Distribution and Rate of Microbial Processes in an Ammonia-Loaded ...

APPLIED AND ENVIRONMENTAL MICROBIOLOGY, June 2009, p. 3705–3713 0099-2240/09/$08.00⫹0 doi:10.1128/AEM.02612-08 Copyright © 2009, American Society for Microbiology. All Rights Reserved.

Vol. 75, No. 11

Distribution and Rate of Microbial Processes in an Ammonia-Loaded Air Filter Biofilm䌤 Susanne Juhler,* Niels Peter Revsbech, Andreas Schramm, Martina Herrmann,† Lars D. M. Ottosen, and Lars Peter Nielsen Department of Biological Sciences, Microbiology, Aarhus University, Bd. 1540, DK-8000 Aarhus C, Denmark Received 14 November 2008/Accepted 30 March 2009

The in situ activity and distribution of heterotrophic and nitrifying bacteria and their potential interactions were investigated in a full-scale, two-section, trickling filter designed for biological degradation of volatile organics and NH3 in ventilation air from pig farms. The filter biofilm was investigated by microsensor analysis, fluorescence in situ hybridization, quantitative PCR, and batch incubation activity measurements. In situ aerobic activity showed a significant decrease through the filter, while the distribution of ammonia-oxidizing bacteria (AOB) was highly skewed toward the filter outlet. Nitrite oxidation was not detected during most of the experimental period, and the AOB activity therefore resulted in NO2ⴚ, accumulation, with concentrations often exceeding 100 mM at the filter inlet. The restriction of AOB to the outlet section of the filter was explained by both competition with heterotrophic bacteria for O2 and inhibition by the protonated form of NO2ⴚ, HNO2. Product inhibition of AOB growth could explain why this type of filter tends to emit air with a rather constant NH3 concentration irrespective of variations in inlet concentration and airflow. Emissions of NH3, odorous organic gasses, and dust from pig facilities cause significant problems for neighbors and the surrounding natural environment. In Denmark, swine production accounts for 34% of the total atmospheric NH3 emission, of which 50% originates from pig house emissions (19). Furthermore, multiple volatile and very odorous organic compounds are emitted with animal house exhaust air and constitute a severe nuisance in residential areas (16, 31). While biofilters based on wood chips, compost, and peat have proven efficient in removing complex mixtures of volatile organic compounds (VOC) from piggery exhaust air, biotrickling filters have been shown to be efficient in both odor and NH3 removal (30). In these filters, airborne VOC and NH3 are taken up by an irrigated biofilm and oxidized by organoheterotrophic and nitrifying bacteria, respectively, resulting in the production of CO2, NO2⫺ or NO3⫺, and microbial biomass (43). The nitrification process, which comprises the two-step oxidation of NH3 via NO2⫺ to NO3⫺, is catalyzed by ammonia-oxidizing bacteria (AOB) and ammonia-oxidizing archaea and by nitriteoxidizing bacteria (NOB), respectively (22, 23). However, in many biotrickling filters, NOB activity seems to be absent, resulting in NO2⫺ being the end product of nitrification (29). Waste products often accumulate in biotrickling filters, as the water is recycled many times to minimize wastewater discharge. In particular, NO2⫺ may accumulate to concentrations above 100 mM (29), resulting in high levels of free nitrous acid (FNA, or HNO2), which is inhibitory to many microorganisms (2, 40, 50). As a result of an overall countercurrent air-water flow, FNA, VOC, and NH3 concentrations are expected to

decrease from the filter air inlet toward the outlet, potentially promoting a gradient of microbial processes and species distribution through the filter. Also, on a smaller scale, physical and chemical heterogeneity results in microgradients of substrates and O2 within the biofilm that may affect the spatial distribution and rate of microbial processes. While considerable research effort has been given to the physicochemical aspects of filter optimization (11, 44), very little is known about the distribution, activity, and potential interactions of bacterial processes in the biofilm. Problems frequently arise, such as highly variable odor reduction, failed NH3 removal, and filter clogging due to excess biofilm growth and slime excretion. Improved insight into the dynamics of the microbial activities may help control the biological processes in order to optimize and stabilize filter performance. In this study, the in situ activity and distribution of organoheterotrophic bacteria and AOB and the identity and NH3 oxidation potential of the AOB were investigated in a twosection, full-scale biological air filter treating waste gas from a pig facility. Biofilms in different filter sections along a gradient of decreasing VOC and NH3 load and NO2⫺ accumulation were characterized. The goal was to attain a basic understanding of the microbial processes and thus filter function. MATERIALS AND METHODS Sampling site: the biotrickling filter. Biofilm sampling and in situ measurements were performed in a biological air filter (SKOV A/S, Roslev, Denmark) installed at a piglet-raising facility in northern Jutland, Denmark. The filter is composed of two sequentially arranged, 15-cm-thick walls of corrugated cardboard supporting the active biofilm (sections 1 and 2) (Fig. 1). Proofing renders the cardboard resistant to disintegration by water and microbial degradation. Air and water are conducted in a partly countercurrent, partly cross-current flow. Air from the stable is sucked in a horizontal direction through both filter sections by ventilators installed at the filter outlet. To keep the biofilm moist and remove biofilm metabolites, the two filter sections are irrigated with water, which is recycled from a reservoir beneath each filter section. Conductivity is monitored automatically as a measure of accumulated nitrogen salts, and when it exceeds a fixed threshold value corresponding to about 200 mM ion equivalents, water is

* Corresponding author. Mailing address: Institute of Biological Sciences, Microbiology, Aarhus University, Bd. 1540, DK-8000 Aarhus C, Denmark. Phone: 45 89423318. Fax: 45 8942 2722. E-mail: susanne [email protected]. † Present address: Leibniz University Hannover, Institute for Microbiology, Schneiderberg 50, D-30167 Hannover, Germany. 䌤 Published ahead of print on 10 April 2009. 3705

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FIG. 1. Directions of airflow (broken lines) and water flow (solid arrows) in the biological air filter. (Modified with permission from SKOV A/S.)

drained from the front filter section (section 1) to the manure tank. Drained and evaporated water is replaced by water from section 2, which is again supplied with fresh water. Due to the net countercurrent and evaporation of water, concentrations of metabolites increase from section 2 to section 1. Sampling and in situ measurements in the air filter were performed 3, 15, and 41 weeks after filter startup. Samples were collected from the front and back of each filter section (section 1in, 1out, 2in, and 2out, respectively) (Fig. 1) by cutting out pieces of biofilm on substratum material with clean scissors and tweezers. In each filter site, the biofilm was quite heterogeneous, but the thickness, texture, and appearance of collected samples represented biofilm covering more than 50% of the investigated filter area. Air and water samples. Once within each sampling period, water samples were collected from both filter sections, filter sterilized (pore size, 0.2 ␮m), and stored at ⫺20°C for later analysis. In the laboratory, NOx⫺ concentrations ([NO2⫺ ⫹ NO3⫺]) were measured with a NOx⫺ macrobiosensor (Unisense), NO2⫺ was analyzed colorimetrically using a modified Griess-Ilosvay reaction (17), the NH4⫹ concentration was determined by a salicylate-hypochlorite method (5), and the concentrations of other cations were determined with an ICP-OES element analyzer (2000DV; Perkin-Elmer, Waltham, MA). Daily, water temperature and pH in section 1 were recorded directly by a sensor-based automatic monitor (dTRANS Lf 01 transmitter/controller; JUMO, Viby Sj., Denmark), while water salinity was measured by a refractometer calibrated with NH4NO3. From these values, the concentrations of HNO2 and NH3 were calculated according to the following equations (2, 20): log[AH]aq ⫺ log[A]aq ⫽ pKa ⫺ pH; pKa(HNO2/NO2⫺) ⫽ 999/(273 ⫹ T); and pKa(NH4⫹/NH3) ⫽ ⫺0.467 ⫹ 0.00113S ⫹ 2887.9/(273 ⫹ T), where AH and A are the protonated and unprotonated forms of a compound, respectively, T is the temperature (°C), and S is the salinity (‰). Before, between, and after the two filter sections, the air temperature was recorded and the NH3 concentration in the air was estimated applying gas detection tubes (Ammonia 2/a; Dra¨ger Safety, Lu ¨beck, Germany). In situ microsensor measurements. Clark-type O2 microsensors with guard cathodes and internal references were prepared and calibrated as described previously (1, 37) and used for profiling of biofilms. The stirring sensitivity was ⬍2%, the tip diameter was ⬍10 ␮m, and the response time was ⬍2 s. To avoid inaccuracies caused by vibrations in the filter pads, cut-out paper support material with intact biofilm was fixed on polystyrene foam with needles and positioned in a stabilized setup firmly attached to a metal stand. Microprofiles were recorded by perpendicularly inserting the microsensor with a motor-driven micromanipulator in step sizes of 20 to 50 ␮m. During profiling, the setup was placed in the air stream inside the full-scale biotrickling filter (samples from sections 1in and 1out in the inlet air, samples from section 2in between the filter sections, and section 2out samples in the outlet air) (Fig. 1). The biofilm surface was facing the air stream while being continuously irrigated with water collected from the corresponding filter section. For measurements in the very thin biofilm of the back of each filter section, the sample was fixed on a piece of polystyrene foam in a petri dish positioned at a 60° angle to the sensor, and depth intervals were subsequently recalculated to correspond to vertical step sizes. Filter paper underneath the sample kept the biofilm moist through capillary transport of filter water applied to the petri dish. The supply of water created by the filter paper simulated the conditions at the back of the filter sections, as convective water

APPL. ENVIRON. MICROBIOL. supply occurred only at the front. All profiles were initiated just above the air-water interface, and depth zero was subsequently determined by a change in the sensor signal at the transition from air to aqueous phase. Furthermore, pH profiles were measured in the biofilm of section 1 by applying a pH microelectrode with a tip diameter of 30 ␮m and a pH-sensitive glass cone 250 ␮m in length (38). Profiles were recorded directly in the filter biofilm (applying a manually controlled micromanipulator positioned at an angle of 45° to the biofilm surface) as well as in a few samples in the setup described above. A commercial reference electrode (REF201; Radiometer Analytical SAS, Lyon, France) was inserted into the biofilm about 1 cm from the point of measurement. The step size was 100 ␮m, and profiling was stopped at a depth of 500 ␮m to avoid sensor damage. Profiles within one filter section were recorded on the same day. Heterotrophic O2 consumption. Oxygen consumption by heterotrophy alone was obtained from microprofiles measured during in situ acetylene inhibition of nitrifiers (3). Applying the setup described above and slim-line O2 microsensors with tip diameters of only 2 to 5 ␮m, two successive O2 profiles were recorded in the same spot of the biofilm, assuring profile stability after repetitive sensor penetration. Acetylene (1%; saturated solution mixed in filter water) was then added dropwise to the biofilm surface as three doses of 1 ml each to inhibit nitrification. After 10 min of incubation with acetylene, an additional O2 profile was recorded in the same spot. Biofilm transplantation. To investigate the degree of substrate limitation near the filter outlet, biofilm samples from section 2out were exposed to inlet air by transplantation to the front filter compartment, where O2 profiles were recorded applying the procedure described above. Biofilm diffusivity. The apparent O2 diffusivity (Dm ⫽ ␾ ⫻ Ds, where Ds is the biofilm diffusion coefficient and ␾ is the porosity) in the biofilm was determined at a temperature of 20°C by use of a microscale diffusivity sensor, as described by Revsbech et al. (39). The microsensor tip was inserted in a biofilm sample embedded in a 1% agar solution containing 1% ZnCl to stop bacterial activity. The sensor measures concentrations of tracer gas, N2O, diffusing from an internal gas reservoir within the sensor tip. The apparent O2 diffusivity of the surrounding media is inversely correlated with the N2O concentration at the sensor tip and was calculated as ␤/S ⫺ ␣, where S is the sensor signal and ␣ and ␤ are constants determined by calibration of the sensor in two media with known Dms: 1% agar and 50-␮m glass beads. The O2 diffusivity in 1% agar corresponds to table values of molecular diffusion in water, while the Dm of the glass beads is reduced by a factor of 0.27 (36). The Dm value found was converted to in situ temperatures and salinities through multiplication with the corresponding fractional change of the diffusion coefficient for O2 in water (table values, http://www.unisense.com/Default .aspx?ID⫽117). Calculations. The biofilm O2 uptake was calculated from the profiles as the flux, J, through the upper layer of the biofilm (from depth 0 to depth step 3). Calculations were made according to Fick’s first law of diffusion (7): J ⫽ Dm ⫻ (␦C(x)/␦x), where C(x) is the concentration of the solute at depth x. The biofilm value of Dm determined by the diffusivity sensor was applied. Assuming the concentration profile to represent a steady-state situation, the rate of O2 consumption as a function of depth was calculated applying the numerical model PROFILE (4). Biodiffusivity and irrigation coefficients were assumed to be zero, diffusivity was expressed as ␾ ⫻ Ds(water), and the porosity (␾) was set at 1. Calculated rates were subsequently multiplied by the factor Dm(biofilm)/Dm(water) determined by use of the microscale diffusivity sensor. As the limited spatial resolution of the diffusivity sensor (about 1 mm) did not allow for a resolution of changes in diffusivity with depth, Dm was set to be constant throughout the biofilm. The level of significance used in the F test to calculate choices of O2 consumption zones were set at 0.001 and the stop criteria for the iterative process at 0.001%. Table values were applied for solubility of O2 (http://www.unisense.com/Default.aspx?ID⫽117). Mass balance calculations. The overall area-specific NH3 oxidation rate within the filter was estimated through mass balance calculations based on water drainage, water NOx⫺ concentrations, and filter surface area. NH3 oxidation potentials. In week 15, potential NH3 oxidation rates of suspended biofilms were measured with a NOx⫺ biosensor (Unisense, Aarhus, Denmark). To eliminate the high in situ level of NO2⫺ interfering with the measurements, biofilm samples were brought back to the laboratory and soaked twice for 7 min each time in tap water and twice for 7 min each time in artificial medium (5 mM NH4Cl, 5 mM NaH2PO4, 3‰ NaCl, 0.3 mM CaCl2, 0.4 mM MgSO4; pH adjusted with NaOH to 7.1). Small pieces of support material covering 0.5 to 1.5 cm2 with biofilm on both sides were incubated with 10 ml medium in vials on magnetic stirrers. The NOx⫺ concentration was measured at 1-h intervals for 2 to 4 h in subsamples of 2 to 3 ml, which were temporarily

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TABLE 1. Characteristics of filter water collected in weeks 3, 15, and 41 Water temp (°C)

Cation (mM)

6.59–6.85 NDa

16.3–17.2 ND

15.5–17.3 ND

42 9

103–116 25

1 2

7.2–7.4 ND

11.7–12.3 ND

15.7–16.2 ND

67 21

1 2

6.2–6.3 ND

17.1–18.2 ND

15.8–17.4 ND

31 13

pH

Week 3

1 2

Week 15 Week 41 a b c

Concn of:

Conductivity (mS cm⫺1)

Filter section

Week

b

⫺

NH4⫹aq (mM)

HNO2aq (␮M)

NH3aq (␮M)

⬍10 ND

61–74 16–40

43–73 ND

72–144 ND

78–84 58

⬍10 ND

17–41 26–37

9–15 ND

76–303 ND

⬍10 ND

116–123 37

85–92 24–40

BDLc ND

45–53 ND

NO2

aq

(mM)

⫺

NO3

aq

(mM)

ND, not determined. Cations other than NH4⫹. BDL, below detection limit.

removed from incubations. Medium with no biofilm added was used as a control. The advantages of using the biosensor were that no filtering or other pretreatment of the sample was necessary and that the analyzed subsample could be transferred back to the incubation vial after each measurement. NH3 oxidation rates were calculated as NOx⫺ production over time per measured sample surface area. An average rate from three replicate experiments was calculated for each filter site. Identification and quantification of nitrifying bacteria. Fixation of biofilm samples, preparation of cryosections, and fluorescence in situ hybridization (FISH) were performed as described previously (42) with minor modifications. In brief, triplicate biofilm samples were paraformaldehyde fixed (4%, 1 h, on ice) immediately after microsensor measurements in week 3 and week 15, washed in phosphate-buffered saline, embedded in an optimum-cutting-temperature compound (Tissue-Tek; Sakura Finetek Europe B.V., Zoeterwoude, The Netherlands) for ⬎4 h, frozen on dry ice, and stored at ⫺80°C. Vertical sections (thickness, 14 ␮m) were prepared in a cryostat (Walter Dittes, Heidelberg, Germany), immobilized on gelatin-coated multiwell slides, dehydrated in an ethanol series, and stored at room temperature for several months. The probes EUB I-III (8), NON (28), NSO1225, NSO190, Nsv443 (32), NEU (47), Nse1472 (21), NIT3 (48), and Ntspa662 (10) were fluorescently labeled with Cy3 (biomers.net) and used for FISH according to published protocols (42); all probe sequences and hybridization conditions can be found at http://www.microbial -ecology.net/probebase (27). After DAPI (4⬘,6-diamidino-2-phenylindole) staining and mounting (42), FISH and DAPI image pairs were recorded as optical sections along 15 to 20 random transects (from biofilm base to surface) per sample on an Axiovert 200 M Apotome epifluorescence microscope (Carl Zeiss, Jena, Germany), resulting in 100 to 200 images per sample; for each transect, biofilm thickness was measured microscopically. The relative abundance of AOB was determined with the image analysis program Daime (9) as the area of FISH-positive cells relative to the area of all bacteria stained by DAPI in the same optical section; autofluorescent noncellular objects were manually deselected, and the final area fractions were calculated after correction with the negative control probe NON. To confirm the FISH quantification with a larger sample size, bacterial ammonia monooxygenase gene (amoA) copy numbers were determined relative to prokaryotic 16S rRNA gene copy numbers by quantitative PCR (qPCR) and converted to relative AOB abundance (18). DNA was extracted in triplicate from week 3 and week 15 samples (0.20 to 0.25 g of pooled biofilm) by a combination of enzymatic and chemical lysis and bead beating (13). DNA concentrations were determined on a NanoDrop ND-1000 (Saveen Werner), and qPCR was run with 10 ng of template DNA and the primer pairs AmoA-1F–AmoA-2R-TC (33) and 907F-1492R (25) as described previously (18). In addition, archaeal amoA genes were assayed by the use of three different primer sets as described previously (13, 14, 22). Statistical analysis. Overall differences in O2 influx, NH3 oxidation potentials, or AOB abundance among the different filter sites within one time period was tested by a Kruskal-Wallis test. Pairwise comparisons were performed by MannWhitney U tests. The effect of time and filter site on AOB qPCR ratios was tested by a two-way analysis of variance (data were log10 transformed). Results for filter sites within one time period were compared pairwise by a Mann-Whitney U test.

RESULTS Air and water measurements. Within each sampling period, which spanned a week, pH was stable, varying with ⱕ0.3 pH units (Table 1). Across the entire experimental period, pH varied from 6.2 to 7.4. In section 1, NOx⫺ and NH4⫹ concentration ranges were 78 to 123 mM and 17 to 92 mM, respectively. In section 2, the levels of both NOx⫺ and NH4⫹ were markedly lower, around 25 to 58 mM and 16 to 40 mM, respectively. In week 3, test strips (Merck KGaA, Darmstadt, Germany) indicated that all NOx⫺ was NO2⫺, while in week 41 it was all converted to NO3⫺. The content of cations other than NH4⫹ was high, and due to water evaporation, concentrations more than doubled between section 2 and section 1. Air-phase NH3 declined from 2 to 5 ppm at the filter inlet to 0 to 1.5 ppm in the outlet air. Biofilm appearance and characteristics. The biofilm appearance differed substantially between the front and the back of each filter section (Fig. 2C to F). The front edges of section 1in and 2in were dominated by a thick and fluffy growth (ⱕ3 mm thick), but biofilm thickness decreased rapidly within 1 cm from the edge. Water was running above and often also below the biofilm. On the back of each filter section, the substratum was covered with a very thin biofilm (ⱕ1 mm deep) which was interrupted by a few dispersed clumps of slightly denser growth. At all filter sites, dipteran larvae and protozoans were grazing on the biofilm, but a weekly wash-down of the biomass at the front of section 1 was necessary to keep airflow resistance low. Although the biofilm was gelatinous, with high water content, the determined apparent diffusivity of 1.38 䡠 10⫺5 cm2 s⫺1 differed substantially from that in water (2.10 䡠 10⫺5 cm2 s⫺1). In situ microscale profiles. (i) Microsensor profiles and in situ aerobic activity. In situ microsensor profiles showed a consumption of O2 in the upper biofilm layers (Fig. 2) and a constant pH throughout the biofilm (data not shown). The pH of the biofilm in section 1 varied between 6.6 and 7.5 depending on the date of the experiment. The aerobic activity of the biofilm was strongly dominated by heterotrophs, which accounted for 78 to 100% and 73 to 100% of the total O2 uptake in the biofilms of sections 1in and 2in, respectively (averages, 93.3% ⫾ 8.7% and 91.0% ⫾ 9.6%). An indication of nitrification was found only in section 2out, in

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FIG. 2. Average O2 profile measured by microsensor (squares) and average simulated O2 consumption (bars) in week 3 sections 1in (A), and 2in (B) and in week 15 sections 1in (C), 1out (D), 2in (E), and 2out (F). Error bars show standard deviations for five (week 3) and three (week 15) replicate biofilm samples. The dotted line in panel A represents the O2 profile in a sixth sample, which showed a markedly different profile (probably caused by faunal activity) and was not included in the calculations. Insets in panels C to F show the appearance of the biofilms in the different filter sections.

which it accounted for 3 to 38% of the O2 uptake (average, 24.8% ⫾ 18.36%). Both week 3 and week 15 measurements showed a decreasing aerobic activity from the filter inlet toward the filter outlet, with the total O2 consumption in section 1 being significantly higher than that in section 2 (Table 2, Pweek3 ⫽ 0.006 and Pweek15 ⫽ 0.05). In week 15, the front and back biofilms of

section 1 (1in and 1out) showed equal O2 consumption rates, while O2 uptake in section 2out was significantly decreased compared to that in section 2in (P ⫽ 0.05). Also, the total O2 consumption in section 1in was markedly increased compared to that in week 3 (Table 2). (ii) Transplantation experiments. Transferring biofilm from filter outlet to filter inlet resulted in a significant increase in

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TABLE 2. Total O2 uptake calculated from in situ microsensor profiles and AOPs from batch incubations of biofilmsa O2 influx (pmol O2 cm⫺2 s⫺1)

AOP (pmol NH4⫹ cm⫺2 s⫺1)

Week 3 1in 2in

139 ⫾ 16 (5) 82 ⫾ 21 (5)

ND ND

Week 15 1in 1out 2in 2out 2out.transp

204 ⫾ 28 (3) 205 ⫾ 36 (3) 107 ⫾ 15 (3) 52 ⫾ 8 (3) 146 ⫾ 42 (3)

9 ⫾ 4 (3) 7 ⫾ 2 (3) 79 ⫾ 40 (3) 33 ⫾ 8 (3) ND

Week 41 2out 2out.transp

74 ⫾ 24 (6) 125 ⫾ 19 (6)

ND ND

Week and biofilm site

a Values are means ⫾ standard deviation (numbers of replicates are in parentheses). ND, not determined; transp., biofilm transplanted to front filter compartment.

biofilm O2 consumption, by a factor of 1.7 to 2.8 (Fig. 3 and Table 2) (Pweek15 ⫽ 0.05; Pweek41 ⫽ 0.006). In week 41, acetylene addition showed that this effect could be explained by an increase in heterotrophic activity alone, while no significant changes in nitrification rates were observed. Identity and distribution of microorganisms. Biofilm thickness varied from 760 to 2,660 ␮m (average, 1,570 ␮m) and from 570 to 2,717 ␮m (average, 1,368 ␮m) at the front of sections 1 and 2, respectively. DAPI staining showed increasing cell density from a loose surface layer consisting mostly of autofluorescent fungal hyphae to a compact, bacterium-dominated zone at the base of the biofilm. In week 3, the fungusdominated zone of section 1in comprised only the upper 57 to 380 ␮m, while this zone extended to a 380- to 2,090-␮m depth in week 15. Section 2in generally contained fewer fungal hyphae but showed a high abundance of fungal spores. Most AOB could be identified as Nitrosomonas eutropha-like phylotypes based on positive hybridization with probes NSO1225 (targeting all AOB), NEU (targeting the N. eutro-

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pha/N. europaea/N. halophila lineage plus Nitrosococcus mobilis), and Nse1472 (targeting N. eutropha, N. europaea, and N. halophila; Fig. 4A and C) and negative results with probe NSO190, which targets most AOB but has a central mismatch to all N. eutropha species (24). Very few cell clusters (⬍5% of all AOB, or ⬍0.04% of all cells) were found to hybridize to probes NSO190 and/or Nsv443 (Fig. 4B), indicating the presence of only a few N. europaea/N. halophila- and Nitrosospiralike AOB. Quantification was therefore done for probe Nse1472 only. In week 15, the area-based fraction of Nse1472positive AOB was significantly higher in section 2in (0.73% ⫾ 0.15% of DAPI-stained cells; range, 0.6 to 0.9%) than in section 1in (0.17% ⫾ 0.12% of DAPI-stained cells; range, 0.1 to 0.3%; P ⫽ 0.046), with AOB being present in 47.8% and 8.1% of the inspected microscopic fields, respectively. The same pattern was found in week 3 but with an overall higher AOB abundance (Fig. 4C). This trend was confirmed by the qPCR results, which also indicated higher relative abundances of AOB in week 3 than in week 15 and in section 2in (week 3, 0.117% ⫾ 0.083%; week 15, 0.036% ⫾ 0.008%) than in section 1in (week 3, 0.010% ⫾ 0.007%; week 15, 0.003% ⫾ 0.001; Pweek3 ⫽ 0.05; Pweek15 ⫽ 0.05). Furthermore, these numbers showed that AOB accounted for less than 0.12% of all bacterial cells. Archaeal ammonia oxidizers were never detected by PCR of archaeal amoA genes. FISH analysis indicated that NOB were present in both filter sections during weeks 3 and 15, but in very low numbers (not quantified), and they belonged exclusively to the genus Nitrobacter, as determined with the probe NIT3 (Fig. 4D). Nitrospira-like cells were never detected. Unfortunately, paraformaldehyde-fixed biofilm samples for FISH were not recovered from week 41. AOP. During the incubations for determination of potential NH3 oxidation rates, the biofilm was suspended by the magnetic stirrer, and O2 measurements never showed ⬍70% air saturation. It can thus be assumed that all AOB were exposed to oxic conditions during the assay and that no denitrification occurred. The measured ammonia oxidation potentials (AOPs) supported the skewed AOB distribution toward section 2 shown by FISH. A nine-times-higher NH3 oxidation

FIG. 3. Week 41. Average measured O2 profile (squares) and average simulated O2 consumption (bars) in section 2out biofilm positioned in the back filter compartment (A) and section 2out biofilm transplanted to the front filter compartment (B). Error bars indicate standard deviations for six replicate biofilm samples.

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FIG. 4. Nitrifying bacteria detected by Cy3-labeled oligonucleotide probes (red). (A) Week 15 section 2in, probe Nse1472; (B) week 3 section 2in, probe NSV443; (C) week 3 section 2in, probe Nse1472; (D) week 15 section 2in, probe NIT3. All bacteria were stained with DAPI (blue). Autofluorescent material is shown in green (FITC channel). Bar, 10 ␮m.

capacity was observed in section 2in than in section 1, indicating a significantly higher AOB abundance in this filter section (Table 2) (P ⫽ 0.05). In sections 1in and 1out, only very low AOPs were observed (Table 2). The average AOP was markedly lower in section 2out than in section 2in (Table 2) (P ⫽ 0.05). At all filter sites, maximum rates of NOx⫺ production were reached immediately after the start of incubation. Whole-filter AOR. Assuming the main AOB activity to be located in section 2, mass balance calculations estimated the area-specific ammonia oxidation rates (AOR) of this filter section to be 24 to 26 and 41 to 46 pmol NH3 cm⫺2 s⫺1 in week 3 and week 41, respectively (Table 3). In this calculation, estimates of denitrification rates based on N2O microscale pro-

files recorded within the biofilm were included as an additional NO2⫺ sink (L.P. Nielsen, S. Juhler, M. Andersen, and K. Sørensen, unpublished data).

DISCUSSION Heterotrophic activity. Overall aerobic activity of the biofilm was strongly dominated by heterotrophs, as shown by the marginal change in O2 profiles upon acetylene inhibition and the much higher total O2 consumption rate of 52 to 205 pmol O2 cm⫺2 s⫺1 (Table 2) compared to the AOR of 12 to 23 pmol NH4⫹ cm⫺2 s⫺1 (Table 3). Taking into account the fact that

TABLE 3. Whole filter AOR in week 3 and week 41 based on mass balance calculations

Week

Week 3 Week 41

Rate (␮mol NOx⫺ s⫺1) of NOx⫺:

Water drainage (m3 d⫺1)

关NOx 兴 (mM)

0.25 0.5

103–116 116–123

⫺

Removal with drainage watera

Consumption by denitrificationb

Production by nitrificationc

298–336 671–712

181–203

479–538 852–914

Total filter surface area (m2)

4,050

Area-specific AOR (pmol NH4⫹ cm⫺2 s⫺1) Whole filterd

Section 2e

12–13 21–23

24–26 42–46

Water drainage multiplied by the NOx⫺ concentration. From denitrification rates measured as N2O production in the biofilm (Nielsen et al., unpublished), assuming denitrifying biofilms to cover the front 1 cm of each filter section. c Total amount of NOx⫺ removed from the filter (NOx⫺ removed with drainage water plus NOx⫺ consumed by denitrification). d Average for the whole filter. e Assuming AOB activity to be restricted to this filter section only. a b

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each NH3 molecule consumes 1.5 O2 molecules, the heterotrophs accounted for, on average, 87% of the O2 consumption in the filter. Affirmatively, AOB constituted only ⬍0.12% of the bacterial cells in the biofilm. Thus, although NH3 oxidation removed 70 to 100% of the inlet NH3, it contributed relatively little to the overall aerobic activity in the filter, indicating that the load of VOC was substantially higher than the NH3 load. With an air load of 10 to 20,000 m3/h, the heterotrophic O2 uptake through the filter corresponded to an inlet VOC concentration of 16 to 44 ppm acetic acid equivalents, assuming respiration to CO2 and 50% growth efficiency. This is orders of magnitude higher than what has been observed in other piggeries (31). It would be very interesting to know the identity of these large amounts of VOC, especially if organic nitrogen compounds could be another significant source of NH3 for nitrification. From the filter inlet toward the outlet, biofilm aerobic activity decreased significantly (Table 2). Transplantation of biofilm from section 2out to the front filter compartment caused an increase in the respiration rate by a factor of 1.7 to 2.8, which exceeded the O2 consumption explainable by the NH3 oxidation capacity of this biofilm (Table 2). Also, acetylene inhibition showed that the enhanced respiration rate could be explained by an increase in heterotrophic activity alone. The general decrease in aerobic activity through the filter could thus be attributed to heterotrophic substrate limitation, caused by the removal of the most soluble organics in the upstream filter sections. Across all time periods, the maximal volume-specific O2 consumption of the biofilm was always 8 to 12 nmol O2 cm⫺3 s⫺1 (Fig. 2), which must represent the maximum possible specific activity and compression of bacterial and fungal cells with an unlimited supply of substrate and O2. When this maximum O2 respiration was sustained throughout the oxic zone of the biofilm (week 15, section 1in) (Fig. 2C), O2 consumption reached 200 pmol O2 cm⫺2 s⫺1 (Table 2), representing the upper limit for aerobic degradation of organic matter within the biofilm. Any excess organic substrate thus had to be degraded by denitrification in anoxic biofilm layers. From Fig. 2C it can also be deduced that even under maximal aerobic respiration, an anoxic zone can develop only in biofilms thicker than about 250 ␮m. In section 1 the load of organic substrates appeared to vary with time. During week 3, the maximum O2 respiration rate was reached only in the upper 150 to 200 ␮m of the biofilm (Fig. 2A), indicating limited penetration of VOC. In week 15, maximum O2 fluxes and high O2 respiration rates in the oxic zone indicated that easily degradable organic substrates penetrated to the anoxic zones of both section 1in and section 1out (Table 2; Fig. 2C and D). AOB distribution and activity. AOB abundance and biofilm AOPs (Table 2) clearly demonstrated a highly skewed distribution of AOB within the air filter, indicating the main NH3 oxidation to be located in section 2. The biofilm of section 2in had a relative AOB abundance and an NH3 oxidation capacity that were 4 to 12 and 8.8 times higher than those in section 1in, respectively. In spite of possible biases on DNA extraction and PCR as well as the fact that samples for qPCR covered larger areas of each filter site while samples for FISH represented only the outermost biofilm, AOB quantification by FISH and


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qPCR showed the same trend. The filter mass balances (Table 1) confirmed the restriction of AOB to section 2, as the increase in NOx⫺ concentration from section 2 to section 1 could be explained solely by evaporative condensation in section 1, as seen for the inert cations. Several previous investigations of mixed biofilm communities of heterotrophic and nitrifying bacteria from wastewater treatment have demonstrated competition for space and O2 between these two groups of microorganisms (12, 35, 51). Due to significantly lower specific growth rate and dependency on O2, the nitrifiers can establish persistent populations only in biofilm strata where the heterotrophs are limited by substrate and not by O2. In cases of high organic loading, such as in section 1 (week 15) (Fig. 2C and D), O2 is depleted within a highly active heterotrophic surface layer, thus outcompeting AOB from the biofilm. In section 2, however, heterotrophs become increasingly substrate limited, allowing O2 to diffuse to the nitrifying zone. As a result, AOB activity is relegated to section 2. Supporting this hypothesis is the markedly higher AOB abundance in week 3, where a deep O2 penetration indicated substantial heterotrophic substrate limitation already in section 1 (Fig. 2A), allowing the establishment of a larger AOB population in the biofilm. Assuming all NH3 oxidation to take place in section 2, AORs obtained from mass balance calculations corresponded to 44 to 48% of the total O2 consumption in section 2in (Tables 2 and 3), thus contradicting results obtained from acetylene inhibition experiments, which showed nitrification to account for, on average, 9% of the O2 uptake. However, the low AOR measured in situ was determined from microsensor measurements performed in the thick biofilm at the very front edge of the filter pad (Fig. 2E). Similar experiments subsequently performed in analogous biotrickling filters (unpublished data) indicated that in section 2in, the nitrifying activity was situated not in this thick biofilm but rather in the thinner biofilm further inside the filter pad, the latter of which was included in measurements of biofilm AOPs and AOB quantifications. Thus, even though the aerobic activity at the very front biofilm was highly dominated by heterotrophs, the main part of section 2 was presumably better represented by the acetylene inhibitions in section 2out, which approached values from mass balance calculations by suggesting NH3 oxidation to account for 3 to 38% of the total O2 consumption. Inhibition. In addition to competition for space and O2, single-cell activity could be affected by chemical inhibition. Free nitric acid is a well-known inhibitor of nitrification (2, 15, 45) and may control AOB activity and in turn also pH in biofilters (34): as NH3 oxidation causes FNA to build up due to NO2⫺ and H⫹ production, AOB activity results in self-inhibition, forcing a slowdown of the process until acid production is balanced by the continuous dissolution of NH3 in the water. As a result, the filter water becomes a solution of ammonium nitrite maintained near neutral pH. Recently, Vadivelu et al. (46) showed that growth of AOB and NOB is much more sensitive to inhibitors than their activity is. In the continuous function of biofilters, the limits of growth should therefore be much more important than the more commonly studied limits of activity. In enrichment cultures, it was found that growth of AOB was more than 95% inhibited around 30 ␮M FNA, while catabolism was repressed

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by only 50% (46). To counteract the slow but inevitable loss of biomass, the average FNA concentration in the biofilters should therefore vary around 30 ␮M to keep a steady-state biomass. With 50% inhibition of activity, this steady-state biomass should represent a potential AOR twice the average realized AOR, which in turn is controlled by the NH3 load. Even though spatial and temporal variations should blur the picture, this hypothesis was very well supported by the observed in situ AOR in section 2 corresponding to 43 to 46% of the NH3 oxidation capacity of the biofilm (Tables 2 and 3). Biotrickling filters, in which water drainage is controlled according to a fixed conductivity threshold value, rarely reduce the NH3 emission by a certain fraction but instead reduce it to a fixed level around or slightly below 1 ppm, disregarding filter inlet NH3 concentrations (SKOV A/S). This observation could be explained through the above-mentioned growth inhibition hypothesis: If the total NO2⫺ and NH4⫹ concentrations are fixed by filter operation and the FNA concentration is fixed by growth kinetics, the pH and the free ammonia (FA; or NH3) concentration are fixed as well, according to the first equation above. In week 15, as an example, the measured values of 58 mM NO2⫺ and 26 to 37 mM NH4⫹ in filter 2 yield a pH of 6.7 and NH3 concentrations of 50 to 70 ␮M, assuming an FNA concentration of 30 ␮M. From the NH3 solubility (41), it is calculated that these concentrations balance with a partial pressure of 0.6 to 0.9 ppm NH3 in the air phase, which indeed represents the typical outlet concentrations. FA also inhibits AOB activity. However, as FA acts as both a substrate and an inhibitor of nitrification, the effect of this compound is more complex. FA has been shown to inhibit NH3 oxidation in concentrations from 0.58 to 8.8 mM (2). With a pKa around 9.5 at the temperatures and salinities prevailing in the filter (20), only a minor fraction of the NH4⫹ was present as NH3 in the water at the measured pH of 6.2 to 7.5 (Table 1), and the concentrations did not reach the previously observed inhibition limits. The fact that the AOB population of the biofilm was heavily dominated by N. eutropha correlates well with the conditions prevailing in the filter. Besides showing a high NH3 tolerance (600 mM NH4Cl at pH 8) as well as a high substrate affinity constant (KS,NH3, 30 to 61 ␮M), N. eutropha tolerates high ionic strength (up to 400 mM NaCl) and is often the dominating AOB in wastewater treatment plants and habitats subjected to eutrophication (23, 24). Heterotrophic activity can also be severely impeded by FNA and FA (26, 40, 50), with distinct bacterial species showing highly varying tolerances for these compounds (6, 49). While FA and FNA therefore most likely influenced the diversity of heterotrophic microorganisms within the air filter, it is presently unclear to what extent their joint activity and hence the VOC removal are affected as well. NOB. Frequently, NOB are not found to be active in the biotrickling filters, which has commonly been explained by FA and FNA inhibition of the NO2⫺-oxidizing process (2, 29). When first established in the biofilm, NOB should promote their own activity by oxidizing NO2⫺, which relieves FNA inhibition not only of the NOB themselves but also of AOB. This allows NH3 oxidation to decrease the pH, resulting in furtherreduced FA inhibition of the NOB. FISH revealed the presence of NOB belonging to the genus Nitrobacter in samples


from both week 3 and week 15, although they were found only as single cells and in very low abundance. The chemical data indicated no NOB activity until week 41, when almost all NO2⫺ was converted to NO3⫺, indicating full NOB activity in the filter (Table 1). According to Vadivelu et al. (46), the above-calculated FA concentration of 50 to 70 ␮M in section 2 corresponded to only 25 to 33% inhibition of NOB growth. However, the assumed FNA level of 30 ␮M by far exceeded 1.6 ␮M, above which no NOB growth was detected (46). Unless the NOB are established together with the AOB during filter startup, they should thus be permanently excluded by FNA inhibition. Other unresolved mechanisms are therefore required to explain the observed emergence of NO2⫺ oxidation in week 41. General functioning of the filters. Our general understanding of the functioning of this type of trickling filter is that NH3 is captured into a large pool of NH3 and NH4⫹ in the recirculating water and then subsequently slowly oxidized by the activity of partly FNA-inhibited AOB. The oxidation of at least some of the VOC constituents seems to be more direct, as a transplantation of biofilm from the back of the filter to the front section resulted in an immediate increase in respiration rate. Advanced methods like microautoradiography combined with FISH should be applied to identify the layers and species of microorganisms responsible for the various degradation processes. ACKNOWLEDGMENTS This research was funded by the Danish Natural Science Research Council (grant no. 272-06-0504) and by the Danish Ministry of Food, Agriculture and Fisheries as part of the project “Function of Biological Airfilters” under the program “Animal Husbandry, the Neighbors and the Environment” (grant no. 3304-VMP-05-0004). We thank Lindhart B. Nielsen for very generous access to the biotrickling filter and Lise B. Guldberg from SKOV A/S for cooperation and highly supportive documentation about the filters. We also acknowledge Preben Sørensen, Aaron Saunders, Bent Lorentzen, and Dorethe Jensen from Aarhus University for their very helpful technical assistance. REFERENCES 1. Andersen, K., T. Kjær, and N. P. Revsbech. 2001. An oxygen insensitive microsensor for nitrous oxide. Sensors Actuators B 81:42–48. 2. Anthonisen, A. C., R. C. Loehr, T. B. S. Prakasam, and E. G. Srinath. 1976. Inhibition of nitrification by ammonia and nitrous acid. J. Water Pollut. Control Fed. 48:835–852. 3. Berg, P., L. Klemedtsson, and T. Rosswall. 1982. Inhibitory effect of low partial pressures of acetylene on nitrification. Soil Biol. Biochem. 14:301– 303. 4. Berg, P., N. Risgaard-Petersen, and S. Rysgaard. 1998. Interpretation of measured concentration profiles in sediment pore water. Limnol. Oceanogr. 43:1500–1510. 5. Bower, C. E., and T. Holm-Hansen. 1980. A salicylate-hypochlorite method for determining ammonia in seawater. Can. J. Fish. Aquat. Sci. 37:794–798. 6. Castellani, A. G., and C. F. Niven. 1955. Factors affecting the bacteriostatic action of sodium nitrite. Appl. Microbiol. 3:154–159. 7. Crank, J. 1975. The mathematics of diffusion, 2nd ed. Clarendon Press, Oxford, United Kingdom. 8. Daims, H., A. Bruhl, R. Amann, K.-H. Schleifer, and M. Wagner. 1999. The domain-specific probe EUB338 is insufficient for the detection of all bacteria: development and evaluation of a more comprehensive probe set. Syst. Appl. Microbiol. 22:434–444. 9. Daims, H., S. Lucker, and M. Wagner. 2006. daime, a novel image analysis program for microbial ecology and biofilm research. Methods Enzymol. 8:200–213. 10. Daims, H., J. L. Nielsen, P. H. Nielsen, K.-H. Schleifer, and M. Wagner. 2001. In situ characterization of Nitrospira-like nitrite-oxidizing bacteria active in wastewater treatment plants. Appl. Environ. Microbiol. 67:5273–5284. 11. Delhomeńie, M.-C., and M. Heitz. 2005. Biofiltration of air: a review. Crit. Rev. Biotechnol. 25:53–72.

VOL. 75, 2009 12. Fdz-Polanco, F., E. Mendez, M. A. Uruena, S. Villaverde, and P. A. Garcia. 2000. Spatial distribution of heterotrophs and nitrifiers in a submerged biofilter for nitrification. Water Res. 34:4081–4089. 13. Foesel, B. U., A. Gieseke, C. Schwermer, P. Stief, L. Koch, E. Cytryn, J. R. de la Torre, J. van Rijn, D. Minz, H. L. Drake, and A. Schramm. 2008. Nitrosomonas Nm143-like ammonia oxidizers and Nitrospira marina-like nitrite oxidizers dominate the nitrifier community in a marine aquaculture biofilm. FEMS Microbiol. Ecol. 63:192–204. 14. Francis, C. A., K. J. Roberts, J. M. Beman, A. E. Santoro, and B. B. Oakley. 2005. Ubiquity and diversity of ammonia-oxidizing archaea in water columns and sediments of the ocean. Proc. Natl. Acad. Sci. USA 102:14683–14688. 15. Fux, C., K. Lange, A. Faessler, P. Huber, B. Grueniger, and H. Siegrist. 2003. Nitrogen removal from digester supernatant via nitrite—SBR or SHARON? Water Sci. Technol. 48:9–18. 16. Gellman, E., E. Hurting, G. Brose, and T. Jungbluth. 2004. Determination of the dynamics of the odour release from a pig house, using an electronic odour sensor. Water Sci. Technol. 50:101–108. 17. Hendry, G. A. F., and J. P. Grime. 1993. Methods in comparative plant ecology: a laboratory manual. Springer, New York, NY. 18. Herrmann, M., A. M. Saunders, and A. Schramm. 2008. Archaea dominate the ammonia-oxidizing community in the rhizosphere of the freshwater macrophyte Littorella uniflora. Appl. Environ. Microbiol. 74:3279–3283. 19. Hutchings, N. J., S. G. Sommer, J. M. Andersen, and W. A. H. Asman. 2001. A detailed ammonia emission inventory for Denmark. Atmos. Environ. 35:1959–1968. 20. Johansson, O., and M. Wedborg. 1980. Ammonia-ammonium equilibrium in seawater at temperatures between 5 and 25°C. J. Solution Chem. 9:37–44. 21. Juretschko, S., G. Timmermann, M. Schmid, K.-H. Schleifer, A. Pommerening-Ro ¨ser, H. P. Koops, and M. Wagner. 1998. Combined molecular and conventional analyses of nitrifying bacterium diversity in activated sludge: Nitrosococcus mobilis and Nitrospira-like bacteria as dominant populations. Appl. Environ. Microb. 64:3042–3051. 22. Ko ¨nneke, M., A. E. Bernhard, J. R. de la Torre, C. B. Walker, J. B. Waterbury, and D. A. Stahl. 2005. Isolation of an autotrophic ammonia-oxidizing marine archaeon. Nature 437:543–546. 23. Koops, H. -P., and A. Pommerening-Ro ¨ser. 2001. Distribution and ecophysiology of the nitrifying bacteria emphasizing cultured species. FEMS Microbiol. Ecol. 37:1–9. 24. Koops, H.-P., U. Purkhold, A. Pommerening-Ro ¨ser, G. Timmermann, and M. Wagner. 2003. The lithoautotrophic ammonia-oxidizing bacteria, p. 778– 881. In M. Dworkin, S. Falkow, E. Rosenberg, K.-H. Schleifer, and E. Stackebrandt (ed.), The prokaryotes, 3rd ed., vol. 5. Springer, New York, NY. 25. Lane, D. J. 1991. 16S/23S rRNA sequencing, p. 115–175. In E. Stackebrandt and M. Goodfellow (ed.), Nucleic acid techniques in bacterial systematics. John Wiley and Sons, New York, NY. 26. Lee, S.-M., J.-Y. Jung, and Y.-C. Chung. 2000. Measurement of ammonia inhibition of microbial activity in biological wastewater treatment process using dehydrogenase assay. Biotechnol. Lett. 22:991–994. 27. Loy, A., F. Maixner, M. Wagner, and M. Horn. 2007. probeBase—an online resource for rRNA-targeted oligonucleotide probes: new features 2007. Nucleic Acids Res. 35:D800–D804. 28. Manz, W., R. Amann, W. Ludwig, M. Wagner, and K.-H. Schleifer. 1992. Phylogenetic oligodeoxynucleotide probes for the major subclasses of proteobacteria—problems and solutions. Syst. Appl. Microbiol. 15:593–600. 29. Melse, R. W., and G. Mol. 2004. Odour and ammonia removal from pig house exhaust air using a biotrickling filter. Water Sci. Technol. 50:275–282. 30. Melse, R. W., and N. W. M. Ogink. 2005. Air scrubbing techniques for ammonia and odor reduction at livestock operations: review of on-farm research in the Netherlands. Trans. ASAE 48:2303–2313.


3713

31. Merchant, J. A., and R. F. Ross. 2002. Iowa Concentrated Animal Feeding Operations Air Quality Study. Iowa State University and The University of Iowa Study Group, Ames. 32. Mobarry, B. K., M. Wagner, V. Urbain, B. E. Rittmann, and D. A. Stahl. 1996. Phylogenetic probes for analyzing abundance and spatial organization of nitrifying bacteria. Appl. Environ. Microbiol. 62:2156–2162. 33. Nicolaisen, M. H., and N. B. Ramsing. 2002. Denaturing gradient gel electrophoresis (DGGE) approaches to study the diversity of ammonia-oxidizing bacteria. J. Microbiol. Methods 50:189–203. 34. Nielsen, L. P., M. L. Nielsen, M. Andersen, and A. M. Nielsen. 2007. Ammonia transformations in a biotrickling filter, p. 263–271. In C. Kennes and M. C. Veiga (ed.), Biotechniques for air pollution control. Universidad da Corun ã, A Corun ã, Spain. 35. Okabe, S., K. Hirata, and Y. Watanabe. 1995. Dynamic changes in spatial microbial distribution in mixed-population biofilms: experimental results and model simulation. Water Sci. Technol 32:67–74. 36. Revsbech, N. P. 1989. Diffusion characteristics of microbial communities determined by use of oxygen microsensors. J. Microbiol. Methods 9:111–122. 37. Revsbech, N. P. 1989. An oxygen microsensor with a guard cathode. Limnol. Oceanogr. 34:474–478. 38. Revsbech, N. P., and B. B. Jørgensen. 1986. Microelectrodes—their use in microbial ecology. Adv. Microb. Ecol. 9:293–352. 39. Revsbech, N. P., L. P. Nielsen, and N. B. Ramsing. 1998. A novel microsensor for determination of apparent diffusivity in sediments. Limnol. Oceanogr. 43:986–992. 40. Rowe, J. J., J. M. Yarbrough, J. B. Rake, and R. G. Eagon. 1979. Nitrite inhibition of aerobic bacteria. Curr. Microbiol. 2:51–54. 41. Sander, R. 1999. Compilation of Henry’s law constants for inorganic and organic species of potential importance in environmental chemistry. http: //www.mpch-mainz.mpg.de/⬃sander/res/henry.html. 42. Schramm, A., D. de Beer, M. Wagner, and R. Amann. 1998. Identification and activities in situ of Nitrosospira and Nitrospira spp. as dominant populations in a nitrifying fluidized bed reactor. Appl. Environ. Microbiol. 64: 3480–3485. 43. Sheridan, B., T. Curran, V. Dodd, and J. Colligan. 2002. Biofiltration of odour and ammonia from a pig unit—a pilot-scale study. Biosyst. Eng. 82:441–453. 44. Swanson, W. J., and R. C. Loehr. 1997. Biofiltration: fundamentals, design and operations principles, and applications. J. Environ. Eng. 123:538–546. 45. Tan, N. C., M. J. Kampschreur, W. Wanders, W. L. van der Pol, J. van de Vossenberg, R. Kleerebezem, M. C. van Loosdrecht, and M. S. Jetten. 2008. Physiological and phylogenetic study of an ammonium-oxidizing culture at high nitrite concentrations. Syst. Appl. Microbiol. 31:114–125. 46. Vadivelu, V. M., J. Keller, and Z. Yuan. 2007. Free ammonia and free nitrous acid inhibition on the anabolic and catabolic processes of Nitrosomonas and Nitrobacter. Water Sci. Technol. 56:89–97. 47. Wagner, M., G. Rath, R. Amann, H. P. Koops, and K.-H. Schleifer. 1995. In-situ identification of ammonia-oxidizing bacteria. Syst. Appl. Microbiol. 18:251–264. 48. Wagner, M., G. Rath, H. P. Koops, J. Flood, and R. Amann. 1996. In situ analysis of nitrifying bacteria in sewage treatment plants. Water Sci. Technol. 34:237–244. 49. Weon, S. Y., C. W. Lee, S. I. Lee, and B. Koopman. 2002. Nitrite inhibition of aerobic growth of Acinetobacter sp. Water Res. 36:4471–4476. 50. Yarbrough, J. M., J. B. Rake, and R. G. Eagon. 1980. Bacterial inhibitory effects of nitrite—inhibition of active-transport, but not of group translocation, and of intracellular enzymes. Appl. Environ. Microbiol. 39:831–834. 51. Zhang, T. C., Y. C. Fu, and P. L. Bishop. 1994. Competition in biofilms. Water Sci. Technol. 29:263–270.