INTRODUCTION

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Petroleum hydrocarbons are among the most common environmental contaminants (8). Much work has been done to understand the aerobic and anaerobic pathways of microbial hydrocarbon degradation (9, 29). Although desirable for active hydrocarbon biodegradation, completely aerobic conditions are hard to implement in the field because of the low solubility of oxygen in water (5, 42). Hydrogen peroxide can provide more oxygen (20). However, hydrogen peroxide may oxidize cellular components by forming singlet oxygen and hydroxyl radicals (29), and using hydrogen peroxide requires special operational design and handling. Moreover, peroxide can form precipitates with soil components, such as metallic compounds, thus decreasing aquifer permeability (11, 30, 40).

It is plausible that uneven distribution of water flow, nutrients, and microbial populations creates a dynamic spectrum of aerobic, microaerobic, and anaerobic conditions. The ability of microorganisms to degrade hydrocarbons under strictly anaerobic conditions is very limited. It is practically absent for aliphatics compounds; there are extremely rare exceptions to this (10, 13, 36). The ability to degrade aromatic compounds under anaerobic conditions is limited to a few strains, and the degradation is typically much slower than aerobic degradation (4, 15, 24). The initial anaerobic transformation of aromatic compounds has been shown to be oxidative hydroxylation, with the oxygen being derived from water (1, 22), which is a much less thermodynamically favorable reaction than oxidation by O₂.

The limitation described above indicates the importance of molecular oxygen (O₂) as a direct reactant in hydrocarbon catabolism (e.g., in the initial conversion to alcohol) (29). Although other oxidants, such as NO₃ or NO₂ (in denitrification), Fe(III), Mn(IV), sulfate, and CO₂ (in methanogenesis) (17, 29), can be used by many microorganisms as terminal electron acceptors that replace O₂ in respiration, they cannot replace O₂ as a direct reactant. Consequently, most anaerobic biodegradation is observed with oxygenated derivatives (e.g., alcohols, aldehydes, acids, phenolic compounds, and amino acids) (3, 16).

Therefore, petroleum bioremediation should address (i) the dual roles of O₂ in biodegradation (i.e., for respiration and as a direct reactant) and (ii) the interactions of microbial activities in different environments (aerobic, microaerobic, and anaerobic environments). Sequential biodegradation of hydrocarbons probably occurs in nature; the initial O₂-requiring oxidation occurs under aerobic or microaerobic conditions, and the subsequent mineralization may proceed even under anaerobic conditions. The same strategy may be used as an advanced bioremediation approach. The initial oxidation (transformation) is effected by aerobic resting cells in a first remediation stage without promoting active growth that may cause undeliverable oxygen demand and plugging of soil pores by overpopulated microbes. The oxidized metabolites are then degraded under the conditions of stimulated microbial activities where the respiration needs are met by adding a highly water-soluble alternative electron acceptor, such as nitrate.

In this work we studied sequential metabolism of n-hexadecane by Pseudomonas aeruginosa, which is among the most commonly isolated microorganisms in petroleum-contaminated soils and groundwater (34). It is known that P. aeruginosa mineralizes aliphatic compounds. P. aeruginosa strains that attack aromatic and polyaromatic hydrocarbons have also been isolated, including a pyrene-degrading strain that was recently isolated in our lab (33). P. aeruginosa strains are typically active denitrifiers and produce biosurfactants (rhamnolipids) when they are grown on hydrophobic substrates (18, 41). Compared to other anaerobic respiration mechanisms, denitrification is more favorable energetically, and benign nitrogen (N₂) is the predominant product of this process (17, 29). The rhamnolipids produced are also very beneficial to bioremediation, because they solubilize and mobilize hydrocarbons and other non-aqueous-phase liquids into the aqueous phase, where they can be biodegraded or removed by advective transport (2, 37).

Because they are hydrophobic, hydrocarbons in the environment are often adsorbed or trapped in pores by capillary action; thus, they are not readily accessible to microbes (26). Some microorganisms produce extracellular biosurfactants that solubilize and facilitate the penetration of hydrocarbons through the hydrophilic cell wall; then the hydrocarbons can be degraded by enzymes integrated in the cytoplasmic membrane (39). The most important examples of such organisms are the rhamnolipid-producing Pseudomonas species (28, 44) and the sophorolipid-producing Torulopsis species (21). The biosurfactants of these species are extremely effective and create low surface tensions at much lower concentrations than those required for synthetic surfactants (14).

It should be noted that rhamnolipids are actively synthesized by stationary-phase or resting P. aeruginosa cultures (44). Therefore, they are also produced during the first stage of the proposed sequential bioremediation process and contribute to mobilizing and solubilizing the contaminants during the subsequent mineralization stage. Because P. aeruginosa has these versatile, unique metabolic characteristics, it is the most suitable species for initial studies of sequential hydrocarbon metabolism.

	MATERIALS AND METHODS

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Microorganism and medium. A pure culture of P. aeruginosa ATCC 10145 was maintained on tryptic soy agar slants (30 g of tryptic soy broth [TSB] per liter, 1.5% agar) and was subcultured regularly. To prepare an inoculum, a loopful of cells from an agar slant was first transferred into a test tube containing 10 ml of TSB and incubated at 35°C for 24 h. For studies in which TSB or glucose medium (basal medium [see below] containing 2 g of glucose per liter) was used, the cells were transferred and grown in 50 ml of TSB at 30°C for 2 days with magnetic stirring at 300 rpm before they were used as the inoculum. For the experiments in which n-hexadecane was the carbon substrate, the cells in 10 ml of TSB were harvested by centrifugation and added to a 500-ml Erlenmeyer flask containing 100 ml of basal medium supplemented with 5 ml of hexadecane; the pH of this medium was adjusted to 7.0. The culture was incubated for 7 days with magnetic stirring at 30°C and then subcultured again in the hexadecane-containing medium for another 4 days before it was used as the inoculum. The basal medium contained (per liter) 4.0 g of NH₄Cl, 2.5 g of K₂HPO₄, 0.5 g of NaCl, 0.3 g of MgSO₄ · 7H₂O, 0.03 g of FeCl₃ · 6H₂O, 0.01 g of CaCl₂, and 0.01 g of MnCl₂ · 4H₂O.

Experimental setup and procedures. (i) Microaerobic denitrification and hexadecane metabolism: effect of oxygen concentration. To study the full spectrum of aerobic, microaerobic, and anaerobic conditions in soil, we performed experiments to confirm the existence of microaerobic denitrification and the effects of low oxygen concentrations on hexadecane metabolism by P. aeruginosa. Different volumes of an inoculated growth medium containing hexadecane, nitrate, and ammonium were added to nine 50-ml vials. An excess of ammonium (0.65 g of NH₄⁺-N per liter) was present to ensure that the nitrate was consumed only for respiration, because the assimilatory nitrate reductases are repressed by ammonium (27). The vials were stirred at the same speed (100 rpm) by using a multipoint electromagnetic stirrer. The caps on vials 1 through 5 were loosened slightly in order to allow gas phase diffusion. Vials 6 through 9 were tightly capped. Since the gas-liquid interfacial areas were the same, the increasing medium volume from vial 1 to vial 5 resulted in a decreasing trend in dissolved oxygen (DO) concentration. Vials 6 through 9 were the most anaerobic vials; the headspaces in these vials were very small, and there was no gas replenishment. Resazurin, a common redox indicator, was added in order to determine the extent of anaerobiosis; this compound is colorless under anaerobic conditions and pink under aerobic conditions (29). The resultant color confirmed the decreasing DO concentration trend. Samples were taken periodically and used for nitrate analysis.

(ii) Effects of nitrate and nitrite concentrations. Two sets of experiments were performed to study the effects of nitrate and/or nitrite concentrations on the growth of P. aeruginosa under aerobic conditions and under anaerobic denitrifying conditions. The experiments were conducted at room temperature (22 ± 1°C). TSB containing various initial concentrations of nitrite (0, 0.05, 0.10, 0.15, 0.20, 0.25, 0.30, and 0.35 g of NO₂-N per liter) were used for the aerobic experiments. The denitrifying experiments were performed with glucose media containing either nitrate alone (0.5, 1, 2, 3, or 4 g of NO₃-N per liter) or 0.5 g of NO₃-N per liter plus 0.1, 0.2, or 0.3 g of NO₂-N per liter. Very pure sodium nitrate (>99.9% pure; Sigma) was used to minimize the accompanying residual nitrite content. The aerobic experiments were performed with shallow medium (4 ml of medium) that was vigorously shaken (~250 rpm) in culture tubes capped with stainless steel closures that allowed good gas exchange. On the other hand, the tubes used for the anaerobic denitrifying experiments were tightly capped, filled with 10 ml of medium, and shaken gently at ~60 rpm. The broth optical density at 460 nm was measured with a Spectronic 20 colorimeter (Bausch & Lomb) in order to determine cell concentrations.

(iii) Hexadecane metabolism in the sequential aerobic and anaerobic denitrifying process. Experiments were carried out at 30°C in 2-liter Erlenmeyer flasks containing 800 ml of basal medium supplemented with 0.7% (vol/vol) hexadecane; the medium was magnetically stirred at 400 rpm. Initially, fermentation was carried out with surface aeration; a slow flow of filter-sterilized air through the reactor headspace resulted in cell growth and stationary-phase hydrocarbon oxidation to oxygenated metabolites, including rhamnolipids. The pH was controlled at 6.5 to 6.7 by automatic addition of 2 N NaOH during this period. Surface aeration was used instead of submerged air sparging in order to avoid severe foaming due to biosurfactant production. The culture was then driven to anaerobic denitrifying conditions by purging nitrogen (N₂) and adding NaNO₃. Ammonium (0.08 g of NH₄Cl per liter) was also added to ensure that nitrate was consumed only for respiration and not for assimilation. Before the N₂ was introduced into the fermentor, it was passed through an oversaturated NaSO₃ solution in order to minimize the residual oxygen content. N₂ purging was performed periodically during anaerobic cultivation, especially after each sample was removed. Because denitrification led to an increase in pH, the pH of the anaerobic culture was controlled by automatic addition of 2 N HCl.

Analytical methods. The cell protein and rhamnolipid concentrations were measured by standard Lowry and anthrone methods, respectively, as described previously (44). A total organic carbon (TOC) analyzer (model TOC-5000A; Shimadzu) was used to measure the TOC content in the aqueous phase after a sample (10 ml) had been extracted with hexane to remove hexadecane. Hexadecane contents were analyzed by high-performance liquid chromatography by using a normal-phase Supelcosil LC-Si column (25 cm by 4.6 mm by 5 µm) and a refractive index detector (model HP 1047A). The mobile phase was n-hexane at a flow rate of 1.0 ml per min.

	RESULTS AND DISCUSSION

Top Abstract Introduction Materials and Methods Results and Discussion References

Microaerobic denitrification and hexadecane metabolism: effect of oxygen concentration. The nitrate consumption profiles obtained with vials containing different DO concentrations are summarized in Fig. 1. Denitrification clearly occurred under the microaerobic conditions studied. This finding is consistent with the previous report that the bacteria in a groundwater microcosm ignored O₂ at levels that were less than 8% of air saturation and switched to other electron acceptors (6). Denitrification in the presence of even higher DO levels, termed aerobic denitrification, has also been reported for several species and strains (32, 38, 43).

View larger version (22K): [in this window] [in a new window]   FIG. 1.   Increase of the denitrification rate with increasing DO concentration during n-hexadecane metabolism by P. aeruginosa under microaerobic conditions. Symbols: , vial 1; , vial 2; , vial 3; , vial 4; , vial 5; , vial 6.

View larger version (29K): [in this window] [in a new window]   FIG. 2.   Proposed dependence of the denitrification rate of P. aeruginosa on DO concentration for different substrates. In addition to the well-known oxygen repression and inhibition, the denitrification rate is also subject to the limitation of electron donation from O2-requiring hydrocarbon oxidation at very low DO concentrations.

Effects of nitrate and nitrite concentrations. Denitrification by P. aeruginosa is known to involve sequential reduction of nitrate (NO₃) to nitrite (NO₂), nitric oxide (NO), nitrous oxide (N₂O), and nitrogen (N₂) (23). Accumulation of intermediate NO₂ during the initial denitrifying period has been observed (7), because nitrate reductases are first induced by the nitrate added and then the nitrite formed from nitrate reduction induces expression of nitrite reductases. Before nitrite reductases are fully synthesized, transient nitrite accumulation is likely to occur. Nitrite is, however, highly toxic (31) and may cause cell death or metabolic inhibition when denitrification is used for sequential bioremediation. Therefore, we examined P. aeruginosa growth in the presence of various nitrite and/or nitrate concentrations.

View larger version (22K): [in this window] [in a new window]   FIG. 3.   Effect of nitrite concentration on aerobic growth of P. aeruginosa in TSB. OD460, optical density at 460 nm.

View larger version (24K): [in this window] [in a new window]   FIG. 4.   Effects of nitrite and/or nitrate concentrations on growth of P. aeruginosa in a glucose-based medium under anaerobic denitrifying conditions. OD460, optical density at 460 nm.

Hexadecane metabolism in sequential aerobic and anaerobic denitrifying processes. The results obtained in a typical experiment carried out under sequential aerobic and anaerobic denitrifying conditions are shown in Fig. 5. The cell growth profile corresponded very well to the ammonium consumption profile, and the culture reached the stationary phase when the N source was depleted. The rates of hexadecane consumption and aqueous-phase TOC production under aerobic conditions were higher in the exponential growth phase than in the stationary phase. The increase in the TOC level confirmed that hydrocarbon was converted to oxygenated organic compounds by cell metabolism. Rhamnolipids were also produced, and the presence of rhamnolipids was accompanied by reduced medium surface tension (Fig. 5).

View larger version (33K): [in this window] [in a new window]   FIG. 5.   Results of a typical experiment to examine n-hexadecane metabolism by P. aeruginosa under sequential aerobic and anaerobic denitrifying conditions.

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View larger version (27K): [in this window] [in a new window]   FIG. 6.   Cell growth, TOC consumption, and transient nitrite accumulation profiles obtained in experiments performed with the high-TOC medium containing different initial nitrate concentrations. Symbols: , 2.7 g of NO3-N per liter; , 0.8 g of NO3-N per liter; , 0.4 g of NO3-N per liter.

View larger version (32K): [in this window] [in a new window]   FIG. 7.   Cell growth, TOC consumption, and transient nitrite accumulation profiles obtained in experiments performed with the low-TOC medium containing different initial nitrate concentrations. Symbols: , 3.3 g of NO3-N per liter; , 1.3 g of NO3-N per liter; , 0.7 g of NO3-N per liter; , 0.35 g of NO3-N per liter.

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