Present addresses: Lydie Herfort, Environmental & Biomolecular Systems, OGI School of Science and Engineering, Oregon Health & Science University, 20000 NW Walker Rd, Beaverton, OR 97006, USA. Marco J. L. Coolen, Woods Hole Oceanographic Institution, Department of Chemistry and Geochemistry, 360 Woods Hole Rd, MA 02543, USA.
Variations in spatial and temporal distribution of Archaea in the North Sea in relation to environmental variables
Article first published online: 6 NOV 2007
FEMS Microbiology Ecology
Volume 62, Issue 3, pages 242–257, December 2007
How to Cite
Herfort, L., Schouten, S., Abbas, B., Veldhuis, M. J. W., Coolen, M. J. L., Wuchter, C., Boon, J. P., Herndl, G. J. and Sinninghe Damsté, J. S. (2007), Variations in spatial and temporal distribution of Archaea in the North Sea in relation to environmental variables. FEMS Microbiology Ecology, 62: 242–257. doi: 10.1111/j.1574-6941.2007.00397.x
Editor: Michael Wagner
- Issue published online: 6 NOV 2007
- Article first published online: 6 NOV 2007
- Received 28 February 2007; revised 17 June 2007; accepted 13 August 2007.First published online November 2007.
- North Sea;
- Top of page
- Materials and methods
- Results and discussion
- Summary and conclusions
The spatial and temporal distribution of pelagic Archaea was studied in the southern North Sea by rRNA hybridization, sequencing and quantification of 16S rRNA gene and membrane lipid analyses and related to physical, chemical and biological parameters to determine the factors influencing archaeal biogeography. A clear temporal variability was observed, with marine Crenarchaeota (Group I.1a) being relatively more abundant in winter and Euryarchaeota dominating the archaeal assemblage in spring and summer. Spatial differences in the lateral distribution of Crenarchaeota were also evident. In fact, their abundance was positively correlated with the copy number of the gene encoding the α subunit of crenarchaeotal ammonia monooxygenase (amoA) and with concentrations of ammonia, nitrate, nitrite and phosphorus. This suggests that most Crenarchaeota in the North Sea are nitrifiers and that their distribution is determined by nutrient concentrations. However, Crenarchaeota were not abundant when larger phytoplankton (>3 μm) dominated the algal population. It is hypothesized that together with nutrient concentration, phytoplankton biomass and community structure can predict crenarchaeotal abundance in the southern North Sea. Euryarchaeotal abundance was positively correlated with chlorophyll a concentrations, but not with phytoplankton community structure. Whether this is related to the potential of Euryarchaeota to perform aerobic anoxygenic phototrophy remains to be shown, but the conspicuous seasonal distribution pattern of Crenarchaeota and Euryarchaeota suggests that they occupy a different ecological niche.
- Top of page
- Materials and methods
- Results and discussion
- Summary and conclusions
For decades, Archaea were thought to only dwell in extreme environments hostile to Eukarya and Bacteria, but in the early 1990s, using rRNA hybridization and cloning and sequencing approaches, Fuhrman et al. (1992) and DeLong (1992) detected planktonic Archaea in the marine-oxygenated water column. Since then, Euryarchaeota and Crenarchaeota, the two main groups of marine Archaea, have been found in many different oceanic regions (DeLong et al., 1994; Murray et al., 1999; Crump & Baross, 2000; Pernthaler et al., 2002). In fact, Crenarchaeota seem ubiquitous and abundant, constituting about 20–30% of total prokaryotic abundance in the aphotic zone of the global ocean, while Euryarchaeota are dominating the archaeal assemblage in surface waters (Massana et al., 2000; Karner et al., 2001; Herndl et al., 2005). These differences in the relative contribution of Crenarchaeota and Euryarchaeota on the total archaeal biomass between surface and deeper waters suggest that Euryarchaeota and Crenarchaeota probably occupy different ecological niches.
Crenarchaeotal chemoautotrophy, whereby dissolved inorganic carbon (DIC) is used as a carbon source, was first suggested from compound-specific stable isotope and radiocarbon analyses of their specific membrane lipids (Hoefs et al., 1997; Pearson et al., 2001) and was confirmed by in situ labeling (Wuchter et al., 2003) and microautoradiography experiments (Herndl et al., 2005). However, interestingly, evidence for the uptake of dissolved organic substrates, including amino acid, also exists (Herndl et al., 2005; Ingalls et al., 2006; Teira et al., 2006), suggesting a chemoheterotrophic or at least mixotrophic capability. In fact, based on the natural distribution of radiocarbon in their membrane lipids, Ingalls et al. (2006) were able to quantitatively partition autotrophy and heterotrophy in pelagic Crenarchaeota of the subtropical North pacific Gyre, with crenarchaeotal autotrophy representing 83% at 670 m. Recent findings have also suggested that Crenarchaeota are involved in nitrification. Crenarchaeotal genes for putative ammonia monooxygenase A (amoA) catalyzing the oxidation of ammonia to hydroxylamine, which is then further converted to nitrite, have been found to be present in seawater from widely different geographic regions (Venter et al., 2004; Francis et al., 2005). The actual ability of, at least, some Crenarchaeota to oxidize ammonia to nitrite was clearly established in pure (Könneke et al., 2005) and enriched cultures of Crenarchaeota and in field samples (Hallam et al., 2006; Wuchter et al., 2006, Mincer et al., 2007). At present, the metabolic requirements of Euryarchaeota remain enigmatic. It is clear that they are also able to assimilate amino acid (Herndl et al., 2005; Teira et al., 2006), thus suggesting heterotrophy. However, in North Atlantic deep waters up to 23% of euryarchaeotal cells took up DIC (Herndl et al., 2005). Thus, in common with Crenarchaeota, there is evidence for chemoautotrophy and heterotrophy also for Euryarchaeota. Furthermore, very recently, genes encoding proteorhodopsin, a protein that catalyzes light-driven proton transfer across the cell membrane, have been detected in Euryarchaeota living in the photic zone but not in the deep waters of the North Atlantic Subtropical Gyre (Frigaard et al., 2006), hence indicating a potential for phototrophy.
Seasonal dynamics in archaeal abundance have been described for several marine environments (Murray et al., 1998; Church et al., 2003; Wuchter et al., 2006). An extensive time-series data set obtained from the coastal southern North Sea surface waters showed that Crenarchaeota dominated the archaeal community from autumn to early spring, while Euryarchaeota were more abundant in summer and early autumn (Wuchter, 2006; Wuchter et al., 2006). In addition, peaks in crenarchaeotal cell numbers coincided with high concentrations of ammonium in seawater and also with high copy numbers of the crenarchaeotal genes encoding the alpha subunit of the putative ammonia monooxygenase (amoA). This distinct seasonal distribution pattern indicates that marine Crenarchaeota may be involved in nitrification in the coastal southern North Sea (Wuchter et al., 2006). Because the study of Wuchter et al. (2006), which focused on Crenarchaeota, was only carried out at one site in the southern North Sea located in the Marsdiep tidal inlet between the southern North Sea and the western Wadden Sea, a more extensive sampling in the southern North Sea was performed to fully assess temporal and spatial differences in the archaeal community in this region. The distribution of Euryarchaeota and Crenarchaeota in surface and bottom waters was examined at eight different stations in the southern North Sea and this was related to physical, chemical and biological parameters. The sampling time was carefully chosen to best represent the seasonality in the region; February characterizing the winter time, with low water temperatures, and low chlorophyll and nutrient concentrations; April typifying the peak of the algal bloom, with still relatively low water temperatures, high phytoplankton abundances and lower nutrient concentrations; and August exemplifying the late summer period, with elevated water temperatures, phytoplankton abundances varying with sites and low nutrient concentrations.
Materials and methods
- Top of page
- Materials and methods
- Results and discussion
- Summary and conclusions
Sampling in the southern North Sea was carried out at eight stations (Fig. 1) during three cruises, with the R.V. Pelagia in February 2003 and April 2004 and with the R.V. Alkor in August/September 2004.
All sampling was conducted following the time-tested and standardized instructions of experienced NIOZ personnel. All analyses were conducted at the NIOZ by the same experienced personnel who routinely perform these measurements at the institute. Seawater salinities and temperatures were obtained using a conductivity–temperature–depth sensor. To determine nutrient concentrations, 3 mL of freshly collected seawater was passed through a 0.2-μm Acrodisc filter and kept frozen until processed. These were later analyzed colorimetrically using a segmented continuous flow analyzer (TRAACS 800 autoanalyzer, Bran & Luebbe). To evaluate field precision, 10 nutrient replicates bottles were collected and measured during the initial winter cruise at the first station (Dutch Coast). This showed that the variability was <4% for all nutrients. Given the typical low nutrient concentrations found in summer, all samples were collected and measured in duplicate in August. For quantification of dissolved organic nitrogen and phosphorus (DON and DOP), 25 mL of seawater was filtered through a 0.2-μm Acrodisc filter and kept frozen until analysis. Total dissolved phosphorus was estimated by acidified persulfate UV-destruction, while total dissolved nitrogen was analyzed by alkaline persulfate UV-destruction. DON and DOP were calculated as the difference between total and inorganic nitrogen and phosphorus, respectively. For dissolved organic carbon (DOC) determinations, 8 mL of seawater was directly added to a combusted glass ampoule and immediately acidified with a few drops of 45% H3PO4, sealed and kept at −20 °C until analysis. DOC concentrations were determined by high-temperature catalytic oxidation on a modified Total Organic Carbon Analyzer (Shimadzu TOC-5000A) linked to an external infrared cell (LiCor Model LI-6252) for CO2 detection. As quality control of the DOC determinations, the Consensus Reference Material (Batch 4) provided by Hansell and Chen (University of Miami) was used. Duplicate measurements of DOC were performed. For DIC measurements, seawater was collected into 2-mL glass bottles and stored at 4 °C until analysis. DIC concentrations were determined on the TRAACS 800 autoanalyzer (Bran & Luebbe). For chlorophyll a measurements, 100–500 mL of seawater was filtered through a GF/F filter (Whatman, 25 mm filter diameter) and kept frozen until analysis. Following extraction in 90% acetone in the dark at −20 °C for 24 h, chlorophyll a concentrations were determined on a fluorometer F-2000 (Hitachi). Flow cytometry analyses of phytoplankton were performed in duplicate and performed on a Coulter XL-MCL as described in Veldhuis & Kraay (2004).
Crenarchaeotal membrane lipids
Suspended particulate organic matter for lipid analysis was collected by filtering 40 L of seawater onto preashed 0.7-μm pore-size glass fiber filters (47 mm, GF8 Schleicher & Schuell). These were kept frozen at −20 °C until extraction. During the first cruise, seawater was sequentially filtered onto a 0.7-μm Glass Fiber filter and a 0.2-μm Cellulose Acetate filter. This showed that <5% of the total amount of archaeal membrane lipids was recovered with the 0.2-μm filter. Hence, given that the cellulose acetate filters tend to cause problems with the HPLC/MS analysis, the 0.2-μm filtration stage was subsequently omitted. Freeze-dried filters were processed and measured as described in Herfort et al. (2006). Only peak areas of the isoprenoid glycerol dialkyl glycerol tetraethers found in the membrane of Crenarchaeota were integrated (m/z 1302.3, 1300.3, 1298.3, 1296.3, 1292.3) and used to give an estimate of crenarchaeotal membrane lipid concentrations. Peak areas were always at least one order of magnitude higher than background noise. Calibration was performed by running along with the samples known concentrations of purified crenarchaeol.
Catalyzed reported deposition FISH (CARD-FISH)
Samples for CARD-FISH and 4′,6′-diamidino-2-phenylindole (DAPI) counts were only taken during the April and August cruises. Seawater (15 mL) was fixed with formaldehyde (final concentration 4%) at room temperature for 1 h and filtered onto 0.2-μm pore-size polycarbonate filters (Millipore, 25 mm diameter) using 0.45-μm pore-size cellulose nitrate supporting filters. Archaea were stained using the improved CARD-FISH protocol described by Teira et al. (2004). Probes specific for Group I.1a Crenarchaeota, Cren537 (5′-TGACCACTTGAGGTGCTG-3′) and Group II Euryarchaeota, Eury806 (5′-CACAGCGTTTACACCTAG-3′), were used (Teira et al., 2004). Both probes were tested for their specificity before this study. Picoplankton cells were counterstained with DAPI (Teira et al., 2004). Archaea and Bacteria were counted using an epifluorescence microscope (Zeiss Axioplan 2) equipped with a 100 W Hg lamp as well as the appropriate filter sets for DAPI and Alexa488 fluorescence. To determine archaeal and total cell abundances, a minimum of 200 cells were counted per filter. The average counting error in cell abundance, expressed as a percentage of SE, was 26% for DAPI staining and Euryarchaeota and 40% for Crenarchaeota. The larger average counting error for Crenarchaeota is associated with extremely low cell numbers at the North Frisian Front in April.
Nucleic acids extraction
For all DNA analyses, 1 L of seawater was filtered onto 0.2-μm pore-size polycarbonate filters (47 mm, Schleicher & Schuell) and stored at −80 °C until extraction. Total DNA was extracted according to Wuchter (2006). This extraction yielded a cell lysis efficiency of about 90% (see Wuchter et al., 2006 for more details).
Archaeal 16S rRNA gene
Partial archaeal 16S rRNA genes (420 bp) were amplified using the general archaeal PCR primers Parch 519f (5′-CAG CCGCCGCGGTAA-3′) and Arch915r (5′-GTGCTCCCCCG CCAATTCCT-3′) with the protocol described previously by Coolen et al. (2004). Partial archaeal amoA genes (256 bp) were amplified by PCR using the specific primers developed by Wuchter et al. (2006), Arc-amoA-for (5′-CTGAYTGGGC YTGGACATC-3′) and Arch-amoA-rev (5′-TTCTTCTTTGT TGCCCAGTA-3′) and using the same thermo cycling conditions described in Wuchter et al. (2006). All amplicons were separated according to the GC content and secondary structure by denaturing gradient gel electrophoresis (DGGE) using a linear denaturing gradient of 30–60% for 6 h at 200 V for Archaea according to Coolen et al. (2004) and of 10–50% at 200 V for 3 h for archaeal amoA DGGE (Wuchter et al., 2006). Gels were stained for 20 min with SYBR gold and documented with a Fluor-S Multi Imager (BioRad). DGGE bands were excised and each one was eluted in sterile 10 mM Tris-HCl (pH 8.0) at 4 °C for 24 h. Sequencing reactions were performed as described in Wuchter et al. (2006).
Following DNA quantification with PicoGreen, reamplified DGGE bands were processed as described Wuchter et al. (2003). Nucleotide sequences were determined by automated sequencing using a 310 Genetic Analyzer capillary sequencer (Applied Biosystem).
Sequence data were compiled using arb software (Ludwig et al., 2004) and aligned with complete length sequences of closest relatives obtained from the National Centre for Biotechnology Information (NCBI) database (http://www.ncbi.nlm.nih.gov/) using the arb FastAligner utility. Using arb, the phylogenetic trees of Figs 3 and 5 were first generated with the aligned, almost complete length sequences of closest relatives from the NCBI database using the neighbor-joining method (Saitou & Nei, 1987) and the Jukes & Cantor (1969) correction. Then, the short-aligned DGGE sequences were added to the trees using the maximum parsimony option implemented in arb. The sequences obtained in this study have all been deposited in NCBI (accession numbers EU239431 to EU239463 and EU244456 to EU244461).
Real-time quantitative PCR
Copy numbers of the archaeal amoA gene were quantified using the above-mentioned primer sets. For the quantification of crenarchaeotal 16S rRNA gene, the specific primers used by Wuchter et al. (2006) (122 bp) were employed: MCG-1 391F (5′-AAGGTTARTCCGAGTGRTTTC-3′) and MCG-1 554R (5′-TGACCACTTGAGGTGCTG-3′). qPCR was carried out in an iCycler system (BioRad, Hercules, CA) as described by Wuchter et al. (2006). Calibration was performed by running a qPCR along with the samples of known copy numbers (between 10−2 and 107) of PCR-amplified crenarchaeotal 16S rRNA gene from the North Sea enrichment culture (Wuchter et al., 2006) or archaeal amoA also originating from the North Sea enrichment culture (Wuchter et al., 2006). Negative controls and blanks were also subjected to qPCR. Aliquots of these qPCR products were run on an agarose gel in order to identify unspecific PCR products such as primer dimers or bands with unexpected fragment lengths. Additional proof for the specificity of the amoA primers used during qPCR came from sequence analysis of amoA-DGGE bands (as described above) that were generated during PCR using the same primer set.
Data were divided into two main groups, surface and bottom waters, within which statistical analysis was carried out over the whole data set and for each month. Abundances of Crenarchaeota and Euryarchaeota were compared with each other and with other environmental variables by Spearman's rank correlation coefficient (rs) analyses. Only significant correlations (P<0.05) are reported.
Results and discussion
- Top of page
- Materials and methods
- Results and discussion
- Summary and conclusions
Archaeal seasonal and spatial distribution
The southern North Sea is a highly dynamic shallow shelf sea (maximum depth 50 m) characterized in most parts by a mixed water column throughout the year. This is reflected in the DGGE banding pattern, which was very similar at all depths for all sites, indicating a similar archaeal community composition in surface and bottom waters (Fig. 2). This was even true for the Oyster Grounds in August where the water column was thermally stratified (Tables 1 and 2). It is, however, important to note that the lack of saline stratification (Tables 2 and 3) suggests that the thermal stratification was a recent event probably due to the heavy storms that occurred in the southern North Sea in August 2004. However, DGGE fingerprinting of PCR-amplified archaeal 16S rRNA gene clearly indicated a seasonal variation in archaeal distribution in the southern North Sea. Most recovered and sequenced DGGE bands from February 2003 belonged to the Group I.1a Crenarchaeota, while in April and August 2004, the Group II Euryarchaeota dominated the archaeal assemblage in this region (Figs 2 and 3). This overall seasonal pattern is consistent with the results of the time-series analysis of Wuchter (2006) and Wuchter et al. (2006), who found a similar succession in archaeal phylotypes in the coastal southern North Sea. This also agrees with the FISH results obtained in the German Bay of the North Sea by Pernthaler et al. (2002). DGGE bands affiliated to the Group II Euryarchaeota were also retrieved in February at the stations Dutch Coast and Breeveertien (Figs 2 and 3). Several water masses converge in this part of the southern North Sea, with Continental Coastal and Channel waters present at the Dutch Coast and Breeveertien, respectively (Lee, 1980). In February, salinity values of 29.7 and 33.7 for the Dutch Coast and Breeveertien (Table 1), respectively, indicate that at the Dutch Coast station Continental Coastal water mass was present, while at the Breeveertien station Channel and Continental Coastal water masses were mixed. Euryarchaeota Group II were only recovered at these two sites in February 2003, illustrating the low contribution of Euryarchaeota to the coastal archaeal community in winter. Interestingly, in contrast to the results of Wuchter (2006) and Wuchter et al. (2006), DGGE bands affiliated to the Group I.1a Crenarchaeota were obtained in April and August 2004 at the Central Southern Bight and the Frisian Front (Figs 2 and 3), indicating spatial heterogeneity in the distribution of Crenarchaeota.
|Archaeal membrane lipids (ng L−1)||12.4||NA||13.5||2.1||6.5||1.2||4.0||2.0||0.6||1.4||10.9||3.0||8.3||0.7||0.3||0.3||1.1||0.5||2.7||0.9||3.3||0.9||0.2||0.7|
|DAPI (105 cells mL−1)||NA||NA||NA||NA||NA||NA||NA||NA||8||14||5||7||4||3||2||6||19||12||5||14||6||7||8||12|
|Euryarchaeota (103 cells mL−1)||NA||NA||NA||NA||NA||NA||NA||NA||18||18||9||25||7||2||1||1||17||4||7||27||6||9||2||1|
|Crenarchaeota (103 cells mL−1)||NA||NA||NA||NA||NA||NA||NA||NA||ND||ND||1.8||ND||4.7||0.4||ND||ND||ND||ND||1.6||ND||1.4||ND||ND||ND|
|Crenarchaeotal 16S rRNA gene (103 copies mL−1)||2.1||1.5||12.1||2.0||6.6||12.8||7.4||0.3||0.02||0.3||5.1||0.01||11.7||1.1||0.1||0.3||0.01||0.1||1.6||0.03||8.0||0.4||0.01||0.03|
|Crenarchaeotal amoA (103 copies mL−1)||7.6||5.4||53.0||7.7||18.1||32.8||20.2||1.0||0.1||0.6||13.6||0.1||55.0||10.5||1.3||0.2||0.04||0.1||5.6||0.1||29.2||0.2||0.1||0.1|
|Chlorophyll a (μg L−1)||0.3||0.5||0.2||0.1||0.5||0.2||0.2||0.5||2.3||1.6||1.1||2.3||0.3||0.2||0.8||0.4||4.1||0.9||1.7||2.5||1.2||1.0||0.4||0.5|
|DOC (μmol L−1)||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||140||97||103||107||94||90||93||95|
|DON (μmol L−1)||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||10||48||7||4||7||7||5||10|
|DOP (μmol L−1)||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||0.18||2.43||0.16||ND||0.08||0.13||0.16||0.01|
|Ammonium (μmol L−1)||3.0||0.7||0.1||2.0||1.4||0.3||0.2||0.3||0.9||0.4||0.4||0.3||0.7||0.3||0.3||0.2||0.3||0.2||1.3||0.3||2.4||0.4||0.2||0.1|
|Nitrate (μmol L−1)||45.8||18.2||10.6||40.1||31.2||11.3||8.1||5.0||12.4||2.6||10.0||4.8||8.2||4.0||3.1||0.03||0.1||0.1||3.8||0.1||0.8||0.1||0.02||0.02|
|Nitrite (μmol L−1)||0.42||0.22||0.07||0.40||0.30||0.12||0.05||0.08||0.22||0.06||0.17||0.12||0.23||0.13||0.17||0.01||0.01||0.00||0.39||0.00||0.23||0.01||0.00||0.00|
|Phosphate (μmol L−1)||0.85||0.57||0.66||0.84||0.74||0.60||0.55||0.45||0.03||0.02||0.46||0.21||0.54||0.39||0.37||0.15||0.14||0.05||0.16||0.06||0.31||0.18||0.07||0.06|
|DIC (mmol L−1)||2.16||2.10||2.08||2.16||2.13||2.09||2.08||2.07||2.12||2.14||2.18||2.13||2.19||2.19||2.19||2.17||2.14||2.13||2.13||2.11||2.12||2.14||2.11||2.12|
|Archacal membrane lipids (ng L−1)||NA||NA||17.1||5.9||12.3||1.4||4.7||2.3||4.5||3.5||13.5||4.3||16.2||1.9||5.8||0.9||2.9||0.7||5.5||1.9||22.0||2.2||3.2||0.4|
|Crenarchaeotal 16S rRNA gene (103 copies mL−1)||7.4||10.7||0.3||12.8||NA||0.8||20.8||3.4||0.1||0.2||8.3||0.2||3.7||1.7||2.1||0.02||0.1||0.1||3.2||0.2||5.6||0.4||2.4||0.05|
|Crenarchaeotal amoA (103 copies mL−1)||28.5||37.8||1.4||50.0||NA||2.2||73.0||11.3||1.0||0.8||32.2||0.9||19.9||10.7||9.1||0.1||0.3||0.1||9.0||0.3||15.0||0.4||2.4||0.1|
|Chlorophyll a (μg L−1)||0.6||0.7||0.2||0.3||0.2||0.2||0.2||0.6||6.6||1.6||1.8||6.3||0.4||0.2||1.1||0.4||4.7||1.0||2.4||3.2||1.1||0.6||0.4||0.3|
|DOC (μmol L−1)||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||130||96||95||104||97||93||88||91|
|DON (μmol L−1)||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||55||10||7||12||6||3||3||4|
|DOP (μmol L−1)||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||NA||0.33||0.03||0.18||0.29||0.37||ND||0.01||0.03|
|Ammonium (μmol L−1)||0.8||0.7||0.1||0.3||0.2||0.2||0.1||0.3||0.5||0.6||0.7||0.5||0.6||0.8||1.1||0.2||0.3||0.3||1.3||0.4||2.9||0.7||1.7||0.2|
|Nitrate (μmol L−1)||18.8||18.7||10.3||11.7||11.4||7.4||8.5||4.9||2.0||2.7||10.1||4.9||8.0||4.4||4.8||0.03||0.1||0.04||3.3||0.1||1.3||0.1||0.4||0.03|
|Nitrite (μmol L−1)||0.22||0.22||0.06||0.15||0.10||0.07||0.05||0.07||0.08||0.06||0.18||0.12||0.22||0.14||0.19||0.01||ND||ND||0.30||ND||0.36||0.01||0.06||ND|
|Phosphate (μmol L−1)||0.56||0.57||0.63||0.62||0.63||0.58||0.57||0.46||0.08||0.02||0.48||0.21||0.55||0.44||0.51||0.11||0.12||0.05||0.13||0.07||0.57||0.21||0.36||0.14|
|DIC (μmol L−1)||2.09||2.10||2.08||2.10||2.10||2.08||2.08||2.06||2.09||2.15||2.19||2.16||2.19||2.21||2.21||2.17||2.14||2.13||2.14||2.11||2.13||2.14||2.20||2.12|
|Archaeal membrane lipids (ng L−1)||0.71**||0.63**||ND||0.73*||0.85*|
|DAPI (cells mL−1)||NA||NA||NA||NA||NA||NA|
|Crenarchaeota 16S rRNA gene (copies mL−1)||ND||ND||ND||ND||ND||ND||ND||ND|
|Crenarchaeotal amoA (copies mL−1)||0.97**||0.97**||0.97**||0.90**||0.95**||1**||0.88**||0.97**|
|Chlorophyll a (μg L−1)||−0.61*||0.79*||0.88**|
|DOC (μmol L−1)||NA||NA||NA||NA||NA||NA||NA||NA|
|DON (μmol L−1)||NA||NA||NA||NA||NA||NA||NA||NA|
|DOP (μmol L−1)||NA||NA||NA||NA||NA||NA||NA||NA|
|Ammonium (μmol L−1)||0.78*||0.79*||0.95**|
|Nitrate (μmol L−1)||0.65*||0.76**||0.85*||0.90**|
|Nitrite (μmol L−1)||0.56*||0.73*||0.65**||0.85*||0.83**|
|Phosphate (μmol L−1)||0.78*||0.71**||0.76*|
|DIC (mmol L−1)||0.76*|
This temporal and spatial distribution pattern was confirmed by quantitative analyses of crenarchaeotal membrane lipids and 16S rRNA gene, and by direct cell counts with CARD-FISH. Overall, higher concentrations of crenarchaeotal membrane lipids (Sinninghe Damstéet al., 2002) were measured in February than in April and August (Table 1). Crenarchaeotal 16S rRNA gene (copies mL−1) was also two to four times more abundant in February than in April and August at the Central Southern Bight (Table 1). This seasonal occurrence of Crenarchaeota is illustrated by the inverse correlation found between Crenarchaeota abundance and seawater temperature when all three seasons are considered together.
However, in good agreement with the fingerprinting data, all three quantitative approaches revealed a higher abundance of Crenarchaeota during spring and summer in the surface waters of the Central Southern Bight and the Frisian Front than at the other stations (Table 1 and Fig. 4). These temporal and spatial variations were also observed in bottom waters (Table 2). Because at all sites the water column is relatively shallow (<42 m) and well mixed (apart from the Oyster Grounds in August), these data at depth constitute a confirmation of the surface findings of temporal and spatial variations in crenarchaeotal abundances. No rRNA-based analysis was carried out on the underlaying sediments, and hence it cannot be ruled out that Crenarchaeota living in the sediment may have resuspended in the water column at these two sites leading to the observed horizontal heterogeneity. The authors do not, however, believe that it is the case here because water column depths (34–47 m) and sediments characteristics (organic matter content and silk %) are similar at the Oyster Grounds, North Frisian Front, Frisian Front and Central Southern Bight (Herfort et al., 2006) while Crenarchaeota was only detected in the water column at the later two sites. Future studies analyzing together the pelagic and benthic marine archaeal population are nevertheless still necessary to test this. Euryarchaeotal cells were abundant in April and August, amounting up to 2 × 104 cells mL−1 at the Frisian Front and representing up to 4% of the total prokaryotic community estimated by DAPI counts (Table 1). As mentioned above, the seasonal distribution of Crenarchaeota and Euryarchaeota in the surface waters of the North Sea has been described before (Pernthaler et al., 2002; Wuchter, 2006; Wuchter et al., 2006), but to the authors' knowledge, this is the first time that small-scale horizontal heterogeneity in crenarchaeotal abundance is reported. Furthermore, this study has shown that Crenarchaeota can in fact be more abundant in spring and summer than in winter (e.g. Frisian Front, Fig. 4). It is at present uncertain whether high crenarchaeotal abundance is a characteristic feature of the water column at the Frisian Front and Central Southern Bight, because a detailed time-series analysis would be necessary to verify this. However, the observed horizontal heterogeneity in crenarchaeotal abundance gives a unique opportunity that previous studies did not provide to examine factors influencing their distribution.
It is also important to note that all the crenarchaeotal sequences recovered using general archaeal primers during PCR were marine Crenarchaeota, now classified as Group I.1a Crenarchaeota (DeLong, 1998), and shared 94% of sequence similarity, whereas the euryarchaeotal sequences clustered in the Group II Euryarchaeota and shared only 85% sequence similarity. This agrees very well with recent data obtained in the coastal southern North Sea, with a sequence similarity of 97% for Crenarchaeota and 85% for Euryarchaeota (Wuchter, 2006). These results are also consistent with other studies carried out in different oceanic regions indicating that Group II Euryarchaeota are phylogenetically more diverse than Group I.1a Crenarchaeota (Massana et al., 2000; Bano et al., 2004). Moreover, the present data support the conclusions drawn by Massana et al. (2000), who analyzed more than 2000 archaeal rRNA gene clones and showed that only a few phylotypes dominate the oceanic planktonic archaeal assemblage worldwide. Nevertheless, supporting the idea of Dolan (2005) of emerging patterns in microbial biogeography, as explained above, the results demonstrate the existence of small-scale horizontal heterogeneity in archaeal community structure in terms of DGGE-banding patterns, CARD-FISH counts and crenarchaeotal lipids in the southern North Sea. This is consistent with recent studies that have investigated bacterial assemblages within a few kilometers (Suzuki et al., 2001; Pinhassi et al., 2003; Ghiglione et al., 2005). Here, it is shown that these variations are also occurring among archaeal assemblages. Hence, the present data support the emerging view that prokaryotic distribution patterns vary over different spatial scales (Seymour et al., 2005).
To investigate the potential for nitrification among the Crenarchaeota in the eight study sites in the southern North Sea, functional gene analysis of crenarchaeotal amoA was carried out by DGGE and sequencing for all samples that showed high crenarchaeotal abundance, i.e. at all sites in February and at the Central Southern Bight and Frisian Front in April and August. This yielded four highly similar amoA nucleotide sequences (99% sequence homology) that were assigned to the marine Crenarchaeota. The four amoA nucleotide sequences were closely related to the crenarchaeotal amoA sequence detected at one location in the coastal southern North Sea (Wuchter, 2006) (Fig. 5). At the amino acid level (data not shown), the resemblance is even more striking because the sequences and that from the coastal southern North Sea were in fact identical, indicating that nucleotide differences occur on the third base pair of each codon. The crenarchaeotal amoA nucleotide sequences were all closely related, with 94% and 92% sequence homology, to that of the only pelagic Crenarchaeota thus far in pure culture, Candidatus‘Nitrosopumilus maritimus’ (Könneke et al., 2005), and with the enriched culture of a crenarchaeote from the coastal southern North Sea (Wuchter et al., 2006) (Fig. 5). Considering that these cultured crenarchaeotes have been clearly characterized as nitrifiers (Könneke et al., 2005; Wuchter et al., 2006), it seems likely that at least some of the Crenarchaeota of the southern North Sea obtain their energy through nitrification. Interestingly, amoA-like sequences were retrieved at all stations where elevated crenarchaeotal abundances were found, i.e. at all sites in February and at the Central Southern Bight and the Frisian Front in April and August. In addition, quantification of crenarchaeotal amoA by qPCR showed exactly the same seasonal dynamics and spatial heterogeneity as that observed for crenarchaeotal 16S rRNA gene (Tables 1 and 2; Fig. 4). These two qPCR-based data sets were highly correlated (P<0.01) (Table 3), with a slope of 3.4 (r2=0.96, n=48). This indicates that in this study each crenarchaeotal cell contained 3.4 copies of the amoA gene. This estimation should, however, be considered with caution as it is at present unknown whether different ecotypes of Group I.1a Crenarchaeota carry identical numbers of amoA copies per cell. It is nonetheless noteworthy that the estimate is only slightly higher than the two to three copies per cell reported previously by Wuchter et al. (2006) for the coastal North Sea. This slight difference may reflect inaccuracy in the qPCR approach or more probably a variability in amoA copies in Crenarchaeota similar to what has been observed for ammonia-oxidizing bacteria (AOB) (Norton et al., 2002). A good correlation was also found between nutrient concentrations and crenarchaeotal abundances (P<0.05 for nitrate, nitrite and phosphate when all data from the different seasons were pooled) (Table 3). This is in agreement with data obtained by Murray et al. (1999), who found a good correlation between Crenarchaeota and nitrite concentrations in the Santa Barbara Channel. Crenarchaeota and ammonium concentrations were also significantly correlated in August in surface waters and in February and April in bottom waters (Table 3), and the high levels of crenarchaeotal amoA gene copy number measured in August in surface waters were associated with elevated nutrient concentrations (Fig. 6). In August, in the stratified water column of the Oyster Grounds the elevated crenarchaeotal abundances found in bottom waters were also associated with higher nutrient concentrations (Tables 1 and 2). Taken together, all the above-presented data suggest that most of the Crenarchaeota detected in the southern North Sea gain their energy by converting ammonium into nitrite and that this ability to perform nitrification explains their distribution in the southern North Sea. It is important to note that the presence of amoA genes is only an indication of a potential for nitrification and does not alone establish that this process is taking place. Only process-oriented studies, such as mRNAs analysis, can alone prove that crenarchaeotal nitrification is carried out in the marine environment. Unfortunately, the samples were not preserved in such a way that makes amoA transcript analysis possible. However, the authors' reasoning is not only based on the presence of amoA genes but also on the good correlation between amoA genes and crenarchaeotal cell counts and with nutrient concentrations. Archaeal nitrification is now a well-established idea and these results show that this ability to nitrify may explain their seasonal and spatial (horizontal) distribution. It may be that marine Crenarchaeota do have other alternative sources of metabolic energy because for instance concentrations of nitrate, nitrite and even phosphate were more consistently correlated with crenarchaeotal abundance than ammonium (Table 3), but taken together the data suggest that the substrate for nitrification appears to be an important determinant of crenarchaeotal distribution. The inverse correlation observed between temperature and crenarchaeotal abundances when all three seasons are pooled (Table 3) is thus likely the result of the low nutrient concentrations usually found during spring and summer.
There are, however, some exceptions to the general observation made above. In April, elevated ammonium concentrations in surface waters were measured at the stations Dutch Coast, Central Southern Bight and Frisian Front, but Crenarchaeota were only abundant at the two latter sites (Table 1 and Fig. 6). This discrepancy may be explained by the high chlorophyll a concentrations present at the station Dutch Coast (Table 1). A significant inverse relation (P<0.05) was found in surface waters between chlorophyll a concentrations and crenarchaeotal abundances when data from all seasons were pooled (Table 3). Possibly, the predominant phytoplankton bloom present at the Dutch Coast station in April prevented Crenarchaeota from flourishing despite elevated levels of ammonium. An inverse correlation between phytoplankton and Crenarchaeota is a well-established idea because contrasting seasonality between chlorophyll a and Crenarchaeota has been reported previously for different oceanic regions including the coastal southern North Sea (Murray et al., 1998; Wuchter, 2006), but this is the first time that this inverse correlation has been observed in space. Hence, the horizontal heterogeneity in crenarchaeotal abundance observed in this study gave a unique opportunity to indicate that crenarchaeotal abundance is directly or indirectly related to the presence of phytoplankton and nutrient availability in the southern North Sea. This is in agreement with the recent findings of Ward (2005), who studied bacterial nitrification in the Pacific off Monterey Bay and showed that biological interactions, including phytoplankton, had the most impact in determining the composition of the nitrifying bacterial assemblage.
In addition to total phytoplankton biomass, its composition may also be affecting crenarchaeotal abundance. Flow cytometry analyses revealed that in the southern North Sea, a considerable variability in phytoplankton composition existed between sites and seasons (Fig. 7). For example, in April at the Frisian Front, where ammonium concentrations were high and Crenarchaeota was abundant, picophytoplanktons (Synechoccocus, phytoplankton <1.5 μm and phytoplankton of 1.5–3 μm; Fig. 7) represented 40% of all photosynthetic organisms, while at the Dutch Coast, where nutrient levels were high but crenarchaeotal cell numbers were low, the larger size-classes (5 μm) dominated the phytoplankton assemblage. In August, picophytoplankton were abundant at all sites, constituting, on average, 46% of the photosynthetic community. Apparently, Crenarchaeota were thus abundant only at sites with high nutrient levels where larger phytoplankton were not dominating the assemblage. Whether this observation indicates that phytoplankton community structure directly or indirectly influences crenarchaeotal abundance will need to be examined by future experiment-based studies. This observation that Crenarchaeota and picoplankton abundance are correlated is nonetheless surprising because the high surface to volume ratio of picoplankton (prokaryotes' and eukaryotes' component of the phytoplankton community <3 μm) enables very efficient uptake of nutrients (Veldhuis et al., 2005), suggesting a more pronounced competition for nutrients between prokaryotes and picoplankton than with larger phytoplankton. While the direct influence of the phytoplankton community composition on prokaryotic community composition is not yet well known, field observations and experimental evidence suggest that a shift in the phytoplankton community induces changes in the bacterial assemblage (Arrieta & Herndl, 2002; Pinhassi et al., 2004). When Spearman's rank correlation coefficient analysis was performed on pooled April and August data, an inverse correlation (α=0.05) was found between DAPI counts and the copy numbers of crenarchaeotal 16S rRNA gene (rs=−0.626, n=16). Bacterial population may thus, at least in part, influence crenarchaeotal distribution in the southern North Sea. Crenarchaeota might constitute the steady component of the microbial assemblage while Bacteria form the dynamic one and as such control the relative contribution of Crenarchaeota to the total prokaryotic community. Recently, using high-resolution molecular fingerprinting, Fuhrman et al. (2006) have not only demonstrated that the temporal distribution and abundance of the marine microbial taxa of the southern California coast is strongly patterned but also that these patterns can be predicted from abiotic and biotic factors. In this context, based both on the time-series analysis of Wuchter (2006) conducted at one site in the coastal southern North Sea and on the present data set obtained at different locations in the southern North Sea, it is suggested that high nutrient concentrations and relatively low phytoplankton biomass (with perhaps also low percentage of large phytoplankton) may predict for high crenarchaeotal abundance.
Euryarchaeotal physiology and environmental gradients
In contrast to the Crenarchaeota, factors influencing euryarchaeotal distribution are less clear. Several indirect and direct evidences suggest that Euryarchaeota are capable of heterotrophy and/or autotrophy (Herndl et al., 2005; Teira et al., 2006; Wuchter, 2006). In this study, euryarchaeotal abundance was related to chlorophyll a concentrations (P<0.05) (Table 3 and Fig. 8). Such a relationship with phytoplankton has been described previously for the coastal southern North Sea and has been interpreted as an indication of euryarchaeotal heterotrophy (Wuchter, 2006). Euryarchaeotal abundance does not seem to be associated with any given phytoplankton assemblage because, for example, phytoplankton larger than 5 μm were the dominant group at the Dutch Coast station in April but not at the South Frisian Front in August, while Euryarchaeota were abundant at both sites (Figs 7 and 8). This suggests that other factors probably determine the covariance between euryarchaeotal abundance in the surface seawater and phytoplankton biomass. Recently, proteorhodopsin has been detected in Euryarchaeota living in the photic zone of the North Atlantic Subtropical Gyre (Frigaard et al., 2006). Their ability to use photons to generate energy may explain the covariance between Euryarchaeota and phytoplankton, i.e. both need sufficient light and nutrients. The exact function of this proteorhodopsin as well as its potential contribution to the overall energy requirements of proteorhodopsin-harboring cells are still uncertain. Some proteorhodopsins have also been described as having sensory capacity rather than being an energy source (Wang et al., 2003) and experimental evidence showed that darkness had little impact on bacterial populations even when cyanobacteria were strongly affected (Schwalbach et al., 2005). Nevertheless, the present study lends indirect support to the emerging notion that aerobic anoxygenic phototrophy may be important among prokaryotes and potentially also in Euryarchaeota (Beja et al., 2000; Kolber et al., 2000; Venter et al., 2004). However, more work is needed to better understand the physiological requirements of pelagic Euryarchaeota and, thus, to elucidate their ecological role in marine systems.
Summary and conclusions
- Top of page
- Materials and methods
- Results and discussion
- Summary and conclusions
This study has shown that pelagic Archaea constitute a dynamic component of the prokaryotic community in the southern North Sea. A clear temporal variability was observed, with Crenarchaeota being relatively more abundant in February and Euryarchaeota dominating the archaeal assemblage in April and August. Spatial differences in the lateral distribution of Crenarchaeota were, however, evident because over all seasons they remained abundant at two sites: the Frisian Front and the Central Southern Bight. This constitutes the first report of small-scale horizontal heterogeneity in Crenarchaeotal abundance in the marine environment. The abundance of Crenarchaeota was positively correlated with the copy number of the gene encoding the α subunit of crenarchaeotal ammonia monooxygenase (amoA) and with concentrations of ammonia, nitrate, nitrite and phosphorus, suggesting that most Crenarchaeota are nitrifiers and consequently that nutrient concentrations affect their distribution. The data also suggested that large phytoplankton abundance is negatively related to crenarchaeotal abundance, most probably via the influence of the bacterial community associated with large phytoplankton blooms. This implies that nutrient concentration as well as phytoplankton abundance and perhaps also community structure may predict crenarchaeotal abundance. In spring and summer, even though Euryarchaeota dominated the archaeal assemblage they constituted <4% of the total prokaryotic community. Their abundance also showed spatial differences in lateral distribution, correlating with chlorophyll a concentrations but interestingly not with phytoplankton community structure.
- Top of page
- Materials and methods
- Results and discussion
- Summary and conclusions
The authors thank the officers, crew and scientific party of the R.V. Pelagia and R.V. Alkor for their support during the cruises, Marianne Baas (NIOZ) for her assistance provided before and during the cruises, Santiago Gonzalez (NIOZ) for performing the DOC measurements, the team of the NIOZ nutrient lab (Jan van Ooijen, Karel Bakker and Evaline van Weerlee) for determining the concentrations of nutrient, DIC, DON and DOP, Anna Noordeloos (NIOZ) for her help with the phytoplankton sampling and data analysis and Ellen Hopmans (NIOZ) for assistance and advice with the HPLC/MS analyses. The authors are also grateful to Arjan Smit (NIOZ) and Elena Stoica (NIOZ) for their helpful support with the CARD-FISH analyses and to Govert van Noort (NIOZ) for his assistance with the microscopy. Special thanks are due to colleagues from the NIOZ molecular lab: Elda Panoto, Judith van Bleijswijk and Harry Witte for their invaluable advices, assistance and enthusiasm.
This study is part of the LOICZ project supported by the Research Council for Earth and Life Science (ALW), with financial aid from the Netherlands Organization for Scientific Research (NWO) (grant number 014.27.003 to J.S.S.D.).
- Top of page
- Materials and methods
- Results and discussion
- Summary and conclusions
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