Archaea Dominate the Ammonia-Oxidizing Community in the Rhizosphere of the Freshwater Macrophyte Littorella uniflora

Abstract

Archaeal and bacterial ammonia monooxygenase genes (amoA) had similar low relative abundances in freshwater sediment. In the rhizosphere of the submersed macrophyte Littorella uniflora, archaeal amoA was 500- to >8,000-fold enriched compared to bacterial amoA, suggesting that the enhanced nitrification activity observed in the rhizosphere was due to ammonia-oxidizing Archaea.

In shallow aquatic ecosystems, rhizosphere-associated nitrogen transformations are central to understanding nutrient cycling (29). Oxygen release from the roots of freshwater macrophytes like Littorella uniflora, Lobelia dortmanna, and Glyceria maxima stimulates nitrification and coupled nitrification-denitrification in freshwater sediments (4, 29, 34). However, the nitrifying community responsible for the increase in the rate has rarely been investigated (4, 21). The recent discovery of ammonia-oxidizing Archaea (AOA) (19, 41), the widespread distribution of these organisms (2, 3, 9, 14, 31, 35), and their predominance in soils (24) and oceans (22, 27, 42) led to the hypothesis that AOA might also be important for nitrification in freshwater environments, including in the rhizosphere of freshwater macrophytes. The goals of this study were therefore (i) to test for the occurrence of AOA in freshwater sediment and in the rhizosphere of the macrophyte Littorella uniflora, (ii) to compare AOA diversity and abundance to the diversity and abundance of ammonia-oxidizing bacteria (AOB), and (iii) to compare the ammonia oxidizer communities in bulk and rhizosphere sediments.

Sampling and chemical analysis.

Sediment cores were obtained in triplicate from within monospecies stands of L. uniflora and from unvegetated sediment that was ≥5 m from the plant stands at a water depth of 20 to 30 cm in oligomesotrophic Lake Hampen, Jutland, Denmark (7). The distances between replicate cores were 2 to 70 m for vegetated sediment and 10 to 40 m for unvegetated sediment. For molecular (September 2005 and June 2006) and pore water (June 2006) analyses, the upper 1.5 cm of unvegetated sediment and L. uniflora rhizosphere sediment (depth, 1 to 6 cm) were transferred in the field to sterile 50-ml Falcon tubes and kept on ice during transport. In the laboratory, pore water was extracted within 24 h for analyses of pH, NO₂⁻ plus NO₃⁻ (6), NO₂⁻ (15), and NH₄⁺ (5), and samples for DNA extraction were frozen at −80°C.

To determine potential nitrification rates (June 2006), intact sediment cores were transported to the laboratory at ambient temperature (22 to 24°C). Rhizosphere and surface sediment samples were obtained and homogenized separately for each replicate core, and incubations at room temperature and 120 rpm were immediately set up with 20-g (fresh weight) subsamples in 40 ml of sterile lake water containing 100 μM NH₄⁺. Apparent potential nitrification rates were calculated from the linear increase in concentrations of NO₂⁻ plus NO₃⁻ during the first 6 h, and the values were about eight times higher for the rhizosphere than for the unvegetated sediment. This difference was reflected by higher concentrations of nitrate and lower concentrations of ammonium in the rhizosphere (Table 1), confirming previous reports of enhanced nitrification in the L. uniflora rhizosphere (29). The actual nitrification rates might be slightly higher as the possibility of denitrification and thus loss of NO₂⁻ plus NO₃⁻ cannot be fully excluded even in strictly oxic incubations.

TABLE 1.

Pore water characteristics and potential nitrification rates in unvegetated sediment and rhizosphere sediment of L. uniflora at Lake Hampen, Denmark (June 2006)^a

Sediment	pHb	NH4+ concn (μM)	NO2− concn (μM)	NO3− concn (μM)	Potential nitrification rate (nmol NO2− + NO3− g [fresh wt]−1 h−1)
Unvegetated	6.8	84.7 ± 63.0	0.4 ± 0.1	1.3 ± 0.2	1.4 ± 0.4
Rhizosphere	6.8	20.0 ± 7.25	0.5 ± 0.1	58.2 ± 22.8	10.5 ± 0.8

^aThe values are means ± standard deviations for triplicate sediment cores.

^bTriplicate pore water extracts were pooled for pH measurement.

Diversity of AOA and AOB.

For each replicate core, DNA was extracted in triplicate from 200 mg sediment by combining enzymatic and chemical cell lysis with a Fast DNA Spin kit for soil (Qbiogene Inc.) (13). Archaeal and bacterial amoA genes were amplified using a HotStar Taq Mastermix kit (Qiagen), previously described protocols, and primer sets Arch-AmoAF/Arch-AmoAR (14) and AmoA-1F/AmoA-2R-TC (28). Purified PCR products from triplicate DNA extracts and triplicate sediment cores were pooled for subsequent cloning (pGEM-T cloning kit; Promega), which yielded one rhizosphere clone library and one sediment clone library per gene. In addition, an archaeal 16S rRNA gene library was constructed from the rhizosphere samples using primers Arch-21F (10) and 1492R (23). Between 14 and 94 clones were analyzed for the libraries. Clones were sequenced by Macrogen (Korea). Alignment, translation into amino acids, and phylogenetic analyses were done in ARB (25), utilizing the SILVA database (33) for the 16S rRNA gene sequences. The coverage of the libraries (39) was 93 to 100% based on 97% similarity grouping on the amino acid (AmoA) or nucleic acid (16S rRNA genes) level in DOTUR (36).

Bacterial AmoA sequences were affiliated with groups commonly found in freshwater habitats (20), including the Nitrosospira, Nitrosomonas marina/Nitrosomonas oligotropha, and Nitrosomonas europaea lineages (Fig. 1A). The distinct occurrence of most AmoA sequence types in either unvegetated or rhizosphere sediment (only one, Nitrosospira-like AmoA sequence type was found in both sediments) may indicate adaptation of certain AOB to the special environment of the macrophyte rhizosphere. Archaeal AmoA sequences were affiliated with the “water column/sediment” cluster (designated cluster A I) and “soil/sediment” cluster (cluster B) originally defined by Francis et al. (14), as well as with a third major archaeal AmoA lineage (designated cluster A II), which previously contained only a few AmoA sequences from estuarine samples or soil (Fig. 1B). Again, most AmoA sequence types were specific for either unvegetated or rhizosphere sediment; only two AmoA sequence types occurred in both sediments. For the rhizosphere samples, the phylogeny of archaeal AmoA is partially mirrored by that of archaeal 16S rRNA genes (Fig. 1C); however, the congruence is not perfect, and the detection of more 16S rRNA types than AmoA sequence types may indicate that not all Crenarchaeota in the rhizosphere are actually ammonia oxidizers; alternatively, the primers used might not target all archaeal amoA or 16S rRNA genes.

FIG. 1.

Phylogenetic affiliation of bacterial (A) and archaeal (B) AmoA and archaeal 16S rRNA genes (C). Sequences obtained in this study are indicated by bold type with the prefix AS_ (sequences from unvegetated sediment) or LAR_ (sequences from L. uniflora rhizosphere). For AmoA, protein distance trees are displayed with bootstrap values (>50%) obtained from a parsimony analysis. Branches not supported by both methods are drawn as multifurcations; the major lineages were also supported by maximum likelihood analysis with a limited number of sequences. For the archaeal 16S rRNA gene, a strict consensus tree obtained by using neighbor joining, parsimony, and maximum likelihood methods is shown, in which branches not supported by all three methods are drawn as multifurcations. Scale bars = 5% estimated sequence divergence.

Abundance of AOA and AOB.

Copy numbers of 16S rRNA and bacterial or archaeal amoA genes were determined in triplicate for each sediment sample by quantitative PCR (qPCR) with Brilliant SYBR Green qPCR Master Mix (Stratagene) using an Mx3005P instrument (Stratagene), the universal 16S rRNA gene primers 907F and 1492R (23), and the amoA primers described above. The PCR conditions were 95°C for 10 min, followed by 50 cycles of 1 min at 95°C, 30 s at 53°C (for archaeal amoA) or at 57°C (for bacterial amoA and 16S rRNA genes), 60 s at 72°C, and data capture for 30 s at 78°C (archaeal amoA) or for 20 s at 80°C (bacterial amoA and 16S rRNA genes). Standard curves were prepared using serial dilutions of Bacteroides fragilis ATCC 25285 genomic DNA for 16S rRNA genes and serial dilutions of plasmids containing an archaeal or bacterial amoA gene; the data were linear for 10⁷ to 10² 16S rRNA gene copies and for 5 × 10⁸ to 5 × 10¹ amoA gene copies, and the detection limit was 5 gene copies. The efficiencies of the qPCR were 88 to 97% for amoA and 75 to 83% for 16S rRNA genes; the specificity of PCR products was confirmed by melting curve analysis. The abundance of AOA or AOB was expressed relative to 16S rRNA gene copy numbers, assuming that there were 2.5 and 1 amoA gene copies per AOB and AOA, respectively (24), one 16S rRNA gene copy per AOA, and 3.6 copies of the 16S rRNA gene per average prokaryotic cell (18). Relative abundances were preferred to absolute values for this study as they are less sensitive to the differences in DNA extraction efficiencies that can be expected when samples as different as sandy surface sediment and organic matter-rich rhizospheres are used. Variations in qPCR amplification and primer biases may introduce further uncertainties, especially when target gene abundances are low, and the exact numbers (absolute or relative) obtained by qPCR should be interpreted cautiously (11, 40). Indeed, the variation between data obtained for triplicate DNA extracts, qPCR, and environmental replicates was considerable. However, the method-based variation was several orders of magnitude lower than the differences in relative abundance between AOA and AOB and between bulk and rhizosphere sediments.

In unvegetated sediment, the relative abundances of AOB and AOA were (with one exception) similar and in general were <0.02% of the total prokaryotic community (Fig. 2). While similar or even lower relative abundances of AOB were observed in the rhizosphere of L. uniflora, AOA were strongly enriched, accounting for 0.5 to 5% of the prokaryotic community. Consequently, AOA outnumbered AOB by 500- to 8,000-fold in the macrophyte rhizosphere (Fig. 2). Similar values for AOA/AOB ratios and abundances have been found in certain soils (24) and for crenarchaeotal abundance (approximately 1%) in the rhizosphere of terrestrial plants (37, 38). Despite some spatial and seasonal variation (i.e., differences between replicate cores and higher relative abundances of both AOA and AOB in September [Fig. 2A] than in June [Fig. 2B]), AOA always dominated the ammonia-oxidizing community in the rhizosphere of L. uniflora; this general trend suggests that AOA might be better adapted than AOB to microaerophilic conditions in the rhizosphere (4, 12) or may profit from root exudates, as suggested for terrestrial Crenarchaeota (38); this hypothesis is supported by the mixotrophic traits detected in the genome of the putative AOA Cenarchaeum symbiosum (16) and the uptake of amino acids by marine Crenarchaeota (17, 30).

FIG. 2.

Relative abundances of AOA and AOB in unvegetated sediment (AS I to AS III) and in the rhizosphere of L. uniflora (LAR I to LAR III) in September 2005 (A) and June 2006 (B). The values are means ± standard deviations of three qPCR with triplicate (A) or single (B) DNA extracts. The columns represent maximum estimates when amoA gene copy numbers remained below the quantification limit of 50 copies (filled circles) or the detection limit of 5 copies per reaction (open circles); archaeal and bacterial amoA genes were present in all the samples, as confirmed with higher template concentrations with the cloning approach. The numbers in boxes are the minimum estimates for AOA/AOB ratios when the level of AOB was below the quantification limit or are maximum estimates when the level of AOA was below the quantification limit. No ratios are shown when the levels of both groups were below the quantification limit.

Potential AOA activity.

Higher nitrification rates in the rhizosphere than in unvegetated sediment (Table 1) coincided with higher relative abundances of AOA but not AOB (Fig. 2B), suggesting that AOA were indeed actively oxidizing ammonia. The numbers of AOB in the rhizosphere can be estimated from the qPCR data and total cell counts for freshwater sediments (1, 12) to be 0.4 × 10⁴ to 4 × 10⁴ cells g⁻¹. To explain the potential nitrification rates (Table 1) by AOB activity alone would therefore require cell-specific ammonia oxidation rates of 200 to 2,000 fmol NH₃ h⁻¹, which are 1 to 2 orders of magnitude greater than the highest rates reported for cultured AOB and wastewater treatment systems (8, 26, 32). Evidently, AOA activity is needed to reconcile the rates. If all estimated 1.5 × 10⁷ to 2.5 × 10⁷ AOA g⁻¹ were equally active, the cell-specific rates for AOA would be around 0.5 fmol NH₃ h⁻¹; this specific activity is in the lower range reported for AOB but is three to six times higher than that estimated for marine AOA (42). Hence, AOA activity is the most likely explanation for the increased nitrification rates in the rhizosphere. In contrast, at such low cell-specific activity AOA would contribute very little to total ammonia oxidation in the unvegetated sediment samples, where AOA abundance is low and AOB alone can theoretically account for most of the observed activity.

Conclusion.

AOA outnumber AOB and likely are responsible for the enhanced nitrification activity in the rhizosphere of the freshwater macrophyte L. uniflora. To our knowledge, this is the first study indicating a potential role for AOA in freshwater lakes, particularly in rhizosphere sediment. The contribution of AOA to ammonia oxidation in unvegetated sediment and under in situ conditions and the factors regulating the occurrence and abundance of AOA versus the occurrence and abundance of AOB remain to be determined.

Nucleotide sequence accession numbers.

Nonredundant sequences determined in this study have been deposited in the GenBank database under accession numbers EU309859 to EU309918.