Nitrification in activated sludge batch reactors is linked to protozoan ...

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Nitrification in activated sludge batch reactors is linked to protozoan grazing of the bacterial population Penny Petropoulos and Kimberley A. Gilbride

Abstract: Protozoa feed upon free-swimming bacteria and suspended particles inducing flocculation and increasing the turnover rate of nutrients in complex mixed communities. In this study, the effect of protozoan grazing on nitrification was examined in activated sludge in batch cultures maintained over a 14-day period. A reduction in the protozoan grazing pressure was accomplished by using either a dilution series or the protozoan inhibitor cycloheximide. As the dilutions increased, the nitrification rate showed a decline, suggesting that a reduction in protozoan or bacterial concentration may cause a decrease in nitrification potential. In the presence of cycloheximide, where the bacterial concentration was not altered, the rates of production of ammonia, nitrite, and nitrate all were significantly lower in the absence of active protozoans. These results suggest that a reduction in the number or activity of the protozoans reduces nitrification, possibly by limiting the availability of nutrients for slow-growing ammonia and nitrite oxidizers through excretion products. Furthermore, the ability of protozoans to groom the heterotrophic bacterial population in such systems may also play a role in reducing interspecies competition for nitrification substrates and thereby augment nitrification rates. Key words: nitrification, activated sludge, protozoan grazing, ammonia-oxidizing bacteria, cycloheximide. Résumé : Les protozoaires s’alimentent à partir de bactéries aquatiques libres et de particules en suspension, entraînant une floculation et augmentant le taux de renouvellement des nutriments dans des communautés complexes mixtes. Dans cette étude, nous avons examiné l’impact du broutage des protozoaires sur la nitrification dans de la boue activée sous forme de cultures discontinues maintenues sur une période de 14 jours. Une réduction de la pression de broutage des protozoaires fut réalisée soit en utilisant des dilutions en série ou avec l’inhibiteur de protozoaires, la cycloheximide. Le taux de nitrification a décliné à mesure que les dilutions augmentaient, indiquant qu’une réduction des concentrations de protozoaires ou de bactéries pourrait entraîner une diminution du potentiel de nitrification. En présence de cycloheximide, où la concentration de bactéries n’était pas altérée, les taux de production d’ammoniac, de nitrite et de nitrate était significativement inférieurs en l’absence de protozoaires actifs. Ces résultats indiquent qu’une réduction du nombre ou de l’activité des protozoaires diminue la nitrification probablement en limitant la disponibilité des nutriments utilisés par les oxydants d’ammoniac et de nitrite à croissance lente alimentés par les produits d’excrétion. Par ailleurs, la capacité des protozoaires à entretenir la population bactérienne hétérotrophe dans de tels systèmes pourrait également jouer un rôle dans la diminution de la compétition inter-espèces pour les substrats de nitrification et ainsi augmenter les taux de nitrification. Mots clés : nitrification, boues activées, broutage de protozoaires, bactéries oxydant l’ammoniac, cycloheximide. [Traduit par la Rédaction]

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Introduction Although molecular nitrogen is abundant, constituting about 79% of the earth’s atmosphere, it is chemically inert and therefore not a suitable source of the element for most living forms that depend on a source of combined nitrogen (i.e., ammonia, nitrate, and organic compounds) or fixed nitrogen Received 8 November 2004. Revision received 3 June 2005. Accepted 6 July 2005. Published on the NRC Research Press Web site at http://cjm.nrc.ca on 21 October 2005. P. Petropoulos and K.A. Gilbride.1 Department of Chemistry and Biology, Ryerson University, 350 Victoria Street, Toronto, ON M5B 2K3, Canada. 1

Corresponding author (e-mail: [email protected]).

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for their nutrition. The cyclic transformation of nitrogenous compounds is therefore of great importance in supplying required forms of nitrogen to the various nutritional classes of organism in the biosphere. The conversion of ammonia to nitrate is a two-step process brought about by two highly specialized groups of obligatory aerobic chemoautotrophic bacteria. In the first step, ammonia is oxidized to nitrite and carried out by a wide variety of beta proteobacterial ammonia oxidizers such as Nitrosomomas and Nitrosococcus (Wagner and Loy 2002). In the second step, nitrite is oxidized to nitrate and is carried out by nitrite oxidizers such as Nitrobacter, Nitrospira, and Nitrospira-like bacteria (Juretschko et al. 1998; Daims et al. 2001). Much of the difficulty in evaluating the role of protozoans in natural ecosystems lies in monitoring the small and rapid changes in nutrient balance in which they play their role. For

doi: 10.1139/W05-069

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this reason, microcosms have been developed that enable both a closer and a more complete monitoring of nutrient cycling than is possible in the field and that also permit more exact measurement of the different components. Although microbial activity plays a key role in the functioning of natural ecosystems (soil and marine and fresh water), they also have a pivotal role in engineered systems (bioremediation and wastewater treatment). Activated sludge is at present the most widely employed biological treatment process for both domestic and industrial wastewaters. The process is dependent on active biomass (e.g., microbial populations), which is mixed in suspension with the wastewater under aerobic conditions. In the presence of adequate nutrients and oxygen, high rates of microbial growth and respiration are achieved. This results in the utilization of organic matter, resulting in the production of oxidized end products, such as CO2, NO3–, SO4–2, and PO4–3, and (or) the biosynthesis of new biomass. Activated sludge treatment removes from the wastewater the biodegradable organics as well as the nonsettleable suspended solids and other constituents, which can be absorbed onto or entrapped by the activated sludge floc (Viessman and Hammer 1998). The microbial community of activated sludge consists of bacteria, protozoans, fungi, algae, and filamentous organisms. Protozoans, filamentous organisms, and bacteria have been shown to actively participate in the biological treatment of wastewater in the activated sludge system. The protozoans are responsible for “grooming” bacterial mass by grazing on it and are, in turn, consumed by rotifers and other higher organisms to establish a complex food web. In this way, a food chain is created, and at each stage of the food chain, a fraction of the original material is removed from the system. Seventy percent of the protozoan species in activated sludge are ciliates (Curds 1975), such as Vorticella, Opercilarias, and Epistylis. Ciliated protozoans of activated sludge processes are associated with the production of clear effluents of good quality. Ciliate protozoans are important predators that have been shown to graze on mostly freefloating bacteria and suspended particles within waste treatment systems (Curds et al. 1968; Wheale and Williamson 1980). In this way, protozoans appear to influence floc formation, which is important for a clear effluent and satisfactory settling. However, protozoans can graze flocculated bacteria when the concentration of suspended bacteria is below a certain threshold. To prevent extinction, they shift their diet from suspended to flocculated bacteria. Crawling ciliates can graze the outside of the flocs and can take up bacteria loosely bound to the floc (Curds 1982; Albright et al. 1987). There have been attempts to relate the physicochemical parameters of effluent or the activated sludge to the species of ciliated protozoans present (Al-Shahawani and Horan 1991; Esteban et al. 1991; Madony et al. 1993). Several hypotheses have been advanced to explain the role of protozoans in activated sludge systems in the enhancement of the mediation of organic compounds. This may be attributed to substrate availability through the excretion of mineral nutrients (phosphorus as phosphate and nitrogen as ammonia or nitrate) by protozoans, resulting in an accelerated usage of organic sources (e.g., carbon and nitrogen) by bacteria (Clarholm 1985; Coleman et al. 1978; Varma et al. 1975). Also, proto-

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zoans have been known to excrete growth-stimulating compounds, which enhance bacterial activity (e.g., flocculation and nutrient mediation) (Nisbet 1984; Ratsak et al. 1996). Lastly, the grazing of bacteria can lead to the selection of species that grow fast and inefficiently (Sherr et al. 1988) and so increase the use of organic sources, since inefficient species must dissimilate more materials to form the same amount of biomass. All this amounts to the loss of energy in the resulting food chain in the activated sludge system (Ratsak et al. 1996). However, it is questionable whether protozoan grazing has a definite impact on bacterial utilization of organic matter, especially within an activated sludge system, since up to now, there has been no direct evidence. The goal of this study was to investigate how nitrification rates are influenced by protozoan predation. The role of protozoans was examined in batch reactors containing activated sludge from a municipal treatment facility that represented a very diverse mixed community. Traditional methods were used to determine the microbial abundances (i.e., bacterial and protozoan numbers) and the physical and chemical parameters, including the concentrations of total nitrogen and the intermediates produced during nitrification. A dilution/extinction method and cycloheximide, a compound that inhibits protozoan metabolism, were used to reduce protozoan grazing pressure to study the effect that protozoans had on the nitrifying ability of the bacterial community.

Materials and methods Sample collection The wastewater sample was obtained from the Ashbridges Bay Treatment Plant in Toronto, Ontario. The sample consisted of a mixed liquor sample obtained from tank 2 (this is approximately one-third the way along the treatment system). The inoculum was transported to the laboratory and used immediately or prepared to be used as diluent (see below). Setup of laboratory microcosms The microcosms consisted of 250-mL Erlenmeyer batch flasks containing 100 mL of sample. They were continuously stirred at room temperature on a bench-scale shaker (VWR-Canlab, Toronto, Ontario) at 150 r·min–1. For the dilution/extinction of protozoans experiments, the original mixed liquor sample was diluted with mixed liquor that had been homogenized at full force for 2 min in a standard blender and filtered through a prewetted 0.45-µm glass-fiber filter (Whatman; VWR-CanLab) to remove active biomass. Flasks were set up in triplicate containing undiluted or fivefold dilutions of the sample (the dilution series was 100.5, 101, 101.5, and 102). For the inhibition of protozoan grazing experiments, cycloheximide (Sigma Co., Oakville, Ontario, Canada) was added to the flasks at a final concentration of 100 mg·L–1. Physical parameter measurements The pH measurement was done on a daily basis with a hand-held pH meter (CorningTM, pH-30 portable pH meter). Temperature measurements were taken using a mercury-filled Celsius thermometer (VWR-CanLab) with a scale marked © 2005 NRC Canada

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for every 0.1 °C. Dissolved oxygen (DO) measurements were taken using a portable DO meter (Extech Instruments, Waltham, Massachusetts). The mixed liquor suspended solids (MLSS) were measured according to standard methods (American Public Health Association 1998). Collected samples (5 mL) were well mixed and filtered through a preweighed 0.45-µm glass-fiber filter (Whatman) (VWR-CanLab). The residue was dried for 1 h at 103 °C. It was then cooled in a desiccator and weighed. It was necessary to repeat the cycle of drying, cooling, desiccating, and weighing until a constant weight was obtained. Chemical oxygen demand The closed reflux colorimetric method was used to measure chemical oxygen demand (COD). This procedure was adapted from standard methods (American Public Health Association 1998). The acclimatized mix liquor sample was filtered through 0.45-µm pore size filter paper (Whatman; VWR-Can Lab). The filtered 2.5-mL sample was placed in culture tubes with Teflon-coated caps (Hach Co., Loveland, Colorado). Then, 1.5 mL of the digestion solution (0.02 mol K2Cr2O7·L–1, 0.1 mol H2SO4·L–1, 0.001 mol HgSO4·L–1 in reagent-grade water) and 3.5 mL of the sulfuric acid reagent (0.02 mol Ag2SO4·L–1 in 0.5 mol H2SO4·L–1) were added. The culture tubes were then placed in a COD block heater (Hach COD reactor, model 45600-00) (Hach Co.) and refluxed at 150 °C for 2 h. The cooled samples were then measured spectrophotometrically (Beckman UV-VIS spectrophotometers) at 600 nm along with potassium hydrogen phthalate standards ranging from 0 to 500 mg O2·L–1). The COD value was calculated as follows: COD (mg O2·L–1) = (mg O2 in the final volume × 1000) / sample volume (mL) Biological oxygen demand The 5-day biological oygen demand (BOD) test was adapted from standard methods (American Public Health Association 1998). Two-millilitre samples were collected and diluted with well-oxygenated and nutrient-containing water (American Public Health Association 1998). The initial DO concentration was determined, and the BOD bottles containing the samples were then stored in the dark at 20 °C for 5 days. The difference in oxygen concentration between the beginning and end of the test period gave the 5-day BOD value. Measurement of ammonia, nitrite, and nitrate concentrations The phenate method modified from standard methods (American Public Health Association 1998) was used to measure ammonia concentrations in the reactors. In brief, the diluted acclimatized mix liquor sample was filtered through 0.45-µm pore size filter paper.(Whatman; VWR-CanLab). To a 2.5-mL sample, 100 µL of phenol solution (11.1% v/v in 95% ethanol), 100 µL of sodium nitroprusside solution (0.5% m/v in reagent-grade water), and 250 µL of oxidizing solution (10 mL of alkaline citrate, 2.5 mL of sodium hypochlorite) were added and mixed thoroughly. The samples were covered and placed in subdued light at room temperature where

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the colour was allowed to develop for a minimum of 1 h. The samples were measured spectrophotometrically (PerkinElmer Lambda 20 spectrophotometer) (Perkin-Elmer Inc., Wellesley, Massachusetts) at 640 nm. A standard curve was prepared by plotting absorbance readings of standards against ammonia concentrations of ammonium chloride standards (0–25 mg NH3-N·L–1). A semimicrocolorimetric method was used for the determination of both nitrite and nitrate using a testing kit purchased from Roche Co. (Laval, Quebec). According to the manufacturer’s procedure, filtered samples were mixed with 770 µL of reagent-grade water and allowed to incubate at room temperature for 30 min. The initial absorbance was measured at 540 nm (Perkin-Elmer Lambda 20 spectrophotometer). After the initial absorbance was measured, 250 µL of sulfanilimide (colour reagent I) and 250 µL of N-(1-naphthyl)-ethylenediamine (colour reagent II) were added. The mixture was then allowed to stand in the dark for 10– 15 min. The final absorbance reading was measured at 540 nm. For the determination of nitrate, samples were mixed with 20 µL of lyophilized nitrate reductase solution (4 U of nitrate reductase dissolved in 700 µL of distilled water) and 250 µL of NADPH buffer solution (0.5 mg of NADPH dissolved in 3 mL of 1 mol K2PO4 ·L–1 buffer, pH 7.5). After the 30-min incubation period, the initial absorbance was measured at 540 nm. Then, 250 µL of sulfanilimide and N-(1-naphthyl)-ethylenediamine, the two colour reagents, were added. After a 10- to 15-min incubation period in the dark, the final absorbance reading was taken. The samples were measured spectrophotometrically at 540 nm against standards of sodium nitrite (0.05–5 mg nitrite·L–1) and potassium nitrate (0.05–5 mg nitrate·L–1) for nitrite and nitrate determination and standard curves were prepared. Measurement of total Kjeldahl nitrogen concentrations The determination of total nitrogen levels was performed using a Nitrogen, Total, Hach Test‘N’TubeTM according to the manufacturer’s instructions (Hach Co.). In brief, a 2-mL sample was collected and added to a reagent vial containing a mixture of 1 mol nitrogen hydroxide·L–1 and 1 mol nitrogen persulfate·L–1. The vials were shaken vigorously for approximately 30 s and then placed in a preheated COD reactor for 30 min of digestion. To the cooled digested vials, 2 mL of 1 mol sodium metabisulfite·L–1 was added to each vial. The vials were shaken for 15 min and 2 mL of 1 mol chromotropic acid·L–1 was added. After 2 min, both the sample and a blank were added to reagent vials containing sodium hydroxide. The vials were incubated at room temperature for 5 min and then measured spectrophotometrically at 410 nm in addition to standards of ammonium p-toluenesulfonate (0–25 mg N·L–1). A standard curve was prepared by plotting the absorbance readings of standards against nitrogen concentrations. Identification and enumeration of protozoans Samples of activated sludge were examined under a phase-contrast microscope and protozoan types were manually assigned into groups based on morphology. The ammoniacal silver carbonate method was used to enumerate the protozoans. The method was adapted from a technical note by Dimas Fernandes-Galiano (Galiano 1994). To 2 mL © 2005 NRC Canada

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of sample, 150 µL of formalin (0.25 mol·L–1), 100 µL of Tween (5% v/v), 100 µL of bacteriological peptone solution (1 mol·L–1), 250 µL of pyridine (1 mol·L–1), 2 mL of ammoniacal silver carbonate solution (2.5 mol·L–1), and 30 mL of distilled water were added. The mixture was shaken and immersed in a water bath (65 °C) until the liquid darkened to a brown–black, usually an incubation period of 10 min. The contents were then poured into a glass evaporation dish containing approximately 50 mL of distilled water. Once the impregnated organisms settled, they were pipetted onto a slide covered with a coverslip and observed under the microscope for computational enumeration using the Northern Eclipse manual count software (Empix Imaging Inc., Mississauga, Ontario). Enumeration of bacteria Molecular Probes’ LIVE/DEAD® BacLight™ bacterial viability kit (Molecular Probes, Eugene, Oregon) was used to enumerate the bacteria. It provides a two-colour fluorescence assay that differentiates between viable and nonviable bacterial cells. This kit utilizes a mixture of SYTO® 9 green fluorescent nucleic acid stain and the red fluorescent nucleic acid stain propidium iodide. These stains differ both in their spectral characteristics and in their ability to penetrate healthy bacterial cells. Equal volumes of SYTO 9 and propidium iodide were combined in a microfuge tube and mixed thoroughly. For each 1 mL of the bacterial suspension, 3 µL of the dye mixture was added. The reagent mixture of 0.3% v/v DMSO solution was added to the staining solution. It was mixed thoroughly and incubated at room temperature in the dark for 15 min. Five microlitres of the stained bacterial suspension was placed between a slide and covered with a coverslip and observed under a fluorescent inverted microscope equipped with a 485-nm excitation filter and a 500-nm emission filter (Axiovert 200) (Carl Zeiss, Gottingen, Germany). The computational enumeration of viable (stained green) and nonviable (stained red) bacteria was performed with the Northern Eclipse manual count software (Empix Imaging Inc.). Data analysis Statistical analysis compared values obtained with reference microcosms with both diluted and cycloheximide microcosms using the Student t distribution. All statistical calculations were done using a statistical add-on in Microsoft Excel (Microsoft Office 2000 for Windows 98). Growth rates (k) were calculated for both the protozoan and the bacterial population growth using the equation k = ln 2/g, where g is the generation time (Madigan et al. 2000). The production rates of ammonia, nitrite, and nitrate were calculated by the difference (∆) in concentration (C) of ammonia, nitrite, and nitrate over time (t): ∆C/t = ∆(C2 – C1)/t. Nitrification rate was calculated per cell over time.

Results and discussion Composition of the protozoan community The most common predators of bacteria in activated sludge are protozoans (Curds 1982). Examination of the activated sludge from the Ashbridges Bay Treatment Plant indicated that an abundant community of protozoans was present.

Can. J. Microbiol. Vol. 51, 2005 Table 1. Composition of the protozoan community. Protozoan type Amoebas Ciliates Flagellates Rhizopods

Number (cells·mL–1) 2.65 3.68 1.25 3.25

× × × ×

4

10 106 103 105

Percent abundance 0.65 91 0.03 8.1

Enumeration of the protozoans indicated that the total density was approximately 4 × 106 protozoans·mL–1 and was composed primarily of ciliates followed by rhizopods, flagellates, and amoebas (Table 1). Other studies have also identified ciliated protozoans as being predominant in both domestic and industrial treatment plants (Curds et al. 1968; Curds 1975; Al-Shahwani and Horan 1991). It is assumed, then, that ciliates were the primary bacterial grazers in the activated sludge system at the Ashbridges Bay Treatment Plant. Effect of protozoan grazing on nitrification Although batch reactors are closed systems, they can be self-sustaining over a short period, and as such, nutrients are required to be recycled within the microcosm. Therefore, they are useful systems to study nutrient cycling and the conditions that affect the efficiency of these reactions. Limiting protozoan grazing was carried out under two different conditions in our batch reactors: dilution of the predator/prey densities and chemical inhibition of protozoan grazing. Dilution/extinction experiments The batch reactors were prepared using undiluted or fivefold dilutions of the activated sludge ranging from 100.5 to 102. Previous research has shown that dilutions up to 102 will not remove functional richness from the bacterial portion of the sample from this treatment plant (Victorio 1995; Victorio et al. 1996). Furthermore, Franklin et al. (2001) have shown that dilution of a complex microbial community does not change the overall diversity of a mixture until the size of the community is decreased so much that the number of individuals in the mixture approximates the original number of species. Since the effluent that was being used in this study contained approximately 108 bacteria·mL–1 and 106 protozoans·mL–1, neither community should have been diluted to extinction or lost its overall diversity in the dilution range that was used. However, in this study, we hoped that this procedure would effectively reduce the grazing pressure on the bacterial community without removing the nitrification potential. Using serial dilutions of a sewage microbial community to define the relationship between dilutions and the effect on diversity of the populations, Franklin et al. (2001) implied that the predation pressure on bacterial abundance would be very small. However, their system contained substantially fewer protozoans (1.5 × 102 protozoans·mL–1) than ours, and for this reason, we expected that an effect on bacterial abundances owing to reduced protozoan grazing pressure would be observed in the range of dilutions that we used. The reactors were monitored for operational parameters to ensure that conditions would be appropriate for nitrification. The parameters measured were pH, temperature, DO, MLSS, © 2005 NRC Canada

0.041

106 106 106 106 × × × × 1.29 1.46 1.78 2.30

0.008

106 106 106 106 × × × ×

0.029 0.008 0.021 0.007 0.024 0.007 0.023 0.007

*Growth rate per day.

k*

No cyclo

1.29 1.35 1.39 1.43 106 106 106 106 × × × ×

With cyclo

1.32 1.63 1.79 1.99 106 106 106 106 × × × ×

No cyclo

1.32 1.39 1.43 1.47 107 107 107 107 × × × ×

With cyclo

1.19 1.25 1.44 1.63 107 107 107 107 × × × ×

No cyclo

1.19 1.21 1.28 1.32 107 107 107 107 × × × ×

With cyclo

1.40 1.68 1.85 2.01 107 107 107 107 × × × ×

No cyclo

1.40 1.48 1.52 1.56 108 108 108 108 × × × ×

With cyclo

1.27 1.53 1.63 1.72 108 108 108 108 × × × × 1.27 1.31 1.35 1.41

No cyclo Time (days)

1 5 10 14

102 101.5 101 100.5 100

Batch reactor dilution

Table 2. Bacterial viable counts in the batch reactors of the dilution series in the presence and absence of cycloheximide (cyclo).

COD, and BOD. The pH, temperature, and DO content of the batch reactors were carefully monitored, since fluctuations in these parameters could indicate a breakdown in the reactors (Curds 1975; Stout 1980). The DO concentration remained constant at 5.5 mg·mL–1 and the temperature was maintained at 25 ± 2 °C throughout the 14-day period. Since nitrification can occur at DO levels above 2 mg·mL–1 and temperatures above 5 °C (Blackall and Burrell 1999; Princic et al. 1998), it was assumed that the conditions in the reactors were suitable for microbial activity and nitrification. The pH of the system was found to drop gradually from 7.5 to 6.4. Since for every milligram of nitrogen nitrified, 7.2 mg of alkalinity is consumed (Princic et al. 1998), a drop in pH is expected. However, since the pH range for the growth of ammonia oxidizers is 5.8–8.5 and the pH range for growth of nitrite oxidizers is 6.5–8.5, nitrification should be able to be carried out in the reactors in the pH range observed. Therefore, the conditions present in the reactors were able to meet the conditions necessary for nitrification during the period that they were studied. The COD, BOD, and MLSS values were also monitored (data not shown) and found to be reduced accordingly by the dilution series; however, they remained constant within individual reactors over the time monitored. Both bacterial and protozoan numbers in the batch reactors showed an increase in all reactors over the 14-day period (Tables 2 and 3). The growth rates (k) for the bacterial community were calculated and the dilution series did not appear to have affected their overall growth rates (Table 2). The dilution series, however, appeared to allow the protozoan community to increase its growth rate slightly (Table 3), possibly by removing interspecies competition for food. To test for nitrification activity, ammonia, nitrite, and nitrate concentrations were monitored. Dilution of the effluent resulted in reduced concentrations of ammonia, nitrite, nitrate, and total nitrogen in the subsequent reactors in the dilution series at the start of the experiments. The concentrations of all of the compounds, however, increased over time in all of the reactors, resulting in slight increases in the accumulation of the products. Taking into consideration the bacterial numbers present in the dilutions, the rates of production of the compounds were calculated. The rate of ammonia production actually declined as the dilutions increased. Similar results were found with both nitrite and nitrate production (Fig. 1). This suggested that the nitrification potential of the population was reduced owing to the dilution effect, possibly by limiting the availability of nutrients for nitrifying bacteria. Since significant levels of ammonia, nitrite, and nitrate were present in all of the reactors, it was unlikely that nitrogen was limiting for nitrification. However, the reduction in interspecies competition in the reactor in the dilution series may allow a selective environment for fast-growing bacteria to proliferate. Fast-growing heterotrophs can compete with the ammonia-oxidizing bacteria for ammonia and therefore reduce the amount of ammonia available for nitrification. Second, since fewer protozoans were present in the subsequent reactors, a reduction in grazing pressure may also add to the proliferation of fastgrowing bacteria, resulting in a decrease of nutrients available for the slow-growing ammonia oxidizers. Nevertheless,

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Can. J. Microbiol. Vol. 51, 2005 Table 3. Protozoan viable counts in the batch reactors of the dilution series in the absence of cycloheximide*. Batch reactor dilution Time (days)

100

100.5

1 5 10 14

9.50 9.51 9.60 9.60

× × × ×

k†

0.002

6

10 106 106 106

1.75 1.79 1.84 1.87

101 × × × ×

5

10 105 105 105

0.005

5.50 5.54 5.74 5.85

101.5 × × × ×

0.005

4

10 104 104 104

2.60 2.80 2.85 2.85

102 × × × ×

0.006

4

10 104 104 104

1.05 1.09 1.12 1.15

× × × ×

104 104 104 104

0.006

*In the presence of cycloheximide, no protozoans were observed to be active; therefore, final counts were the same as initial counts. † Growth rate per day.

Fig. 1. Effect of diluting the microbial community on the production of (a) ammonia, (b) nitrite, and (c) nitrate in the batch reactors. Mean values are shown with error bars indicating the standard deviation.

since there was not a corresponding increase in bacterial growth rates with the lower protozoan numbers, it was hard to conclude whether a reduction in the grazing pressure had

been achieved. Therefore, a second series of reactors was set up to better address this. Cycloheximide experiments The protozoan metabolic inhibitor cycloheximide has the ability to inhibit protozoan metabolism and reduce their ability to graze (Sanders and Porter 1986). Therefore, in addition to using the dilution/extinction technique, each batch microcosm was supplemented with cycloheximide at a final concentration of 100 mg·L–1 to study the effect of a static protozoan community on nitrification. Again, temperature, DO, and pH were measured to monitor the stability of the system. The temperature (25 °C) and DO (5.5 mg·mL–1) remained constant and the pH dropped gradually to 6.4, similar to the reactors without cycloheximide. The BOD and COD also remained constant. Therefore, the microcosm systems were considered stable. Microscopic analysis of the activated sludge was done before and after the addition of the cycloheximide to view the activity of the protozoans. In the presence of cycloheximide, the protozoan community was clearly nonmotile and considered inactive. Furthermore, there was no change in total protozoan density during the experiment in the reactors containing the inhibitor. The bacterial community, however, showed a large increase in numbers in the presence of cycloheximide. The elevated bacterial growth rates in the presence of cycloheximide was assumed to be due to the release from predation pressure (Table 2). The amounts of ammonia, nitrate, and nitrite present in the reactors were found to be significantly less in the presence of cycloheximide than in its absence by the end of the 14-day period (Figs. 2a–2c). The concentration of total nitrogen was also less in the presence of cycloheximide (Fig. 2d). Overall, less nitrification was occurring in the absence of protozoan grazing (Fig. 3) and concurs with the theory that the presence of protozoans enhances nitrification (Clarholm 1985; Verhagen and Laanbroek 1992). How protozoans play a role in this phenomenon may be explained by suggesting that the lack of grazing pressure allowed an increase in bacterial numbers, which in turn consumed more available resources and lowered the regeneration and availability of nutrients. However, the bacteria involved in nitrification may not be part of this phenomenon, since they are slow-growing bacteria (Hagopian and Riley 1998). Possibly, the increase in the bacterial growth rate in the reactor owing to the reduced protozoan grazing represented an increase in © 2005 NRC Canada

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Fig. 2. Effect of protozoan inhibition on the production of (a) ammonia, (b) nitrite, (c) nitrate, and (d) total nitrogen in the batch reactors. Mean values are shown with error bars indicating the standard deviation.

the numbers of fast-growing heterotrophic bacteria. These bacteria have been shown to outcompete nitrifying bacteria for ammonia, and therefore, an increase in their numbers relative to the nitrifying bacteria could cause a decrease in nitrification potential. Second, the absence of protozoan grazing could also decrease the ammonia production rate because protozoans are no longer excreting ammonia as a byproduct of metabolic activity. An ungrazed population is often controlled by some limiting supply that causes individual cells to lower levels of activity and therefore reduce the efficiency of the whole community. These ideas may be applied to this system to suggest that the presence of protozoans and the predation pressure that they apply stimulate nutrient turnover and the nitrification process. Finally, the total nitrogen concentration was significantly less in the absence of protozoans than in their presence, and overall, the concentration of nitrogen declined in the absence of protozoans over time. This suggests that the system was losing nitrogen in the absence of protozoans, possibly through denitrification and emission of nitrogen gas. If this is the case, then the absence of active protozoans in the system could lead to nitrogen-limiting conditions. Relevance of protozoan grazing for the activated sludge process This study has suggested that in the absence of protozoan grazing, nitrification occurs at a lower rate than in the presence of protozoans. This finding is supported by several other studies that also found enhanced nitrogen cycling in the presence of protozoan grazing in activated sludge and other environments (Clarholm 1985; Verhagen and Laanbroek 1992; Strauss and Dodds 1997; Petropoulos 2003). What is still not clear is the exact mechanism by which protozoans affect nitrification. The possible role of protozoans can be summarized in two main ideas: the grazing effect and the cycling effect. The grazing effect entails the theory that protozoans may affect nitrification by controlling populations of heterotrophic bacteria that normally compete with nitrifying bacteria for ammonia. It has been shown that the assimilation of ammonia by heterotrophs happens in preference to nitrification and in fact reduces the available ammonia for nitrification. Thus, it follows that when heterotrophic bacterial numbers are controlled by protozoan grazing, more ammonia will be available for nitrifying bacteria, and nitrification will be more efficient. Several studies have shown that the presence of protozoan grazing in fact selects for fast-growing heterotrophic bacteria and that slow-growing nitrifying bacteria are eliminated (Sherr et al. 1988; Sinclair and Alexander 1989); however, those studies did not involve bacterial populations that are highly flocculated, such as those present in activated sludge used in this study. In activated sludge, the results of many studies have indicated that nitrifying bacteria are located within the interior of activated sludge flocs (Wagner et al. 1995, 1996, 1998; Mobarry et al. © 2005 NRC Canada

798 Fig. 3. Nitrification rate in the batch reactors (a) over the dilution series and (b) with and without cycloheximide. Mean values are shown with error bars indicating the standard deviation.

1996; Daims et al. 2001), while heterotrophic bacteria are located in all areas of the floc (Wagner et al. 1995). As such, nitrifying bacteria are not available to protozoans for predation under normal circumstances. Thus, the theory that protozoan grazing reduces numbers of slow-growing nitrifying bacteria is not applicable in activated sludge systems. Heterotrophic bacteria in activated sludge are also located in floc, and those that are not in flocs are consumed by protozoans, hence the basis of a self-regulating activated sludge system that selects for flocculating bacteria. In the absence of protozoan activity, bacterial populations are not controlled, and fast-growing heterotrophic bacteria proliferate and subsequently outcompete the nitrifying bacteria for ammonia. The net result of this is a decrease in nitrification. The cycling theory of the effect of protozoans on nutrient cycling follows that protozoan grazing results in the excretion of mineral nutrients such as ammonia and phosphate, decreasing the carbon:nitogen:phosphorus ratio and resulting in an accelerated use of carbon resources by bacteria (enhanced carbon mineralization). A study by Sherr et al. (1988) found that in aquatic environments, protozoans that consume living cells have the highest biomass specific excretion rates of inorganic nitrogen of all zooplankton groups. Protozoan grazing changes the carbon:nitogen:phosphorus ratio by increasing the concentrations of mineral nitrogen (excreted by protozoans as ammonia) and phosphorus (excreted by protozoans as phosphate). The excretion of ammonia by protozoans provides more ammonia for oxidation by nitrifying bacteria and also for assimilation by heterotrophs. One problem with the cycling theory applied to municipal activated sludge is that in such a system, ammonia is often in abundant supply and is rarely limiting. Thus, the protozoan ex-

Can. J. Microbiol. Vol. 51, 2005

cretion of ammonia may have little effect on enhancing nitrification, since it is probably already occurring at a high rate. However, it is possible that the influence of protozoan grazing on the carbon:nitogen:phosphorus ratio is somewhat underestimated because the composition of wastewater (i.e., influent nitrogen and carbon loadings) is highly variable (Ratsak et al. 1996). For example, the nitrogen concentration may be lowered by processes such as denitrification and assimilation of ammonia by heterotrophic bacteria. Furthermore, some types of wastewater (such as pulp and paper) contain much higher levels of carbon, in which case nitrogen may become limiting (Ratsak et al. 1996). Previously, the presence of protozoans has been correlated with good floc formation and settling characteristics (Curds 1982). This study shows that their grazing activity can also have an effect on the nitrification rate in activated sludge. Since nitrification can be the weak link in activated sludge systems, the results of this research imply that maintaining a healthy protozoan community is beneficial for sustaining nitrification rates and producing good-quality effluent. In addition, reduced nitrification rates can also augment toxicity problems by raising the ammonia concentrations. Since the health of the protozoan community is susceptible to environmental conditions such as temperature, pH, and toxicological events, it follows that a maintenance scheme that supports the protozoan community is important to the biological process within the activated sludge system. Overall, this study suggests that protozoan grazing of bacteria is an important factor in nitrogen cycling within this system. It is likely that both the grazing effect and the cycling effect play a role in how protozoans affect the nitrification rate. Future work will focus on establishing whether or not protozoan grazing increases the nitrification rate by increasing the number of nitrifying bacteria or by increasing per-cell nitrification rates.

Acknowledgements This research was supported through a Natural Sciences and Engineering Research Council of Canada research grant to K.A. Gilbride. The authors wish to thank D. Wright for his valuable input.

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