Bacterium-Based NO2- Biosensor for Environmental Applications

A sensitive NO₂^– biosensor that is based on bacterialreduction of NO₂^– to N₂O and subsequent detection of theN₂O by a built-in electrochemical N₂O sensor was developed.Four different denitrifying organisms lacking NO₃^– reductaseactivity were assessed for use in the biosensor. The relevantphysiological aspects examined included denitrifying characteristics,growth rate, NO₂^– tolerance, and temperature and salinityeffects on the growth rate. Two organisms were successfullyused in the biosensor. The preferred organism was Stenotrophomonasnitritireducens, which is an organism with a denitrifying pathwaydeficient in both NO₃^– and N₂O reductases. AlternativelyAlcaligenes faecalis could be used when acetylene was addedto inhibit its N₂O reductase. The macroscale biosensors constructedexhibited a linear NO₂^– response at concentrations upto 1 to 2 mM. The detection limit was around 1 µM NO₂^–,and the 90% response time was 0.5 to 3 min. The sensor signalwas specific for NO₂^–, and interference was observed onlywith NH₂OH, NO, N₂O, and H₂S. The sensor signal was affectedby changes in temperature and salinity, and calibration hadto be performed in a system with a temperature and an ionicstrength comparable to those of the medium analyzed. A broadrange of water bodies could be analyzed with the biosensor,including freshwater systems, marine systems, and oxic-anoxicwastewaters. The NO₂^– biosensor was successfully usedfor long-term online monitoring in wastewater. Microscale versionsof the NO₂^– biosensor were constructed and used to measureNO₂^– profiles in marine sediment.

INTRODUCTION

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Introduction

Nitrite (NO₂^–) has a central position in the global nitrogen(N) cycle and is involved in many important biological N transformations.With an intermediate oxidation state, NO₂^– acts as anelectron donor in nitrification and serves as an electron acceptorin denitrification, dissimilative nitrite reduction to ammonium,and anammox.

In natural ecosystems NO₂^– is of interest because of itstoxicity for microorganisms and higher organisms (20, 37). NO₂^–concentrations in natural bulk water bodies are usually verylow, and in freshwater systems the average worldwide NO₂^–concentration has been estimated to be about 1 µg of Nliter^–1 ( $~$ 0.07 µM NO₂^–) (28). Recently, however,there have been several reports of NO₂^– accumulation toconcentrations of 5 to 140 µM in European eutrophic riversand estuaries (3, 8, 15, 19, 44). Studies have indicated thatNO₂^– accumulations can be related to imbalances in NO₂^–production and consumption rates for either aerobic sedimentprocesses (nitrification) or anaerobic sediment processes (denitrificationand dissimilatory nitrate reduction to ammonia). In marine systemsNO₂^– concentrations are normally negligible, but concentrationsof 0.3 to 2.5 µM have been reported near the oxic-anoxicboundary of stratified marine water bodies (5, 22). The distributionof NO₂^– in sediments is largely uncharacterized, exceptfor a few studies that have documented NO₂^– accumulationsin freshwater sediments (19, 38, 39, 40). The recent discoveryof high anammox rates in marine sediments (6) has caused increasedinterest in NO₂^– in sediments.

Generally, relatively little attention has been directed towardsdescriptions of NO₂^– concentrations in wastewaters; themain emphasis has been on the status of ammonium (NH₄⁺) andnitrate (NO₃^–). In wastewater treatment N removal is traditionallyperformed by degradation of the organic compounds, followedby oxidation (nitrification) of the liberated NH₄⁺ to NO₃^–and subsequent reduction (denitrification) of the NO₃^–produced to free nitrogen (N₂). During this type of oxic-anoxiccycling, transient NO₂^– accumulations have been foundand were recognized to be a result of imbalanced nitrificationand denitrification processes (29). Inhibitory effects of highNO₂^– concentrations have been observed for different reactorprocesses, like nitrification, denitrification, and biologicalphosphate uptake (2, 13, 27, 43). Accumulation of NO₂^–is also presumed to promote release of the greenhouse gas nitrousoxide (N₂O) from wastewaters (12, 16). Recently, the researchfocus has been on development and control of partial nitrificationto NO₂^– (14, 33, 42) instead of full nitrification toNO₃^–. Compared to full nitrification-denitrification processes,the implementation of partial nitrification-anammox processesis expected to significantly decrease energetic and economiccosts of N removal from high-ammonium wastewaters (17, 36).

Several different types of NO₂^– sensors have been developed.Examples include a liquid ion-exchange sensor (7) and a sensorbased on an immobilized enzyme (35). However, as tools for onlinemeasurement of NO₂^– in wastewater or microscale analysisof NO₂^– in marine sediment, these types of sensors havedistinct limitations. A specific NO_x^– biosensor for environmentalanalysis, including wastewater monitoring, has been describedpreviously (24, 26, 29); this biosensor is based on bacterialreduction of NO₂^– and NO₃^– to N₂O, followed by electrochemicalN₂O detection. The strain used in the NO_x^– biosensor hasa truncated denitrifying pathway with termination at N₂O insteadof N₂. If we could identify a suitable bacterium that also lacksa nitrate reductase, it should be possible to make an NO₂^–biosensor. Apparently, very few bacterial strains have sucha restricted denitrifying pathway, and only three such isolateshave been described (10). In this paper we describe a new NO₂^–biosensor based on one of these strains and compare this sensorwith a sensor based on Alcaligenes faecalis, which may be usedwhen acetylene is added to block its N₂O reductase (45).

MATERIALS AND METHODS

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Materials and Methods

Macroscale biosensor construction.
The macroscale NO₂^– biosensor functioned according tothe same principles as those previously described for a microscaleNO_x^– biosensor (23) and was in most respects identicalto the commercially available NO_x^– biosensor (Unisense,Aarhus, Denmark). The sensor consisted of two major compartments(Fig. 1). A Clark-type N₂O sensor (Unisense) with a tip diameterof about 0.275 mm was positioned behind a reaction chamber withan inner diameter of 0.3 mm. The bacterial chamber was drilledin a 0.35-mm-thick polyester plate that was subsequently gluedonto a 6-mm-inside-diameter glass tube. The two compartmentswere glued together so that the N₂O sensor partly protrudedinto the hole of the plate, creating a small, 0.15- to 0.3-mmdeep ( $~$ 0.01- to 0.02-mm³) reaction chamber in front of the sensortip. Bacterial biomass for the sensor was obtained by aerobicgrowth on a tryptic soy broth (TSB) agar plate. The culturewas collected from the plate with a sterile glass rod and subsequentlyinoculated into the reaction chamber from the front of the sensorthrough a very thin plastic tube (diameter, 0.1 to 0.2 mm) madeby heat drawing a plastic syringe (125-µl Distritip syringe;Gilson). To help retain the culture in the reaction chamber,a 1.3% agarose solution was placed behind the bacterial biomass.To separate the bacterial biomass from the outside environment,a 50-µm-thick ion-permeable membrane (Unisense) was attachedto the front plate by using double-adhesive tape. A large mediumreservoir ( $~$ 1,000 mm³) supplied the culture with all essentialgrowth constituents except the electron acceptor. The standardgrowth medium inside the sensor contained 4 g of sodium acetateliter^–1, 5 g of sodium citrate · 2H₂O liter^–1,0.5 g of TSB (Difco) liter^–1, 0.1 g of (NH₄)₂SO₄ liter^–1,and 3.3 g of Na₂WO₄ liter^–1 ( $~$ 10 mM). The pH was adjustedto 6.9. Tetracycline (20 mg liter^–1) was added to thegrowth medium when Stenotrophomonas nitritireducens was used.Acetylene-saturated growth medium was used when the organismwas A. faecalis. A sterilization procedure in which propyleneoxide was the sterilizing agent was used to minimize the frequencyof contamination with other bacteria, as follows: (i) dropletsof propylene oxide were added to the bacterial chamber and allowedto evaporate; (ii) pieces of ion-permeable membrane were bathedin a solution of 10% propylene oxide in water for 24 h beforeuse; and (iii) the glue surface of the sensor tip was exposedto a 10% propylene oxide solution for 4 to 6 h while it wascoated with a 5-µm-thick polyethylene film. All work withthe highly toxic and volatile compound propylene oxide was performedin a fume hood.

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FIG. 1. Macroscale NO₂^– biosensor based on bacterial reduction of NO₂^– to N₂O and subsequent detection by a built-in electrochemical N₂O sensor. The entire sensor is shown on the left. An enlargement of the sensor tip region is shown on the right.

Microscale biosensors.
Microscale biosensors were constructed as described by Larsenet al. (23) and were made with tips as large as 60 µmto enable analysis of low NO₂^– concentrations (<5 µM).They were operated with electrophoretic sensitivity control(ESC) (21), which means that a positive tip potential of 0.5V versus a standard calomel reference electrode was appliedto facilitate entry of the negatively charged NO₂^– ionsinto the sensor. The bacterial chambers of the microscale biosensorswere not sterilized, and the sensors usually functioned foronly a few days.

Sensor principle.
The detection of NO₂^– by the biosensor is based on bacterialreduction of NO₂^– to N₂O and subsequent detection of thisN₂O by the built-in electrochemical N₂O sensor. NO₂^– diffusesfreely through the ion-permeable membrane into the bacterialbiomass of the reaction chamber, where it is denitrified toN₂O and subsequently reduced to N₂ at the cathode of the N₂Osensor. There is a linear correlation between the amount ofNO₂^– reduced and the magnitude of the sensor signal. TheN₂O sensor is sensitive to O₂, but bacterial respiration inthe outer oxic parts of the chamber prevents O₂ from reachingthe cathode. Bacterial biomass in the deeper, anoxic parts ofthe chamber is responsible for the reduction of NO₂^– toN₂O, and the denitrifying capacity of the biomass determinesthe linear range of the sensor.

Physiological characteristics of bacterial strains.
Three organisms (S. nitritireducens, Luteimonas mephitis, andPseudoxanthomonas broegbernensis) that can be obtained fromDeutsche Sammlung von Mikroorganismen und Zellkulturen are allcharacterized by a denitrifying pathway deficient in both NO₃^–and N₂O reductases (10). These three organisms were tested foruse in an NO₂^– biosensor together with the denitrifyingorganism A. faecalis, which is deficient in an NO₃^– reductasebut has an active N₂O reductase (31, 32). Batch cultures withcomplex media containing 10 g of TSB liter^–1 plus 1 mMNO₃^– and batch cultures with 10 g of TSB liter^–1plus 1 mM NO₂^– incubated under anoxic conditions wereused to test the capacity for NO₃^– and N₂O reduction,respectively, in order to confirm previously described denitrifyingpathways. An N₂O sensor (1) was used to measure accumulationsof N₂O in the batch cultures. NO₂^– tolerance was examinedby anoxic incubation of batch cultures (10 g of TSB liter^–1)in the presence of 0, 2, 5, and 10 mM NO₂^–. The pH growthrange was evaluated by oxic incubation on agar plates at pH5 to 9. Detailed physiological studies of growth-temperatureand growth-salinity relationships were performed with S. nitritireducensand A. faecalis. The growth rate was examined at a temperaturerange of 15 to 42.5°C and at a salinity range of 0 to 90g of NaCl liter^–1 (in addition to the salts present inTSB). The growth rates of L. mephitis and P. broegbernensiswere determined only at 30°C and with no NaCl. For eachgrowth condition three replicates of 500-ml Erlenmeyer bottlescontaining 200 ml of complex medium (10 g of TSB liter^–1)were used. After addition of a small inoculum, the bottles wereplaced in an incubator shaker (Innova 4330 refrigerated incubatorshaker; New Brunswick Scientific) and kept at a constant temperaturewith continuous stirring (120 rpm). The bacterial density (opticaldensity at 600 nm) was measured regularly to determine the growthrate. Prior to salinity experiments the cultures used for inoculawere acclimatized to the experimental conditions to preventa long lag phase due to osmotic stress.

Sensor characteristics.
Macrosensor characteristics and function were examined mostextensively for biosensors with S. nitritireducens, but mostcharacteristics were also tested for sensors constructed withA. faecalis. An experimental setup in which several biosensorscould be tested simultaneously was used. The setup consistedof a plastic container holding 2 liters of water with controlof the temperature, O₂ level, turbulence, and concentrationsof various chemical species. Calibrations were performed todetermine the linearity, linear range, response time, and sensitivity.The effects of various parameters on the sensor signal wereevaluated; these parameters included temperature, O₂ level,salinity, turbulence, and pH. Interference with the NO₂^–signal was tested for a range of environmentally relevant compounds,including NO₃^–, NO, N₂O, NH₄⁺, NH₂OH, and H₂S. Exceptfor a linearity check, no investigations of microsensor performancewere conducted as similar characteristics were expected.

Biosensor method versus standard analytical method.
Activated sludge from a municipal wastewater treatment plantwas placed in a 2-liter laboratory-scale reactor and exposedto oxic-anoxic cycles. Two NO₂^– biosensors, one NO_x^–biosensor (Unisense), and one Clark-type O₂ minisensor (Unisense)supplied online information about chemical species in the sludge.Sensor signals were recorded continuously by using a data logger(ADC-16; Pico Technology) connected to a laptop computer. Calibrationsof NO₂^– and NO_x^– biosensors were performed in 100ml of reactor water that was previously separated from the biomassfraction by centrifugation and filtration. NO₂^– and NO₃^–were added from 100 mM stock solutions to final concentrationsof 100 µM. During the oxic-anoxic cycle, sludge sampleswere obtained from the reactor at 5-min intervals, rapidly filteredthrough 0.22-µm-pore-size filters (CAMEO 25AS; Osmonics),and immediately frozen. NO₂^– concentrations in the sampleswere subsequently determined with a biosensor and also by spectrophotometry(30). Sensor measurements of NO₂^– levels were obtainedat a controlled temperature in a small glass container withconstant stirring. To increase the small volumes of filtrateobtained, 2-ml samples were mixed with 2 ml of a 1-g liter^–1NaCl solution. Sensor calibration was performed at a similarsalinity by using 2 ml of sterile filtered wastewater (withoutNO₂^–) and 2 ml of an NaCl solution.

Environmental monitoring.
Macroscale NO₂^– biosensors were used to monitor NO₂^–concentrations in a 1,000-liter pilot-scale wastewater treatmentplant with activated sludge during a 3-week period. The plantwas operated with intermittent oxic-anoxic cycles. Microscaleversions of the NO₂^– biosensor were used to analyze theNO₂^– distribution in sediment from the Wadden Sea (Germany)in a laboratory setup.

RESULTS AND DISCUSSION

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Results and Discussion

Our research goal was to develop a long-term stable macroscaleNO₂^– biosensor with a lifetime on the order of monthsthat is suitable for laboratory analysis and online monitoringof wastewater. In addition, we also wanted to use the technologyfor construction of an NO₂^– microscale biosensor for profilingbiofilms and sediments. A successful outcome of our effortswas expected due to the availability of a stable electrochemicalN₂O sensor (1) that has previously been used in well-functioningNO_x^– biosensors.

Physiological characteristics of bacterial strains.
In a preliminary evaluation of the four bacterial strains availablefor the NO₂^– biosensor, the two most important propertieswere denitrification characteristics and growth rate. The objectivewas to identify the most suitable strain, ideally a strain witha very restricted denitrifying pathway limited to the reductionof NO₂^– to N₂O. Alternatively, successful use of a strainwith N₂O reductase activity should also have been possible whenacetylene was added to the sensor interior to block N₂O reductaseactivity (45). Tests with batch cultures supplemented with NO₂^–or NO₃^– confirmed previously described denitrificationpathways (Table 1). A high respiration rate of the culture usedwas equally important to ensure complete consumption of O₂ andto give the sensor a wide linear range for NO₂^–. S. nitritireducensand A. faecalis (Table 1), which were characterized by growthrates that were significantly higher than those of the othertwo organisms, were selected as the most promising candidatesfor the biosensor. The linear range for similar NO_x^– biosensorsdoes not exceed 2 mM at room temperature, so the NO₂^–tolerance of up to 5 mM observed for S. nitritireducens wasmore than sufficient for any relevant measuring range. A. faecalisexhibited even higher NO₂^– tolerance.

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TABLE 1. Physiological characteristics of four bacterial strains that were tested for the NO₂^– biosensor

Temperature and salinity effects on the growth rate were criticalaspects of the bacterial physiology, as these two key parameterslimit the range of environmental applications of a biosensor.The growth constants that were obtained showed that both S.nitritireducens and A. faecalis are typical mesophilic organisms(Fig. 2A), with temperature optima around 33 and 37°C, respectively.The growth range was broader in the case of A. faecalis, whichhad higher upper and lower temperature limits for growth. Thesalinity-growth curves showed that both organisms had the highestgrowth rate under no-salt conditions, but there was a significantlyhigher salt inhibition factor for S. nitritireducens than forA. faecalis. At a sea strength concentration (34 g liter^–1)the growth rate of S. nitritireducens was reduced to less thanone-third the maximum rate when no salts were added, and theupper limit for growth was about 45 g liter^–1. A. faecalisshowed only a 20% reduction in the growth rate at a sea strengthconcentration, and the upper limit for growth was about 80 gliter^–1.

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FIG. 2. Temperature-growth curves (A) and salinity-growth curves (B) for S. nitritireducens ( ${circ}$ ) and A. faecalis ( ${square}$ ). The growth rates (µ_max) are the means for three replicates, and the error bars indicate standard deviations.

In summary, both of the organisms characterized could grow undera broad range of conditions. However, the physiological dataobtained also indicated that under certain environmental conditions,biosensor application would be suitable only when A. faecaliswas used. In particular, for S. nitritireducens a comparativelylow growth rate (and respiration rate) at a low temperatureand high salinity might limit the applicability of sensors madewith this organism.

Sensor characteristics.
The sensor characteristics described below refer to macroscalebiosensors containing S. nitritireducens, but most characteristicswere also observed for sensors with A. faecalis. The biosensorhad a linear response to NO₂^– (Fig. 3), and dependingon individual sensor characteristics the linear range extendedto between 0.5 and 2 mM NO₂^– at 20°C. A negative correlationbetween the linear range and the sensor response time was found,with 90% response times ranging from 0.5 to 3 min (20°C),indicating that the sensors with the longest diffusion pathbetween the tip membrane and the internal electrochemical N₂Osensor and thus the largest active biovolume had the largestmeasuring range. The NO₂^– response was about 4 to 6 pAfor 1 µM NO₂^–, and the sensor zero signal rangedfrom 100 to 300 pA (20°C). The detection limit was around1 µM NO₂^– and depended primarily on the stabilityof the zero signal from the N₂O sensor

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FIG. 3. Calibration curves for an NO₂^– biosensor (S. nitritireducens) at three temperatures, 10°C ( ${triangleup}$ ), 20°C ( ${circ}$ ), and 30°C ( ${square}$ ).

The effects of temperature, salinity, turbulence, pH, and O₂level on the sensor characteristics were investigated. Therewas a substantial temperature effect as a result of the temperatureinfluence on diffusion coefficients and the bacterial respirationrate (Fig. 3). Several sensor characteristics were affected,including the linear range, the slope of the calibration curve,the sensor zero signal, and the response time. In the temperaturerange from 10 to 37°C the linear range of the sensor increasedwith an increase in the temperature. At temperatures below 8to 9°C several sensors ceased to work and became sensitiveto O₂ due to an insufficient respiration rate. This observationcorrelated well with the temperature effect on the growth ratefound for S. nitritireducens in batch culture (Fig. 2A). A significantincrease in the NO₂^– response corresponding to an increasein the slope of the calibration curve of $~$ 2.5% per degree Celsiuscorresponded well to the calculated increase in diffusion coefficients(4, 25) at 20°C, which was 2.7% per degree Celsius. Highertemperatures also increased the zero signal, presumably by increasedhydrogen production at the cathode surface by electrolysis ofwater. The response time decreased with increasing temperatureas a result of the increased diffusion speed, and the 90% responsetime for a typical sensor thus decreased from about 115 s at15°C to about 80 s at 30°C. This is in agreement withthe theoretical change in response time calculated by the Einstein-Smulochowskiequation (18), which states that the diffusion time is inverselyproportional to the diffusion coefficient. According to thisequation there should be a 1.46-fold increase in the responsetime when the temperature is lowered from 30 to 15°C.

The sensor signal was influenced by changes in the salinityand turbulence of the medium analyzed (Fig. 4). An increasedNO₂^– signal was observed with an increased salt concentration(Fig. 4A). This effect was most pronounced at low salt concentrations,and at salinities above about 0.5 to 1.0 g liter^–1 therelative signal change due to moderate variations in salinitywas small. However, the salinity effect was also affected bythe ionic composition of the internal sensor medium. Additionof extra salt (NaCl) to the internal medium reduced the sensitivityto variations in the external salinity, particularly in thelower-salinity range (Fig. 4A). Both external salinity and internalsalinity also affected the sensitivity to turbulence in theexternal medium. Generally, sensors exhibited the greatest sensitivityto turbulence at a low external salinity (Fig. 4B), and at anNaCl concentration of 0.1 g liter^–1 the signal in stagnantmedium was up to 18% higher than the signal in vigorously stirredmedium. At salinities higher than 0.5 g liter^–1, the sensitivityto stirring was negligible (±2%). The signal differencebetween low turbulence and high turbulence was much less thanthe difference between totally stagnant and vigorously stirredconditions illustrated in Fig. 4B.

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FIG. 4. Responses of two NO₂^– biosensors (sensor 1 [ ${square}$ and ${blacksquare}$ ] and sensor 2 [ ${circ}$ and •]) to variations in salinity (A) and water turbulence (B) of a medium containing 100 µM NO₂^–. Data are shown for two types of internal sensor media that differed in ionic strength, standard medium (open symbols) and standard medium containing 10 g of NaCl liter^–1 (solid symbols). The turbulence effects (B) are expressed as the relative difference between the sensor signals under stagnant and stirred conditions.

No marked effect of either pH or O₂ level on sensor characteristicswas observed. Variation in the external pH did not affect thesensor signal in the range from pH 5 to 9. However, a shiftfrom oxic to anoxic conditions caused a small decrease in thesensor zero signal that for the sensors tested correspondedto an NO₂^– concentration of 0.1 to 4 µM. This deviationcan be explained by the relatively higher CO₂ production inthe bacterial biomass during oxic conditions that affected theN₂O sensor signal. High CO₂ concentrations have previously beenshown to elevate the zero signals from N₂O sensors (1). Furthermore,a slight increase ( $~$ 1%) in the relative NO₂^– response wasobserved under anoxic conditions. This effect may be explainedas a change in the location of the denitrifying zone insidethe reaction chamber in the absence of O₂. Unexpectedly, noeffect of O₂ status on the linear range for NO₂^– was observed.A broader range was expected under anoxic conditions due toa potentially larger NO₂^–-respiring biomass, and a widerrange has previously been observed for NO_x^– biosensors(23).

Interference.
Ideally, the biosensor developed should respond only to NO₂^–,and it was thus important to examine interference with the NO₂^–signal by relevant chemical species. As expected, the sensorresponded to any N₂O present in the system. Theoretically, equalconcentrations of N₂O and NO₂^– should result in a 2.5-fold-highersignal for N₂O as it takes two NO₂^– molecules to produceone N₂O molecule and the diffusion coefficient for NO₂^–is about 0.8 times that of N₂O (4, 25). However, variationsin the tip potential of the sensor may change this value, andfor several sensors tested a factor of about 3 was observed.For NO, which was reduced to N₂O by the bacteria inside thesensor, the equivalent factor was about 1.5. Low to moderate(100 µM) concentrations of hydrogen sulfide (H₂S) at pH7.2 had no effect on the sensor zero signal, but only 1 µMH₂S caused a 15% decrease in the NO₂^– sensitivity, andat a concentration of 100 µM less than 5% of the signalremained. Fortunately, the decrease in sensitivity caused byH₂S is reversible (1)

Interference from NH₄⁺ and NH₂OH was expected for sensors basedon A. faecalis because this organism has been described as aheterotrophic ammonium oxidizer and also is known to oxidizeNH₂OH with production of N₂O (31). However, NH₄⁺ interferencewas not observed for sensors containing either of the two organisms.Interference from NH₂OH was observed when both organisms wereused, and the interfering signal was further enhanced by a highNH₄⁺ concentration. The relative response to NH₂OH was highlyvariable, corresponding to 1 to 10% of the signal for an equivalentconcentration of NO₂^–, and the response was much slowerthan the responses to NO₂^–, N₂O, and NO. The effect wasmost pronounced with biosensors containing S. nitritireducens.No interference from NO₃^– was observed with freshly preparedsensors, but during the first 1.5 years of our work with theNO₂^– biosensor NO₃^– interference developed afterperiods varying from hours to weeks. Analysis of biomass fromNO₃^–-sensitive sensors confirmed the presence of NO₃^–-reducingcontaminants. Addition of tungstate (Na₂WO₄) to the internalgrowth medium at a final concentration of 10 mM alleviated thecontamination problem with sensors containing S. nitritireducens.Sensors made with A. faecalis have not been tested with tungstate.Tungstate is placed in the NO₃^– reductase of contaminantsinstead of molybdenum, resulting in a nonfunctional enzyme (9).Not all bacterial NO₃^– reductases are completely inhibitedby even 10 mM tungstate (11), but for the present applicationthe inhibitory effect seemed to be sufficient.

The most relevant interfering agent is probably N₂O. There havebeen several reports which described release of significantamounts of N₂O from wastewaters, and the rate of N₂O productionhas been correlated with various parameters, like low pH, highNO₂^– levels, and low chemical oxygen demand/N ratios (12,16, 41). We have, however, never observed detectable N₂O orNO (>0.5 µM) during normal operation with nitrifying-denitrifyingactivated sludge, nor have we detected the oxidized nitrogengases (concentrations, >0.5 µM) in marine sediments.By use of microsensors it is very easy to check for the presenceof N₂O plus NO, as a negative ESC potential of 0.5 V preventsentry of NO₂^– plus NO₃^– into the sensor (21) anda signal can then be assumed to be due to N₂O plus NO. We arecurrently trying to develop membranes that allow use of ESCalso with a macroscale sensor so that similar interference tests(and zero control) can be performed. H₂S may be a problem insome systems, but the low concentrations present in, for example,activated sludge have not affected the sensor response. Formost field applications, changes in sensitivity and zero currentcaused by changes in temperature are a much larger source oferror than chemical interference.

Stability of NO₂^– biosensor.
Once bacterial biomass was successfully introduced, the biosensorimmediately responded to NO₂^–. The stability of the sensorsignal was examined during online monitoring in a pilot-scalewastewater treatment plant and in different laboratory setups.Generally, the characteristics of the biosensors constructedchanged gradually, and there was an observed increase in thesensor zero current (0 to 100%) and also a decrease in the relativeNO₂^– response (0 to 30%) during a 2-month period in alaboratory setup. For most sensors the linear NO₂^– rangeremained unchanged without a loss of bacterial activity. Itis likely that sensors may monitor NO₂^– for up to 3 months,as this is the case for the equivalent NO_x^– biosensors,but due to previous problems with contamination of cultureswithin biosensors we have not tested NO₂^– biosensors inwastewater for more than 6 weeks. None of four biosensors constructedwith a tungstate-containing inner medium that were used in wastewaterfor 3 weeks showed any sign of deterioration of characteristicsin this period.

Although addition of tungstate seems to be efficient in termsof avoiding NO₃^– reduction to NO₂^– within the sensor,contamination should be kept at a low level to minimize therisk of contamination with an N₂O-reducing bacterium that wouldrender the sensor insensitive to all nitrogen species. Threeroutes of contamination were recognized: (i) proliferation ofbacteria already present on various sensor parts, (ii) introductionof contaminants during the inoculation procedure, and (iii)contamination with bacteria from the external environment causedby insufficient sealing at the glue-membrane interface. Sterilizationof various sensor parts, including the interior of the reactionchamber, the membrane, and the glue surface, by using propyleneoxide as the sterilization agent proved to be very efficient.Addition of tetracycline at a concentration of 20 mg liter^–1to the growth medium was used for sensors containing S. nitritireducensto limit proliferation of contaminants that might enter alongthe glue-membrane interface. However, like S. nitritireducens,many other bacterial strains are resistant to tetracycline (34).Analysis of biomass from several contaminated sensors to whichtetracycline was added revealed resistance for all contaminantsexamined.

Biosensors containing A. faecalis.
Functional biosensors were successfully constructed by usingA. faecalis, although this organism has a denitrifying pathwaywith termination at N₂ instead of N₂O. It was possible to solvethis problem through saturation of the growth medium insidethe sensor with acetylene, which inhibited the activity of N₂Oreductase (45). However, in many cases the sensor response tolow NO₂^– concentrations was completely absent (Fig. 5),presumably due to inadequate acetylene saturation in some anoxicparts of the reaction chamber. Apparently, a relatively loosefit between the sensor tip and the reaction chamber was criticalto ensure sufficient diffusive flux of acetylene from the mediumreservoir to the reaction chamber. Acetylene inhibition wasimproved if an N₂O sensor with a smaller tip diameter was usedand when the usual 0.35-mm front plate was replaced with a 0.2-mmplate. However, such sensors suffered from slower response dueto diffusion of N₂O in the gap between the N₂O sensor and thefront plate wall; when acetylene could be supplied to the tip,N₂O could also escape to the bulk medium reservoir. In agreementwith the growth-versus-temperature relationships (Fig. 2A) biosensorsconstructed with A. faecalis functioned well at temperatureslower than the temperatures at which sensors containing S. nitritireducensfunctioned. At 5°C, the A. faecalis-based sensors were stillworking and had a linear range to about 0.5 mM NO₂^–.

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FIG. 5. Calibration curves for two NO₂^– biosensors containing the bacterium A. faecalis, showing examples of complete ( ${circ}$ ) and incomplete ( ${square}$ ) inhibition of N₂O reductase activity by acetylene.

Biosensor versus standard analytical method.
An experiment was conducted to examine the performance of theNO₂^– biosensor during online monitoring of wastewater.Fast process dynamics made it difficult to make an unbiasedcomparison of online biosensor readings for the sludge witha standard analysis of NO₂^– in filtered samples, and forcomparison we therefore also analyzed NO₂^– in the sampleswith a biosensor.

A very good correlation was found between online signals fromthe two NO₂^– biosensors, which showed almost identicalNO₂^– concentrations during the complete cycle (Fig. 6).Furthermore, NO₂^– analysis of a filtered sample showedthat there was good agreement between conventional analysisand sensor analysis. There was, however, a marked differencebetween the data from the online sensor analysis and the concentrationsanalyzed in the filtered samples. This discrepancy could beexplained by either the 1- to 2-min response time of the sensorsor the nitrogen conversions that occurred during processingof the filtered samples.

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FIG. 6. Calibrated online sensor outputs for an O₂ sensor, an NO_x^– biosensor, and two NO₂^– biosensors during one cycle consisting of 0.75 h of oxic conditions and 1 h of anoxic conditions in a 2-liter laboratory-scale reactor with activated sludge. The graph includes comparable data for NO₂^– concentrations in filtered samples analyzed with a biosensor or by spectrophotometry.

Environmental monitoring.
Online monitoring of NO₂^–, NO_x^–, and O₂ in activatedsludge exposed to oxic-anoxic cycles (Fig. 7) demonstrated thatNO₂^– accumulated to concentrations of about 65 µMduring operation. NO₂^– accumulated in the early part ofthe oxic phase due to higher rates of NO₂^– production(ammonia oxidation) than of NO₂^– consumption (nitriteoxidation). A second NO₂^– peak was observed during theanoxic phase, which was caused by imbalanced denitrificationwith higher rates of NO₃^– reduction than of NO₂^–reduction.

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FIG. 7. Online sensor outputs for two NO₂^– biosensors [NO₂^– (1) NO₂^– (2)] and one NO_x^– biosensor (NO_x^–) during cycles consisting of 1.5 h of oxic conditions and 1.5 h of anoxic conditions in a pilot-scale wastewater treatment plant with activated sludge. The shaded areas indicate oxygenated periods. Calibrated sensor signals correspond to concentration peaks of about 65 µM NO₂^– and 600 µM NO_x^–. The sensor signals for 100 µM NO₂^– and 100 µM NO_x^– were as follows: biosensor NO₂^– (1), 0.42 nA; biosensor NO₂^– (2), 0.33 nA; and biosensor NO_x^–, 0.79 nA.

Microscale analysis of NO₂^–, NO_x^–, and O₂ in a marinesediment (Fig. 8) showed that there was NO₂^– accumulationto concentrations of about 3 to 6 µM in the oxic partof the sediment. The NO₂^– peak was located at a depthof about 1.5 mm and was found at approximately the same depthas the NO_x^– peak (about 15 to 19 µM). The NO₂^–accumulation was most likely due to a high ammonia oxidationrate in the oxic part of the sediment. The NO₂^– concentrationgradients indicated that there was a flux down to the deeperanoxic layers ( $~$ 2.4 to 3.2 mm) and that there was also net releaseof NO₂^– to the overlying water phase.

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FIG. 8. Concentration profiles for NO₂^– and NO_x^– in a marine sediment (Wadden Sea) measured during incubation in the dark in a laboratory setup. The shaded area indicates the location of the oxic zone in the sediment.

Perspectives.
The new biosensor may be used for sensitive analysis of NO₂^–at high spatial and temporal resolution. This sensor can beused in basic research on processes, but it may also be a valuabletool for online monitoring and feedback control in wastewatertreatment. The preferred organism for the biosensor is S. nitritireducens.However, for some purposes it maybe more suitable to use sensorscontaining A. faecalis, due to this organism's higher toleranceto extremes of temperature and salinity. We are currently tryingto isolate psychrotrophic bacteria that enable NO₂^– analysisat temperatures down to those of freezing seawater, so thatwe can perform analyses under most relevant environmental conditions.

ACKNOWLEDGMENTS

This study was performed as part of the Icon project under theFifth Framework Programme of the European Commission (EVK1-CT-2000-00054).

FOOTNOTES

* Corresponding author. Mailing address: Department of Microbiology, University of Aarhus, Bd. 540, Ny Munkegade, 8000 Aarhus C, Denmark. Phone: 45 89423329. Fax: 45 86127191. E-mail: mnielsen{at}biology.au.dk.

REFERENCES

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