Abstract
Methanol is the predominant oxygenated volatile organic compound in the troposphere, where it can significantly influence the oxidising capacity of the atmosphere. However, we do not understand which processes control oceanic concentrations, and hence, whether the oceans are a source or a sink to the atmosphere. We report the first methanol loss rates in seawater by demonstrating that 14C-labelled methanol can be used to determine microbial uptake into particulate biomass, and oxidation to 14CO2. We have found that methanol is used predominantly as a microbial energy source, but also demonstrated its use as a carbon source. We report biological methanol oxidation rates between 2.1 and 8.4 nmol l−1 day−1 in surface seawater of the northeast Atlantic. Kinetic experiments predict a Vmax of up to 29 nmol l−1 day−1, with a high affinity Km constant of 9.3 nM in more productive coastal waters. We report surface concentrations of methanol in the western English channel of 97±8 nM (n=4) between May and June 2010, and for the wider temperate North Atlantic waters of 70±13 nM (n=6). The biological turnover time of methanol has been estimated between 7 and 33 days, although kinetic experiments suggest a 7-day turnover in more productive shelf waters. Methanol uptake rates into microbial particles significantly correlated with bacterial and phytoplankton parameters, suggesting that it could be used as a carbon source by some bacteria and possibly some mixotrophic eukaryotes. Our results provide the first methanol loss rates from seawater, which will improve the understanding of the global methanol budget.
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Introduction
Methanol is the predominant oxygenated volatile organic compound in the troposphere, and is thought to have an important role in the chemistry of the atmosphere by sequestering nitrogen oxides, and by providing HOx radicals through photolysis (Singh et al., 1995; Wennberg et al., 1998), thus influencing the oxidising capacity and ozone formation (Singh et al., 2001). However, the role of the oceans in acting as a source (Singh et al., 2000; Heikes et al., 2002) or sink (Carpenter et al., 2004; Williams et al., 2004) of methanol (and other oxygenated volatile organic compounds) to the atmosphere is uncertain. Oceanic deposition of methanol has been inferred from several modelling studies (Galbally and Kirstine, 2002; Heikes et al., 2002; Singh et al., 2003), and by Carpenter et al. (2004), who demonstrated an anti-correlation between atmospheric methanol concentrations and wind speed at Mace Head (Ireland), which suggested a net air to sea flux of methanol over the northeast Atlantic Ocean. However, air sea flux estimates are uncertain because of the analytical difficulties in reliably determining the concentration of methanol, due to its high solubility in seawater. The only published dataset with methanol measurements in seawater and air also suggest an influx of methanol to the ocean in the tropical Atlantic (Williams et al., 2004). Little is known about how methanol is produced and/or consumed in seawater. Biological production of methanol by phytoplankton cultures during exponential and stationary phase growth has been proposed (Milne et al., 1995; Heikes et al., 2002), as have bacterial sources via transformation of algal carbohydrates within the aerobic upper ocean, (Sieburth and Keller, 1989) but no measurements have been made to support these hypotheses. An early gas chromatography mass spectrometry study of water surrounding intact macroalgal samples showed that methanol, amongst others oxygenated volatile organic compounds, were present at much greater levels than ambient seawater (Nightingale, 1991). Possible oceanic loss processes include photochemical destruction and microbial uptake, although methanol is thought to be long lived in seawater with respect to photochemical oxidation (Heikes et al., 2002). Kiene (1993) suggested methanol uptake as a carbon and energy source by oceanic methylotrophic organisms, but no rate measurements have ever been reported. Methylotrophic organisms, by their definition, must synthesise all their cellular components from one carbon (C1) compounds, such as methanol, methylated sulphur species, methylated amines and so on, wherein formaldehyde is the common central metabolite (Murrell and McDonald, 2000). Most organisms that are capable of using methanol are thought to contain an enzyme called methanol dehydrogenase for oxidising methanol to formaldehyde in the cell (Smith and Murrell, 2009). From molecular studies, a diverse range of both prokaryotes and eukaryotes are thought able to use methanol for growth (Murrell and McDonald, 2000). However, there are no published measurements of methanol uptake rates in the marine environment.
The aim of this study was to make the first ever measurements of the biological consumption of methanol in marine waters, and to determine what proportion of the uptake was attributable to either cell growth or energy use.
Materials and methods
We have modified a radiochemical technique that uses the addition of 14C-labelled methanol (14CH3OH) to seawater to determine the microbial conversion of methanol into particulate cell biomass, and oxidation of methanol to carbon dioxide (used as an energy source).
Sample collection
Surface water samples (⩽10 m) for time course, kinetic and addition experiments (Table 1) were collected from a long term monitoring station, situated approximately 10 nautical miles south–west of Plymouth, called L4 (Figure 1: 50.25 N, 04.22 W, water depth ∼55 m). Samples were pumped directly into acid-washed quartz Duran bottles and stored in the dark for the 2–3 h transit back to the laboratory. Surface seawater samples were also collected from the northeast Atlantic (Figure 1) during a research cruise carried out as part of the UK Surface Ocean Lower Atmosphere Study aboard the Royal Research Ship Discovery (D320, 16 June–17 July 2007). Samples were either collected using 20 l Niskin bottles (General Oceanics, Miami, FL, USA) deployed on a rosette equipped with a Seabird (Sea-Bird Electronics, Inc., Bellevue, WA, USA) conductivity, temperature and depth sensors, or were pumped while underway from an approximate depth of 5 m using a peristaltic pump. The seawater was collected directly into acid washed polycarbonate bottles and experiments were started typically within 1 h of sampling. The labelled 14C methanol was purchased from American Radiolabeled Chemicals Inc. (Saint Louis, MO, USA) with a specific activity of 57.1 mCi mmol−1. Primary stocks were diluted 1:10 with 18 MΩ milli Q water (Millipore, Billerica, MA, USA) (0.1 mCi ml−1) and were stored in gas tight vials in the dark at 4 °C. Stability and storage trials suggest a loss in activity of <10% over 12 months. Addition volumes of 14C methanol to seawater samples were always <1% of the sample volume and typically ⩽12% of the label was used during incubations ⩽24 h.
Linear time course experiments
Seawater samples were treated with between 10 and 100 nM 14CH3OH final added concentration (1494–13 434 disintegrations per minute per ml) depending on the experiment (Table 1). Samples were incubated in acid washed polycarbonate bottles in the dark for between 0.1 and 120 h at in situ temperature. At selected times, triplicate sub-samples to assess uptake into particulate cell biomass and total oxidation to 14CO2 were taken. For uptake into particles during L4 experiments, 1.7 ml samples were placed in 2 ml screw top sterile polypropylene micro centrifuge tubes and fixed with 100 μl trichloroacetic acid (TCA, 5% final concentration). Samples were centrifuged (16 000 × g) for 10 min, aspirated and washed with 80% ethanol. This process was repeated twice, and after the third aspiration, liquid scintillation fluid (Optiphase HiSafe III; Perkin-Elmer, Buckinghamshire, UK) was added and the sample mixed and placed in the dark overnight to allow chemiluminiscence to subside before counting on a liquid scintillation counter (Tri-carb 3100, Perkin-Elmer). This approach is similar to the leucine incorporation method modified for microcentrifuge tubes (Smith and Azam, 1992).
During the UK Surface Ocean Lower Atmosphere Study research cruise, larger sampling volumes between 50 and 100 ml were used for methanol uptake into particles to increase the sample counts (Table 1). Seawater samples were filtered onto 0.2 μm Supor (Pall Corporation, Portsmouth, Hampshire, UK) (polyethersulfone) filter membranes with a vacuum of <10 cm Hg. The filters were rinsed three times with 2–3 ml of 80% ethanol, placed in a 4 ml scintillation vial and dried overnight in a desiccator, before the addition of scintillation fluid. Radioactivity retained on the filters was measured as disintegrations per minute (DPM) onboard ship using a liquid scintillation counter as above. Total oxidation of 14CH3OH to 14CO2 was determined in triplicate by pipetting 1 ml samples into 2 ml microcentrifuge tubes and adding 0.5 ml of SrCl2.6H2O (1 M), to precipitate the 14CO2 as Sr14CO3, 20 μl of NaOH (1 M), to neutralise the HCl produced, and 100 μl of Na2CO3 (1 M), to ensure adequate pellet formation (Connell et al., 1997; Goodwin et al., 1998). After centrifugation the supernatant was aspirated, the pellet washed twice with ethanol (80%), resuspended in 1 ml of concentrated NaOH solution (∼10 nM) that had been adjusted to a pH of 11.7, before addition of Optiphase HiSafe III to create a slurry. The samples were vortex mixed and stored in the dark for >24 h before being analysed on the scintillation counter. This period ensures that any chemiluminescence arising from interactions between the NaOH added and the Optiphase scintillant subside (Kiene and Hoffmann Williams, 1998).
Kinetic determinations
Microbial methanol kinetics were investigated at L4 using 1.7 and 1.0 ml seawater samples to determine uptake into particulate biomass and total oxidation, respectively. Individual samples received an addition of 14CH3OH, and a series of tubes for both uptake into particles and total oxidation were treated to yield a range of added 14CH3OH between 5 and 100 nM. Samples were incubated in screw topped, O-ring sealed microtubes in the dark at in situ temperature (Table 1). Triplicate replicates from each methanol concentration were processed, as detailed above, after incubation periods of 3 and 24 h.
The kinetics of methanol uptake into the particulate phase only was determined at stations 2 and 3 in the northeast Atlantic, where 50 ml sample aliquots were incubated in the dark at in situ temperature, after additions of 14CH3OH between 1 and 250 nM, with more emphasis on the lower range (1–50 nM). Incubations were terminated after approximately 1 h at station 2, and 24 h at station 3 by gentle vacuum filtration onto Supor filters. These filters have been reported to give the best retention of particulate material >0.2 μm, compared with Nuclepore or Anopore filters (Whatman plc, Maidstone, Kent, UK), with minimal blanks (Kiene and Hoffmann Williams, 1998). Vmax and Km (Michaelis constant) were calculated using the Lineweaver–Burk transformation.
Substrate addition and inhibitor experiments
Addition experiments
The effects of adding trimethylamine (TMA; 10–100 μM) and dimethylsulphoxide (10 and 100 μM) on rates of methanol oxidation and uptake into particles were assessed on surface seawater samples from the coastal station L4. Triplicate samples for methanol oxidation (1.0 ml sample volume) and uptake into particulate cellular biomass (1.7 ml sample volume), were incubated in the dark, at in situ temperature, over a time series varying between 1 and 120 h. All the additions were made immediately before adding the 14C-labelled methanol.
Cycloheximide and chloroamphenicol inhibitor experiments
The extent to which prokaryotes (assumed to be mainly bacteria) and eukaryotes (phytoplankton) use methanol as an energy and carbon source was investigated on surface seawater samples collected during a dinoflagellate bloom of Prorocentrum minimum at the coastal station L4. Chloroamphenicol (20 μg ml−1) and cycloheximide (483 μg ml−1) were used to inhibit prokaryotic and eukaryotic protein synthesis, respectively (Goodwin et al., 1998). Methanol oxidation and uptake into particulate cellular biomass was measured in triplicate 1 ml and 1.7 ml samples, respectively, and incubations terminated after 24 h. Seawater samples were processed as described previously.
Surface uptake in the northeast Atlantic
At a series of stations in the northeast Atlantic, rates of methanol uptake into particulate cellular biomass were determined using 50 ml seawater samples collected from the surface (<10 m; Table 1) and processed as detailed previously. The data is reported as apparent rate constants, k (day−1) calculated from a ratio of the 14C counts collected in either the particulate phase or as 14CO2 (nCi ml−1 day−1) divided by total 14CH3OH added to the sample (nCi ml−1). All rate constants were corrected by subtracting killed sample (TCA, 5% final concentration) counts to correct for passive surface sorption. Rates of bacterial protein production and the numbers of heterotrophic bacteria, Prochlorococcus spp, Synechococcus spp and picoeukaryotes were also determined to investigate any trends. Bacterial protein production was determined by measuring the incorporation of 3H-leucine (20 nM final concentration) into bacterial protein on 1.7 ml seawater samples following the method of Smith and Azam (1992). The numbers of bacterioplankton cells were determined by flow cytometry on SYBR green I DNA-stained cells from 1.8 ml seawater samples fixed in paraformaldehyde (5%, final concentration), flash frozen in liquid nitrogen immediately after fixation, and stored frozen at −80 °C (Marie et al., 1997). Numbers of Prochlorococcus spp, Synechococcus spp and picoeukaryotes were analysed on unstained samples by flow cytometry (Zubkov et al., 2000). Surface concentrations of chlorophyll a were estimated using a composite Aqua-MODIS chlorophyll a remotely sensed image for the cruise duration (16 June–16 July 2007), as unfortunately the continuous surface fluorometer onboard the ship was not working correctly. Chlorophyll a samples from L4 were determined by fluorometric analysis of acetone-extracted pigments (Holm-Hansen et al., 1965).
Results
Linear time course experiments
When nanomolar concentrations of 14CH3OH were added to coastal seawater samples from L4, and oceanic samples from the northeast Atlantic, radioactive carbon was incorporated into particulate cellular material and expired to 14CO2 (Figure 2), and thus used as a carbon and an energy source.
At the coastal station L4, methanol oxidation was linear for up to 48 h after the addition of 10 nM 14CH3OH (only data up to 6 h incubation is shown in Figure 2a). The rate of methanol oxidised to 14CO2 was 0.01 nCi ml−1 h−1 (R2=0.999, n=9), after which 33% of the added substrate had been oxidised after 48 h incubation. After the addition of 100 nM 14CH3OH, methanol oxidation was linear for up to 5 days at an increased rate of 0.02 nCi ml−1 h−1 (R2=0.999, n=10), after which 20% of the added label had been oxidised.
Methanol uptake into particles after the addition of 10 nM 14CH3OH was also linear up to 48 h, but with a lower rate of 0.005 nCi ml−1 h−1 (R2=0.997, n=6), after which 15% of the added substrate had been used. However, after the addition of 100 nM 14CH3OH, there was no increase in added label counted in the particulate phase over time. Approximately 0.2% of the added substrate was associated with particles >0.2 μm after only 10 min incubation, which could either suggest very rapid uptake and saturation, or substrate contamination. Evaluation of control samples (killed with 100% TCA final concentration) suggest that up to 0.3% of the added 14C-labelled methanol (after addition of 100 nM) is recovered on the filters, which is attributed to either nonspecific adsorption to surfaces (including the filter), or stock contamination.
Methanol oxidation an the shelf station (station 1, Figure 1) was linear for ∼18 h at a rate of 0.007 nCi ml−1 h−1 (R2=0.996, n=6) after which 8% of the added label (10 nM final concentration) had been used, which increased to 12% after 24 h (Figure 2b). Uptake into particles (>0.2 μm) was linear for the same time period at a lower rate of 0.002 nCi ml−1 h−1 (R2=0.989, n=6). After 18 h 2.3% of the added label was counted in the particles, which increased to 3.5% after 24 h.
At the more oceanic, low chlorophyll a stations (station 2 and 3, Table 1), rates of incorporation of added label into particles were very low. At station 2 only ∼0.4% of the added label was found in the particles after 26 h. At station 3, uptake into particles was linear for up to 12 h at a very low rate of 0.001 nCi ml−1 h−1 (R2=0.994 n=5), after which only 0.07% of the added label (100 nM final concentration) had been used.
Rates of methanol oxidation in nmol l−1 day−1 were calculated from the product of the apparent rate constant, k × C, where C=concentration (nM) of 14CH3OH (Connell et al., 1997), and are summarised in Table 2. The in situ concentrations of methanol are not known, however, we have recently determined that the surface concentration of methanol at L4 was 97±8 nM (n=4) between May and June 2010, and for the wider temperate north Atlantic waters was 70±13 nM (n=6). Analysis was performed using a membrane-inlet system coupled to detection by proton-transfer mass spectrometry. These concentrations are within the range quoted by Williams et al. (2004), for surface waters of 118±48 nmol l−1 for the tropical north Atlantic. This remains the only published seawater methanol dataset. We have therefore estimated methanol turnover times of 12–23 day at the coastal station, 24 day at the shelf break and between 11 and 33 day at a lower productivity offshore oceanic site (station 7a) in the northeast Atlantic (Figure 1).
Uptake kinetics
The microbial uptake of 14CH3OH typically displayed non-saturation type kinetics for rates of methanol oxidation, and uptake onto particles for additions of methanol up to 250 nmol l−1. However, when the data was plotted using the Lineweaver–Burk transformation (double reciprocal plot), linear relationships were found to explain between 74 and 99% of the variance in the data (Figure 3), enabling the first estimates of Vmax and Km to be determined (Table 3). Total microbial methanol oxidation Vmax of between 1.0 and 1.2 nmol l−1 h−1 were found for the coastal station L4, with Km between 9.3 and 86 nmol l−1. Methanol uptake into particles at L4 showed a lower range for Vmax between 0.20 and 0.26 nmol l−1 h−1, with Km values between 6.9 and 43 nmol l−1.
The range in Vmax found for methanol uptake into particles for stations 2 and 3 in the northeast Atlantic was approximately three times lower compared with the coastal station ranging between 0.06 and 0.07 nmol l−1 h−1. The Km value at station 3 was notably high at 974 nmol l−1, suggesting that the microbial population at this station had a much lower affinity for incorporating methanol into cellular biomass.
Addition and inhibitor experiments
The addition of TMA (10–100 μM) to surface coastal seawater samples (L4) did not significantly change the rate of uptake of 14C-labelled methanol into particles (0.006±0.001 nCi ml−1 h−1, Figure 4a) during the first 24 h of the incubation. Thereafter, the addition of TMA increased uptake rates, with a maximum of 0.01 nCi ml−1 h−1 found after the addition of 50 μM. In contrast, the addition of TMA did slightly increase the rate of oxidation of 14CH3OH to 14CO2 during the first 24 h (Figure 4b) from 0.03 nCi ml−1 h−1 (no addition) to 0.04±0.003 nCi ml−1 h−1. The maximum rate of methanol oxidation (with TMA) was 0.09±0.004 nCi ml−1 h−1 after addition of 50–100 μM TMA (between 1 and 5 days incubation), which suggests that the addition of TMA increased methanol oxidation approximately threefold compared with samples without TMA. Methanol uptake rates (into particles or oxidation) were not measurable when samples were killed with TCA (5% final concentration).
The addition of dimethylsulphoxide resulted in rates of methanol uptake into particles and its oxidation of 0.004±0.0004 and 0.03±0.005 nCi ml−1 h−1, respectively, (average of 10 and 100 μM additions). These rates were sustained for up to 5 days compared with unamended samples, whose utilisation rates decreased after 48 h incubation. The addition of prokaryotic (chloroamphenicol) and eukaryotic (cycloheximide) inhibitors reduced the uptake of methanol into biomass by >95% and its oxidation by between 67 and 83% (Figure 5).
Latitudinal trends in the northeast Atlantic
The uptake rates of 14CH3OH into particulate biomass from surface seawater samples, (shown as k) were plotted with corresponding rates of bacterial protein synthesis according to the latitude of sampling location (Figure 6a). Numbers of heterotrophic bacteria, picoeukaryotes (Figure 6b), Synechococcus and Prochlorococcus cells are also shown (Figure 6c). Surface constants for methanol uptake into particles were typically <0.002 day−1 at the low chlorophyll a (<0.2 μg l−1, Table 1) stations located ∼43°N. There was an exception to this at station 4 (43.7 °N, Table 1), where an anomalously higher methanol uptake rate constant of 0.012 day−1 was coincident with an unusually high bacterial protein synthesis of 531 pM Leu day−1, compared with an average of 137 pM Leu day−1 in this area.
Methanol uptake rates into particles and bacterial leucine incorporation generally increased in a northerly direction up 0.017 day−1 and 684 pM Leu day−1, respectively, at 54°N. Surface concentrations of chlorophyll a also increased northerly from <0.2 μg l−1 at 43 °N to 0.7–1.0 μg l−1 at 54 °N (Table 1). Linear regression analysis shows positive correlations between methanol uptake into particles and numbers of heterotrophic bacteria (r=0.8796, n=10, P⩽0.001), picoeukaryotes (r=0.8398, n=10, P⩽0.01), and bacterial protein production (r=0.7153, n=11, P⩽0.02). Although it is acknowledged that the numbers of heterotrophic bacteria, as determined by flow cytometry, could potentially include members of the Archaea, some of which could be autotropic (Meng et al., 2009). The numbers of Synechococcus or Prochlorococcus cells showed no statistically significant relationships with uptake of methanol into particles. This could suggest that they are not using methanol as a carbon substrate, although this is not a definitive conclusion as their cellular activities were not determined, and total numbers could include live and dead cells.
Discussion
This study is the first to show that radiolabelled methanol (14CH3OH) can be successfully used to determine microbial uptake into biomass and oxidation in seawater samples. We have found that methanol is used predominantly as a microbial energy source, but also demonstrated that it is used as a carbon source.
For the first time, we report biological methanol oxidation rates between 2.1–8.4 nM day−1 in surface seawater of the northeast Atlantic, although kinetic experiments predict a Vmax of up to 29 nmol l−1 day−1 in more productive coastal waters. The lower range in Km values found for methanol uptake in particles compared with its oxidation rates suggests that the microbial enzymes used in converting methanol into cellular carbon (14C-macromolecular incorporation) have a slightly higher affinity for the substrate, compared with the enzyme systems involved in using methanol as an energy source, and its ultimate respiration to 14CO2. It is useful to distinguish between total microbial use of methanol, and the uptake into particulate fractions so that we can estimate how significant methanol is as a carbon source for cell growth, in addition to its use as an energy source. Methlyotrophic bacteria must use C1 compounds more reduced than CO2 as sole energy sources, and to assimilate carbon into cell biomass (Murrell and McDonald, 2000). Both pathways initially oxidise methanol to formaldehyde in the cell using the enzyme methanol dehydrogenase. Thereafter, incorporation into cell material is thought to proceed either via the RuMP or serine cycle into 3-carbon compounds and ultimately to cell material (Murrell and McDonald, 2000). If methanol is used as a source of energy, then formaldehyde is further oxidised in the cell to formic acid with the enzyme formaldehyde dehydrogenase, before being respired to CO2 using the enzyme formate dehydrogenase. Our kinetic experiments determine the amount of 14C retained in the particulate phase and the amount respired to 14CO2, which we assume is representative of cellular incorporation (cell growth) and energy use respectively. However, both end processes measured will have inevitably been the resultant actions of more than one enzyme.
The calculated Vmax rates and Km constants derived from Figure 3 assume saturation of a single enzyme (Michaelis–Menten kinetics), which unfortunately was not confirmed by our additions of 14C-labelled methanol up to 250 nmol l−1. These non-saturation kinetics could imply that there is no active enzyme transport system for methanol. However, when samples were killed with TCA (5% final concentration), methanol uptake did not increase over time, which suggests that despite a possible lack of an active transport system, the uptake was nevertheless because of microbial metabolic activity. Jennings (1963) points out that if the absorption of a solute, for example, methanol is the result of metabolic activity, the rate of adsorption will inevitably be describable by Michaelis–Menten kinetics, even though no specific transport system is involved, lending validity to our derived Vmax and Km results. Furthermore, it is highly likely that methanol uptake by natural microbial communities in seawater results not only from bacteria, but also from mixotrophic and heterotrophic phytoplankton. This is evident from the multiphasic uptake kinetics particularly shown by methanol uptake onto particles for L4 and stations 2 and 3 (Figure 7). Wright and Hobbie (1966) suggested that at very low concentrations of substrate, most glucose was incorporated because of the active transport systems in bacteria, whereas at higher concentrations diffusion across algal cells dominated. Therefore, our methanol uptake onto particles (Figure 7) could suggest that bacterial transport processes dominated for added concentrations ⩽20–30 nmol l−1, whereas at higher concentrations most of the incorporation was because of diffusion.
For rates of microbial methanol oxidation in nearshore coastal waters (L4), which increased linearly with substrate concentration (y=0.006 × +0.03, n=5, R2=0.99 for 24 h incubations, data not shown in this format) a diffusion constants (Kd) can be calculated from the slope of the linear relationship (Wright and Hobbie, 1965). This constant assumes that the organisms are incorporating the methanol as rapidly as it diffuses in (Wright and Hobbie, 1965). A Kd of 0.006 h−1 is equivalent to a turnover time of 7 days (Wright and Hobbie, 1965). This estimated turnover time is not dependant on an assumed in situ surface seawater concentration of methanol, and suggests a faster turnover compared with the previous estimates of 12–23 days (as in Table 2 calculations). It is also closer to the ∼3 day estimate, which was based on the turnover of other compounds, such dimethylsulphide from the literature (Heikes et al., 2002). The seawater sampled for the kinetic experiments contained high numbers of Prorocentrum minimum, who are mixotrophic species (Stoecker, 1999), so could be contributing via methanol diffusing across their surface membranes, and thus contributing to a faster turnover time than predicted from Table 2. These phytoplankton could also be consuming bacterioplankton, who have previously incorporated methanol (Zubkov and Tarran, 2008).
The kinetic characteristics of microbial methanol utilisation can be compared with those of other substrates commonly used by bacteria, so that the ecological significance of methanol to marine microbial metabolism can start to be evaluated (Table 3). Proteolytic enzyme Vmax are generally significantly greater than those of methanol, and have correspondingly higher Km concentrations, suggesting much lower enzyme substrate affinity for proteins compared with methanol. This is perhaps to be expected considering that this enzyme activity reflects the hydrolysis of a broad range of proteins and peptides, whose concentrations in seawater are thought likely to exceed those of methanol by at least one order of magnitude. The range found in Vmax for the hydrolysis of carbohydrates associated with particles is comparable with those of methanol uptake into particles in surface waters. The range in Km constants for cellulose in particular, is higher than that calculated for methanol in more coastal waters, but is more similar to methanol Km constants in particles sampled from more remote Atlantic surface waters.
The addition of TMA, as an example of a methylated amine compound, increased both methanol uptake into particles and oxidation at the coastal L4 station. This suggests that TMA induced methylotrophic pathways, indicating that the bacteria utilising methanol (presumably via methanol dehydrogenase) were also capable of incorporating methylated amines, and thus were likely to contain methylated amine dehydrogenases and methylamine oxidase (Murrell and McDonald, 2000). Interestingly, the predominance of Methylophaga spp in the consumption of both methanol and methylamine at L4 has been reported by Neufeld et al. (2007). The addition of dimethylsulphoxide, as an example of a methylated sulphur compound, suggested that sulphur utilising methylotrophs in surface seawater at L4 also incorporated methanol as a carbon and energy source, and can perhaps outcompete non-sulphur utilising methlyotrophs over longer incubation times, when presumably nutrients deplete. Some methylotrophic bacteria, such as Hyphomicrobium, are known to be capable of using compounds, such as dimethylsulphoxide and so on, via the serine pathway (Murrell and McDonald, 2000). We have consistently identified Hyphomicrobium spp from all the DNA samples collected between 35oN and 30oS in the Atlantic Ocean from 200 m depth (S Sargeant, personal communication). These addition experiments suggest that microbes use methanol mainly as an energy source, and only ∼12% of the uptake is attributable to meeting cellular carbon requirements.
The identification of the organisms involved in C1 uptake and linking this specifically to function can be further achieved using DNA-stable-isotope probing (Neufeld et al., 2007, 2008). Using these techniques at L4, and in the western English Channel, Neufeld et al. (2007, 2008) in addition to Methylophaga spp, have unexpectedly found sequences from undescribed gammaproteobacteria that use methanol, methylamines and dimethylsulphide. These bacteria have no cultured representatives and suggest ecological adaptation of particular methylotrophs to specific C1 compounds in the coastal marine environment (Neufeld et al., 2007).
The reduction in the uptake of methanol into the particulate phase and its oxidation with additions of chloroamphenicol and cycloheximide suggests that not only prokaryotic bacteria, but also perhaps eukaryotic cells use methanol for carbon and energy. In contrast, Goodwin et al. (1997) reported that cycloheximide had no effect on dibromomethane oxidation rates in seawater. We have also found positive linear correlations between uptake of methanol into particulate material and the numbers of heterotrophic bacteria and picoeukaryotes and heterotrophic bacterial production. Additionally, we demonstrated a clear pattern of increasing methanol uptake into particles with increasing chlorophyll a concentrations.
This research now offers methanol loss rates from seawater, for the first time, which will improve the understanding of the global methanol budget and may have implications for bacterial productivity.
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Acknowledgements
We wish to thank all the scientists and crew onboard RRS Discovery during the D320 cruise. In addition, I would like to thank Stephanie Sargeant for the DNA sequencing, Glen Tarran for his help with the flow cytometer, and Claire Widdicombe for the phytoplankton and chlorophyll a analysis at L4, which is provided by the Plymouth Marine Laboratory Western Channel Observatory (http://www.westernchannelobservatory.org.uk), and is funded by the NERC Oceans 2025 Sustained Observatories theme. Satellite data were processed by the NERC Earth Observation Data Acquisition and Analysis Service (NEODAAS) at Plymouth Marine Laboratory (http://www.neodaas.ac.uk). This work was funded by the Surface Ocean Lower Atmosphere Study UK project NE/C517192/1 and by OCEANS 2025, Plymouth Marine Laboratories NERC funded core research programme.
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Dixon, J., Beale, R. & Nightingale, P. Microbial methanol uptake in northeast Atlantic waters. ISME J 5, 704–716 (2011). https://doi.org/10.1038/ismej.2010.169
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DOI: https://doi.org/10.1038/ismej.2010.169
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