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Applied and Environmental Microbiology, January 2006, p. 575-584, Vol. 72, No. 1
0099-2240/06/$08.00+0     doi:10.1128/AEM.72.1.575-584.2006
Copyright © 2006, American Society for Microbiology. All Rights Reserved.

Influence of Nutrient Inputs, Hexadecane, and Temporal Variations on Denitrification and Community Composition of River Biofilms

M. R. Chénier,1,2 D. Beaumier,1 N. Fortin,1 R. Roy,1,{dagger} B. T. Driscoll,2 J. R. Lawrence,3 and C. W. Greer1*

Biotechnology Research Institute, National Research Council of Canada, 6100 Royalmount Avenue, Montreal, Quebec, Canada H4P 2R2,1 Department of Natural Resource Sciences, McGill University, 21 111 Lakeshore Road, Ste-Anne-de-Bellevue, Quebec, Canada H9X 3V9,2 National Water Research Institute, Environment Canada, 11 Innovation Boulevard, Saskatoon, Saskatchewan, Canada S7N 3H53

Received 10 May 2005/ Accepted 17 October 2005


   ABSTRACT
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Biofilms were cultivated on polycarbonate strips in rotating annular reactors using South Saskatchewan River water during the fall of 1999 and the fall of 2001, supplemented with carbon (glucose), nitrogen (NH4Cl), phosphorus (KH2PO4), or combined nutrients (CNP), with or without hexadecane, a model compound representing aliphatic hydrocarbons used to simulate a pollutant. In fall 1999 and fall 2001, comparable denitrification activities and catabolic potentials were observed in the biofilms, implying that denitrifying populations showed similar activity patterns and catabolic potentials during the fall from year to year in this river ecosystem, when environmental conditions were similar. Both nirS and nirK denitrification genes were detected by PCR amplification, suggesting that both denitrifying bacterial subpopulations can potentially contribute to total denitrification. Between 91.7 and 99.8% of the consumed N was emitted in the form of N2, suggesting that emission of N2O, a major potent greenhouse gas, by South Saskatchewan River biofilms is low. Denitrification was markedly stimulated by the addition of CNP, and nirS and nirK genes were predominant only in the presence of CNP. In contrast, individual nutrients had no impact on denitrification and on the occurrence of nirS and nirK genes detected by PCR amplification. Similarly, only CNP resulted in significant increases in algal and bacterial biomass relative to control biofilms. Biomass measurements indicated a linkage between autotrophic and heterotrophic populations in the fall 1999 biofilms. Correlation analyses demonstrated a significant relationship (P ≤ 0.05) between the denitrification rate and the biomass of algae and heterotrophic bacteria but not cyanobacteria. At the concentration assessed (1 ppb), hexadecane partially inhibited denitrification in both years, slightly more in the fall of 2001. This study suggested that the response of the anaerobic heterotrophic biofilm community may be cyclic and predictable from year to year and that there are interactive effects between nutrients and the contaminant hexadecane.


   INTRODUCTION
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Denitrification is an anaerobic respiratory process defined as the consecutive reduction of nitrate (NO3–) to nitrite (NO2–), nitric oxide (NO), nitrous oxide (N2O), and finally to dinitrogen gas (N2) (29). This pathway is of ecological significance because it is the major biological process by which fixed nitrogen is returned from freshwater, marine, and soil ecosystems to the atmosphere, offsetting the fixation of N2 in the global nitrogen cycle (33). Denitrifying organisms, although generally slow growers, reduce inorganic N and oxidize organic matter and have a higher energy yield than fermentation processes (2), giving them with a competitive advantage in low-oxygen environments. Because the ability to denitrify is a widespread trait across different physiological and taxonomic bacterial groups (68), it may also have a significant role in the utilization of organic matter in anaerobic aquatic habitats (6).

In lotic ecosystems, microorganisms can be found in three states: as free-living planktonic cells, as flocs of aggregated microbial cells, or attached to various surfaces, where these sessile cells form biofilms (57) and function as a closely integrated community (41). It is known that, in some river ecosystems, biofilm bacteria can constitute 90% or more of the total bacterial populations (14) and account for the majority of the bacterial activity (18). Within biofilms, the utilization of nutrients, such as those arising from agricultural, industrial, and municipal activities, leads to an increase in microbial biomass and activity, resulting in an increased demand for oxygen. Thus, anaerobic zones may form due to oxygen depletion, potentially allowing the proliferation of strict or facultative anaerobic microorganisms, such as denitrifying bacteria (36).

It is generally accepted that, in aquatic ecosystems, bacterial production and activity are mainly limited by temperature (20, 63) and nutrients (44, 61). In lotic ecosystems, these parameters vary considerably with the season (28). Industrial, municipal, and agricultural activities introduce nutrients such as organic carbon, inorganic nitrogen, and phosphate into rivers (17). This can lead to eutrophication and alteration of the distribution and diversity of aquatic species (58). Moreover, the effects of combined environmental stresses on biofilm microorganisms may be independent or additive or may mask the effects of each other. Despite the importance of these ecosystems, little is known about lotic biofilm communities or the effect that changing natural and anthropogenic conditions has on them.

Pollutants such as those arising from industrial activities can deleteriously impact anaerobic heterotrophic processes like denitrification. Hexadecane (C16H34), a major component of diesel fuel (45), is used here as a model compound, representative of the aliphatic hydrocarbons found in crude oil (2). Petroleum hydrocarbons such as hexadecane may find their way into rivers after refining processes (27), via leakage from storage tanks and distribution systems (42), and from accidental spills (55) and may affect the microbial activity and composition of river ecosystems.

Although a number of studies conducted recently provided insights into the chemical and biological composition of river biofilms (1, 32, 37, 38), rivers are still understudied compared to lake, estuarine, and marine environments (52). The technical difficulties in controlling environmental conditions and in sampling river and stream biofilms, due to the rapid movement of water, make the use of model systems highly desirable (35, 39). Therefore, a laboratory system was used to study the effects of specific variables (nutrient and pollutant inputs) on the denitrification activity and microbial composition of river biofilms, while other variables (temperature, water flow, shear forces, exchange rates, light conditions, etc.) were maintained constant, which is usually not possible in situ. The objectives of the present work were to assess the impact of carbon (C), nitrogen (N), and phosphorus (P), added alone or in combination, and of hexadecane on South Saskatchewan River biofilms grown in rotating annular bioreactors and to compare community responses in the fall of 1999 and 2001. The impacts of nutrients, hexadecane, and temporal variations on biofilm denitrification and biomass composition were evaluated by using a combination of microcosm assays, microscale imaging, and molecular biology techniques.


   MATERIALS AND METHODS
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Biofilm cultivation.
Water samples were taken from the South Saskatchewan River (Saskatoon, Saskatchewan, Canada) during the fall of 1999 and the fall of 2001, as previously described (46), and were used as inocula for the experiments. General water analyses were carried out according to the procedures described in reference 19.

Continuous-flow stirred-tank reactors (CFSTRs), which are open systems in which no concentration gradients exist in the liquid phase, are appropriate and provide the following significant advantages for cultivating and sampling river biofilms and analyzing their biomass and processes (10): (i) the bulk liquid phase is uniform, which makes chemical analysis and biofilm sampling simple; (ii) the steady-state condition is reproducible; (iii) the shear forces are fairly uniform.

The study of river biofilms requires that the physical integrity of the microbial community be preserved (38). Accordingly, the CFSTR geometry that was selected in the context of this work was the rotating annular bioreactor (RAB), since, in addition to the advantages cited above, RABs (10) (i) maintain the physical and taxonomic structure of the community during growth and sampling; (ii) provide a turbulent flow, as in rivers; (iii) provide a completely mixed bulk liquid phase and as such, represent CFSTRs (59); and (iv) are compact and easy to operate.

The reactor was operated at 132 rpm (a surface velocity of 0.44 m s–1). The reactor volume was 500 ml, and flow through the reactors was 1 liter day–1. The water in the reservoir was replaced with fresh river water throughout the experimental period on an 8-day schedule (days 1, 3, 5, 8, etc.). Biofilms were grown on polycarbonate strips in rotating annular bioreactors, as described previously. In brief, the reactors were constructed using a rotating inner solid polycarbonate cylinder machined to allow insertion of 12 removable polycarbonate slides; the outer cylinder was a commercially available 1-liter glass jar with a threaded mouth (38). Mixing in the reactor was via the rotation of the inner cylinder and the circulation of water through the reactor from top to bottom. The experimental setup included a temperature-controlled (17 ± 2°C) 10-liter reservoir for the raw water, a peristaltic pump for circulating the water, and the rotating annular reactor with motor and control for adjusting the speed of the rotating inner cylinder. All reactor components were surface disinfected with 1.2% sodium hypochlorite for 1 h prior to use.

The biofilms were grown for 8 weeks at the dates described in Table 1. Each fall, two replicate bioreactors were operated for each of the following conditions, in the absence and in the presence of hexadecane (1 ppb): river water alone (no nutrient addition, control); river water amended with glucose at 67 µmol/liter of carbon; river water amended with NH4Cl at 80 µmol/liter of nitrogen; river water amended with KH2PO4 at 5 µmol/liter of phosphorus; and river water amended with a combination of all three nutrients at the concentrations described above (44).


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  		TABLE 1. Results of analysis on South Saskatchewan River waters

 
Microcosm preparation.
After the 8-week period of growth in the bioreactors, fresh biofilm samples on polycarbonate strips were used to assess the impact of nutrients, hexadecane, and temporal variations on biofilm activity in microcosms. To prepare the microcosms, the polycarbonate strips were aseptically cut (2 cm2) and each piece with its associated biofilm was transferred into a 27-ml glass vial with 5 ml of the same river water supplemented with the same nutrient concentrations used in the bioreactors, as described previously (11). For each experimental group, four replicate microcosms (two replicate bioreactors, two replicate microcosms/bioreactor) were prepared. All microcosms were incubated statically, in the dark, at room temperature, under the conditions described below. Some of the polycarbonate strips collected from the bioreactors were frozen immediately at –80°C for subsequent molecular analyses.

Denitrification.
Microcosms prepared as above, with or without nutrient amendment, received 5 mM NaNO3 (BDH, Toronto, Ontario, Canada). The microcosms were capped with 20-mm rubber stoppers (Bellco Glass Inc., Vineland, NJ), and their atmosphere was made anaerobic by evacuating (5 min each time) and refilling to 1 atmosphere three times with N2 (prepurified, 99.998%; Praxair, Mississauga, Ontario, Canada) in order to eliminate dissolved gases in the water. For each treatment, C2H2 was added to the headspace of half of the microcosms at a final partial pressure of 2 kPa (50) to inhibit N2O reductase and prevent the reduction of N2O to N2 (66). Denitrification was measured by determination of N2O production in the presence of C2H2. Periodically, gas samples (0.2 ml) were withdrawn from the headspace of the microcosms and analyzed for N2O on an 8610C gas chromatograph (SRI, Torrance, CA) with a thermal conductivity detector and an electron capture detector in parallel, as previously described (49). The amount of N2O was calculated based on a standard curve (49).

Calculations.
Considering the quantity of NO3– initially added to each microcosm (25 µmol) and considering that 2 µmol of NO3– is required to produce 1 µmol of N2O, the molar percentage of N consumed through denitrification by the biofilms at the end of the incubation period (41 days) was estimated as follows: 100 · quantity of N2O produced in the flask (µmol)/maximum quantity of N2O possible to produce (12.5 µmol) (49). Also, the molar ratio of N2/N2O produced through denitrification by the biofilms at the end of the incubation period was estimated as follows: quantity of N2O produced in the flask in the presence of C2H2 (µmol)/quantity of N2O produced in the flask in the absence of C2H2 (µmol) (49).

To determine if duplicate microcosms from one bioreactor were equivalent to duplicate microcosms from the other bioreactor operated under the same experimental conditions, maximum rates of denitrification for individual microcosms were computed by linear regression of the amount of N2O produced over time using Excel (49). The time frame for the regression was different between fall 1999 and fall 2001 experiments. To determine maximum denitrification rates, 1999 data from day 0 to day 12 showing maximum rates were used, whereas 2001 data from day 0 to day 30 showing maximum rates were used. Individual maximum denitrification rates for one set of duplicate microcosms were compared with individual maximum denitrification rates for the other set of duplicate microcosms using the Mann-Whitney U test (double-sided, n1 = 2, n2 = 2) (53). Results of this statistical test, which was performed for all experimental groups, revealed that the two sets of duplicate microcosms are not significantly different (P > 0.05). Thus, results are presented as the average of four microcosms (two replicate bioreactors, two replicate microcosms/bioreactor). Maximum denitrification rates were also compared with the biomass of algae, cyanobacteria, bacteria, and exopolymers of the biofilms as freshly sampled from the bioreactors.

Total community DNA extraction.
For each bioreactor, a frozen polycarbonate strip was aseptically cut (2 cm2) and transferred into a 50-ml polypropylene tube (Falcon; Becton Dickinson, Franklin Lanes, NJ). Cells from the frozen biofilm samples were lysed by enzymatic treatment as performed by Fortin et al. (21) with the following modifications: the volumes of reagents used were fivefold reduced, agitation during lysozyme treatment was at 200 rpm, and proteinase K was added to a final concentration of 200 µg ml–1. DNA extraction was performed by phenol-chloroform treatment as described by Ausubel et al. (3), except that incubation after cetyltrimethylammonium bromide addition was at 65°C for 30 min. The DNA was precipitated overnight at –20°C with an equal volume of cold 100% isopropanol and then centrifuged for 30 min at 4°C (17,400 x g). The pellets were washed three times with 2 ml of 70% cold ethanol, centrifuged for 5 min at 4°C (17,400 x g), and air dried. The DNA was dissolved in 300 µl of Tris (10 mM)-EDTA (0.1 mM) (TE; pH 8) with gentle agitation at room temperature for 1 h, followed by 1 h on ice. The DNA was precipitated overnight at –20°C with 1/10 volume of 3 M sodium acetate (pH 5.2) and 2.5 volumes of cold 100% ethanol. The precipitated DNA was recovered by centrifugation for 15 min at 4°C (16,060 x g), washed three times with 500 µl of 70% ethanol at room temperature, air dried, and dissolved in 30 µl of TE (11). Genomic DNA from Pseudomonas stutzeri ATCC 14405 and Pseudomonas aureofaciens ATCC 13985 was prepared by the method described by Ausubel et al. (3).

PCR amplification.
For the reduction of NO2–, which is the first reaction of denitrification sensu stricto, two types of mutually exclusive enzymatic systems are found in bacteria, i.e., bacteria harbor either the nirS or the nirK gene, but not both (67). To assess the metabolic potential of the biofilm by studying both denitrifying bacterial genotypes, the total DNA extracts were analyzed by PCR amplification for the following target genes: the nirS gene encodes cytochrome cd1 nitrite reductase, which reduces nitrite to nitric oxide during denitrification (6), and the nirK gene encodes copper-containing nitrite reductase, which also reduces nitrite to nitric oxide during denitrification (6). For nirS amplification, the sequence of the forward primer was 5'572-601-CGG CTA CGC GGT GCA TAT CTC GCG TCT GTC-3', whereas the sequence of the reverse primer was 5'864-893-GAT GGA CGC CAC GCG CGG CTC GGG GTG GTA-3' (base positions according to Pseudomonas stutzeri ATCC 14405) (7, 8, 26). For nirK amplification, the sequence of the forward primer was 5'560-589-GGG CAT GAA CGG CGC GCT CAT GGT GCT GCC-3', whereas the sequence of the reverse primer was 5'906-935-CGG GTT GGC GAA CTT GCC GGT GGT CCA GAC-3' (base positions according to Pseudomonas aureofaciens ATCC 13985) (7, 8, 26).

The PCR amplification procedure was modified from that of references 4 and 30. The PCR mixture consisted of an aliquot of total DNA extract (1 µl of undiluted DNA and 1/10- and 1/100-diluted DNA), which was added to a final volume of 50 µl of reaction mixture (Amersham Pharmacia Biotech Inc., Piscataway, NJ) containing 10 mmol/liter Tris-HCl (pH 9); 50 mmol/liter KCl; 2.5 mmol/liter MgCl2; 200 µmol/liter each dATP, dTTP, dGTP, dCTP; and 1 µmol/liter of each primer (forward and reverse). The tubes were placed in a DNA thermal cycler (Perkin-Elmer Cetus, Montreal, Quebec, Canada) and heated to 96°C for 5 min. The temperature was then brought down to 80°C, and 2.5 U of Taq DNA polymerase (Amersham Pharmacia Biotech Inc., Piscataway, NJ) in 10x PCR buffer (100 mmol/liter Tris-HCl [pH 9.0], 500 mmol/liter KCl, 15 mmol/liter MgCl2) was added. PCR conditions were 30 cycles of 1 min at 94°C (denaturing), 1 min at 63°C for nirS or 65°C for nirK (annealing), and 1 min at 72°C (extension), followed by a final extension of 3 min at 72°C. DNA was electrophoresed in 1.4% agarose (Wisent, St-Bruno, Quebec, Canada) gels with 1x Tris-acetate-EDTA buffer (51) using 250 ng of the GeneRuler 100-bp DNA ladder (MBI Fermentas, Vilnius, Lithuania) as the molecular weight marker. Gels were stained with ethidium bromide and photographed under UV light.

Community composition.
Examination of all stained and control materials was carried out using an MRC 1024 confocal laser scanning microscope (CLSM; Bio-Rad, Hemel Hempstead, United Kingdom) attached to a Microphot SA microscope (Nikon, Tokyo, Japan). Slides from each of the replicate reactors were cut into ca. 1-cm2 pieces and mounted in small petri dishes using Dow Corning 3140 acid-free silicone coating (WPI Inc., Sarasota, FL) and then stained and analyzed according to the procedures that follow. For observation the following water-immersible lenses were used: 63x, 0.9 numerical aperture (NA) (Zeiss, Jena, Germany), and 40x, 0.55 NA (Nikon, Tokyo, Japan). The biofilms were observed using a double-labeling procedure and three-channel recording: bacteria were stained with the fluorescent nucleic acid stain SYTO 9 (excitation, 488 nm, emission 522 to 532 nm), a lectin probe (Triticum vulgaris; tetramethyl rhodamine isothiocyanate) (excitation, 568 nm; emission, 605 to 632 nm) was used to visualize exopolymer, and autofluorescence (excitation, 647 nm; emission, 680 to 732 nm) was used to detect algal and cyanobacterial cells. Digital image analysis of the CLSM optical thin sections in each of the three channels was used to determine such parameters as biofilm depth, bacterial cell area (biomass), exopolymer biomass, cyanobacterial biomass, and total photosynthetic biomass at various depths. Image analyses were performed using NIH Image version 1.61 (http://rsb.info.nih.gov/nih-image/) with macros written for semiautomated quantification as described by Manz et al. (40).

Statistical analyses.
The experimental design consisted of two identical replicate reactors randomly assigned to each nutrient and hexadecane treatment. Each analysis was done on subsamples from randomly selected biofilm coupons from among the 12 identical coupons in each replicate reactor. CLSM imaging was done at five random locations at five positions in a transect across the 1-cm2 piece of the biofilm coupon. Analysis of variance (ANOVA) was used to detect significant differences among sample means at a P of <0.05. ANOVA and correlation analyses were carried out using the MiniTab (State College, PA) commercial package.


   RESULTS
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
River water analyses.
Table 1 summarizes the chemistry of the South Saskatchewan River water used to feed the bioreactors. The river water temperature was higher in fall 2001 than in fall 1999, whereas the nutrient concentrations (dissolved organic carbon [DOC], N, PO43–) were higher in fall 1999 than in fall 2001. Heterotrophic plate counts and chlorophyll a levels were 5.9 times and 6.5 times higher, respectively, in fall 1999 than in fall 2001.

Denitrification.
In both years of study, the combined nutrients (CNP) stimulated denitrification by the biofilms (Fig. 1 and 2). Hexadecane partially inhibited denitrification in the presence of CNP, with a slightly more marked effect in fall 2001. When nutrients were absent or added separately, hexadecane addition had no detectable impact on biofilms from fall 1999 but moderately reduced denitrification in those from fall 2001. When nutrients were absent or added separately, activities were similar between fall 1999 and fall 2001 biofilms but were higher in fall 2001 in the absence of hexadecane. CNP biofilms were more active in fall 1999 than in fall 2001.


Figure 1
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  		FIG. 1. Effects of nutrients and hexadecane on N2O production during denitrification by fall 1999 (upper panel) and fall 2001 (lower panel) biofilms (2 cm2). Biofilms were grown without nutrient amendments (Ctrl) or with glucose (C), NH4Cl (N), and KH2PO4 (P) separately or in combination (CNP), in the absence and presence of hexadecane. Results of a Mann-Whitney U test (double sided, n1 = 2, n2 = 2) performed for all experimental groups revealed that the two sets of duplicate microcosms made from duplicate bioreactors operated under the same experimental conditions are not significantly different (P > 0.05). Each datum point is the average of four replicate microcosms, and the bars represent one standard error of the mean.

 

Figure 2
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  		FIG. 2. Effects of nutrients and hexadecane on N consumption during denitrification by fall 1999 and fall 2001 biofilms. Values are expressed as the molar percentages of N consumed through denitrification by the biofilms (%, mol/mol) after 41 days of incubation. Biofilms (2 cm2) were grown without nutrient amendments (Ctrl) or with glucose (C), NH4Cl (N), and KH2PO4 (P) separately or in combination (CNP), in the absence and presence of hexadecane. Results of a Mann-Whitney U test (double sided, n1 = 2, n2 = 2) performed for all experimental groups revealed that the two sets of duplicate microcosms made from duplicate bioreactors operated under the same experimental conditions are not significantly different (P > 0.05). Each column is the average of four replicate microcosms, and the bars represent one standard error of the mean.

 
In fall 1999, no lag in the denitrification activity was observed before N2O accumulated with CNP biofilms, whereas with separate nutrients or in their absence, activity was very low or absent for the first 3 to 4 days (Fig. 1). In all the experimental groups, denitrification activity reached a plateau at the end of the incubation period (41 days). In the absence of C2H2, small amounts of N2O accumulated temporarily only with CNP biofilms. In fall 2001, denitrification could not be detected for at least 2 days, whatever the nutrients and hexadecane status. The control biofilms and the biofilms incubated with individual nutrients were still active after 41 days, whereas denitrification by the CNP biofilms reached a plateau after 1 month.

The proportion of NO3– that was consumed by denitrification after 41 days of incubation in the presence of C2H2 was estimated based upon the addition of an initial amount of 25 µmol of N to each microcosm in the form of NO3– (Fig. 2). When nutrients were added separately or absent, in the absence of hexadecane, 1.5 to 2 times more NO3– was consumed in fall 2001 than in fall 1999. In the presence of hexadecane and in both fall 1999 and fall 2001, very similar proportions of N were consumed (3.6 to 5.2%) when nutrients were added separately or absent. Hexadecane negatively affected NO3– consumption in fall 2001 but had no impact in fall 1999. When CNP were added, the presence of hexadecane negatively affected NO3–-N consumption, and more NO3–-N was consumed in fall 1999 than in fall 2001. In the absence of C2H2, the percentage of NO3–-N consumed and retrieved in the form of N2O-N was always less than 1% regardless of nutrient, hexadecane, and seasonal status (data not shown).

The molar ratio of N2 over N2O (a greenhouse gas) produced by denitrification after 41 days of incubation was estimated. Table 2 shows that N2 emissions were between 12 and 576 times higher than N2O emissions. Hexadecane generally decreased the N2/N2O ratio both in fall 1999 and fall 2001 but had no impact when N alone was added. The N2/N2O ratio was higher in fall 2001 than in fall 1999 for the same experimental conditions, but ratios were similar when only P and hexadecane were present in combination.


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  		TABLE 2. Effects of nutrients and hexadecane on N2/N2O production during denitrification by fall 1999 and fall 2001 biofilms

 
Detection of nirS and nirK.
The results of the molecular analyses agreed with those of the microcosm assays. The genes nirS (Fig. 3) and nirK (Fig. 4) were detected by PCR amplification in all experimental groups (but not all dilutions), with the exception of nirS from the fall 2001 biofilm grown without hexadecane in the presence of N alone. There was also some denitrification activity in all experimental groups, independent of the nutrient, hexadecane, or seasonal status. Figure 3 shows that, in fall 1999, the nirS band intensity was higher for undiluted and diluted extracts of biofilms grown with P amendments (P and CNP biofilms), whether hexadecane was present or not, and that the presence of hexadecane did not affect band intensities. In fall 2001, the nirS band intensity was also higher for undiluted and diluted extracts of biofilms grown with P amendments (P and CNP biofilms) and hexadecane, which was not observed for biofilms grown in the absence of this petroleum hydrocarbon. Figure 4 shows that the nirK band intensity was moderately higher in undiluted and diluted extracts from CNP biofilms in both years, whether hexadecane was present or not, and that hexadecane did not have a detectable impact on the band intensity. There was no PCR-inhibitory material present in the biofilm extracts, since nirS and nirK were detected in undiluted extracts and, in some cases, at lower levels in diluted extracts.


Figure 3
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  		FIG. 3. PCR amplification of the nirS gene from fall 1999 (upper panels) and fall 2001 (lower panels) biofilms. Biofilms were grown without nutrient amendments (CTRL) or with glucose (C), NH4Cl (N), and KH2PO4 (P) alone or in combination (CNP), in the absence (no hexadecane) or presence (with hexadecane) of hexadecane (H). Total community DNA extracts were undiluted (0) or diluted 10–1 (–1) or 10–2 (–2) prior to PCR amplification using the nirS consensus primers. Lane M, molecular weight marker, with the asterisks indicating the 500-base pair marker; lane +, positive control (50 ng of Pseudomonas stutzeri ATCC 14405 DNA template); lane –, negative control (no DNA). The arrows indicate the position of the 320-bp nirS gene amplicon.

 

Figure 4
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  		FIG. 4. PCR amplification of the nirK gene from fall 1999 (upper panels) and fall 2001 (lower panels) biofilms. Biofilms were grown without nutrient amendments (CTRL) or with glucose (C), NH4Cl (N), and KH2PO4 (P) alone or in combination (CNP), in the absence (no hexadecane) or presence (with hexadecane) of hexadecane (H). Total community DNA extracts were undiluted (0) or diluted 10–1 (–1) or 10–2 (–2) prior to PCR amplification using the nirK consensus primers. Lane M, molecular weight marker, with the asterisks indicating the 500-base pair marker; lane +, positive control (50 ng of Pseudomonas aeruginosa ATCC 13985 DNA template); lane –, negative control (no DNA). The arrows indicate the position of the 380-bp nirK gene amplicon.

 
Community composition.
The total biofilm biomasses were higher in fall 1999 than in fall 2001 for the same experimental conditions, and cyanobacteria were a minor component of the biofilm communities under all experimental conditions (Fig. 5).


Figure 5
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  		FIG. 5. Effects of nutrients, hexadecane, and temporal variations on the community composition in fall 1999 (A and B) and fall 2001 (C) biofilms (1 cm2) as determined by in situ probe binding and CLSM analyses. Bacteria were stained with SYTO 9, exopolymer (polymer) was visualized with tetramethyl rhodamine isocyanate-labeled lectin, and autofluorescence was used to detect algal and cyanobacterial cells. Biofilms were grown without nutrient amendments (control) or with glucose (C), NH4Cl (N), and KH2PO4 (P) separately or in combination (CNP), in the absence and presence of hexadecane. Asterisks indicate statistically significant differences (P < 0.05) relative to the control treatment.

 
In fall 1999, the addition of CNP to the bioreactors had a dramatic impact on the biofilm community by significantly (P < 0.05) increasing the cyanobacterial, bacterial, and polymer biomasses (Fig. 5) relative to control values. When comparing biofilms grown under the same nutrient status, the addition of hexadecane increased the total biomass in control biofilms and in biofilms grown with only C or N but decreased it when P was present (P and CNP biofilms). Polymers were the major component when nutrients were absent or added separately, whether hexadecane was present or not. When CNP was present, the algal biomass and polymers were dominant and represented similar biofilm component amounts, whereas when CNP and hexadecane were added, the biomass was evenly distributed between the four measured biofilm components.

In fall 2001, the positive impact of CNP was not as strong as that observed in fall 1999, and the effect of hexadecane was also different than in fall 1999. When comparing biofilms grown under the same nutrient status, the presence of hexadecane increased the total biomass of P and CNP biofilms, had no impact on control and N biofilms, and decreased the biomass of C biofilms. Except for cyanobacteria, which always represented a minor fraction of the biofilm biomass, the community composition was generally evenly distributed when nutrients were absent or added separately, whether hexadecane was present or not. Bacteria dominated when CNP were added in the presence and in the absence of hexadecane.

A statistical treatment of the denitrification rates for the first 12 days of microcosm incubation and of the community composition indicated that the strongest correlation (Pearson correlation; P < 0.05) was between the algal biomass and the denitrification rates (Table 3). Denitrification rates and other biofilm components were correlated to a lesser extent, whereas cyanobacteria, which represented a minor fraction of the biofilm biomass under most experimental conditions, were generally not correlated to any biofilm component or activity.


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  		TABLE 3. Correlation between biofilm components and the denitrification rates (days 0 to 12)

 

   DISCUSSION
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Temporal variations.
The microcosm assays and PCR analyses demonstrated that the biofilms that developed from the South Saskatchewan River water in fall 1999 and in fall 2001 had similar denitrification activities (Fig. 1 and 2) and catabolic potentials (Fig. 3 and 4), unless combined nutrients were added to the bioreactors. This implies that facultative heterotrophs performing specific activities such as denitrification have either stable or cyclic but predictable activities and genetic potential from year to year in this ecosystem. Since we previously showed that denitrification and PCR amplification of the nirK gene from biofilms from this river varied seasonally (11), it is likely that denitrifying populations show similar patterns in the same season from year to year, when environmental conditions are similar.

The availability of nitrate and organic carbon and low levels of O2 are recognized as having direct stimulatory effects on denitrification (47, 56). In the South Saskatchewan River water, the NO2–-plus-NO3– concentration was slightly higher in fall 1999 than in fall 2001, whereas the DOC concentrations were almost identical over the two growth periods (Table 1). Despite differences in the river water temperature, denitrification activities were comparable between fall 1999 and fall 2001. The variations observed in the river water chemistry were not sufficient to induce significant changes in biofilm denitrification, unless an exogenous source of nutrients (CNP) was provided.

Total biofilm biomass was higher in fall 1999 than in fall 2001 and was generally evenly distributed between algal, cyanobacterial, bacterial, and polymer biomasses in fall 2001, whereas polymers generally dominated in fall 1999 (Fig. 5). In fall 1999, autotrophic populations (algae and cyanobacteria) and heterotrophs were much more abundant when CNP were added to the bioreactors. Biomass measurements provide an indicator of the state of river biofilm populations, and these results indicate the linkage or coupling that occurs between autotrophs and heterotrophs in river biofilms, as was previously observed in biofilms from the South Saskatchewan River (34). Studies of alga-bacterium interactions in mountain streams showed that excreted inorganic compounds synthesized by algae via CO2 fixation were taken up by heterotrophic bacteria (24) (25). Another study showed that algal productivity and bacterial consumption of organic compounds synthesized by algae were the highest in late July and that the activity of both communities became undetectable in early September (62).

Nutrients.
In a previous study we showed that the addition of CNP to the bioreactors had an important stimulatory effect on the biofilm activities, including denitrification, and influenced their composition (11). This suggested that a major factor limiting bacterial activity in the South Saskatchewan River water was the presence of available carbon and other utilizable nutrients. In the present study, results revealed that combined nutrients stimulated all measured parameters and that individual nutrients had no stimulatory impact on the biofilms. Based on these experimental observations, we could not determine which nutrient was limiting for the biofilm microbial populations. In another report, we showed that P was the limiting nutrient for nitrifying bacteria of South Saskatchewan River biofilms, but the amounts of various biofilm polymers were not increased (12). In the present work, despite the prevalence of the nirS gene in biofilms amended with P alone, denitrification was not stimulated, suggesting that another factor may be limiting for the activity of nirS-harboring denitrifiers. Similarly, it was recently reported that Baltic Sea cyanobacteria contained denitrification genes but that, despite optimal environmental conditions and genetic potential, had negligible denitrification activity (60). In the present study, the addition of CNP resulted in significant increases in cyanobacterial biomass (P < 0.05) in both years, especially in the presence of hexadecane. P and N limitations were also reported for biofilm microorganisms in other Canadian rivers (44, 48).

Organic compounds have different effects on denitrification by natural aquatic communities. We observed that the addition of glucose alone as a carbon source to the bioreactors had no impact on denitrification or on the detection of the nirS and nirK genes. In contrast, in a study of the impact of organic C compounds commonly found in agricultural pollutants on N cycling by sediments of a river in Ireland, a 2.3-fold increase in denitrification was observed when glucose was added to sediment cores, whereas denitrification was 1.5 times higher in the presence of glycine, was partially inhibited by acetate, and was totally inhibited by formate (31).

Hexadecane.
The microcosm analyses showed that, at the concentration assessed (1 ppb in the aqueous phase), hexadecane partially inhibited denitrification (Fig. 1) and reduced the N2/N2O ratio (Table 2). These results show that petroleum hydrocarbons like hexadecane, even when present at low concentrations, can alter the flux of N in river ecosystems. Since petroleum hydrocarbons are hydrophobic, they can accumulate in lipid bilayers of bacterial cytoplasmic membranes or within the outer membranes of gram-negative bacteria (54). This causes alterations of membrane structure (43) and function (54), which have a negative impact on cellular activity. The low concentration of hexadecane applied to the South Saskatchewan River biofilms may explain why only partial inhibition of denitrification was observed in both fall 1999 and 2001. Partial inhibition of bacterial activity by alkanes has also been reported elsewhere (5, 64, 65).

Denitrification.
This study showed that the consumption of N through denitrification by South Saskatchewan River biofilms may be significant (up to 18.5% of NO3– initially added; Fig. 2). In the Elbe River (Czech Republic), it was found that the losses of N through denitrification varied between 6 and 23% depending on the season (23). In a hill pasture catchment (New Zealand), in situ denitrification measurements following in-stream NO3– additions revealed that only about 1% of N losses could be accounted for by denitrification (13). Our results also showed that 91.7 to 99.8% of the consumed N is emitted in the form of N2 (N2/N2O ratios between 12 and 576; Table 2). This suggests that emissions of N2O, which is a major potent greenhouse gas (16) known to destroy the stratospheric ozone layer (15, 26), by the South Saskatchewan River biofilms are low, since the process of denitrification (consecutive reduction of NO3– to NO2–, NO, N2O, and finally N2) is more complete.

For the reduction of NO2–, which is the first reaction of denitrification sensu stricto (67), two types of mutually exclusive enzymatic systems are found in bacteria, i.e., bacteria harbor either the nirS or the nirK gene but not both (29, 67). Depending on the source, it is estimated that between two-thirds (29) and three-fourths (22) of denitrifying bacterial strains possess the tetraheme cytochrome protein cd1 (encoded by nirS) as the respiratory NO2– reductase, and the remaining strains possess the copper-containing NO2– reductase (encoded by nirK). In this study, both nir genotypes were detected in the biofilm DNA extracts by PCR amplification and thus could potentially contribute to total denitrification. Different denitrification genes were predominant in the presence of CNP (nirK) and in the presence of P or CNP (nirS).

Community composition and selection of the experimental model.
The experimental model used in this work is most relevant to mixed phototrophic/heterotrophic biofilms on rocks, plants, and river bed surfaces in rivers but does not address those events occurring within the sediments. The strong correlation observed between the algal biomass and the denitrification rates in this study (Table 3) is in agreement with observations of the positive response of heterotrophic bacteria to metabolites of phototrophic populations in lotic ecosystems (25), for example, through the flux of soluble algal products to the bacterial populations and the exchange of inorganic and organic carbon among these two biofilm populations (24). The absence of correlation between cyanobacteria and algae or denitrification rates indicates that the significant increase in cyanobacterial biomass within biofilms grown in the presence of CNP (with or without hexadecane) in fall 1999 was not directly related or linked to the increase in denitrification or in algal biomass.

The denitrification rates for the first 13 days of microcosm incubation were strongly correlated with the total photosynthetic (algal and cyanobacterial) and bacterial biomasses of the biofilms as freshly sampled from the bioreactors (Table 3). Since a strong correlation between microcosm denitrification and bioreactor biomass was not observed for any other time interval, the biofilm community composition in the microcosms may have evolved relative to that of the bioreactor biofilms after this initial period of incubation. Since the biofilms were cultivated in open system bioreactors within a turbulent flow and were subsequently incubated statically in closed microcosms, the experimental conditions may have resulted in potential changes in the microcosm biofilm community composition. As the changes occurred only after about 2 weeks of incubation, measurements of maximum denitrification rates in the initial phase of microcosm incubation represent accurate measurements of the potential activity of bioreactor biofilms. In another study, denaturing gradient gel electrophoresis analyses of the microbial communities of lake microcosms also revealed changes in the community composition over time, depending on experimental conditions (9). The authors suggested that stochastic events in the preparation of the microcosms, such as filtration of their water samples, may have resulted in alterations in the microbial numbers and in interactions between grazing protozoa and pollutant-degrading bacteria, which subsequently affected community succession.

Conclusions.
This study revealed that the denitrification activities and catabolic potentials of South Saskatchewan River biofilms were similar in fall 1999 and fall 2001 and that both nirS and nirK bacterial denitrification genes could be detected in the biofilms. Denitrification, the occurrence of nir genes, and algal and bacterial biomass were increased by the presence of CNP. Biomass measurements showed the linkage between autotrophic and heterotrophic populations in fall 1999. At the concentration assessed (1 ppb), hexadecane partially inhibited denitrification to similar extents in both years. Emissions of N2O (a greenhouse effect gas) via denitrification by the biofilms represented less than 9% of consumed N.


   ACKNOWLEDGMENTS
 
We gratefully acknowledge Suzanne Labelle, Anca Mihoc, Claude Masson, Danielle Ouellette, Sylvie Sanschagrin, and Gesine Wisse for their excellent technical assistance; George Swerhone for biofilm cultivation; Alain Corriveau for performing high-pressure liquid chromatography analyses; and Kerry Peru for hexadecane analyses.

This research was supported by Health Canada through the Toxic Substances Research Initiative (J.R.L. and C.W.G.), by the Biotechnology Research Institute, by the National Water Research Institute of Environment Canada, and by FCAR-MRST (M.R.C.).


   FOOTNOTES
 
* Corresponding author. Mailing address: Biotechnology Research Institute, National Research Council of Canada, 6100 Royalmount Avenue, Montreal, Quebec, Canada H4P 2R2. Phone: (514) 496-6182. Fax: (514) 496-6265. E-mail: charles.greer{at}cnrc-nrc.gc.ca Back

{dagger} Present address: Department of Biology, University of Victoria, P.O. Box 3020, Stn CSC, Victoria, British Columbia, Canada V8W 3N5. Back


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 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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Applied and Environmental Microbiology, January 2006, p. 575-584, Vol. 72, No. 1
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