Use of ¹³C Nuclear Magnetic Resonance To Assess Fossil Fuel Biodegradation: Fate of [1-¹³C]Acenaphthene in Creosote Polycyclic Aromatic Compound Mixtures Degraded by Bacteria

Abstract

[1-¹³C]acenaphthene, a tracer compound with a nuclear magnetic resonance (NMR)-active nucleus at the C-1 position, has been employed in conjunction with a standard broad-band-decoupled ¹³C-NMR spectroscopy technique to study the biodegradation of acenaphthene by various bacterial cultures degrading aromatic hydrocarbons of creosote. Site-specific labeling at the benzylic position of acenaphthene allows ¹³C-NMR detection of chemical changes due to initial oxidations catalyzed by bacterial enzymes of aromatic hydrocarbon catabolism. Biodegradation of [1-¹³C]acenaphthene in the presence of naphthalene or creosote polycyclic aromatic compounds (PACs) was examined with an undefined mixed bacterial culture (established by enrichment on creosote PACs) and with isolates of individual naphthalene- and phenanthrene-degrading strains from this culture. From ¹³C-NMR spectra of extractable materials obtained in time course biodegradation experiments under optimized conditions, a number of signals were assigned to accumulated products such as 1-acenaphthenol, 1-acenaphthenone, acenaphthene-1,2-diol and naphthalene 1,8-dicarboxylic acid, formed by benzylic oxidation of acenaphthene and subsequent reactions. Limited degradation of acenaphthene could be attributed to its oxidation by naphthalene 1,2-dioxygenase or related dioxygenases, indicative of certain limitations of the undefined mixed culture with respect to acenaphthene catabolism. Coinoculation of the mixed culture with cells of acenaphthene-grown strain Pseudomonas sp. strain A2279 mitigated the accumulation of partial transformation products and resulted in more complete degradation of acenaphthene. This study demonstrates the value of the stable isotope labeling approach and its ability to reveal incomplete mineralization even when as little as 2 to 3% of the substrate is incompletely oxidized, yielding products of partial transformation. The approach outlined may prove useful in assessing bioremediation performance.

Microbial degradation of polycyclic aromatic compounds (PACs) is increasingly being considered for bioremediation applications and has been proposed as an attractive approach to remediation technologies dealing with fossil fuel wastes (10, 11). However, biodegradation of PACs of creosote and of related hydrocarbons of petroleum by defined and undefined mixed bacterial cultures may result in elevated rates of formation and even accumulation of organic end products (3, 5, 20). Some of these products are toxic to certain test organisms (1, 3, 20). Such information may be relevant in determining, for example, why little change in toxicity is observed when creosote undergoes biodegradation in groundwater (11). Therefore, chemical and toxicological characterization of any biodegradation process is required before the method is applied.

A realistic assessment of biodegradation efficacy requires delineation of a pollutant’s fate and effects beyond its depletion, as revealed by sensitive and exact analytical methods. For complex mixtures of chemicals, such as those found in coal- and petroleum-derived materials, this is not so readily accomplished. Assessment of efficacy cannot be based solely on rates of mineralization inferred from data obtained by trapping CO₂ released after the addition of a radioactively labeled tracer compound. Although this method is a convenient laboratory indicator of limited aspects of mineralization, it fails to provide information about the presence, nature, and distribution of organic end products or their interactions with soil and sedimentary matter.

¹³C-nuclear magnetic resonance (NMR) spectroscopy, combined with ¹³C labeling, however, offers an approach whereby details of the chemical structures of individual components of PAC mixtures undergoing biodegradation can be investigated. With the ¹³C-labeled tracers available, ¹³C-NMR has been successfully applied to an investigation of the interaction of 2,4-dichlorophenol with humic matrixes (8) and to a study of the alteration of jet fuels undergoing thermal stress (9). For a more complete study of its utility, various specifically ¹³C-labeled constituents of fossil fuels are required for evaluation of this technique in assessing the biodegradation of coal- and petroleum-derived wastes.

The applicability of this approach to the biodegradation of hydrocarbon mixtures is examined in the present study by using synthetic [1-¹³C]acenaphthene (99% ¹³C) introduced as a tracer compound into creosote PAC mixtures degraded by different undefined, mixed, and axenic bacterial cultures. The initial choice of acenaphthene labeled with ¹³C at a benzylic carbon as a tracer is based on the abundance of acenaphthene in creosote and the straightforward route of its synthesis. It also represents an extension of published work on acenaphthene oxidation catalyzed by naphthalene dioxygenase (16), biotransformation reactions catalyzed by bacteria and fungi (12, 14), and catabolism of acenaphthene by certain soil bacteria (19), with the consequent availability of suitable acenaphthene-utilizing microbial cultures.

(Preliminary accounts of aspects of this work have appeared elsewhere [2, 17, 18].)

MATERIALS AND METHODS

Chemicals.

PACs were purified from creosote P2 (Aristech Chemical Corp., Clairton, Pa.) by a modification of a previously described procedure (6, 13). Characterization of the composition of the PAC fraction and quantitative analysis of the principal PAC components, using as a reference a standard coal tar material, SRM1597 (National Institute of Standards and Technology, Gaithersburg, Md.) (21), were reported previously (6). Authentic acenaphthenone and cis- and trans-acenaphthene-1,2-diols were available from a previous study (16). All the commercial chemicals were of the highest purity available.

[1-¹³C]Acenaphthene was obtained by a four-step synthesis K¹³CN (670 mg, 99% ¹³C; Aldrich Chemical Co., Milwaukee, Wis.) was added to a stirred solution of 2.1 g of 1-(chloromethyl)naphthalene in 20 ml of aqueous ethanol (70%, vol/vol) to give 1′-naphthyl-[1-¹³C]acetonitrile (not isolated). After the mixture was refluxed for 1 h, 20 ml of a 30% (wt/vol) solution of NaOH in water was added. Refluxing was continued until evolution of ammonia ceased (approximately 6 h). The reaction mixture was cooled and washed three times with 50 ml of diethyl ether. The aqueous layer was separated and acidified to pH 2.0 to 2.5 by the addition of concentrated HCl. Precipitated 1′-naphthyl-[1-¹³C]acetic acid was extracted into ethyl acetate and recrystallized from diethyl ether (1.62 g, 92% yield, based on K¹³CN). Intramolecular acylation of 1′-naphthyl-[1-¹³C]acetic acid (0.8 g) was performed by addition of 30 ml of hot 85% polyphosphoric acid (110 to 130°C) with vigorous stirring for 4 to 5 min. The reaction was stopped by the rapid addition of ice-water (50 ml). The solution was adjusted to pH 8 to 9 by the addition of NaOH, and crude [1-¹³C]acenaphthenone was extracted into diethyl ether. [1-¹³C]acenaphthenone was purified by sublimation to remove colored dimeric products. When performed repeatedly, the cyclization reaction gave [1-¹³C]acenaphthenone with an overall yield in the range of 42 to 63%. Reaction yields were strongly dependent on the temperature of the polyphosphoric acid, the incubation time, and the stirring regimen.

[1-¹³C]Acenaphthenone was converted to [1-¹³C]acenaphthene by Clemmensen reduction. [1-¹³C]acenaphthenone (400 mg) was added to 30 ml of 10% aqueous HCl mixed with 0.05% (vol/vol) toluene and 3 g of amalgamated zinc wool. The mixture was refluxed for 4 h. [1-¹³C]acenaphthene was recovered from the reaction mixture by extraction into n-hexane and was purified by sublimation to yield 337 mg (92% yield). The structures of the product and precursors were confirmed by mass spectrometry and NMR spectroscopy; the spectra were compared to those of authentic unlabeled materials. The sublimed [1-¹³C]acenaphthene (99% ¹³C-1) was of 98% purity, as determined by gas chromatography (GC), with a single impurity identified as [1-¹³C]acenaphthylene (2%).

Microorganisms.

An undefined mixed culture of bacteria (CREOMIX) was obtained from creosote-contaminated soil from the American Creosote Works site in Pensacola, Fla. (3). The creosote PAC fraction, as the sole carbon and energy source in a mineral salts medium (7), was used to establish the enrichment culture. The culture was maintained at 20 to 25°C and transferred biweekly. This culture has been previously shown to completely deplete all constituents with three or fewer rings in the PAC mixture within 14 days of incubation. No significant degradation of compounds with four or more rings was observed in this culture, and no further depletion of PACs occurred during periods of incubation extending beyond 2 weeks. Complex mixtures of low-molecular-weight aromatic compounds were accumulated by this culture as end products of the PAC degradation (3). The CREOMIX culture was maintained for more than 3 years without loss of performance before it was used in this study.

Two individual strains, BR and BC, were isolated from the CREOMIX culture and tentatively identified as Pseudomonas spp. Both of these strains grew with either naphthalene or phenanthrene, but not on acenaphthene, as the sole carbon source and were used in this study to determine their contribution in the accumulation of biodegradation products from acenaphthene.

Attempts to isolate from the CREOMIX culture any individual strains of bacteria able to utilize acenaphthene as the sole carbon and energy source were not successful. The acenaphthene-catabolizing Pseudomonas sp. strain A2279 used in this study was obtained from the same creosote-contaminated soil by enrichments with acenaphthene as the sole carbon and energy source. This organism also grows with acenaphthylene or naphthalene as the sole carbon and energy source. Acenaphthene and acenaphthylene are degraded by this organism via naphthalene-1,8-dicarboxylic acid and 2-hydroxybenzene-1,3-dicarboxylic acid (15).

Biodegradation experiments.

All biodegradation experiments involving the use of [¹³C]acenaphthene and growth of all inoculum cultures were performed in 125-ml Erlenmeyer flasks with 25 ml of mineral salts medium (7) and with the pH adjusted to 7.2. Aliquots of stock solutions of the test compounds in methylene chloride were added to empty sterile flasks. Sterilized medium was added to the flasks after evaporation of the solvent (usually after 2 to 4 h). Uninoculated flasks were used as controls for extraction efficiency, losses by volatilization, and abiotic degradation.

The individual naphthalene-degrading strains BR and BC were grown in 25 ml of the liquid mineral salt medium with 1 g of naphthalene per liter until the late exponential phase (approximately 36 h). The acenaphthene-degrading strain A2279 was grown similarly with 1 g of acenaphthene per liter. Aliquots (1 ml) of the cultures were used to inoculate a series of replicate flasks, which contained either (i) a mixture of 10 mg of [1-¹³C]acenaphthene (previously diluted with unlabeled acenaphthene to give 20% ¹³C) and 10 mg of naphthalene or (ii) a mixture of 25 mg of creosote PACs and 1 mg of [¹³C]acenaphthene (99% ¹³C).

To examine the action of the undefined CREOMIX mixed culture on acenaphthene, the culture was first grown on 1 g of creosote PACs per liter for 2 weeks. Aliquots (1 ml) of this culture were then used to inoculate replicate flasks containing a mixture of 25 mg of creosote PACs and 1 mg of [¹³C]acenaphthene (99% ¹³C). A separate series of replicate flasks coinoculated with 1 ml of the CREOMIX culture and 1 ml of an acenaphthene-grown culture of strain A2279 was used to elucidate the effects of the latter organisms.

The flasks were incubated at 25°C on a rotary shaker with shaking at 200 rpm in the dark. In time course experiments, flasks were collected and their contents were acidified to pH 2 to 3 by the addition of 5 N HCl. The entire flask contents were extracted three times with 10 ml of methylene chloride. The extracts were dried over anhydrous sodium sulfate and concentrated under a stream of N₂ to 25 ml in Kuderna-Danish evaporative tubes without application of heat or reduced pressure. Aliquots (1 ml) were taken for GC analysis with flame ionization detection (FID). The remainder of each extract was evaporated at reduced pressure and redissolved in 0.7 ml of deuterochloroform for ¹³C-NMR analysis.

Analytical methods.

Capillary GC-FID analysis of the PACs remaining in the cultures and GC-mass-spectrometry analysis of metabolites were conducted as described previously (5, 6).

Broad-band-decoupled ¹³C-NMR spectroscopy experiments were carried out with a GE-QE 300Plus or a Nicolet NT-300-WB NMR spectrometer at a resonance frequency of 75.61 MHz. The ¹³C-NMR spectra in experiments with labeled acenaphthene were acquired in 2,400 scans, using a 31° pulse width and a 1-s recycle delay time. ¹H- and ¹³C-NMR spectra for the identification of synthetic materials and comparison with those of authentic compounds were recorded on the same spectrometers. Deuterochloroform was used as the solvent. Tetramethylsilane (0.00 ppm) and the central signal of the CDCl₃ (77.0) ppm were used as reference signals.

RESULTS AND DISCUSSION

Pure-culture incubations with naphthalene.

Experiments on degradation of [1-¹³C]acenaphthene in the presence of naphthalene by the individual naphthalene- and phenanthrene-degrading strains BR or BC were used to establish the experimental and analytical parameters needed to carry out ¹³C-NMR spectroscopy studies under more diverse chemical and microbiological scenarios. Although neither strain BR nor BC could grow on acenaphthene as the sole carbon and energy source, acenaphthene was oxidized by these strains when naphthalene was added as a cosubstrate. In the presence of naphthalene, [1-¹³C]acenaphthene was oxidized by strains BR and BC, with naphthalene no longer detectable after 24 h. Only partial (30 to 40%) depletion of acenaphthene occurred during the first 5 days of incubation. No further degradation of acenaphthene was observed in longer incubations.

¹³C-NMR spectroscopy analyses showed that oxidation of [1-¹³C]acenaphthene under these conditions was accompanied by accumulation of several extractable ¹³C-labeled biotransformation products, as evidenced by the appearance of new resonances (Fig. 1; Table 1). Based on chemical shift data and comparison with ¹³C-NMR data for authentic unlabeled compounds, these resonances were assigned to the acenaphthene oxidation products marked B through G in Fig. 2. Resonance B was tentatively assigned to [2-¹³C]acenaphthen-1-ol or [2-¹³C]acenaphthen-1-one, both of which had δ_C-2 resonance frequencies at approximately 42.1 ppm. Resonances E (δ_C-1 = 74.4 ppm) and D (δ_C-1 = 73.2 ppm) were assigned to [1-¹³C]acenaphthen-1-ol and [1-¹³C]acenaphthen-1,2-diol, respectively. The stereochemistry of [1-¹³C]acenaphthen-1,2-diol could not be unequivocally assigned since the authentic cis- and trans-diols produce nearly coincidental signals for each of the sec-alcohol carbon atoms. Resonance F (169.4 ppm) was indicative of a compound with a labeled carboxyl group and was assigned to the anhydride of [1-¹³C]naphthalene-1,8-dicarboxylic acid (formed from the free acid under the acidic conditions of extraction). Resonance G (δ_C-1 = 203.0 ppm) was due to the ¹³C-labeled carbonyl carbon in [1-¹³C]acenaphthen-1-one. As indicated by the intense resonance A, [1-¹³C]acenaphthene (δ_C-1 = 30.3 ppm) was still present in the cultures. With respect to the chemical nature of the compound responsible for resonance C (δ = 51.1 ppm), a definitive assignment cannot be made at present. None of the known acenaphthene metabolites identified as products of initial reactions catalyzed by bacteria or fungi (12, 14, 16, 19) can account for a ¹³C chemical shift value of a carbon atom derived from the labeled benzylic atom of acenaphthene. One possible explanation for resonance C is that it is from an unidentified compound formed by bacterial oxidation, and possibly cleavage, of one of the aromatic rings of [2-¹³C]acenaphthen-1-one. A chemical shift of 51.1 ppm could be due to a ¹³C-labeled sp³ carbon located between a carbonyl group and an aromatic ring or between two carbonyl groups of a β-diketone.

FIG. 1.

Representative ¹³C-NMR spectra of extractable compounds obtained in biodegradation experiments with Pseudomonas sp. strain BR after 2 days of incubation of [1-¹³C]acenaphthene (20% 1-¹³C, 10 mg/25 ml) and naphthalene (10 mg/25 ml) (A) and after 7 days of incubation of creosote PACs (25 mg) spiked with 1 mg of [1-¹³C]acenaphthene (B). For assignment of resonances A to G, see Fig. 2. T, tetramethylsilane; S, solvent (CDCl₃); N, signals due to the natural abundance of aromatic [¹³C]carbon atoms in acenaphthene and creosote PACs.

TABLE 1.

Broad-band-decoupled ¹³C-NMR analysis of [1-¹³C]acenaphthene and extractable labeled metabolites formed by the aromatic hydrocarbon-degrading microorganisms in time course biodegradation experiments

Microorganism	Carbon sources	Incubation time (days)	% Abundance of observed label-related 13C-NMR signals (δ, ppm)a in:
			A (30.3)	B (42.1)	C (51.1)	D (73.2)	E (74.4)	F (169.4)	G (203.0)
Pseudomonas sp. strain BC	[13C]Acn + Nahb	0	142	—c	—	—	—	—	—
		2	104	14.5	4.6	—	15.8	7.0	3.8
		5	95	29.4	7.0	—	9.4	5.8	12.5
Pseudomonas sp. strain BR	[13C]Acn + Nah	0	142	—	—	—	—	—	—
		2	107	12.0	—	22.0	12.3	7.3	3.5
		5	92	10.7	—	7.1	4.5	9.5	2.9
Pseudomonas sp. strain BC	[13C]Acn + PACs	0	282	—	—	—	—	—	—
		7	127	5.9	4.8	6.0	—	4.2	3.1
		14	110	5.2	2.6	5.1	—	5.6	2.6
Pseudomonas sp. strain BR	[13C]Acn + PACs	0	282	—	—	—	—	—	—
		7	163	9.1	6.0	—	—	6.2	3.6
		14	136	16.0	2.6	—	—	8.1	4.2
CREOMIX	[13C]Acn + PACs	0	276	—	—	—	—	—	—
		3	196	14.0	15.1	—	—	9.2	6.0
		7	172	7.2	10.2	—	—	7.0	3.0
		14	76	15.1	6.2	—	4.9	10.5	5.5
Pseudomonas sp. strain A2279	[13C]Acn + PACs	0	276	—	—	—	—	—	—
		3	264	—	—	—	—	—	—
		7	84	—	—	—	—	—	—
		14	—	—	—	—	—	—	—
CREOMIX + Pseudomonas sp. strain A2279	[13C]Acn + PACs	0	276	—	—	—	—	—	—
		3	97	—	—	—	—	—	—
		7	11	—	—	—	—	—	—
		14	—	—	—	—	—	—	—

^aRelative peak height (intensity of CDCl₃ signal at 77.0 ppm was used as 100%). 

^bA to G are identified in Fig. 2. Acn, acenaphthene; Nah, naphthalene. 

^c—, below detection limit (<2% of the CDCl₃ signal at 77.0 ppm). 

FIG. 2.

Assignment of ¹³C-NMR signals to detected metabolites of the pathway for acenaphthene degradation by CREOMIX culture and individual strains BR and BC. •, ¹³C-labeled carbons.

Recently, a key enzyme of bacterial naphthalene catabolism, naphthalene 1,2-dioxygenase, has been shown to catalyze monooxygenation of benzylic methylenic groups of acenaphthene. A series of acenaphthene oxygenation products are formed by strains that express genes encoding this enzyme (16). These products include acenaphthen-1-ol, acenaphthenone, both cis- and trans-acenaphthene-1,2-diols, and naphthalene-1,8-dicarboxylic acid (recovered as its anhydride). The same range of products formed from [¹³C]acenaphthene by naphthalene-grown cells of strains BR and BC (Fig. 2) indicates that these labeled metabolites are likely to be formed in reactions initiated via monooxygenation of acenaphthene by the naphthalene dioxygenases of these strains.

When acenaphthene-grown cells of strain A2279 were incubated with [1-¹³C]acenaphthene in the presence of naphthalene, GC analysis revealed that both hydrocarbons were completely depleted within 24 h. None of the metabolites formed by strains BC and BR was detected by ¹³C-NMR spectroscopy or GC-mass spectrometry (data not shown). Evidently, enzymes specific for acenaphthene catabolism convert this substrate to central cell metabolites without product accumulation.

Pure-culture incubations with creosote PACs.

Individual bacterial strains BC and BR, in the presence of creosote PACs, produced essentially the same set of oxidation products from [1-¹³C]acenaphthene as those observed in experiments with [1-¹³C]acenaphthene in the presence of naphthalene. The relative amounts of products were different, however (Fig. 1B; Table 1). Despite the complexity of the substrate mixture introduced by using creosote PACs, the biodegradation reactions performed by these strains led to the accumulation of the same “dead-end” oxidation products. Formation of the same oxidation end products in this series and in the experiments described above implies that limited oxidation of acenaphthene by naphthalene dioxygenase, or a closely related oxygenase system, may also occur when acenaphthene is biodegraded in the presence of PAC mixtures such as are encountered in creosote. Substantial amounts of acenaphthene (40 to 50% of the initial levels) remained even after 2 weeks of incubation, indicating the relatively inefficient oxidation of this compound under these conditions.

The action of the individual acenaphthene-catabolizing strain A2279 on [1-¹³C]acenaphthene in the presence of creosote PACs was also examined. Although the ability of this organism to degrade PACs other than acenaphthene and naphthalenes is limited (Table 2), no accumulation of ¹³C-labeled acenaphthene biodegradation products was detected by NMR spectroscopy analysis during growth on creosote PACs supplemented with [1-¹³C]acenaphthene (Table 1). As discussed previously (3), the limited microbial diversity of cultures used in degradation of hydrocarbon mixtures may be a factor leading to elevated levels of biodegradation end products. For example, biodegradation of an artificially weathered oil by defined cocultures of microbial isolates resulted in the accumulation of larger amounts of organic acidic and neutral products than the amounts accumulated by more diverse undefined mixed cultures established by an enrichment on oil (3).

TABLE 2.

Concentration of selected PACs during biodegradation experiments with creosote PACs spiked with [1-¹³C]acenaphthene

Compound	Recovery of PACs (μg/ml)a from different cultures grown in mineral medium with the PACs
	Undefined mixed culture (CREOMIX)				Pseudomonas sp. strain A2279				CREOMIX + Pseudomonas sp. strain A2279				Control (day 14)
	Day 0	Day 3	Day 7	Day 14	Day 0	Day 3	Day 7	Day 14	Day 0	Day 3	Day 7	Day 14
Naphthalene	70.2	—c	—	—	66.0	5.0	—	—	73.5	—	—	—	9.1
2-Methylnaphthalene	45.1	5.3	—	—	44.4	29.3	1.1	—	46.5	5.4	—	—	25.3
1-Methylnapthalene	21.3	5.4	—	—	21.0	13.8	1.0	—	21.9	3.6	—	—	13.0
Biophenyl	11.7	3.7	—	—	11.6	10.2	7.2	4.7	12.0	3.9	—	—	9.9
2,6-Dimethylnaphthalene	8.6	5.6	—	—	8.5	7.7	5.5	3.7	8.8	5.1	—	—	7.4
2,3-Dimethylnaphthalene	2.8	1.9	—	—	2.8	2.6	1.4	—	2.8	1.7	—	—	2.5
Acenaphthylene	7.0	4.4	1.8	1.4	6.6	6.1	1.0	—	6.9	2.6	1.0	—	6.5
Acenaphtheneb	109.5	82.8	45.9	22.8	109.9	99.3	32.8	1.1	112.2	44.6	1.8	—	104.7
Dibenzofuran	45.0	16.9	1.9	1.2	45.2	43.2	24.1	14.3	46.5	14.8	1.6	—	43.5
Fluorene	44.5	27.7	1.6	1.2	45.0	44.3	31.8	30.0	46.3	25.5	1.2	—	39.0
Dibenzothiophene	12.7	9.5	1.6	1.3	12.8	12.9	11.9	9.2	13.0	8.7	1.8	—	13.5
Phenanthrene	138.6	95.0	1.8	1.4	140.0	141.2	127.7	114.1	142.1	90.2	2.1	1.3	136.9
Anthracene	15.1	11.9	—	—	14.9	15.1	13.6	12.4	15.5	11.7	—	—	15.2
2-Methylanthracened	3.4	3.2	11.7	8.5	3.3	3.4	3.3	2.9	3.4	3.3	5.7	3.7	3.5
2-Phenylnaphthalene	8.5	6.8	—	—	8.5	8.6	8.1	7.6	8.7	6.5	4.6	1.7	7.8
Fluoranthene	90.9	91.9	82.6	76.6	91.5	92.6	88.9	87.8	92.7	92.2	93.1	70.3	90.7
Pyrene	69.0	70.1	66.0	59.4	69.5	70.3	68.0	68.4	71.4	70.2	70.9	56.7	72.0
Benzo[b]fluorene	11.6	12.0	11.2	9.8	12.1	12.7	12.2	11.4	11.8	12.1	12.6	8.2	14.1
Benzo[a]anthracene	13.3	15.2	14.6	13.1	13.6	14.0	16.1	15.0	13.7	14.3	16.4	11.2	14.9
Chrysene + triphenylene	11.1	12.3	5.5	5.1	11.0	11.6	5.6	5.2	11.5	11.8	6.3	4.9	10.0
Benzo[b]fluoranthened	4.5	6.7	8.0	7.7	4.7	4.9	8.2	7.6	4.5	5.5	9.5	7.6	5.0
Benzo[a]pyrened	1.8	3.0	3.6	3.4	1.8	1.9	3.8	3.5	1.8	2.2	4.4	3.5	2.0

^aQuantified by GC-FID analysis. Data reported are means of duplicate samples. 

^bIncludes both [1-¹²C]- and [1-¹³C]acenaphthene. The concentration of acenaphthene due to P-2 creosote PACs was determined to be 75.6 μg/ml. [1-¹³C]acenaphthene was added at 40.3 μg per ml of culture. 

^c—, Below the detection limit of 100 ppb. 

^dThe apparent increase in the concentration of certain compounds is due to accumulation of coeluting biodegradation products (6). 

Undefined mixed-culture incubations with creosote PACs.

A series of experiments with the undefined mixed bacterial culture CREOMIX was undertaken to establish whether the formation of acenaphthene oxidation products is due to the limited catabolic versatility of the individual strains used or is observable when PACs are degraded by more complex cultures. Biodegradation of creosote PACs, spiked with [1-¹³C]acenaphthene, by the CREOMIX culture resulted in degradation of all aromatic compounds with three or fewer rings (Table 2). Significant volatility of naphthalene, monomethylnaphthalenes, and biphenyl had also contributed to the observed depletion of these three compounds, as evidenced by the data shown for the 14-day control incubation. However, effective biodegradation of these compounds by the CREOMIX culture was evident, since their levels were reduced severalfold within first 3 days and completely depleted by day 7 of incubation. Only limited degradation of two PACs with four rings, fluoranthene and pyrene, was observed. ¹³C-NMR spectroscopy analysis of the total extractable compounds showed the presence of resonances corresponding to several of the acenaphthene oxidation products previously detected in experiments with the individual strains BR and BC (Table 1; Fig. 3A to C). The ¹³C-NMR spectra showed the presence of [1-¹³C]- and [2-¹³C]acenaphthen-1-ones, naphthalene 1,8-dicarboxylic acid anhydride, and the same unidentified product described above with a labeled methylenic group (resonance C at 51.1 ppm). In addition to these products, traces of carboxyl-labeled carboxylic acids (minor resonances with δ around 170 ppm) and of a product with a labeled methylenic group (δ ∼ 41.7 ppm) were observed after 14 days of incubation (Fig. 3C).

FIG. 3.

Representative ¹³C-NMR spectra (expanded baseline region) of extractable compounds obtained in biodegradation experiments with creosote PACs (25 mg) spiked with 1 mg of [1-¹³C]acenaphthene. (A) Day zero; (B) CREOMIX culture, 3 days of incubation; (C) CREOMIX culture, 14 days of incubation; (D) CREOMIX culture plus Pseudomonas sp. strain A2279, 3 days of incubation; (E) CREOMIX culture plus Pseudomonas sp. strain A2279, 14 days of incubation. For assignment of the signals, see Fig. 2 and the legend to Fig. 1.

The range of accumulated ¹³C-labeled products in experiments with the CREOMIX culture indicates that biodegradation of acenaphthene in a creosote PAC mixture also occurs via oxidation of benzylic methylenic groups but is not limited to these reactions. Additional reactions responsible for the formation of more extensively oxidized metabolites, such as aromatic ring oxidation products, that were not observed in experiments with individual strains may have occurred due to the action of other strains present in the CREOMIX culture. However, even in these biodegradation experiments with mixed cultures presumed to provide an extended biochemical diversity, acenaphthene biodegradation was not complete after 14 days of incubation. Because acenaphthene-utilizing strains could not be isolated from the CREOMIX culture by subculturing with acenaphthene as the sole carbon source (see Materials and Methods), this compound is evidently degraded to only a limited extent via biotransformations initiated by naphthalene dioxygenase (or phenanthrene dioxygenase) rather than by a more complete growth-based process.

Coinoculation of the acenaphthene-grown strain A2279 with the CREOMIX culture resulted in accelerated biodegradation of creosote PACs supplemented with [1-¹³C]acenaphthene. Nearly complete depletion of all aromatic compounds with three or fewer rings was achieved within the first 7 days of incubation (Table 2). In contrast to the experiments where the CREOMIX culture was acting alone (Fig. 3B and C), resonance A, due to the ¹³C-labeled methylenic group of acenaphthene, was no longer detectable by ¹³C-NMR spectroscopy after 14 days of PAC biodegradation by a coculture of CREOMIX and strain A2279 (Table 1; Fig. 3E). At all sampling times, the ¹³C-NMR spectra of the compounds extracted from the coinoculated cultures revealed no distinct resonances due to the labeled acenaphthene metabolites described above and therefore demonstrated that these products do not accumulate under this biodegradation scenario (Table 1; Fig. 3D and E).

The results obtained demonstrate that mixed bacterial cultures, established in the laboratory by enrichments on creosote PACs, have a restricted biochemical diversity with respect to furnishing an efficient pathway for acenaphthene degradation and catabolism. It would appear that the enrichment technique with creosote PACs used here does not provide favorable conditions for establishing bacterial populations which contain bacterial strains capable of mineralizing acenaphthene. Acenaphthene-utilizing strains, such as A2279 and the Alcaligenes strains reported previously (19), could readily be isolated from soils contaminated with coal-derived products by enrichments with acenaphthene. Although biochemical pathways for acenaphthene catabolism have not been established in detail, they are clearly distinct from those of naphthalene and phenanthrene utilization and require enzyme systems for the conversion of key intermediates such as naphthalene-1,8-dicarboxylic acid. It is possible that the presence of large quantities of naphthalene and phenanthrene in creosote PACs provides more favorable conditions for the selection and dominance of bacterial strains that effectively catabolize these constituents, rather than the less abundant acenaphthene. This, however, does not completely explain why acenaphthene-utilizing strains are not evident in the CREOMIX culture, since acenaphthene is not completely removed even after long incubation times. Regardless of the explanation, this study demonstrates that the variety of substrates present in complex mixtures of fossil fuel-derived materials demands a biochemically diverse and versatile microbial flora for their extensive biodegradation.

Another set of conclusions can also be drawn from comparison of the ¹³C-NMR (Table 1) data with those obtained by GC-FID analysis (Table 2), a method more routinely used to quantify the depletion of key PAC analytes. Although no special efforts were made at this point to acquire quantitative NMR data on the compounds of interest, the decrease in the area of the resonance due to the ¹³C-labeled carbon of [1-¹³C]acenaphthene (resonance A) is correlated with acenaphthene depletion as measured by GC-FID, with an error not exceeding 10%. At the same time, however, changes in concentrations of accumulated metabolites together with structural information could also be shown by NMR spectroscopy, while a typical GC-FID protocol is not satisfactory for this purpose. It is also important to note that, based on the semiquantitative estimates of this study, as little as 2 to 3% of the introduced ¹³C label was readily detected in intermediates or biodegradation end products by ¹³C-NMR spectroscopy. This observation is significant because it points out that very little ¹³C-labeled material is needed to obtain useful structural information.

The results of this study demonstrate the utility of ¹³C labeling in combination with standard NMR spectroscopy experiments for studies of the biodegradation of complex mixtures of compounds, such as coal-derived wastes. The approach allows for direct analysis of stable-isotope-labeled products and, in this case, an assessment of the course of the biodegradation process. NMR spectroscopy is a recognized structural analytical tool, and when it is used in combination with site-specific ¹³C labeling, the enhanced sensitivity provided by the labeled carbon(s) serves as a means to track the fate of a chosen substrate and the chemical changes which occur at, or in the vicinity of, the label. Further successful application of the method to the biodegradation of a diverse spectrum of fossil-derived materials depends on the availability of relevant ¹³C tracer compounds. When chosen in accordance with their abundance in the mixtures of interest and labeled in positions which are relevant to available information on their biodegradation pathways, such labeled compounds can serve as a diversified collection of specific tracers for a variety of fossil fuel materials. The use of this collection, in conjunction with NMR spectroscopy and other sensitive methods such as isotope ratio MS, can provide an invaluable tool for the identification of the biochemical limitations of biodegradation processes, for the evaluation of bioremediation performance, and for the analysis of pollutant and metabolite fate in complex organic matrixes such as soils and sediments.