplankton and its ecological implications remain basically unknown. Here we studied the potential influence of H2O2 on prokaryotic heterotrophic production and extracellular enzymatic activities (β-glucosidase, leucine aminopeptidase and alkaline phosphatase) in the water column of the subtropical Atlantic. Our results suggest that H2O2 can have substantial effects on prokaryotic metabolism from the epipelagic to bathypelagic waters, which in turn might have profound implications on marine carbon cycling.


Ethics Statement

No specific permits were required for the described field studies. Sampling locations are not privately-owned or protected and sampling did not involve endangered or protected species.

Study Site and Experimental Setup

Two different sets of experiments were conducted with water collected in the subtropical northeast Atlantic Ocean during the MEDEA-I cruise with the RV Pelagia in October-November 2011 (Fig. 1). Seawater was collected from the bathypelagic in the core of the North Atlantic Deep Water (NADW, 2700–2800 m depth), the mesopelagic in the oxygen minimum layer (OML, 500–990 m depth) and the epipelagic at the base of the euphotic layer (100 m depth) using 25-L Niskin bottles mounted on a CTD (conductivity-temperature-depth) rosette sampler (Table S1).

Figure 1. Locations where hydrogen peroxide enrichment experiments were carried out during the MEDEA cruise in October-November 2011.

In the first set of experiments, 100, 500 and 1000 nM of H2O2 was added to seawater in duplicate 50 ml sterile conical tubes (Greiner Bio One). Blanks with no H2O2 addition were simultaneously done. On these samples, triplicate measurements of extracellular enzymatic activity (EEA) were performed immediately after H2O2 addition as explained below. These experiments were conducted at 9 stations (Stn. 6–16; see Fig. 1). The second set of experiments was carried out at 3 consecutive stations (Stn. 18–21; see Fig. 1) by adding different concentrations of H2O2 (50, 250, 500 nM at Stn. 18, and 10, 50, 100 nM at Stn. 20–21) to duplicate 1-L bottles containing water from the OML and 100 m depth. Amended samples were incubated together with duplicate unamended controls, in the dark at in situ temperature for 24 h. In this second set of experiments, EEAs were measured in triplicates at the beginning of the experiment (0 h) and after 24 h. Additionally, prokaryotic heterotrophic production (PHP) was measured at the start of the experiment, after 6 h and 24 h. The concentration of H2O2 stock solutions was determined daily prior to adding H2O2 to the samples using a spectrophotometer and the molar absorptivity of 38.1±1.4 M−1 cm−1 at 240 nm [34]. Measurements of in situ H2O2 concentrations were attempted but failed due to an error in the preparation of the working solutions that were brought to the cruise.

Measurements of Prokaryotic Extracellular Enzymatic Activity (EEA)

The hydrolysis of the fluorogenic substrate analogs 4-methylcoumarinyl-7-amide (MCA)-L-leucine-7-amido-4-methylcoumarin, 4-methylumbelliferyl (MUF)-phosphate and MUF-β-D-glucoside was measured to estimate the potential hydrolytic activity of leucine aminopeptidase (LAPase), alkaline phosphatase (APase) and β-glucosidase (BGase), respectively [35]. All chemicals were obtained from Sigma. The procedure was followed as described previously [19], [36]. Briefly, EEA was determined after substrate addition and incubation using a spectrofluorometer (Fluorolog-3) with a microwell plate reader (MicroMax 384, Horiba) at an excitation and emission wavelength of 365 and 445 nm, respectively. Samples (300 µl) were incubated in the dark at in situ temperature for 3–24 h. The linearity of the increase in fluorescence over time was checked on sets of samples incubated for 24 to 48 h, resulting in the same hydrolytic rates h–1. Subsamples without substrate additions served as blanks to determine the background fluorescence of the samples. Previous experiments showed insignificant abiotic hydrolysis of the substrates [37], [38], [39]. The fluorescence obtained at the beginning and the end of the incubation was corrected for the corresponding blank. This increase in fluorescence over time was transformed into hydrolysis rates using standard curves established with different concentrations of the fluorochromes MUF and MCA added to 0.2 µm filtered sample water. A final substrate concentration of 10 µmol l–1 for BGase, 100 µmol l–1 for APase and 500 µmol l–1 for LAPase was used. These concentrations have been previously determined as saturating substrate concentrations [36], i.e., resulting in maximum hydrolysis rates. Consequently, the EEAs given throughout the paper represent potential hydrolysis rates. The substrates used in this study were previously shown to be unaffected by H2O2 [40] thus, excluding the possibility of abiotic artifacts due to the added H2O2.

Prokaryotic Heterotrophic Production (PHP)

Bulk PHP was measured following the centrifugation method [41]. Triplicate 1 ml live-samples and TCA-killed blanks (5% final concentration) with 10 nM [3H]-leucine (final concentration, specific activity 140 Ci mmol−1; Perkin Elmer) were incubated in temperature-controlled incubators in the dark at in situ temperature for 3–24 h. Incubation times were consistent for EEA and PHP incubations, and similar to those used in previous studies in the same region [18], [19], [36]. Incubations were terminated by adding TCA to a final concentration of 5% and live-samples and blanks were centrifuged at 12,000 g for 10 min. After aspirating the water, 1 ml of 5% TCA was added followed by a second round of centrifugation. Subsequently, the water was aspirated again, the vials dried and scintillation cocktail (1 ml Canberra-Packard Ultima-Gold) was added. After 18 h, the samples and blanks were counted in a liquid scintillation counter (Tri-Carb 3100TR, Perkin Elmer). The mean disintegrations per minute (DPM) of the TCA-fixed blanks were subtracted from the mean DPM of the respective samples, and the resulting DPM converted into leucine incorporation rates.


Effect of H2O2 Concentration on EEA in the Northeast Atlantic Water Column

In the first set of H2O2 enrichment experiments (i.e., 100, 500 and 1000 nM), LAPase exhibited the highest rates among the EEA measured followed by APase and BGase (Wilcoxon rank sum test, p<0.0001) (Table 1 and Table S2). Of all the extracellular enzymes tested, LAPase decreased most pronouncedly with depth from the 100 m horizon to the NADW (ca. >4 fold) while APase decreased only by about 50% from 100 m depth to the NADW (Table 1). In contrast to LAPase and APase, BGase remained stable with depth. Throughout the study area, a negative effect of H2O2 on all the measured EEAs was observed in the epi-, meso- and bathypelagic waters; BGase was more inhibited by H2O2 than APase and LAPase (Wilcoxon rank sum test, p = 0.0002) (Table 1). On average, H2O2 reduced BGase rates to 40.3 to 61.9% at all depths, depending on the H2O2 concentration added. The strongest BGase reduction was found in the 500 nM H2O2 treatment at all depths. The lowest concentration of H2O2 used in this first set of experiments (i.e., 100 nM) reduced BGase rates between 20.7–50.5% (Table 1). In contrast, APase and LAPase were much less affected by H2O2. The strongest APase reduction occurred at 100 m depth, where the addition of 500 and 1000 nM H2O2 inhibited APase by 19.7 and 23%, respectively (Table 1). In deeper waters, APase was reduced by less than 12.7%. On average, LAPase was never inhibited more than 11% in all depth layers (Table 1).

Table 1. Average (±SD) response in the microbial extracellular enzymatic activities (EEAs), to different H2O2 concentrations in the experiments done with epi- meso- and bathypelagic waters at Stn. 6–16 (n = 9).

Temporal Variation of the H2O2 Impact on PHP and EEA

Since short-term impacts of H2O2 on enzyme activities were detected (Table 1), time course experiments were performed at Stn. 18, 20 and 21 to determine the temporal variations of this inhibitory effect of H2O2 on both PHP and EEA (Figs. 2, 3, 4). Since already the lowest H2O2 concentration used in the first set of experiments substantially inhibited the microbial metabolic rates, we applied lower concentrations of H2O2 in these experiments.

Figure 2. Average (±SD) temporal variation in the percentage of PHP inhibited under different H2O2 concentrations as compared to the unamended control.

PHP was estimated at 100 m (A, B, C) and the oxygen minimum layer (D, E, F) in the three experiments done at Stn. 18 (A, D), 20 (B, E) and 21 (C, F).

Figure 3. Average (±SD) temporal variation in leucine incorporation rates and extracellular enzymatic activity under different H2O2 concentrations at 100 m depth.

Leucine incorporation rates (A, D, G) were estimated at the start of the experiment, after 6 h and 24 h. The percentages of extracellular enzymatic activities inhibited under different H2O2 concentrations as compared to the unamended control were estimated at time zero (B, E, H) and after 24 h incubation (C, F, I). Experiments were performed at station 18 (A, B, C), station 20 (D, E, F), and station 21 (G, H, I). BGase: β-glucosidase, APase: alkaline phosphatase, LAPase: leucine aminopeptidase.

Figure 4. Average (±SD) temporal variation in leucine incorporation rates and extracellular enzymatic activity under different H2O2 concentrations at the oxygen minimum layer (OML).

Leucine incorporation rates (A, D, G) and the percentage of EEA inhibited under different H2O2 concentrations as compared to the unamended control were estimated at time zero (B, E, H) and after 24 h incubation (C, F, I) in experiment done at St. 18 (A, B, C), St. 20 (D, E, F), and St. 21 (G, H, I). BGase: β-glucosidase, APase: alkaline phosphatase, LAPase: leucine aminopeptidase.

PHP was reduced by 36–100% depending on the H2O2 concentration and incubation time in both the 100 m layer and OML waters (Fig. 2). This inhibitory effect on PHP augmented with increasing H2O2 concentrations and incubation time with the strongest impact on PHP at the highest H2O2 concentrations (i.e., 100–500 nM) after an incubation time of 24 h and the lowest at time zero with 10 nM H2O2. Significantly stronger (Wilcoxon rank sum test, p<0.005) inhibition of PHP rates towards higher H2O2 concentrations was found in the second (0 and 6 h incubation) and third (0, 6 and 24 h incubation) experiments in the 100 m layer, and in the first (24 h incubation), second (0 and 24 h incubation) and third (0 h incubation) experiments in the OML waters (Fig. 2). The highest H2O2 concentration of this set of experiments (i.e., 500 nM) initially reduced PHP by 70% and 97% in the OML and 100 m waters, respectively, and almost completely inhibited PHP at both depths after 24 h (Fig. 2A, D). Interestingly, PHP increased after 24 h only in the unamended treatment (i.e., where H2O2 was not added) in the 100 m waters experiments (see Fig. 3A, D, G).

In these experiments, the EEA inhibition pattern at time zero was similar to that found in the first set of experiments performed with waters collected at Stn. 6–16 (compare Fig. 3 and 4 to Table 1). Also in the second set of experiments, BGase was generally most strongly inhibited and LAPase was the least inhibited EEA. As for PHP, the inhibition of EEAs due to H2O2 was stronger after 24 h than at the initial time in all the 100 m experiments (Wilcoxon rank sum test, p<0.001) (Fig. 3). This increased H2O2-induced inhibition after 24 h was not only found for BGase but also for APase and LAPase, the latter EEAs being reduced by 20–80% (Fig. 3C, F, I). In contrast, less inhibition was found in the OML experiments after 24 h than in the 100 m layer (Wilcoxon rank sum test, p<0.005). In fact, only in one out of the three OML experiments (St. 21; Fig. 4I), the inhibition of APase and LAPase (but not BGase) was stronger after 24 h (Wilcoxon rank sum test, p<0.005) (Fig. 4).


In this study, a major inhibiting effect of H2O2 on prokaryotic metabolism as deduced from measurements of EEA and PHP was found throughout the Northeast Atlantic water column. Few other studies have investigated the effect of H2O2 on prokaryotes in aquatic environments, and to the best of our knowledge none have been done in open ocean waters. In our study region, H2O2 concentrations range between 75–220 nM in surface waters but decrease with depth to 5–10 nM below 100 m depth [6], suggesting that the relative contribution of in situ H2O2 should be very different between samples collected in the upper mixed layer and the ones from the deep ocean. Previous findings suggest that oxidative stress caused by H2O2 may become important above the 100 nM level in surface seawater [29], [30]. Our results suggest that H2O2 concentrations lower than 100 nM still significantly retard prokaryotic metabolism in the subtropical North Atlantic water column reducing EEA and PHP by about 1–85% and 38–96%, respectively (Figs. 2, 3, 4).

We also found that the detrimental impact of H2O2 on prokaryotic metabolism varies with time depending on the parameter investigated. For instance, although APase and LAPase were not strongly reduced by H2O2 initially, they were greatly inhibited after 24 h of exposure to H2O2 (Fig. 3). Recently, it was found that even short-term (∼4 h) exposure to another reactive oxygen species (singlet oxygen) has profound effects on the prokaryotic community composition in a German lake [42]. This implies that reactive oxygen species can have a dual effect on the prokaryotic community: i) directly reducing the metabolic activity of the in situ community, and ii) altering the community structure and thus, indirectly affecting metabolic rates. We cannot confirm that a shift in community composition occurred in our experiments since we did not measure it.

It is also noteworthy that H2O2 did not reduce all EEAs equally but preferentially inhibited BGase, thereby shaping the spectrum of EEAs. Although there are no previous studies investigating the effect of H2O2 on EEA rates, the inhibition observed in the EEAs reflects the potential importance of reactive oxygen species such as H2O2 in affecting the initial step in heterotrophic processing of polymeric organic matter in the ocean. This is critical since the activity of extracellular enzymes largely determines the supply of low molecular weight substrates for direct bacterial uptake [43]. The preferential inhibition of BGase over LAPase (Fig. 3) would result in an increase in the LAPase:BGase ratio. This can be interpreted as an enhanced degradation of protein over polysaccharides [44] which, in turn, ultimately affects the quality and quantity of the organic matter pool available to microbial communities.

Studies investigating the role of H2O2 in lakes report contrasting results. Studying the effect of photodegradation of humic substances on prokaryotic metabolism in a Swedish lake, Anesio et al. [31] found that H2O2 concentrations of about 2000–3000 nM were inhibitory for PHP. This range was similar to that formed during UV exposure of aquatic macrophyte leachates (4000–8000 nM H2O2), suggested to inhibit growth of prokaryotes [45]. However, those H2O2 levels are around one order of magnitude higher than reported by Xenopoulos and Bird [22] for a Canadian humic lake. These authors found that small amounts of added H2O2 (<50 nM) inhibited PHP in that lake, and that 100 nM H2O2 inhibited the PHP by as much as 40%. Although the photochemical production of low-molecular-weight substances from recalcitrant dissolved organic matter can stimulate prokaryotic activity and growth [46], [47], photochemical reactions are known to inhibit prokaryotic activities [48], [49], [50] by the formation of inhibitory substances such as H2O2 [51], [52], [53], but also by polymerization and condensation of labile compounds [54], [55] and direct mineralization of dissolved organic matter [56], [57]. Thus, these complementary effects of the photolysis of dissolved organic matter on prokaryotic activity might mask the direct effect of H2O2 on prokaryotic metabolic rates in surface waters. This could potentially explain why in those photochemical studies where H2O2 concentrations were raised by exposing samples to UV radiation, the inhibitory H2O2 concentrations were higher compared to experiments where H2O2 was directly added without exposure to UV radiation.

H2O2 concentrations have been shown to follow a diel periodicity in oceanic surface waters in our study region, ranging between ∼50–220 nM with increasing concentrations until the mid afternoon [6]. According to our data and assuming that surface water prokaryotes would behave similarly to those at 100 m depth, the range in H2O2 concentrations in surface waters could provoke a considerable PHP inhibition. Although speculative, our results could then suggest the possibility that the periodicity of PHP observed in the surface waters of the ocean (e.g. [58]), commonly interpreted as UV-induced [59], could be due, at least partly, to the accumulation of H2O2 in surface waters during daytime.

Taken together, our results suggest a potentially relevant role of oxidative stress affecting bacterioplankton metabolism, reducing prokaryotic growth and the hydrolysis of specific components of the organic matter pool. Reactive oxygen species such as H2O2 can significantly reduce the amount of organic carbon being channeled through prokaryotes, as well as alter the bioavailability of organic carbon and nutrients via the reduction of PHP and EEA rates and shifts in the EEA spectrum. This influence of oxidative stress could be potentially important throughout the entire oceanic water column where the dark biological production of H2O2 has been recently shown to be significant [14]. These effects would be even more crucial in specific (micro)environments where elevated concentrations of biologically-produced reactive oxygen species can be expected, like in particle-attached communities. The recent finding of considerable concentrations of hydroperoxides in sinking particles collected in the Arctic Ocean [15] supports this idea. Due to the assumed preferential particle-associated way of life of dark ocean prokaryotes [16], [17], [18], [19], the impact of oxidative stress on this community should be particularly significant. Taking into account the paramount role of open-ocean prokaryotes in the marine biogeochemical cycles, reactive oxygen species such as H2O2 can have relevant repercussions in marine carbon fluxes.

Supporting Information

Table S1.

Physicochemical characteristics of the stations where hydrogen peroxide enrichment experiments were carried out during the MEDEA cruise in October–November 2011.


Table S2.

Response in the microbial extracellular enzymatic activities to all the different H2O2 concentrations (0, 100, 500, 1000 nM) used in the experiments done with epi- meso- and bathypelagic waters at Stn. 6–16. LAPase: leucine aminopeptidase, APase: alkaline phosphatase, BGase: β-glucosidase.


Author Contributions

Conceived and designed the experiments: FB GH. Performed the experiments: FB TR. Analyzed the data: FB TR. Contributed reagents/materials/analysis tools: FB JP GH. Wrote the paper: FB JP GH TR.


  1. 1. Lesser MP (2006) Oxidative stress in marine environments: biochemistry and physiological ecology. Annu Rev Physiol 68: 253–278.
  2. 2. Cooper WJ, Zika RG (1983) Photochemical formation of hydrogen peroxide in surface and ground waters exposed to sunlight. Science 220: 711–712.
  3. 3. Clark CD, De Bruyn WJ, Jakubowski SD, Grant SB (2008) Hydrogen peroxide production in marine bathing waters: implications for fecal indicator bacteria mortality. Marine pollution bulletin 56: 397–401.
  4. 4. Abele D, Ferreyra GA, Schloss I (1999) H2O2 accumulation from photochemical production and atmospheric wet deposition in Antarctic coastal and off-shore waters of Potter Cove, King George Island, South Shetland Islands. Antarctic Science 11: 131–139.
  5. 5. Yuan J, Shiller AM (2004) Hydrogen peroxide in deep waters of the North Pacific Ocean. Geophysical research letters 31: L01310.
  6. 6. Obernosterer I, Ruardij P, Herndl GJ (2001) Spatial and diurnal dynamics of dissolved organic matter (DOM) fluorescence and H2O2 and the photochemical oxygen demand of surface water DOM across the subtropical Atlantic Ocean. Limnol Oceanogr 46: 632–643.
  7. 7. Petasne RG, Zika RG (1987) Fate of superoxide in coastal sea water. Nature 325: 516–518.
  8. 8. Micinski E, Ball LA, Zafiriou OC (1993) Photochemical oxygen activation: Superoxide radical detection and production rates in the eastern Caribbean. Journal of geophysical research 98: 2299–2306.
  9. 9. Herut B, Shoham-Frider E, Kress N, Fiedler U, Angel DL (1998) Hydrogen peroxide production rates in clean and polluted coastal marine waters of the Mediterranean, Red and Baltic Seas. Marine pollution bulletin 36: 994–1003.
  10. 10. Palenik B, Morel FMM (1988) Dark production of H2O2 in the Sargasso Sea. Limnol Oceanogr 33: 1606–1611.
  11. 11. Yuan J, Shiller AM (2001) The distribution of hydrogen peroxide in the southern and central Atlantic ocean. Deep Sea Res II 48: 2947–2970.
  12. 12. Avery GBJ, Cooper WJ, Kieber RJ, Willey JD (2005) Hydrogen peroxide at the Bermuda Atlantic Time Series Station: temporal variability of seawater hydrogen peroxide. Mar Chem 97: 236–244.
  13. 13. Yuan J, Shiller AM (2005) Distribution of hydrogen peroxide in the northwest Pacific Ocean. Geochemistry Geophysics Geosystems 6: Q09M02.
  14. 14. Vermilyea AW, Hansard SP, Voelker BM (2010) Dark production of hydrogen peroxide in the Gulf of Alaska. Limnology and Oceanography 55: 580–588.
  15. 15. Rontani J, Charriere B, Forest A, Heussner S, Vaultier F, et al. (2012) Intense photooxidative degradation of planktonic and bacterial lipids in sinking particles collected with sediment traps across the Canadian Beaufort Shelf (Arctic Ocean). Biogeoscience 9: 4787–4802.
  16. 16. DeLong EF, Preston CM, Mincer T, Rich V, Hallam SJ, et al. (2006) Community genomics among stratified microbial assemblages in the ocean’s interior. Science 311: 496–503.
  17. 17. Arístegui J, Gasol JM, Duarte CM, Herndl GJ (2009) Microbial Oceanography of the dark ocean’s pelagic realm. Limnology and Oceanography 54: 1501–1529.
  18. 18. Baltar F, Arístegui J, Gasol JM, Sintes E, Herndl GJ (2009) Evidence of prokaryotic metabolism on suspended particulate organic matter in the dark waters of the subtropical North Atlantic. Limnology and Oceanography 54: 182–193.
  19. 19. Baltar F, Arístegui J, Gasol JM, Sintes E, Aken HM, et al. (2010) High dissolved extracellular enzymatic activity in the deep central Atlantic Ocean. Aquatic Microbial Ecology 58: 287–302.
  20. 20. Abele-Oeschger D, Buchner T, Theede H (1994) Biochemical adaptations of Nereis diversicolor (Polychaeta) to temporarily increased hydrogen peroxide levels in intertidal sandflats. Mar Ecol Prog Ser 106: 101–110.
  21. 21. Imlay JA (2008) Cellular defenses against superoxide and hydrogen peroxide. Annual review of biochemistry 77: 755.
  22. 22. Xenopoulos MA, Bird DF (1997) Effect of acute exposure to hydrogen peroxide on the production of phytoplankton and bacterioplankton in a mesohumic lake. Photochem Photobiol 66: 471–478.
  23. 23. Fridovich I (1978) The biology of oxygen radicals. Science (New York, NY) 201: 875.
  24. 24. Dean R (1987) Free radicals, membrane damage and cell-mediated cytolysis. The British Journal of Cancer Supplement 8: 39.
  25. 25. Malins DC, Haimanot R (1991) The etiology of cancer: hydroxyl radical-induced DNA lesions in histologically normal livers offish from a population with liver tumors. Aquatic toxicology 20: 123–129.
  26. 26. Halliwell B (1995) How to characterize an antioxidant: an update. 73–101.
  27. 27. Winston GW (1991) Oxidants and antioxidants in aquatic animals. Comparative Biochemistry and Physiology Part C: Comparative Pharmacology 100: 173–176.
  28. 28. Cooke MS, Evans MD, Dizdaroglu M, Lunec J (2003) Oxidative DNA damage: mechanisms, mutation, and disease. The FASEB Journal 17: 1195–1214.
  29. 29. Weinbauer MG, Suttle CA (1999) Lysogeny and prophage induction in coastal and offshore bacterial communities. Aquat Microb Ecol 18: 217–225.
  30. 30. Angel D, Fiedler U, Eden N, Kress N, Adelung D, et al. (1999) Catalase activity in macro-and microorganisms as an indicator of biotic stress in coastal waters of the eastern Mediterranean Sea. Helgoland Marine Research 53: 209–218.
  31. 31. Anesio AM, Graneli W, Aiken GR, Kieber DJ, Mopper K (2005) Effect of humic substance photodegradation on bacterial growth and respiration in lake water. Appl Environ Microbiol 71: 6267–6275.
  32. 32. Drábková M, Admiraal W, Marsálek B (2007) Combined Exposure to Hydrogen Peroxide and Light Selective Effects on Cyanobacteria, Green Algae, and Diatoms. Environmental science & technology 41: 309–314.
  33. 33. Barrington DJ, Ghadouani A (2008) Application of hydrogen peroxide for the removal of toxic cyanobacteria and other phytoplankton from wastewater. Environmental science & technology 42: 8916–8921.
  34. 34. Miller WL, Kester DR (1988) Hydrogen peroxide measurement in seawater by (p-hydroxyphenyl) acetic acid dimerization. Analytical Chemistry 60: 2711–2715.
  35. 35. Hoppe H-G (1983) Significance of exoenzymatic activities in the ecology of brackish water: measurements by means of methylumbelliferyl-substrates. Mar Ecol Prog Ser 11: 299–308.
  36. 36. Baltar F, Arístegui J, Sintes E, van Aken HM, Gasol JM, et al. (2009) Prokaryotic extracellular enzymatic activity in relation to biomass production and respiration in the meso- and bathypelagic waters of the (sub)tropical Atlantic. Environmental Microbiology 11: 1998–2014.
  37. 37. Hoppe HG (1993) Use of fluorogenic model substrates for extracellular enzyme activity (EEA) measurement of bacteria. Handbook of methods in aquatic microbial ecology: 423–431.
  38. 38. Azúa I, Uanue M, Ayo B, Arrtolozaga I, Arrieta JM, et al. (2003) Influence of organic matter quality in the cleavage of polymers by marine bacterial communities. Journal of Plankton Research 25: 1451–1460.
  39. 39. Unanue M, Ayo B, Agis M, Slezak D, Herndl GJ, et al. (1999) Ectoenzymatic activity and uptake of monomers in marine bacterioplankton described by a biphasic kinetic model. Microb Ecol 37: 36–48.
  40. 40. Scully NM, Tranvik LJ, Cooper WJ (2003) Photochemical effects on the interaction of enzymes and dissolved organic matter in natural waters. Limnol Oceanogr 48: 1818–1824.
  41. 41. Smith DC, Azam F (1992) A simple, economical method for measuring bacterial protein synthesis rates in seawater using 3H-leucine. Mar Microb Food Webs 6: 107–114.
  42. 42. Glaeser SP, Grossart HP, Glaeser J (2010) Singlet oxygen, a neglected but important environmental factor: short-term and long-term effects on bacterioplankton composition in a humic lake. Environmental Microbiology 12: 3124–3136.
  43. 43. Chrost RJ (1991) Microbial enzymes in aquatic environments. New York: Springer Verlag. 317 p.
  44. 44. Middelboe M, Søndergaard M, Letarte Y, Borch NH (1995) Attached and free-living bacteria: production and polymer hydrolysis during a diatom bloom. Microb Ecol 29: 231–248.
  45. 45. Farjalla VF, Anesio AM, Bertilsson S, Granéli W (2001) Photochemical reactivity of aquatic macrophyte leachates; abiotic transformations and bacterial response. Aquat Microb Ecol 24: 187–195.
  46. 46. Reitner B, Herzig A, Herndl GJ (1997) Role of ultraviolet-B radiation on photochemical and microbial oxygen consumption in a humic-rich shallow lake. Limnol Oceanogr 42: 950–960.
  47. 47. Jörgensen NOG, Tranvik L, Edling H, Granéli W, Lindell M (2006) Effects of sunlight on occurrence and bacterial turnover of specific carbon and nitrogen compounds in lake water. Fems Microbiology Ecology 25: 217–227.
  48. 48. Benner R, Biddanda B (1998) Photochemical transformation of surface and deep marine dissolved organic matter: effects on bacterial growth. Limnol Oceanogr 43: 1373–1378.
  49. 49. Tranvik L, Kokalj S (1998) Decreased biodegradability of algal DOC due to interactive effects of UV radiation and humic matter. Aquat Microb Ecol 14: 301–307.
  50. 50. Tranvik LJ, Bertilsson S (2001) Contrasting effects of solar UV radiation on dissolved organic sources for bacterial growth. Ecology Letters 4: 458–463.
  51. 51. Zepp RG, Wolfe NL, Baughman G, Hollis RC (1977) Singlet oxygen in natural waters. Nature 267: 421–423.
  52. 52. Baxter RM, Carey JH (1983) Evidence for photochemical generation of superoxide ion in humic waters. Nature 306: 575–576.
  53. 53. Cooper WJ, Zika RG, Petasne RG, Fischer AM (1989) Sunlight-induced photochemistry of humic substances in natural waters: major reactive species. Adv Chem Ser 219: 333–362.
  54. 54. Harvey GR, Chesal LA, Tokar JM (1983) The structure of marine fulvic and humic acids. Mar Chem 12: 119–132.
  55. 55. Kieber RJ, Hydro LH, Seaton PJ (1997) Photooxidation of triglycerides and fatty acids in seawater: implication toward the formation of marine humic substances. Limnol Oceanogr 42: 1454–1462.
  56. 56. Valentine RL, Zepp RG (1993) Formation of carbon monoxide from the photodegradation of terrestrial dissolved organic carbon in natural waters. Environmental science & technology 27: 409–412.
  57. 57. Obernosterer I, Reitner B, Herndl GJ (1999) Contrasting effects of solar radiation on dissolved organic matter and its bioavailability to marine bacterioplankton. Limnol Oceanogr 44: 1645–1654.
  58. 58. Gasol JM, Doval MD, Pinhassi J, Calderon-Paz JI, Guixa-Boixareu N, et al. (1998) Diel variations in bacterial heterotrophic activity and growth in the northwestern Mediterranean Sea. Mar Ecol Prog Ser 164: 107–124.
  59. 59. Kaiser E, Herndl GJ (1997) Rapid recovery of marine bacterioplankton activity after inhibition by UV radiation in coastal waters. Appl Environ Microbiol 63: 4026–4031.