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Confidential Information: Manuscript Submitted to Environ. Microbiol. (31 January 2003)
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In situ Distribution and Activity of Nitrifying Bacteria
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in Freshwater Sediment
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Dörte Altmann 1, Peter Stief 1, Rudolf Amann 1, Dirk de Beer 1,
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and Andreas Schramm 2*
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Max Planck Institute for Marine Microbiology, D-28359 Bremen, Germany
Department of Ecological Microbiology, BITOEK, University of Bayreuth,
D-95440 Bayreuth, Germany
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Running title: In situ analysis of nitrifiers in freshwater sediment
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*Corresponding author
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Mailing address: Department of Ecological Microbiology, BITOEK, University of
Bayreuth, Dr.-Hans-Frisch-Straße 1-3, D-95440 Bayreuth, Germany
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Phone:
+49 (0)921-555642
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Fax:
+49 (0)921-555799
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E-Mail:
andreas.schramm@bitoek.uni-bayreuth.de
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Summary
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Nitrification was investigated in a model freshwater sediment by the combined use of
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microsensors and fluorescence in situ hybridization. In situ nitrification activity was
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restricted mainly to the upper 2 mm of the sediment and coincided with the maximum
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abundance of nitrifying bacteria, i.e. 1.5·107 cells cm-3 for ammonia-oxidizing Beta-
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proteobacteria (AOB) and 8.6·107 cells cm-3 for Nitrospira-like nitrite-oxidizing bacteria
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(NOB). Cell numbers of AOB decreased more rapidly with depth than numbers of
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NOB. For the first time, Nitrospira-like bacteria could be quantified and correlated with
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in situ nitrite oxidation rates in a sediment, and cell-specific nitrite oxidation rates
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were 1.2-2.7 fmol NO2- cell-1 h-1.
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Introduction
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Nitrification in sediments has been shown to occur in narrow zones, i.e., within a few or
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sometimes even less than one mm (Jensen et al., 1993; Jensen et al., 1994; Lorenzen et al.,
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1998). This observation provides a challenge for the exact determination of rates and for the
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quantification and identification of the nitrifying community with sufficiently high spatial
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resolution.
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aggregates, the combined use of microsensors and fluorescence in situ hybridization (FISH)
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has been successfully applied (reviewed by Schramm, 2003).
For the analysis of processes and populations in nitrifying biofilms and
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This approach has so far not been used for the investigation of nitrification in freshwater
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sediments, partially because the low abundance of nitrifiers and the background
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fluorescence of sediment material was assumed to reduce the applicability of FISH (Prosser
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and Embley, 2002). Although numerous studies have addressed the diversity of nitrifying
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bacteria in sediments with molecular methods (e.g., Hastings et al., 1998; Kowalchuk et al.,
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1998; Whitby et al., 2001), information about abundance and fine-scale distribution of
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ammonia-oxidizing bacteria (AOB) in sediments is scarce, and nitrite-oxidizing bacteria
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(NOB) have been widely neglected.
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The objectives of the present study were therefore (i) to evaluate the potentials and
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limitations of the combined use of microsensors and FISH in freshwater sediments, and (ii) to
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obtain first insights into the in situ abundance, vertical distribution, and activity of AOB and
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NOB in a model freshwater sediment.
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Results and Discussion
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Sediment pre-incubation.
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Sediment from a small lowland stream (fine sand, low organic content) that had been sieved
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to remove macrofauna was allowed to settle in cylindrical chambers (Jensen et al., 1994).
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After two weeks of incubations in the dark with stream water close to in situ conditions (O 2
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adjusted to air saturation, 50 µM NH4+, 500 µM NO3-, 15°C) the sediment NH4+ conversions
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had reached steady state which was proven by repeated bulk measurements of NH4+ in the
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overlying water (data not shown), i.e. NH4+ fluxes into the sediment remained stable over
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time.
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disturbance of measurements. However, after this initial perturbation, care was taken to
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maintain near-in situ conditions during re-stratification of the sediment; it is therefore
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anticipated that the original nitrifying community re-established in the sediment columns, and
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that similar communities and distributions of nitrifiers are to be found under similar conditions
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in other freshwater sediments.
Macrofauna had been removed to allow for a stable stratification and to avoid
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Activity of nitrifying bacteria.
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Nitrification activity as determined by microsensor measurements was restricted to the upper
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2 mm of the sediment (Fig. 1A, B). Maximum conversion rates were found at the sediment
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surface, and nitrate production rates (0.2 µmol cm-3 h-1) were comparable to rates obtained in
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other freshwater sediments by microsensor measurements (Jensen et al., 1993; Jensen et
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al., 1994; Lorenzen et al., 1998). Depth-integrated reaction rates of spatially coinciding O2
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consumption, NH4+ consumption, and NO3- production were 1.00  0.04 mmol m-2 h-1, 0.16 
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0.01 mmol m-2 h-1, and 0.34  0.16 mmol m-2 h-1, respectively. The production of NO3- at a
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two-fold higher rate than consumption of NH4+ might be explained by two processes: (i)
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simultaneous NH4+ production by mineralization (de Beer et al., 1991) leads to an
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underestimation of NH4+ consumption by nitrification (Okabe, 1999); and (ii) NO3- reduction in
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anoxic sediment layers might provide additional NO2- for NO3- production in the oxic zone
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independent of NH4+ (Stief et al., 2002). Indeed, NO3- consumption was detected between a
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sediment depth of 2.5 mm (i.e., after O2 had disappeared) and 8.5 mm at rates of 0.02-0.06
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µmol cm-3 h-1, which is at the lower end of rates measured in other freshwater sediments
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(e.g., Jensen et al., 1994; Lorenzen et al., 1998).
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Abundance and vertical distribution of nitrifying bacteria.
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Total cell numbers were 2.4-3.4·109 cm-3, with highest numbers at the sediment surface. Of
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these cells, 80% at the surface and about 50% in the deepest layers could be detected by
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FISH with a combination of probes Eub338 (Amann et al., 1990), Eub338-II, and Eub338-III
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(Daims et al., 1999). Unspecific signals (due to binding of the negative control probe Non338
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(Manz et al., 1992) or autofluorescent cells) were detected for only ≤ 0.04% of all cells, which
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means that theoretically microbial populations with an abundance of less than 8·105 cells cm-
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by probe Nso1225 (Mobarry et al., 1996) (Fig. 2A, B) accounted for only about 0.5% of all
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cells. Their cell numbers decreased even further with depth (Fig. 3). Due to these low
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numbers, further identification of -AOB by FISH was not attempted.
were not detectable by FISH. Ammonia-oxidizing Beta-proteobacteria (-AOB) as detected
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NOB could be identified by hybridization with probe Ntspa662 (Daims et al., 2001a) as
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members of the genus Nitrospira (Fig. 2C, D) which accounted for maximal 2.7% of all cells.
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Identification was confirmed by hybridization with probe Ntspa712 specific for the phylum
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Nitrospira (Daims et al., 2001a) which yielded similar numbers (data not shown). Nitrobacter,
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the most commonly isolated NOB, was not detected by hybridization with probe Nit3
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(Wagner et al., 1996). After identification of Nitrospira as key nitrite oxidizer in engineered
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systems like freshwater aquaria (Hovanec et al., 1998), activated sludge (Juretschko et al.,
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1998; Daims et al., 2001a) and bioreactors (e.g., Schramm et al., 1998; Okabe et al., 1999;
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Gieseke et al., 2001), this is yet another example of the importance of Nitrospira not
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Nitrobacter for nitrite oxidation in situ. Nitrospira-like 16S rRNA gene sequences have also
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been retrieved from several natural aquatic habitats like deltaic mud, caves or lakes
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(summarized in Daims et al., 2001a). However, this is the first report of not just detection but
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also quantification and distribution of Nitrospira sp. in a quasi natural system, and to correlate
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these data with in situ activity measurements. With a size of 0.4 - 0.8 µm (Fig. 2C,D) the
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cells were somewhat larger than previously detected by FISH (e.g., Schramm et al., 1998),
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but still well in the range of cultured Nitrospira (Bock, 1992; Ehrich et al., 1995). Although
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cell numbers of Nitrospira sp. also decreased with depth, their abundance remained
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relatively high (0.4-1.0 % of all cells), even in the deeper, anoxic layers (Fig. 3).
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These results are in agreement with previous studies showing that (i) the abundance of
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nitrifying bacteria decreases with depth in freshwater sediments (Whitby et al., 2001); (ii) the
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numbers of NOB might be 3-30 times higher than that of AOB in sediments (Smorczewski
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and Schmidt, 1991), nitrifying aggregates (Schramm et al., 1999), or biofilms (Gieseke et al.,
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2001); and (iii) NOB of the genus Nitrospira maintain a ribosome content high enough for
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detection by FISH even in the absence of O2 (Schramm et al., 1998; Okabe et al., 1999;
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Schramm et al., 2000; Gieseke et al., 2001). The latter suggests an adaptation of Nitrospira
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sp. to survive or even thrive under anoxic conditions.
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Compared to the enumeration of nitrifying bacteria in freshwater sediments by the most
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probable number method (Hall, 1986; Smorczewski and Schmidt, 1991; Hastings et al.,
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1998; Pauer and Auer, 2000; Whitby et al., 2001), cell numbers determined by FISH in the
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present study are 1-4 orders of magnitude higher. This finding is consistent with the general
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observation that cultivation-dependent methods severely underestimate the actual size of
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microbial populations (Amann et al., 1995), which is also true for nitrifying bacteria (Hall,
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1986).
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Cell-specific nitrification rates.
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Using volumetric conversion rates of NH4+ and NO3-, and cell numbers of -AOB and NOB in
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the nitrification zone, the cell-specific NH4+ oxidation and NO3- production rates were
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estimated to be 1.3-8 fmol NH4+ cell-1 h-1 and 1.2-2.7 fmol NO3- cell-1 h-1, respectively. The
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cell-specific NH4+ oxidation rates are at the lower end of rates found in the literature (1-30
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fmol cell-1 h-1; Prosser, 1989), and are most likely underestimated due to the underestimation
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of NH4+ consumption rates (see above). In contrast, the cell-specific NO3- production rates
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are high compared to 0.01-0.07 fmol cell-1 h-1 calculated for Nitrospira spp. in nitrifying
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aggregates (Schramm et al., 1999) and biofilms (Gieseke et al., 2001). This observation
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might be explained in several ways: (i) different strains of Nitrospira might be present in the
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sediment and in the bioreactors, and, considering the broad phylogenetic diversity of the
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genus (Daims et al., 2001a), the specific NO2--oxidation rates might greatly differ between
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these different strains; (ii) the cell-specific NO2--oxidation rates of Nitrobacter are much
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higher (5-40 fmol cell-1 h-1 ; Prosser, 1989) than those reported for Nitrospira. Even though
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undetectable by FISH in the sediment, i.e. at cell numbers ≤ 8·105 cells cm-3, Nitrobacter
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might significantly contribute to the production of NO3-; and (iii) the occurrence of other NOB,
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e.g., of the genus Nitrococcus or Nitrospina, which have so far only been detected in marine
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environments, cannot completely be ruled out.
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Concluding remarks.
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The combined use of microsensors and FISH proved to be applicable to freshwater
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sediments. For the first time, the in situ abundance of nitrifiers in a sediment could be
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determined with sufficiently high spatial resolution to be combined with fine-scale activity
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measurements. Nitrospira-like bacteria were the dominant if not sole nitrite-oxidizers in the
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sediment. Limitations for the application of the combined approach to sediments are the
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potential occurrence of bioturbating macrofauna that may disturb microsensor measurements
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and interpretation of profiles in a real natural sediment, and the extremely time-consuming
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counting procedure during FISH analysis; despite an enormous counting effort, populations
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with lower abundance (106 cells cm-3 of sediment) will remain undetectable by FISH. While
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the application of automated image analysis (e.g., Daims et al., 2001b) might in the future
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speed up FISH analysis even in difficult samples like sediments, it will not improve its
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detection limit.
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Acknowledgements
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We thank Enrique Llobet-Brossa and Armin Gieseke for their advice and helpful comments
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regarding FISH in sediments. Gabi Eickert, Anja Eggers, and Ines Schröder are
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acknowledged for the construction of the O2 microelectrodes.
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Financial support was provided by the German Research Foundation (STI202/1), by the Max
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Planck Society, Germany, and by an Otto-Hahn Award of the Max Planck Society to Andreas
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Schramm.
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along a macroscale gradient in a nitrifying bioreactor: quantification by in situ hybridization
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Schramm, A., de Beer, D., Wagner, M. and Amann, R. (1998) Identification and activity in
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22
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Stief, P., de Beer, D. and Neumann, D. (2002) Small-scale distribution of interstitial nitrite in
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freshwater sediment microcosms: the role of nitrate and oxygen availability, and sediment
3
permeability. Microb Ecol 43: 367-378.
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Wagner, M., Rath, G., Koops, H.-P., Flood, J. and Amann, R. (1996) In situ analysis of
nitrifying bacteria in sewage treatment plants. Water Sci Technol 34: 237-244.
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ammonia-oxidiser populations in eutrophic and oligotrophic basins of a large freshwater
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lake. Antonie Van Leeuwenhoek 79:179-188.
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Figure legends
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Fig. 1.
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Mean values + standard deviation for concentrations (A) and volumetric conversion rates (B)
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of O2 (n = 5), NH4+ (n = 4), and NO3- (n = 4) along the depth of the freshwater model
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sediment. Dotted line corresponds to the sediment surface.
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Microsensors for O2 , NO3-, and NH4+ were prepared and calibrated according to published
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protocol (Revsbech, 1989; de Beer et al., 1997).
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sediment chambers were recorded using a measuring setup as previously described (Stief et
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al., 2002). Concentration profiles were used to calculate volumetric conversion rates of O2,
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NH4+ and NO3- (de Beer and Stoodley, 1999; Stief and de Beer, 2002).
Vertical concentration profiles in the
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Fig. 2.
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Epifluorescence micrographs of a cluster of ammonia-oxidizing -proteobacteria hybridized
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with probe Nso1225 (A) and Nitrospira-like bacteria hybridized with probe Ntspa662 (C), and
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the respective DAPI counterstain (B, D). Scale bars are 5 µm; arrows indicate Nitrospira-like
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cells.
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Sediment cores (diameter 2.5 cm, taken from sediment columns after microsensor
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measurements) were sectioned horizontally and fixed with paraformaldehyde (Llobet-Brossa
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et al., 1998). Sediment sections were diluted 30-fold in a 1:1 mix of 1 phosphate-buffered
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saline (10 mM sodium phosphate [pH 7.2], 130 mM NaCl) and 96% Ethanol, sonicated ( 3 x
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60 s, 20 % pulse, 109 µm amplitude) with a type UW70 probe (Sonopuls HD70; Bandelin,
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Berlin, Germany), and aliquots of 20-30 µl were immobilized on gelatine-coated microscopic
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slides. FISH with CY3-labeled oligonucleotide probes (Hybaid Interactiva, Ulm, Germany),
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counter staining of all cells with 4´,6-diamino-2-phenylindole (DAPI; 0.5 µg ml-1), and
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microscopic analysis was according to published protocol (Pernthaler et al., 2001).
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1
Fig. 3.
2
Mean values + standard deviation for cell numbers of ammonia-oxidizing -proteobacteria
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hybridized with probe Nso1225 (Mobarry et al., 1996) (n = 3), and nitrite-oxidizing Nitrospira
4
spp. hybridized with probe Nts662 (Daims et al., 2001a) (n = 3), along the depth of the
5
sediment.
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All FISH counts were corrected by subtracting cell numbers obtained with control probe
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Non338. For probe Nts662, 20 randomly chosen microscopic fields (corresponding to 1000
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to 2000 DAPI-stained cells) were investigated. For probes Nso1225 and Non338 200 to 400
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microscopic fields were analyzed to account for the low cell numbers and the uneven
10
distribution in the sample; total cell counts of 20 microscopic fields were then extrapolated to
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the area analyzed for FISH-signals. The absolute numbers of FISH-positive cells were
12
calculated for each probe using the relative FISH-positive counts (as percentage of DAPI-
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stained cells) and the total cell counts from separately conducted DAPI stains of the
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sediment samples on black membrane polycarbonate filters (pore size, 0.2 µm; Osmonics
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Inc., Livermore, California, USA).
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13
1
Fig. 1
2
0
concentration [µmol l-1]
200
400
volumetric conversion rates
-3 -1
[µmol cm h ]
-0,6
-0,3
0,0
0,3
0,6
600
-4
0-1
-2
sediment depth [mm]
1-2
0
2-3
3-4
2
4-5
4
6
8
5-6
6-7
O2
7-8
NH4+
8-9
NO3-
9-10
10
3
4
5
14
O2
NH4+
NO3-
1
Fig. 2
2
3
4
5
6
7
8
A
B
C
D
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1
Fig. 3
2
3
7
0
-3
cells x 10 cm
8
4
sediment depth [mm]
0-2
2-4
4-6
6-8
-AOB
Nitrospira spp.
8-10
4
16
12