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with previous observations that the microbial DMSP lyase pathway accounts for only a small fraction of dissolved DMSP degradation (and therefore acrylate production) in the Gulf of Mexico.
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Concentrations, biological uptake, and respiration of dissolved acrylate and
dimethylsulfoxide in the northern Gulf of Mexico
Inger Marie B. Tyssebotn1,3, Joanna D. Kinsey1,4, David J. Kieber1*, Ronald P. Kiene2, Alison N. Rellinger2, Jessie Motard-Côté2
1
State University of New York, College of Environmental Science and Forestry, Department of Chemistry, Syracuse, New York 13210, USA
2
University of South Alabama, Department of Marine Sciences, Mobile, Alabama, 36688 and Dauphin Island Sea Lab, Dauphin Island, Alabama, 36528, USA
3
Now at: Department of Chemistry, Washington State University, Pullman, Washington 99164, USA
4
Now at: Department of Marine Earth and Atmospheric Sciences, North Carolina State University, Raleigh, North Carolina 27695, USA
*
Corresponding Author: 1 Forestry Drive, Syracuse, New York 13210, USA,
[email protected], Phone: 1-315-470-6951, Fax: 1-315-470-6856
Condensed running head: Acrylate and DMSO biological uptake in seawater
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Abstract
The abundant marine organosulfur compound, dimethylsulfoniopropionate (DMSP) can be degraded to acrylate and dimethylsulfide (DMS), with DMS being further transformed to dimethylsulfoxide (DMSO). Despite intensive study of DMSP and DMS in a variety of marine settings, the processes affecting acrylate and DMSO concentrations in marine waters are poorly known, particularly their loss from the dissolved phase through biological uptake. We measured the concentrations of dissolved acrylate (acrylated) and DMSO (DMSOd) in coastal and open-ocean waters of the northern Gulf of Mexico during non-bloom conditions and quantified the rates and kinetics of their biological uptake using 14C labeled substrates. Acrylated
concentrations and uptake rates ranged from 0.8-2.1 nmol L-1 and 0.07-1.8 nmol L-1 d-1, respectively. Somewhat higher uptake rates were observed for DMSOd (0.27-3.9 nmol L-1 d-1) owing to higher DMSOd concentrations (5.5-14 nmol L-1). Both compounds were taken up by the microbial community with high affinity uptake systems (low nmol L-1 Ks and Vmax) similar to
Ks and Vmax for other well-studied biological substrates including amino acids and
monosaccharides. However, median turnover times for these compounds were relatively slow, 4.8 d for acrylated and 7.4 d for DMSOd. The slow acrylated turnover points to low supply rates of this compound to the dissolved phase, a finding consistent with previous observations that the microbial DMSP lyase pathway accounts for only a small fraction of dissolved DMSP
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Introduction
Acrylate and dimethylsulfoxide (DMSO) are direct and indirect transformation products of dimethylsulfoniopropionate (DMSP), a phytoplankton metabolite with a global production estimated at 59-516 Tmol C year-1, equivalent to 1-14% of global primary production based on carbon (Howard et al. 2006; Galí et al. 2015). Enzymatic cleavage of DMSP by DMSP lyases in both bacteria (e.g., Curson et al. 2011; Moran et al. 2012) and phytoplankton (Alcolombri et al. 2015) produces equimolar quantities of acrylate and dimethylsulfide (DMS) (Cantoni et al. 1956), with some DMS further oxidized to DMSO through either photochemical or biological pathways (Kieber et al. 1996; del Valle et al. 2009; Hatton et al. 2012). Acrylate production may serve as an activated algal defense system (Sieburth 1960; van Alstyne et al. 2001; Evans et al. 2006), and both acrylate and DMSO may function as cellular antioxidants (Sunda et al. 2002). Despite their potential importance, many aspects of acrylate and DMSO cycling in marine waters remain unknown.
Dissolved acrylate (acrylated) has been detected in seawater in the low to high nanomolar range (e.g., Vairavamurthy et al. 1986; Yang et al. 1994; Gibson et al. 1996; Osinga et al. 1996), but overall concentration data are sparse because analysis of field samples has been hindered by high detection limits. Direct analysis of seawater by high performance liquid chromatography (HPLC) with absorbance detection at ~210 nm can quantify, at best, 5-10 nmol L-1 acrylated (Gibson et al. 1996; Evans et al. 2007), and seawater concentrations may often be below this detection limit, especially during non-bloom conditions. In addition to limited concentration data, there are no published reports of the microbial uptake of acrylate in seawater other than in culture studies.
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grown in culture are capable of using acrylate as a substrate for growth. Several strains from the Roseobacter group isolated from coastal Georgia seawater grew with acrylate as the carbon source in the growth medium (Gonzalez et al. 1999). The bacterial strain LFR, isolated from the Sargasso Sea, was similar to an α-proteobacterium and grew when supplied with acrylated
(Ledyard et al. 1993), rapidly metabolizing it to β-hydroxypropionic acid extracellularly (Ansede et al. 2001). Likewise, a Caribbean Sea bacterium isolated during a Trichodesmium bloom oxidized acrylate when grown on DMSP (Diaz et al. 1992). Garren et al. (2014) recently reported that the coral pathogen, Vibrio coralliilyticus, readily degraded millimolar levels of acrylated in culture even though it could not degrade DMSP. Over the course of a Phaeocystis sp. bloom, acrylate-consuming bacteria increased 15-fold, and additional laboratory experiments using a Marinobacter sp. isolate (strain AC-2) showed acrylated uptake rates as high as
4.2 µmol L-1 h-1 (Noordkamp et al. 2000). Given the diversity of environments containing marine bacterial isolates that use acrylate for growth, it is likely that acrylated uptake will be an important process regulating its concentration in seawater.
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longer lifetime for DMSOd in the water column compared to dissolved DMSP (DMSPd) or DMS (Ridgeway et al. 1992). Only a few studies have determined in situ biological uptake rates. del Valle et al. (2007a) determined that the biological uptake rate for DMSOd was less than
0.076 nmol L-1 d-1 in the Ross Sea, Antarctica, based on the loss of DMSOd in dark incubations of unfiltered seawater amended with picomolar levels of 35S-labeled DMSOd. In the North Sea, Simó et al. (2000) determined a net DMSO biological uptake rate of 4 nmol L-1 d-1 and a
turnover time of 2 d, with corresponding DMS and DMSPd turnover times of 0.6 and 1.3 d, respectively.
With very high global production of their precursor, DMSP, acrylate, and DMSO may be important substrates in the marine microbial food web, given that both compounds are used as carbon sources by bacteria in culture. However, understanding of the marine cycling of acrylate and DMSO is limited due to the lack of biological uptake rate data in seawater and, for acrylate, a paucity of dissolved concentration measurements. The objective of this study was to determine concentrations, and rates of total biological uptake (uptake into cells + respiration) of acrylated and DMSOd, and the percentage of total uptake that was respired to carbon dioxide or
assimilated (the subset of cellular uptake incorporated into macromolecular material) in
unfiltered seawater collected at several stations in the northern Gulf of Mexico during non-bloom conditions. Rates of total biological uptake, assimilation, and respiration were determined from dark incubation experiments in which 14C-acrylate or 14C-DMSO were added to unfiltered
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Methods
Hydrographic stations and sampling
Seawater samples were collected from ten stations in the northern Gulf of Mexico during a cruise aboard the R/V Pelican from 12-22 September 2011. Clear skies and calm weather were experienced during most sampling days. Hydrographic data for depth, temperature, salinity, and chlorophyll a (Chl a) were collected for each sampling station.
Seawater was collected in 20 L Niskin sampling bottles attached to a rosette equipped with an Epply PAR irradiance sensor, an Aquatraka III Chl a fluorometer (Chelsea Instruments), and Sea-Bird Electronic SBE conductivity, temperature, and pressure sensors. Unfiltered
seawater was transferred from Niskin bottles directly to sample bottles using silicone tubing. Seawater was also gravity filtered by attaching a pre-cleaned 0.2 µm Nylon Polycap AS filter (Whatman) to the silicone tubing. Polycap filters were cleaned prior to use by alternating rinses of acetonitrile (J.T. Baker) and Milli Q water (> 18.2 MΩ-cm, Millipore Milli Q ultrapure water system), followed by extensive flushing with Milli Q water (Toole et al. 2003). All sampling was performed between 07:30 and 09:00 local time, with the exception of Stn. 20 where samples were collected at 13:00 (Table 1).
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filtered seawater were obtained using both Polycap and 25 mm diameter GF/F filters (Whatman) to compare filtration techniques. Additionally, acrylated concentrations from two depth profiles at stations 4 and 9 were compared following shipboard analysis with freshly collected samples as well as in the laboratory using sample aliquots frozen for several weeks. Due to time constraints, most acrylated analyses were conducted in the laboratory in Syracuse, NY with frozen samples following the cruise.
Unfiltered seawater samples were collected at Stn. 4 to determine total concentrations of acrylate (i.e., dissolved + particulate). Three individual aliquots of approximately ~10 mL of unfiltered water from 44 and 50 m depths were dispensed into pre-cleaned and baked (550°C, 8 h) 20 mL glass scintillation vials (VWR) with Teflon-lined thermoset caps (Qorpak) using silicone tubing directly from the Niskin bottles. These samples were microwaved individually near the center of the rotating plate of a 900 W model SGB8901 microwave oven (Sunbeam), for 5-10 s until boiling (Kinsey and Kieber 2016). Samples for total acrylate concentrations were cooled to room temperature prior to sample preparation and shipboard analysis. Extensive tests showed that microwaving did not cause production or loss of acrylate from sterilized seawater samples, indicating no artifacts associated with the procedure (I. M. B. Tyssebotn unpubl.).
14
C-incubations
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surface seawater incubator to maintain samples at the ambient seawater temperature (~28°C). Dark incubations were performed to avoid photochemical loss of acrylate or DMSO production, or confounding biological light effects.
Concentration-series experiments were performed at eight stations by amending a series of 125 or 250 mL Nalgene polycarbonate bottles containing unfiltered seawater (≤ 20%
headspace) with five to nine different concentrations, ranging from 3.57 to 200 nmol L-1 for 14 C-acrylate, and from 2.00-198 nmol L-1 for 14C-DMSO (60-4,000 and 240-240,00 dpm mL-1, respectively). Incubation times were 1.5-2 h for acrylate and 3-6 h for DMSO. At the end of an incubation, subsamples were prepared for substrate uptake, respiration, and assimilation
measurements (vide infra).
Time-course experiments were performed at ten stations by amending 1 or 2 L
polycarbonate bottles containing unfiltered seawater (≤ 20% headspace) with 7.14 nmol L-114 C-acrylate or 2.00 nmol L-114C-DMSO. Incubation times ranged from 0-3 h for acrylate and 0-8 h for DMSO, with subsamples taken at five or six time points for substrate uptake into cells and respiration. Subsamples for assimilation (the fraction of the uptake into cells that was
incorporated into macromolecules) were only taken at the final time point. Controls consisted of 0.2 µm filtered seawater amended with either 14C-acrylate or 14C-DMSO.
Substrate uptake into cells was determined by filtering 50-200 mL of incubated seawater samples through a 25 mm diameter, 0.22 µm Nylon filter (Tisch Scientific) using a Gelman polysulfone vacuum filtration apparatus. Following filtration, the Nylon filter was rinsed with a small volume of 0.2 µm filtered seawater and transferred to a 6 mL plastic scintillation vial (VWR) containing 4 mL EcoLume scintillation fluid (MP Biomedicals) for subsequent counting.
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80 mL) of incubated seawater were transferred to 160 mL glass serum vials (Wheaton). Serum vials were capped with a rubber stopper containing a 10 mm center-well (Kimble Chase) containing 200 µL β-phenylethylamine (Alpha Aesar) adsorbed onto a fluted 24 mm diameter type 691 glass microfiber filter paper (VWR). Just prior to capping the serum vials, 500 µL of concentrated sulfuric acid (17.8 mol L-1) (EMD) was added to acidify samples and drive the 14
CO2 into the headspace where it adsorbed onto the fluted filter paper and reacted with β-phenylethylamine. Serum vials were shaken for 8-10 h while on a rotating shaker (~100 rpm), after which each filter was removed and placed in a 6 mL plastic scintillation vial containing 4 mL EcoLume.
Assimilation of isotope into macromolecules was determined by filtering 50-200 mL of incubated seawater using the filtration set up previously described for substrate uptake, except that, following filtration, 5 mL of trichloroacetic acid (5% v/v in Milli Q water) was added to the filter and allowed to remain on the filter for 5 min before removal via vacuum filtration. The filter was rinsed with three small volumes of Milli Q water and transferred to a 6 mL scintillation vial containing 4 mL EcoLume.
The total 14C radioactivity of each experimental bottle was determined by pipetting 1 mL of radioisotope-amended seawater into a 6 mL scintillation vial containing 4 mL EcoLume. All 14
C-acrylate and 14C-DMSO amended samples for the determination of total radioactivity, uptake into the cell, respiration, and assimilation were counted for 10 min each using a Wallac 1409 DSA liquid scintillation counter (PerkinElmer).
The maximum rate of total biological uptake (Vmax) and the heterotrophic community
uptake affinity constant (Ks) were determined from the concentration-series experiments using a
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initial substrate concentration. The rate of total biological uptake was calculated from the sum of the rate of substrate uptake into the cells and the rate of respiration. The initial substrate
concentration (So) was calculated from the sum of the added 14C-labelled substrate concentration
and the ambient unlabeled substrate concentration (Sn) initially present in the sample. The
substrate concentration at incubation time t (S) was calculated as So minus the substrate taken up
by the cells and respired. The total biological uptake rate constant (kbio) was determined from the
slope of the first-order plot of ln[S]/[So] versus incubation time. The substrate turnover time was
calculated from the inverse of kbio. The percent respiration (%R) and percent assimilation (%A)
were calculated from time-series experiments using the fraction of 14C-isotope respired to CO2 or assimilated into macromolecules compared to the total biological uptake. The rate of total biological uptake (uptake in cells + respiration) at the ambient substrate concentration (VSn) was
determined by multiplying kbio by Sn.
Leucine incorporation and bacterial production
Bacterial production (BP) was determined through the measurement of 3H-leucine incorporation, as adapted for small volume samples (Smith and Azam 1992). The contribution of acrylated and DMSOd to the bacterial carbon demand (BCD) was calculated using the method outlined by Motard-Côté et al. (2016). In short, leucine incorporation rates were used to estimate bacterial carbon production (BCP) assuming a leucine to carbon conversion factor (LCF) of 1500 g C per mol leucine consumed (Simon and Azam 1989; Ducklow 2000):
BCP=BP×LCF
MM 4.1
where MM is the molar mass of carbon (12 g mol C-1).
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determined using the model of del Giorgio and Cole (1998), and, in combination with BCP, used to estimate BCD:
BGE= 0.037+0.65BCP
1.8+BCP 4.2
BCD= BCP
BGE 4.3
The percent contribution of acrylated or DMSOd to the BCD was calculated from:
% contribution=൬ VSn × #C
BCD ൰*100 4.4
where #C is the number of carbon atoms in acrylate or DMSO.
Analytical procedures
Acrylate – Acrylate concentrations were determined using a pre-column derivatization technique based on the reaction of a chromophoric thiol with the acrylate carbon-carbon double bond through conjugate addition (Tyssebotn 2015). The reagent stock solution was prepared by adding recrystallized o-thiosalicyclic acid to methanol to a final concentration of 20 mmol L-1. A sample (or standard) was derivatized by adding 300 µL of reagent to 3 mL of sample (or
standard) in a 4 mL borosilicate Qorpak vial containing a polypropylene open-top threaded screw cap with a 13 mm dia. Teflon-faced silicone septum insert (National Scientific Company). After the pH was adjusted to 4.0 by addition of 70 µL of 100 mmol L-1 aqueous Ultrex-grade hydrochloric acid (VWR), all samples were screw-capped and reacted for 6 h at 90°C. Once samples cooled to room temperature, they were filtered through 0.2 µm Nylon Acrodisc syringe filters (Pall) prior to analysis. Standards and samples were analyzed using a Shimadzu
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diameter C18 packing (Waters). Derivatized samples and standards were injected and
concentrated on a 2 mm i.d. × 20 mm long stainless steel enrichment column used in lieu of an injection loop. The enrichment column contained 40 µm dia. Waters C18 Sep-Pak cartridge packing, and was rinsed several times with HPLC-grade methanol (Burdick and Jackson) and Milli Q water prior to use. Standards were prepared by standard additions of a 10 mmol L-1 aqueous acrylate stock solution to seawater. The acrylate stock solution was prepared in Milli Q water using sodium acrylate (97%, Aldrich). The limit of detection for the method was
0.2 nmol L-1 for a 1.0 mL injection of derivatized water sample, and the coefficient of variation was 5.3% at 2 nmol L-1 acrylate (Tyssebotn 2015).
DMS and DMSO – To quantify DMS during the cruise, 1-3 mL of seawater was syringe-filtered through a Whatman GF/F filter into a sealed 14 mL serum vial that was attached to a purge-and-trap system used to cryotrap DMS with subsequent introduction into a gas
chromatograph equipped with a flame photometric detector (Motard-Côté et al. 2016).
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Additional analyses – Chromophoric dissolved organic matter absorption spectra were determined in 0.2 µm-filtered samples using an Agilent 8453 UV-Vis spectroscopy system equipped with a microvolume, flow-through 5 cm path length quartz cell according to the procedure outlined in Kieber et al. (2007). The absorption coefficient at 300 nm (a300) was
calculated from the sample absorbance at 300 nm (A300) and path length (l) where a300 =
2.303A300/l.
Samples for Chl a analysis were collected by filtering 250-1000 mL of seawater through a Gelman polysulfone filter holder using a 25 mm diameter Whatman GF/F filter to capture particulate material. Filters were subsequently placed in 10 mL of cold 90% aqueous acetone. The filters were allowed to extract at -4°C for 24 h in the dark before analysis using a Turner Designs TD-700 fluorometer, following the method described by Welschmeyer (1994).
Statistical analyses
Statistical analyses were performed using either SigmaPlot 11.0 with the SigmaStat software package (Systat Software, Inc.) or Minitab 17.2.1 (Minitab, Inc.). All sample sets were tested for normality with a Shapiro-Wilk test prior to correlation analyses with either a Pearson’s product moment correlation (rp) or a Spearman rho correlation (rs). All statistical analyses were
performed at α = 0.05.
Results
Hydrographic stations
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27.4-29.2°C). Sample sites were characterized and differentiated by a300 and salinity (Table 1).
A strong inverse linear relationship was observed between a300 and salinity for the stations
occupied during this study (r2 = 0.969, n = 10), with lower CDOM levels associated with higher salinities, suggesting conservative mixing of CDOM in this region of the Gulf between the rivers and coastal/offshore waters (Fig. 2). Based on the grouping of a300 and salinity data, stations
were qualitatively characterized as coastal (a300 > 1 m-1 and a salinity < 34 ppt) or oceanic (a300 <
1 m-1 and a salinity > 34 ppt) sites. The highest CDOM values were generally closest to shore at the coastal sites (e.g., Stn. 17) with relatively high BCP (Table 2), and strong terrestrial
influences from the Mississippi and Atchafalaya outflows. An exception to this pattern was seen at Stn. 14, which despite being further inshore than Stn. 2, was classified as oceanic based on our operational a300 and salinity criteria. Observed differences between these stations were likely
due to water mass movements during the seven days separating sampling at Stn. 2 and 14. Bacterial carbon production and, by inference, terrestrial influences decreased by a factor of 2 to 5 further from the coast (e.g., Stn. 8, Table 2). Chlorophyll a, a proxy for phytoplankton
biomass, showed a similar trend as BCP, with generally high but variable Chl a at coastal stations (0.18-2.8 µg Chl a L-1) and relatively low Chl a at the offshore stations (0.06-0.27 µg Chl a L-1) (Table 1).
Acrylate and DMSO concentrations
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approximately in steady state in the study region. The lowest acrylated concentrations were detected at Stn. 20 with the highest CDOM absorbance coefficients, whereas the highest acrylate concentrations were found near the Mississippi outflow (Stn. 2) and at one oceanic site (Stn. 23). A comparison of filtration techniques for acrylated showed no significant difference (t4 = -0.302,
p = 0.778) when samples were filtered through a GF/F filter using the Kiene and Slezak (2006) small volume drip filtration technique (1.1 ± 1.2 nmol L-1) compared to concentrations
determined in samples that were filtered through a Polycap AS filter (1.4 ± 1.0 nmol L-1). Dissolved DMSO concentrations in surface waters (5.5-13.8 nmol L-1) were up to an order of magnitude higher than acrylated concentrations (Table 1), and as with acrylated, there were no significant differences in DMSOd concentrations between coastal (9.2 ± 3.2 nmol L-1) and oceanic (7.0 ± 1.6 nmol L-1) waters.
Depth profiles for acrylated, DMSOd, and DMS, along with temperature, salinity, Chl a, and CDOM (a300) were determined at three stations with low (Stn. 8), intermediate (Stn. 14), and
high (Stn. 9) BCP (Figure 3A, D, G; see Table 2 for BCP values). Dissolved acrylate
concentrations were relatively constant throughout the upper water column at all three stations (~1-2 nmol L-1), and were similar to DMS concentrations, but an order of magnitude lower than DMSOd. Unlike acrylated and DMS profiles, DMSOd depth profiles showed a distinct pattern with depth, with the highest concentrations in surface waters (6-14 nmol L-1) and lower, relatively uniform concentrations deeper in the water column (4-5 nmol L-1, Fig. 3). No
significant correlations were observed between DMS, DMSOd, or acrylated and Chl a or CDOM (a300).
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1.1 ± 0.5 nmol L-1 (data from Fig. 3 panel G) in samples that were collected in parallel and stored frozen until they were analyzed in Syracuse, NY. A similar result was obtained for acrylated in the Stn. 4 depth-profile study (shipboard 1.1 ± 0.4 nmol L-1; laboratory 1.3 ± 0.1 nmol L-1). The good agreement between shipboard and home laboratory results with respect to acrylated
indicated that sample storage did not significantly affect acrylated concentrations.
The Stn. 4 unfiltered samples had total acrylate concentrations that ranged from 1.4 to 3.4 nmol L-1 for the two depths in the upper 50 m of the water column, with corresponding dissolved concentrations of 0.87-1.7 nmol L-1. The difference between total acrylate and acrylated concentrations yielded particulate acrylate concentrations of 0.53-1.7 nmol L-1, indicating that a significant fraction of acrylate was present in both the dissolved (39-50%) and particulate phases (50-61%).
Uptake rates, turnover times, respiration, and assimilation
Rates of total biological uptake showed Michaelis-Menten saturation-type behavior with increasing substrate addition (Fig. 4). Rates were corrected for 14C activity in filtered seawater controls, which were indistinguishable from background counts. Maximum rates of total biological uptake, Vmax, for acrylated and DMSOd were at concentrations that were 3 to 10 fold higher than ambient dissolved concentrations (Table 1, Fig. 4). Comparison among stations of Ks and Vmax showed that these parameters varied by more than an order of magnitude, with
oceanic sites generally having lower Vmax than coastal sites (Fig. 5). Maximum rates of total
biological uptake ranged from 0.06 to 1.2 nmol L-1 h-1 for acrylated, and were approximately three times higher than observed for DMSOd (0.023-0.33 nmol L-1 h-1) (Fig. 5, Table 3);
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DMSOd, respectively (Fig. 5, Table 3), with no significant correlations to Chl a, BCP, CDOM or salinity.
Initial substrate concentrations (Sn + 14C-substrate) were similar to Ks for both acrylate
and DMSO, well within the first-order kinetic region. First-order rate plots for acrylated uptake versus incubation time (Fig. 6) showed a significantly slower uptake at the low BCP station (Stn. 4) compared to the high BCP stations (Stns. 2, 9, 20), and all of the high BCP stations were within experimental error of each other. By contrast, DMSOd kinetic plots exhibited a gradient of increasing rates of DMSOd uptake with increasing BCP from Stn. 4 to Stn. 20 (Fig. 6, Table 2). The kbio for acrylated and DMSOd were highly correlated to each other with rs = 0.927 and p
< 0.001 (Table 3).
Microbial respiration (%R) and assimilation (%A) were calculated as a percentage of the total biological uptake of acrylated or DMSOd (Table 3). Respiration accounted for 16-40% of the total uptake of acrylated, whereas %R was much higher for DMSOd (62-75%). Assimilation accounted for 22-60% of total acrylated uptake and only 15-33% of total DMSOd uptake (Table 3). For acrylated, %A and %R showed opposite trends when moving from coastal to oceanic waters (Fig. 7); %A for acrylated was typically lower in oceanic waters (median 35%) than in biologically productive coastal sites (median 49%), whereas %R was higher in oceanic waters (median 36%) compared to coastal stations (median 20%). In contrast, %R for DMSOd was similar for most stations (Table 4), while %A was slightly lower at the oceanic stations (median 25%) than at the coastal Mississippi outflow stations (median 29%). For all samples, means and medians differed by 1-3%.
Rates of total biological uptake at ambient substrate concentrations (VSn) were low for
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biological uptake were observed for DMSOd (0.27-3.9 nmol L-1 d-1), owing to higher DMSOd concentrations in surface waters, with higher rates in coastal waters (e.g., Stn. 2) compared to offshore oceanic waters (e.g., Stn. 8). Despite higher VSn, turnover times for DMSOd were slow,
ranging from 2.2 to 37 d (median 7.4 d), nearly twice as long as the median acrylated turnover time (0.8-22 d, median 4.8 d) (Fig. 8).
Acrylated and DMSOd contribution to bacterial carbon demand
Dissolved acrylate and DMSO each contributed less than 0.2% to the BCD (Table 2), with no significant difference in percent contribution between coastal and oceanic sites.
Nonetheless, a significant correlation was observed between the BCP and kbio for acrylated (rs =
0.786, p = 0.015) and for DMSOd (rs = 0.762, p = 0.021).
Discussion
Dissolved acrylate and DMSO concentrations
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concentrations that were less than 0.5 µg L-1. These concentrations were more than an order of magnitude higher than we detected for acrylated in the northern Gulf of Mexico. The reasons for these differences are unknown, but may be related to differences in sampling, sample storage or analytical methods; Liu et al. (2016) stored GF/F-filtered samples (~40 mL) in glass vials at 4oC, which likely contained bacteria (Lee et al. 1995). In the present study, most samples were gravity filtered through 0.2 µm pore-size filters, which likely removed ~97% of bacteria (Benner and Biddanda 1998), and stored frozen at -20oC or, in a few cases, analyzed immediately
following collection. Acrylated concentrations determined in frozen samples were not significantly different from acrylated concentrations determined in freshly collected Gulf of Mexico samples.
Dissolved acrylate concentrations in the surface mixed layer (0.8-2.1 nmol L-1, median 1.5 nmol L-1) were comparable to DMS (0.9-4.7 nmol L-1, median 2.3 nmol L-1) and DMSPd (0.11-2.7 nmol L-1, median 1.1 nmol L-1) concentrations measured in parallel during the same cruise (DMS and DMSPd surface mixed layer data from Motard-Côté et al. 2016). Although dissolved concentrations were in the low nanomolar range for all three compounds (Tables 1 and 4, Fig. 3), only a modest correlation was observed between acrylated and DMS (rs = 0.496, p =
0.010, n = 26) and no correlation was seen between acrylated and DMSPd (rs = 0.024, p = 0.931,
n = 8) or for DMS and DMSPd (rs = -0.06, p = 0.0.839, n = 8). The correlation between DMS
and acrylated may be expected, since the enzymatic lysis of particulate DMSP should be an important source for both DMS and acrylate. However, differences in the input and removal rates for acrylate and DMS in the dissolved phase likely affected concentrations of these
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charged species at physiological pH and has more complex removal mechanism. Thus, although total acrylate concentrations were low (1.4-3.4 nmol L-1) and only a few samples were analyzed, a significant fraction of total acrylate (50-61%) was present in the particulate phase, consistent with algal culture studies (Tyssebotn 2015; Kinsey et al. 2016). By contrast, DMS should mostly (~100%) be in the dissolved phase (Spiese et al. 2016).
Another important difference between acrylated and DMS was that DMS removal rates from the dissolved phase were faster than for acrylated, for both photolysis (J. D. Kinsey unpubl.) and biological uptake (Motard-Côté et al. 2016). The first-order DMS photolysis rate constant during our cruise (0.04-0.20 h-1, J. D. Kinsey unpubl.) was more than an order of magnitude greater than the rate constant for acrylated photolysis (0.6-1.5 × 10-3 h-1, Tyssebotn 2015). With a lower rate constant and similar DMS and acrylated concentrations, corresponding photolysis rates will be slower for acrylated (Tables 1 and 4).
Biological uptake rates of acrylated at ambient concentrations were also generally slower than for DMS, but mostly at the oceanic stations. For the coastal stations, VSn for DMS,
determined from dark incubations of 35S-DMS-amended unfiltered seawater (Motard-Côté et al. 2016), were comparable to those for acrylated (Tables 2 and 4, Stns. 2, 9, 17). In oceanic waters (Stns. 4, 6, 8, 14, 19) DMS VSn were substantially greater than the corresponding acrylate VSn by
a factor of 4.6-12 (Tables 2 and 4). These large differences were not due to differences in Sn
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Overall, similar acrylated and DMS concentrations (Fig. 3, Tables 1 and 4), but lower acrylate VSn (at the open-ocean sites) and photolysis rates (0.6-1.5 × 10-3 h-1, Tyssebotn 2015),
suggest that rates of acrylate input to the dissolved phase were less than corresponding DMS inputs into northern Gulf of Mexico waters. This might be expected because bacteria that take up DMSPd and cleave it intracellularly, likely metabolize acrylate further while losing DMS to diffusion out of the cell (Ansede et al. 2001; Kiene et al. 2000; Spiese et al. 2016). Similarly, smaller fluxes of acrylate into the dissolved phase from phytoplankton cells might be expected because some acrylate is retained in phytoplankton cells (Tyssebotn 2015; Kinsey et al. 2016), and marine particulate matter (this study), while DMS will rapid diffuse out of phytoplankton cells into the dissolved phase (Spiese et al. 2016). Thus extracellular releases of acrylate will mostly depend on processes such as viral lysis of or grazing on phytoplankton cells, which one would expect to be relatively low under non-bloom conditions. In addition, DMS has other sources that do not result in acrylate production. Three potential sources of DMS that would not lead to acrylate production are DMSO reduction (Fuse et al. 1995; Spiese et al. 2009),
methylation of methanethiol (Kiene and Taylor 1989; Carrión et al. 2015), and the DddD gene-encoded cleavage of DMSP to form DMS and β-hydroxypropionate (Todd et al. 2007).
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significantly greater than the rate of photolysis (ca. 0.01 nmol L-1 d-1), we infer that the net production term must be low, ca. 0.53 nmol L-1 d-1, and therefore conclude, that inputs of acrylated from DMSP cycling and particulate sources must be slow. By comparison, the
biological production of DMS from the biological consumption of DMSPd in the same study area ranged from 1.2 to 5.0 nmol L-1 d-1 (Motard-Côté et al. 2016).
Dissolved DMSO was the dominant dissolved organosulfur compound measured in the Gulf of Mexico, a finding that is consistent with several studies in oceanic waters including the Arabian Sea (Hatton et al. 1996), Pacific Ocean (Bates et al. 1994), Mediterranean Sea (Simó et al. 1995), coastal Atlantic Ocean (Ridgeway et al. 1992), the Ross Sea (del Valle et al. 2009), and Antarctic coastal waters (Gibson et al. 1990). Dissolved DMSO concentrations showed no significant differences between stations or correlations with acrylated, DMSPd, a300, BCP, or
Chl a. However, DMSOd concentrations showed a strong linear correlation to DMS in both surface waters (rp = 0.776, p = 0.024, n = 8) and when depth profile data were included (rp =
0.759, p < 0.001, n = 26). This relationship was expected as DMS is the main source of DMSOd in the surface mixed layer through its photochemical and biological oxidation (Kieber et al. 1996; del Valle et al. 2009), and DMSOd losses are slow. Previous studies have also reported strong correlations between DMS and DMSO concentrations during the spring and summer in a variety of oceanic surface waters. Kiene et al. (2007) collected DMS and DMSOd concentration data from a Southern Ocean transect between New Zealand and the Ross Sea in three separate years, once in the austral summer (2004) and twice in the austral spring (2003 and 2005); they observed a relatively high correlation between DMS and DMSOd (rs = 0.630, p < 0.001, n = 108)
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strong correlation was observed between DMS and DMSOd (r2 = 0.801, p < 0.001) irrespective of geographical location.
Depth profiles of DMSOd concentrations in the Gulf of Mexico (Fig. 3) are consistent with previously reported depth profiles in several oceanic water types ranging from tropical to polar regions (Andreae 1980; Ridgeway et al. 1992; Kiene and Gerard 1994; Hatton et al. 1996; Simó et al. 1997; del Valle et al. 2009). In these studies, the highest DMSOd concentrations were nearly always detected in the surface mixed layer, followed by a rapid decrease in DMSOd with depth. This trend was observed in the present study (Fig. 3) and was likely controlled by photochemical and biological oxidation of DMS to DMSO (Kieber et al. 1996; del Valle et al. 2007a,b; del Valle et al. 2009) weakly offset by the slow biological uptake of DMSOd (vide infra) and weak vertical mixing due to the very calm conditions throughout the cruise (median wind speed 2.4 m s-1).
Biological uptake and transformations of acrylated and DMSOd
The low nanomolar Ks values reported here for the biological uptake of acrylated and DMSOd are consistent with the low nanomolar concentrations observed for these compounds in the northern Gulf of Mexico. For acrylated, these low values suggest that it was taken up
efficiently by the heterotrophic community through high-affinity uptake systems. For DMSOd, it will rapidly diffuse into the cell, reaching equilibrium within seconds (Spiese et al. 2009), and therefore specific cell-membrane transporters are not needed for its uptake. Nonetheless, we observed saturation kinetics for DMSO uptake that likely reflected the heterotrophic
community’s average Ks and Vmax for the enzymes responsible for the assimilation and
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Values of Ks and Vmax obtained for the uptake of acrylated and DMSOd are comparable to
values of these kinetic parameters observed in marine waters for the biological uptake of other low molecular weight organic substrates including dissolved amino acids (Ferguson and Sunda 1984; Suttle et al. 1991), monosaccharides (Skoog et al. 1999), α-keto acids (Kieber et al. 1989), glycolate (Edenborn and Litchfield 1987), glycine betaine (Kiene and Hoffman Williams 1998), methanol (Dixon et al. 2011), and CO (Xie et al. 2005), all of which have Ks in the low
nanomolar range and Vmax between 10-10 and 10-9 mol L-1 h-1. Despite the similarity in the uptake
kinetic constants, turnover times were considerably slower for acrylate and DMSO compared to several other biological substrates including amino acids, as discussed in the ensuing section.
Although heterotrophic bacteria have been shown to take up DMSOd, its cellular transformations are not well understood. The only cellular transformation that has been
identified for DMSO in marine systems is the enzymatic reduction of DMSO to DMS (Fuse et al. 1995; Spiese et al. 2009), but this was not determined in the present study since any DMS
formed from 14C-DMSO through this process would quickly partition to the dissolved phase (Spiese et al. 2016) and not be captured on a filter. Dimethylsulfide produced from DMSO in this way could be further metabolized as well, either back to DMSO or to other mineralized end products (Wolfe and Kiene 1993; del Valle et al. 2007b). Enzymes capable of reducing DMSO to DMS appear to be ubiquitous in marine bacteria (Fuse et al. 1995) and marine algae (Spiese et al. 2009). It is unknown whether enzymatic reduction of DMSO to DMS occurred at an
appreciable rate in the Gulf of Mexico water column, but it has been shown to be important in Antarctic sea-ice brine communities wherein DMSO reduction, as opposed to DMSP cleavage, was the main source of DMS (Asher et al. 2011).
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our results indicate that DMSO is mostly oxidized for energy with a small percentage assimilated for growth. The majority of DMSOd taken up by the heterotrophic community was respired (62-75%, Table 3). This degree of respiration is generally greater than seen for other biological substrates that have been studied in seawater including glucose (e.g., 24-41%, Arctic, Griffiths et al. 1984; 30-60%, equatorial Pacific, Rich et al. 1996), glycine betaine (40-60%, Gulf of Mexico, Kiene and Hoffman Williams 1998), leucine (e.g., < 10%, Sargasso Sea, Suttle et al. 1991; 30-70% Mediterranean Sea, del Giorgio et al. 2011), or dissolved free amino acids (e.g., 30-65%, Kysing Fjord, Jørgensen 1982; 42-68%, Arctic, Griffiths et al. 1984). A few substrates have been found to have similarly high percentages of respiration to those of DMSO, including mannose, which was 60-90% respired in the equatorial Pacific (Rich et al. 1996), and glutamate, which was 60-65% respired in the Kysing Fjord (Jørgensen 1982) and 60-80% respired in the Sargasso Sea (Suttle et al. 1991), or even a higher percentage as observed for CO which is essentially all respired (~99%, Tolli and Taylor 2005). It is interesting to note that for DMSOd, unlike acrylated, there was no significant difference in the percent respiration between the low and high bacterial productivity stations in the Gulf of Mexico; a high percentage of respiration was observed at all stations. This implies that the DMSO carbon moieties were mostly
metabolized for energy, irrespective of the amount or lability of dissolved carbon or nutrient status of the seawater.
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therefore not surprising that there was biological uptake of acrylate in northern Gulf of Mexico waters. However, acrylated exhibited a very different uptake pattern compared to DMSOd. First, acrylate was assimilated into macromolecules at percentages comparable to or greater than its respiration (Table 3). Second, as %R increased, %A decreased for acrylate uptake when going from the high CDOM and bacterial productivity coastal stations to the low CDOM and bacterial productivity oceanic stations (Fig. 7). This indicated that acrylate was metabolized more for energy than assimilated for growth in offshore waters compared to coastal waters, similar to what is generally expected for the BGE when comparing productive nearshore waters to oligotrophic waters (del Giorgio and Cole 1998). The third difference in biological uptake between the two compounds was that a significant percentage (ca. 33% on average) of the total acrylate uptake was not accounted for by either assimilation into macromolecules or respiration, unlike DMSO whose respiration and assimilation accounted for nearly 100% of its total uptake. This implies that, unlike DMSO, some 14C-acrylate uptake remained in the cytosol as acrylate or as a transformed low molecular weight 14C product. One potential transformation is the
conversion of acrylate to acrylyl-CoA, a toxic compound which may be further metabolized to β-hydroxypropionate potentially as a detoxification mechanism (Curson et al. 2014), followed by the accumulation of β-hydroxypropionate in the cytosol (Ansede et al. 1999).
Acrylated and DMSOd turnover times
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reported for acrylate, and despite extensive global measurements of DMSO concentrations, there are very few reported DMSO turnover times in oceanic waters.
The prevailing view in the literature is that DMSOd turnover times are slow in marine waters, and this is certainly true compared to reported turnover times for other simple organic molecules that are generally less than a day (vide infra). This has led to speculation by Hatton et al. (1996) and Ridgeway et al. (1992) that DMSO will serve as a net loss for DMS from the water column. This is likely correct over short time scales (days) because DMSOd turnover times in the Gulf of Mexico were slow (median 7.4 d). However, whether DMSO serves as a net loss for DMS or not, will depend on how much DMSO produced from DMS is enzymatically reduced back to DMS, as shown in culture (Fuse et al. 1995; Spiese et al. 2009), and therefore recycled, versus the amount of DMSO that is respired or assimilated.
Dissolved DMSO turnover times for coastal waters (median 3.2 d) were comparable to published DMSO turnover-time estimates, which are generally on the order of a few days in coastal waters (Kiene and Gerard 1994; Simó et al. 2000; Asher et al. 2011). In contrast, DMSOd turnover times in the Gulf of Mexico oceanic sites were 3 to 5 times longer (median 11.3 d) than previously reported in coastal waters, likely due to the correspondingly lower bacterial productivities (Table 2). These differences indicate that there may be considerable spatial and temporal variability in DMSOd turnover in oceanic waters, warranting a more detailed study to determine the factors controlling the biological consumption of DMSOd.
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acrylated turnover time (4.8 d) is nearly 37 times longer than the median turnover time for DMSPd, and approximately 3 times longer than the median DMS turnover time. Turnover times for DMSOd and acrylated (Table 3, Fig. 8) are also generally much longer than dissolved free amino acids, which are generally less than 1 d (Ferguson and Sunda 1984; Fuhrman and Ferguson 1986; Kiel and Kirchman 1999; Suttle et al. 1991). Monosaccharide turnover times tend to be somewhat longer, on the order of 1-10 d (Suttle et al. 1991; Rich et al. 1996; Skoog et al. 1999), which are comparable to DMS and somewhat shorter than acrylated, but still
substantially shorter than DMSOd. Rich et al. (1996) observed considerable temporal variation in the uptake of glucose in the equatorial Pacific, with faster turnover (~2 d) in the summer months under normal sea surface temperature conditions compared to the following winter (~10 d) during an El Niño event. They also observed a large difference between mannose and glucose, with mannose exhibiting very little temporal variation in its turnover time, which was nearly 50 d on average. This is much slower than observed for acrylate or DMSO, but the reasons for these differences are unknown. Clearly, there will be interacting physical and
biogeochemical factors at play in contributing to these differences, not only between glucose and mannose but also among all the substrates discussed here. Temporal and spatial variations in temperature, light, the quality and quantity of organic matter and nutrients available to the microbial community, and in the makeup of microbial community itself will be important, as no one substrate will likely be taken up by all representative microbial groups. Additional studies are needed to relate substrate uptake to specific environmental conditions and the composition of the microbial community.
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Dissolved acrylate and DMSO contributed 0.013-0.13% and 0.04-0.14% of BCD, respectively (Table 2). Given the slow turnover times and low concentrations of acrylated and DMSOd, it is not surprising that they were not quantitatively important carbon substrates in northern Gulf of Mexico waters for the entire heterotrophic community during non-bloom conditions. Contributions of both compounds to BCD are approximately an order of magnitude lower than observed for DMSPd on the same cruise (1.1-7.0%) (Motard-Côté et al. 2016) or on a previous cruise in the Gulf of Mexico (0.4-6.5%) (Kiene and Linn 2000), and generally much lower than the 10 to ~100% contributions to BCD by dissolved free amino acids or glucose observed in numerous studies (for review see Nagata 2008). The low percent contribution of acrylated and DMSOd to the BCD determined here does not exclude the possibility that acrylated and DMSOd are important sources of carbon to specific bacterial groups. Our finding that acrylated and DMSOd biological uptake at ambient concentrations occurred through high-affinity (i.e., low nmol L-1 Ks) uptake systems indicates that at least some subset of the bacterial
community is poised to efficiently utilize these substrates in both productive coastal waters and under oligotrophic conditions.
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whether acrylate and DMSO make greater contributions to BCP in major DMSP- and DMS-producing blooms, and to assess the diversity of marine microbes that are capable of using acrylated and DMSOd as carbon sources for microbial energy and growth.
Conclusions
Our study significantly advances the field by filling in gaps about the fate of acrylate and DMSO in seawater. These two compounds are very often discussed in the context of DMSP and DMS cycling, but acrylate and DMSO have not previously been investigated at the detailed level of our study. We have shown that the microbial community takes up acrylated and DMSOd with high affinity systems, but biological turnover times are slow. The slow biological loss of
DMSOd, along with its photochemical and thermal stability, suggests that DMSOd serves as a significant short-term (days - weeks) dimethylsulfur storage species in the photic zone. Given the slow turnover rate of acrylate and DMSO, it is possible that these compounds are not
consumed by a wide array of microbes or by the dominant microbes present in seawater. This is in contrast to findings for DMSP, which is used by a wide spectrum of the microbial community (Malmstrom et al. 2004; Vila et al. 2004). Based on the relatively long biological turnover times of acrylated, we also conclude that supply rate of this compound to the dissolved pool is
Accepted Article
Accepted Article
Acknowledgments
The authors thank the officers and crew of the R/V Pelican for their technical support at sea, and our colleagues Lisa Oswald and Bradley Myer for their help with sampling and analysis.
Accepted Article
References
Alcolombri, U., S. Ben-Dor, E. Feldmesser, Y. Levin, D. S. Tawfik, and A. Vardi. 2015.
Identification of the algal dimethylsulfide-releasing enzyme: A missing link in the marine sulfur cycle. Science 348: 1466-1469. doi:10.1126/science.aab1586
Andreae, M. O. 1980. Dimethylsulfoxide in marine and freshwaters. Limnol. Oceanogr. 25: 1054-1063. doi:10.4319/lo.1980.25.6.1054
Ansede, J. H., P. J. Pellechia, and D. C. Yoch. 1999. Metabolism of Acrylate to
β-hydroxypropionate and its role in dimethylsulfoniopropionate lyase induction by a salt marsh sediment bacterium, Alcaligenes faecalis M3A. Appl. Environ. Microbiol. 65: 5075-5081.
Ansede, J. H., P. J. Pellechia, and D. C. Yoch. 2001. Nuclear magnetic resonance analysis of [1-13
C]dimethylsulfoniopropionate (DMSP) and [1-13C]acrylate metabolism by a DMSP lyase-producing marine isolate of the α-subclass of proteobacteria. Appl. Environ. Microbiol. 67: 3134-3139. doi:10.1128/AEM.67.7.3134-3139.2001
Asher, E. C., J. W. H. Dacey, M. M. Mills, K. R. Arrigo, and P.D. Tortell. 2011. High concentrations and turnover rates of DMS, DMSP and DMSO in Antarctic sea ice. Geophys. Res. Lett. 38: L23609. doi:10.1029/2011GL049712
Bates, T. S., R. P. Kiene, G. V. Wolfe, P. A. Matrai, F. P. Chavez, K. R. Buck, B. W. Blomquist, and R. L. Cuhel. 1994. The cycling of sulfur in surface seawater of the northeast Pacific. J. Geophys. Res. 99: 7835-7843. doi:10.1029/93JC02782
Benner R., and B. Biddanda. 1998. Photochemical transformations of surface and deep marine dissolved organic matter: Effects on bacterial growth. Limnol. Oceangr. 43: 1373-1378. doi:10.4319/lo.1998.43.6.1373
Cantoni, G. L., and D. G. Anderson. 1956. Enzymatic cleavage of dimethylpropiothetin by Polysiphonia lanosa. J. Biol. Chem. 222: 171-177.
Carrión, O., A. R. J. Curson, D. Kumaresan, Y. Fu, A. S. Lang, E. Mercade´, and J. D. Todd. 2015. A novel pathway producing dimethylsulphide in bacteria is widespread in soil environments. Nat. Comm. 6: 6579. doi:10.1038/ncomms7579
Curson, A. R. J., J. D. Todd, M. J. Sullivan, and A. W. B. Johnston. 2011. Catabolism of dimethylsulphoniopropionate: microorganisms, enzymes and genes. Nature Rev. Microbiol. 9: 849-859. doi:10.1038/nrmicro2653
Curson, A. R. J., O. J. Burns, S. Voget, R. Daniel, J. D. Todd, K. McInnis, M. Wexler, A. W. B. Johnston. 2014. Screening of metagenomic and genomic libraries reveals three classes of bacterial enzymes that overcome the toxicity of acrylate. PLoS ONE 9: e97660.
doi:10.1371/journal.pone.0097660
del Giorgio, P. A., and J. J. Cole. 1998. Bacterial growth efficiency in natural aquatic systems. Annu. Rev. Ecol. Syst. 29: 503-541. doi:10.1146/annurev.ecolsys.29.1.503
Accepted Article
along a northeastern Pacific inshore-offshore transect. Limnol. Oceanogr. 56: 1-16. doi:10.4319/lo.2011.56.1.0001
del Valle, D. A., D. J. Kieber, J. Bisgrove, and R. P. Kiene. 2007a. Light-stimulated production of dissolved DMSO by a particle-associated process in the Ross Sea, Antarctica. Limnol. Oceanogr. 52: 2456-2466. doi:10.4319/lo.2007.52.6.2456
del Valle, D. A., D. J. Kieber, and R. P. Kiene. 2007b. Depth-dependent fate of biologically-consumed dimethylsulfide in the Sargasso Sea. Mar. Chem. 103: 197-208.
doi:10.1016/j.marchem.2006.07.005
del Valle, D. A., D. J. Kieber, D. A. Toole, J. Bisgrove, and R. P. Kiene. 2009. Dissolved DMSO production via biological and photochemical oxidation of dissolved DMS in the Ross Sea, Antarctica. Deep-Sea Res. I 56: 166-177. doi:10.1016/j.dsr.2008.09.005
Diaz, M. R., P. T. Visscher, and B. F. Taylor. 1992. Metabolism of dimethylsulfoniopropionate and glycine betaine by a marine bacterium. FEMS Microbiol. Lett. 96: 61-65.
doi:10.1016/0378-1097(92)90457-Y
Dixon, J. L., R. Beale, and P. D. Nightingale. 2011. Microbial methanol uptake in northeast Atlantic waters. ISME J 5: 704-716. doi:10.1038/ismej.2010.169
Ducklow, H. W. 2000. Bacterial production and biomass in the oceans, p. 85-120. In D. Kirchman [ed.], Microbial ecology of the oceans. Wiley.
Edenborn, H. M., and C. D. Litchfield. 1987. Glycolate turnover in the water column of the New York Bight apex. Mar. Biol. 95: 459-467. doi: 10.1007/BF00409575
Evans, C., G. Malin, W. H. Wilson, and P. S. Liss. 2006. Infectious titres of Emiliania huxleyi virus 86 are reduced by exposure to millimolar dimethyl sulfide and acrylic acid. Limnol. Oceanogr. 51: 2468-2471. doi:10.4319/lo.2006.51.5.2468
Evans, C., S. V. Kadner, L. J. Darroch, W. H. Wilson, P. S. Liss, and G. Malin. 2007. The relative significance of viral lysis and microzooplankton grazing as pathways of dimethylsulfoniopropionate (DMSP) cleavage: An Emiliania huxleyi culture study. Limnol. Oceanogr. 52: 1036-1045. doi:10.4319/lo.2007.52.3.1036
Fuhrman, J. A., and R. L. Ferguson. 1986. Nanomolar concentrations and rapid turnover of dissolved free amino acids in seawater: Agreement between chemical and
microbiological measurements. Mar. Ecol. Prog. Ser. 33: 237-242. doi: 10.3354/meps033237
Ferguson, R. L., and W. G. Sunda. 1984. Utilization of amino acids by planktonic marine bacteria: Importance of clean technique and low substrate additions. Limnol. Oceanogr. 29: 258-274. doi:10.4319/lo.1984.29.2.0258
Fuse, H., O. Takimura, K. Kamimura, K. Murakami, Y. Yamaoka, and Y. Murooka. 1995. Transformation of dimethyl sulfide and related compounds by cultures and cell extracts of marine phytoplankton. Biosci. Biotech. Bioch. 59: 1773-1775.
Accepted Article
Galí, M., E. Devreda, M. Levasseur, S. Royer, and M. Babin. 2015. A remote sensing algorithm for planktonic dimethylsulfoniopropionate (DMSP) and an analysis of global patterns. Remote Sens. Environ. 171: 171-184. doi: 10.1016/j.rse.2015.10.012
Garren, M., K. Son, J.-B. Raina, R. Rusconi, F. Menolascina, O. H. Shapiro, J. Tout, D. G. Bourne, J. R. Seymour, and R. Stocker. 2014. A bacterial pathogen uses
dimethylsulfoniopropionate as a cue to target heat-stressed corals. ISME J. 8: 999-1007. doi:10.1038/ismej.2013.210
Gibson, J. A. E., R. C. Garrick, H. R. Burton, and A. R. McTaggart. 1990. Dimethylsulfide and the alga Phaeocystis pouchetii in Antarctic coastal waters. Mar. Biol. 104: 339-346. doi: 10.1007/BF01313276
Gibson, J. A. E., K. M. Swadling, and H. R. Burton. 1996. Acrylate and
dimethylsulfoniopropionate (DMSP) concentrations during an Antarctic phytoplankton bloom, p. 213-222. In R. P. Kiene, P. T. Visscher, M. D. Keller, and G. O. Kirst. [eds.], Biological and environmental chemistry of DMSP and related sulfonium compounds. Plenum Press.
Gonzalez, J. M., R. P. Kiene, and M. A. Moran. 1999. Transformation of sulfur compounds by an abundant lineage of marine bacteria in the α-subclass of the class Proteobacteria. Appl. Environ. Microbiol. 65: 3810-3819.
Griffiths, R. P., B. A. Caldwell, and R. Y. Morita. 1984. Observations on microbial percent respiration values in arctic and subarctic marine waters and sediments. Microb. Ecol. 10: 151-164. doi: 10.1007/BF02011422
Hatton, A. D., G. Malin, S. M. Turner, and P. S. Liss. 1996. DMSO: A significant compound in the biogeochemical cycle of DMS, p. 405-412. In R. P. Kiene, P. T. Visscher, M. D. Keller, and G. O. Kirst [eds.], Biological and environmental chemistry of DMSP and related sulfonium compounds. Plenum Press.
Hatton A. D., S. M. Turner, G. Malin, and P. Liss. 1998. Dimethylsulphoxide and other biogenic sulphur compounds in the Galapagos plume. Deep-Sea Res. II 45:1043-1053.
doi:10.1016/S0967-0645(98)00017-4
Hatton, A. D., L. J. Daroch, and G. Malin. 2004. The role of dimethylsulphoxide in the marine biogeochemical cycle of dimethylsulphide. Oceanogr. Mar. Biol. Annu. Rev. 42: 29-56. doi: 10.1201/9780203507810.ch2
Hatton, A. D., D. M. Shenoy, M. C. Hart, A. Mogg, and D. H. Green. 2012. Metabolism of DMSP, DMS and DMSO by the cultivable bacterial community associated with the DMSP-producing dinoflagellate Scrippsiella trochoidea. Biogeochemistry 110: 131-146. doi:10.1007/s10533-012-9702-7
Howard, E. C., J. R. Henriksen, A. Buchan, C. R. Reisch, H. Bürgmann, R. Welsh, W. Ye, J. M. González, K. Mace, S. B. Joye, R. P. Kiene, W. B. Whitman, and M. A. Moran. 2006. Bacterial taxa that limit sulfur flux from the ocean. Science 314: 649-652. doi:
Accepted Article
Jørgensen, N. O. G. 1982. Heterotrophic assimilation and occurrence of dissolved free amino acids in a shallow estuary. Mar. Ecol. Prog. Ser. 8:145-159.
Keil, R. G., and D. L. Kirchman. 1999. Utilization of dissolved protein and amino acids in the northern Sargasso Sea. Aquat. Microbiol. Ecol. 18: 293-300. doi:10.3354/ame018293
Kieber, D. J., J. McDaniel, and K. Mopper. 1989. Photochemical source of biological substrates in sea water: implications for carbon cycling. Nature 341: 637-639.
doi:10.1038/341637a0
Kieber, D. J., J. Jiao, R. P. Kiene, and T. S. Bates. 1996. Impact of dimethylsulfide
photochemistry on methyl sulfur cycling in the equatorial Pacific Ocean. J. Geophys. Res. 101: 3715-3722. doi:10.1029/95JC03624
Kieber, D. J., D. Toole, J. Jankowski, R. P. Kiene, G. Westby, D. del Valle, and D. Slezak. 2007. Chemical “light meters” for photochemical and photobiological studies. Aquat. Sci. 69: 360-376. doi:10.1007/s00027-007-0895-0
Kiene, R. P., and B. F. Taylor. 1989. Metabolism of acrylate and 3-mercaptopropionate. p. 222-230. In E. S. Saltzman and W. J. Cooper [eds.], Biogenic sulfur in the environment, vol. 393. American Chemical Society.
Kiene, R. P., and G. Gerard. 1994. Determination of trace levels of dimethylsulfoxide (DMSO) in seawater and rainwater. Mar. Chem. 47: 1-12. doi:10.1016/0304-4203(94)90009-4 Kiene, R. P., and L. P. Hoffmann Williams. 1998. Glycine betaine uptake, retention, and
degradation by microorganisms in seawater. Limnol. Oceanogr. 43: 1592-1603. doi:10.4319/lo.1998.43.7.1592
Kiene, R. P., and L. J. Linn. 2000. Distribution and turnover of dissolved DMSP and its
relationship with bacterial production and dimethylsulfide in the Gulf of Mexico. Limnol. Oceanogr. 45: 849-861. doi:10.4319/lo.2000.45.4.0849
Kiene, R. P., L. J. Linn, and J. A. Bruton. 2000. New and important roles for DMSP in marine microbial communities. J. Sea Res. 43: 209-224. doi: 10.1016/S1385-1101(00)00023-X Kiene, R. P., and D. Slezak. 2006. Low dissolved DMSP concentrations in seawater revealed by
small-volume gravity filtration and dialysis sampling. Limnol. Oceanogr.: Meth. 4: 80-95. doi:10.4319/lom.2006.4.80
Kiene, R., D. J. Kieber, D. Slezak, D. Toole, D. del Valle, J. Bisgrove, J. Brinkley, and A.
Rellinger. 2007. Distribution and cycling of dimethylsulfide, dimethylsulfoniopropionate, and dimethylsulfoxide during spring and early summer in the Southern Ocean south of New Zealand. Aquat. Sci. 69: 305-319. doi:10.1007/s00027-007-0892-3
Kinsey, J. D., and D. J. Kieber. 2016. Microwave preservation method for DMSP, DMSO, and acrylate in unfiltered seawater and phytoplankton culture samples. Limnol. Oceanogr. Methods 14: 196-209. doi:10.1002/lom3.10081
Kinsey, J. D., D. J. Kieber, and P. J. Neale. 2016. Effects of iron limitation and UV radiation on Phaeocystis antarctica growth and dimethylsulfoniopropionate, dimethylsulfoxide and acrylate concentrations. Environ. Chem. 13: 195-211. doi:10.1071/EN14275
Accepted Article
Ledyard, K. M., E. F. DeLong, and J. W. H. Dacey. 1993. Characterization of a DMSP-degrading bacterial isolate from the Sargasso Sea. Arch. Microbiol. 160: 312-318. doi: 10.1007/BF00292083
Liu, Y., C.-Y. Liu, G.-P. Yang, H.-H. Zhang, and S.-H. Zhang. 2016. Biogeochemistry of dimethylsulfoniopropionate, dimethylsulfide and acrylic acid in the Yellow Sea and the Bohai Sea during autumn. Environ. Chem. 13: 127-139. doi:10.1071/EN15025
Malmstrom, R. R., R. P. Kiene, and D. L. Kirchman. 2004. Identification and enumeration of bacteria assimilating dimethylsulfoniopropionate (DMSP) in the North Atlantic and Gulf of Mexico. Limnol. Oceanogr. 49: 597-606. doi:10.4319/lo.2004.49.2.0597
Moran, M. A., C. R. Reisch, R. P. Kiene, and W. B. Whitman. 2012. Genomic insights into bacterial DMSP transformations. Annu. Rev. Mar. Sci. 4: 523-542. doi:10.1146/annurev-marine-120710-100827
Motard-Côté, J., D. J. Kieber, A. Rellinger, and R. P. Kiene. 2016. Influence of the Mississippi River plume and non-bioavailable DMSP on dissolved DMSP turnover in the northern Gulf of Mexico. Environ. Chem. 13: 280-292. doi:10.1071/EN15053
Nagata, T. 2008. Organic matter-bacteria interactions in seawater, p. 207-241. In D. Kirchman [ed.], Microbial ecology of the oceans, 2nd Edition. Wiley.
doi:10.1002/9780470281840.ch7
Noordkamp, D. J. B., W. W. C. Gieskes, J. C. Gottschal, L. J. Forney, and M. van Rijssel. 2000. Acrylate in Phaeocystis colonies does not affect the surrounding bacteria. J. Sea Res. 43: 287-296. doi:10.1016/S1385-1101(00)00021-6
Osinga, R., R. L. J. Kwint, W. E. Lewis, G. W. Kraay, J. D. Lont, H. J. Lindeboom, and F. C. van Duyl. 1996. Production and fate of dimethylsulfide and dimethylsulfoniopropionate in pelagic mesocosms: The role of sedimentation. Mar. Ecol. Prog. Ser. 131: 275-286. doi:10.3354/meps131275
Raina, J.-B., D. Tapiolas, B. L. Willis, and D. G. Bourne. 2009. Coral-associated bacteria and their role in the biogeochemical cycling of sulfur. Appl. Environ. Microbiol. 75: 3492-3501. doi:10.1128/AEM.02567-08
Rich, J. H., H. W. Ducklow, and D. L. Kirchman. 1996. Concentrations and uptake of neutral monosaccharides along 140°W in the equatorial Pacific: Contribution of glucose to heterotrophic bacterial activity and the DOM flux. Limnol. Oceanogr. 41: 595-604. doi:10.4319/lo.1996.41.4.0595
Ridgeway, R. G., D. C. Thornton, and A. R. Bandy. 1992. Determination of trace aqueous dimethylsulfoxide concentrations by isotope dilution gas chromatography/mass spectrometry: Application to rain and sea water. J. Atmos. Chem. 14: 53-60. doi:10.1007/BF00115222
Accepted Article
Simó, R., J. O. Grimalt, C. Pedrós-Alió, and J. Albaiges. 1995. Occurrence and transformation of dissolved dimethyl sulfur species in stratified seawater (western Mediterranean Sea). Mar. Ecol. Prog. Ser. 127: 291-299. doi:10.3354/meps127291
Simó, R., J. O. Grimalt, and J. Albaigés. 1997. Dissolved dimethylsulphide,
dimethylsulphoniopropionate and dimethylsulphoxide in western Mediterranean waters. Deep Sea Res. II 44: 929-950. doi:10.1016/S0967-0645(96)00099-9
Simó, R., C. Pedrós-Alió, G. Malin, and J. O. Grimalt. 2000. Biological turnover of DMS, DMSP and DMSO in contrasting open-sea waters. Mar. Ecol. Prog. Ser. 203: 1-11. doi:10.3354/meps203001
Simon, M., and F. Azam. 1989. Protein content and protein synthesis rates of planktonic marine bacteria. Mar. Ecol. Prog. Ser. 51: 201-213. doi:10.3354/meps051201
Skoog, A., B. Biddanda, and R. Benner. 1999. Bacterial utilization of dissolved glucose in the upper water column of the Gulf of Mexico. Limnol. Oceanogr. 44: 1625-1633.
doi:10.4319/lo.1999.44.7.1625
Smith, D. C., and F. Azam. 1992. A simple, economical method for measuring bacterial protein synthesis in seawater using 3H-leucine. Mar. Microb. Food Webs 6: 107-114.
doi:10.1037//0893-164X.6.2.107
Spiese, C. E., D. J. Kieber, and C. T. Nomura. 2009. Reduction of dimethylsulfoxide to dimethylsulfide by marine phytoplankton. Limnol. Oceanogr. 54: 560-570. doi:10.4319/lo.2009.54.2.0560
Spiese, C. E., T. Le, R. L. Zimmer, and D. J. Kieber. 2016. Dimethylsulfide membrane permeability, cellular concentrations and implications for physiological functions in marine algae. J. Plankton Res. 38: 41-54. doi:10.1093/plankt/fbv106
Sunda, W., D. J. Kieber, R. P. Kiene, and S. Huntsman. 2002. An antioxidant function for DMSP and DMS in marine algae. Nature 418: 317-320. doi:10.1038/nature00851
Suttle, C. A., A. M. Chan, and J. A. Fuhrman. 1991. Dissolved free amino acids in the Sargasso Sea: uptake and respiration rates, turnover times, and concentrations. Mar. Ecol. Prog. Ser. 70: 189-199. doi:10.3354/meps070189
Todd, J. D., R. Rogers, Y. G. Li, M. Wexler, P. L. Bond, L. Sun, A. R. J. Curson, G. Malin, M. Steinke, and A. W. B. Johnston. 2007. Structural and regulatory genes required to make the gas dimethyl sulfide in bacteria. Science 315: 666-669. doi:10.1126/science.1135370 Todd, J. D., A. R. J. Curson, N. Nikolaidou-Katsaraidou, C. A. Brearley, N. J. Watmough, Y.
Chan, P. C. B. Page, L. Sun, and A. W. B. Johnston. 2010. Molecular dissection of bacterial acrylate catabolism – unexpected links with dimethylsulfoniopropionate catabolism and dimethyl sulfide production. Environ. Microbiol. 12: 327-343. doi: 10.1111/j.1462-2920.2009.02071.x
Tolli, J. D., and C. D. Taylor. 2005. Biological CO oxidation in the Sargasso Sea and in Vineyard Sound, Massachusetts. Limnol. Oceanogr. 50: 1205-1212.
Accepted Article
Toole, D. A., D. J. Kieber, R. P. Kiene, D. A. Siegel, and N. B. Nelson. 2003. Photolysis and the dimethylsulfide (DMS) summer paradox in the Sargasso Sea. Limnol. Oceanogr. 48: 1088-1100. doi:10.4319/lo.2003.48.3.1088
Tyssebotn, I. M. B. 2015. An investigation of the marine acrylate cycle. Ph.D. thesis. State University of New York, College of Environmental Science and Forestry.
Vairavamurthy, A., M. O. Andreae, and J. M. Brooks. 1986. Determination of acrylic acid in aqueous samples by electron capture gas chromatography after extraction with tri-n-octylphosphine oxide and derivatization with pentafluorobenzyl bromide. Anal. Chem. 58: 2684-2687. doi:10.1021/ac00126a023
van Alstyne, K. L., G. V. Wolfe, T. L. Freidenburg, A. Neill, and C. Hicken. 2001. Activated defense systems in marine macroalgae: Evidence for an ecological role for DMSP cleavage. Mar. Ecol. Prog. Ser. 213: 53-65. doi:10.3354/meps213053
Vila, M., R. Simó, R. P. Kiene, J. Pinhassi, J. M. González, M. A. Moran, and C. Pedrós-Alió. 2004. Use of microautoradiography combined with fluorescence in situ hybridization to determine dimethylsulfoniopropionate incorporation by marine bacterioplankton taxa. Appl. Environ. Microbiol. 70: 4648-4657. doi: 10.1128/AEM.70.8.4648-4657.2004 Welschmeyer, N. A. 1994. Fluorometric analysis of chlorophyll a in the presence of chlorophyll
b and pheopigments. Limnol. Oceanogr. 39: 1985-1992. doi:10.4319/lo.1994.39.8.1985 Wolfe, G. V., and R. P. Kiene. 1993. Radioisotope and chemical inhibitor measurements of
dimethyl sulfide consumption rates and kinetics in estuarine waters. Mar. Ecol. Prog. Ser. 99: 261-269.
Xie, H., O. C. Zafiriou, T. P. Umile, and D. J. Kieber. 2005. Biological consumption of carbon monoxide in Delaware Bay, NW Atlantic and Beaufort Sea. Mar. Ecol. Prog. Ser. 290: 1-14. doi: 10.3354/meps290001
Accepted Article
Tables
Table 1 Hydrographic stations, water mass properties, and acrylated and DMSOd concentrations (mean ± SD, n = 3) in surface waters
for the cruise aboard the R/V Pelican in the northern Gulf of Mexico, 12-22 September 2011. ND denotes not determined.
Station Latitude (°N) Longitude (°W) Sampling depth (m) Salinity (ppt) Temp (°C) a300 (m-1) Chl a (µg L-1) Acrylateda (nmol L-1) DMSOda (nmol L-1) Oceanic sites 8 26.023 90.998 15.4 36.0 29.2 0.192 0.06 1.4 ± 0.2 9.8 ± 0.01 6 27.888 91.003 14.6 36.3 29.1 0.269 0.10 1.6 ± 0.1 5.5 ± 0.5 4 28.163 90.482 15.4 36.5 28.5 0.301 0.09 1.3 ± 0.1 5.9 ± 0.2 14 28.671 90.502 15.5 34.9 28.2 0.368 0.25 1.6 ± 0.2 7.3 ± 0.3 23 28.467 92.227 15.0 36.0 28.3 0.436 ND 2.1b 6.1 ± 0.2 19 28.355 90.785 4.0 35.3 28.2 0.651 0.27 1.2 ± 0.2 7.2 ± 0.7 Coastal sites 2 28.554 90.532 16.2 32.8 27.4 1.52 2.8 2.0 ± 0.3 6.4 ± 0.6 9 28.670 89.000 2.2 31.9 28.2 1.68 0.40 1.5b 13.8 ± 0.6 17 28.705 90.007 5.2 31.2 27.9 1.94 0.18 1.5 ± 0.3 8.0 ± 0.9 20 28.957 90.786 0.7 29.9 28.3 2.53 ND 0.8 ± 0.1 8.5 ± 0.7 a
Concentrations were determined from the measurement of three separate samples collected from a Niskin bottle.
b
