Efficacy of Bacterial Bioremediation: Demonstration of Complete

Chemistry, Florida State University, 1800 East Paul Dirac. Drive, Tallahassee, Florida 32310. We present a method and example to establish complete...
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Environ. Sci. Technol. 2000, 34, 535-540

Efficacy of Bacterial Bioremediation: Demonstration of Complete Incorporation of Hydrocarbons into Membrane Phospholipids from Rhodococcus Hydrocarbon Degrading Bacteria by Electrospray Ionization Fourier Transform Ion Cyclotron Resonance Mass Spectrometry RYAN P. RODGERS,‡ ERIN N. BLUMER,† MARK R. EMMETT,† AND A L A N G . M A R S H A L L * ,‡ National High Magnetic Field Laboratory and Department of Chemistry, Florida State University, 1800 East Paul Dirac Drive, Tallahassee, Florida 32310

We present a method and example to establish complete incorporation of hydrocarbons into membrane phospholipids (and their constituent individual fatty acids and polar headgroup) of putatively bioremediative bacteria. Bacteria are grown on minimal media containing a specified carbon source (in this case, C16 and C18 alkanes), either natural abundance (99% 12C) or enriched (99% 13C). After extraction (but no other prior separation) of the membrane lipids, electrospray ionization yields a negative-ion FTICR mass spectrum containing prominent phospholipid parent ions. If 13C-enriched hydrocarbon incorporation is complete, then the mass of the parent ion will increase by n Da, in which n is the number of its constituent carbon atoms; moreover, the 13C isotopic distribution pattern will be reversed. The identities of the constituent fatty acids and polar headgroup are obtained by collisional dissociation (MS/ MS), and their extent of 13C incorporation determined individually. The method is demonstrated for Rhodococcus rhodochrous (ATCC# 53968), for which all 44 carbons of a representative phosphatidylinositol are shown to derive from the hydrocarbon source. Interestingly, although only C16 and C18 alkanes are provided in the growth medium, the bacteria synthesize uniformly enriched C16:0 and C19:0 fatty acids.

Introduction The identification of the fatty acid and polar headgroup constituents of phospholipids synthesized by bacteria, as a function of food source, is of fundamental interest in its own right. In addition, the proposed bioremediation by bacteria of hydrocarbons spilled or leaked into the environment has * Corresponding author phone: (850)644-0529; fax: (850)644-1366; e-mail: [email protected]. † National High Magnetic Field Laboratory. ‡Department of Chemistry. 10.1021/es990889n CCC: $19.00 Published on Web 12/30/1999

 2000 American Chemical Society

created a very important practical interest in bacterial lipid metabolism. Specifically, bacteria could in principle (a) not metabolize hydrocarbons at all; (b) metabolize them incompletely; or (c) “mineralize” hydrocarbons, i.e., break them down completely for biosynthesis and/or respiration to CO2 and H2O. For maximal bioremediative value, it therefore becomes essential to show whether bioremediation candidate bacteria mineralize hydrocarbons. In this paper, we offer a simple, new, rapid method for determining the extent of mineralization of phospholipids (as well as their fatty acid and headgroup constituents) by a bacterial bioremediation candidate, Rhodococcus rhodochrous (ATCC# 53968, strain IGTS8). Leaking underground storage tanks, pipeline ruptures, tanker failures (223 million gallons of oil lost in and around U.S.A. waters from 1973 to 1993 alone), and various other production and transportation accidents produce large volumes of hydrocarbon-contaminated soil and groundwater each year (1-5). In addition, many other contaminated sites are produced from halogenated pesticides (6) and other industrial solvents. Remediation of these sites can be extremely costly. Since its development in the early 1970s by Richard Raymond (7), bioremediation by bacteria has been proposed as a relatively low-cost option (8). Candidate bacteria that could degrade hydrocarbon wastes in the environment are ubiquitous (9), and some may be able to consume hydrocarbons as their sole source of carbon (1019). The discovery of bacteria capable of degrading contaminant and the subsequent ability to accelerate/enhance bioremediation through the use of fertilizers (20-23) and surfactants (24-28) has led to the development of bioremediation as a potentially economically viable in situ remediation technique (29). Prior attempts to establish unequivocally the mineralization of hydrocarbon contaminants have typically relied on radioactive- or stable isotope-labeled growth substrates or on monitoring the ratio of naturally occurring carbon isotopes (3, 30-36). Mass spectrometry is well established as a structural analysis technique for biological membrane lipids (37, 38) and is commonly used for either high sensitivity isotope ratio (δ13C ratio and 14C measurements) (39-42) or low-resolution mass analysis (e.g., total petroleum hydrocarbon monitoring) (43). Incorporation of radiolabeled growth substrates may be measured by separation and detection of the cellular lipids. Lipids are attractive for this purpose because they are easily isolated from the total cellular biomass: simple organic extraction of the bacterial cultures returns high lipid yield for subsequent analysis. However, the complexity of the lipid extract typically requires prior separation by (e.g.) thin-layer chromatography to simplify and isolate the lipids of interest (phospholipids in the present case). In related work, mass spectrometric characterization of bacterial lipids has been used to identify and/or distinguish between various bacterial species (44-46). Fourier transform ion cyclotron resonance mass spectrometry (FT-ICR MS) (47) offers high mass resolving power (>106), high mass accuracy (low ppm, for direct determination of elemental composition), and rapid analysis, making it highly attractive for complex mixture analysis. Over the past decade, electrospray ionization (ESI) mass spectrometry has developed principally for the analysis of proteins and peptides; however, several researchers have recently noted its applicability to lipid analysis (48-50). For example, Lehmann et al. demonstrated picomole-level analysis of biological membrane lipids, allowing the detection of individual classes of lipids in an unprocessed lipid extract VOL. 34, NO. 3, 2000 / ENVIRONMENTAL SCIENCE & TECHNOLOGY

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FIGURE 1. Simulated mass spectrum for an arbitrarily chosen phospholipid (36:2 phosphatidylinositol) isolated from bacteria grown on unenriched hydrocarbon media (top) and 99% 13C enriched media (bottom). Notice the predicted shift to higher mass (equivalent to the number of carbons in the phospholipid) and reversal of the isotopic pattern as a result of complete incorporation. An inset (top, right) shows the expected fragmentation pattern that results from collisionally induced dissociation of the parent lipid anion. (51). Samples were prepared from cultures through simple organic extraction prior to electrospray ionization with no prior chromatographic separation. Coupling ESI sensitivity and selectivity with the high mass spectral performance of FT-ICR MS allows for the analysis of complex biological mixtures at subnanomolar or even picomolar concentration (52). Here we grow Rhodococcus rhodochrous (chosen because it is capable of consuming n-alkanes and because it cleaves organic C-S bonds for our future study of bacterial biodesulfurization) from a minimal medium containing natural abundance (99% 12C) or enriched (99% 13C labeling at all carbons) hexadecane and octadecane substrates as the only carbon source. Lipids are extracted but not separated, and the extract is subjected to ESI negative-ion FT-ICR MS. If the bacteria completely metabolize the hydrocarbon, then the molecular ion of the enriched phospholipid containing n carbon atoms will be shifted n Da higher in mass, and the 13C isotopic distribution pattern will be reversed, for lipids from enriched compared to natural abundance growth media, as shown in Figure 1 for a simulated phosphatidylinositol mass spectrum. Further evidence is provided by isolating a given phospholipid parent negative ion and collisionally dissociating it. Figure 1 (top right) shows a representative phospholipid structure, with collisionally induced dissociation (CID) cleavage sites shown as dotted lines. CID of that phosphatidylinositol generates fragment ions corresponding to each of the two fatty acids, the inositol phosphate, and fragments corresponding to neutral loss of each of those units from the parent phospholipid. Thus, the CID mass spectrum identifies the individual fatty acids and polar headgroup (53). Moreover, as for the parent phospholipid, comparison of the fragment ions from 13C-enriched and natural abundance sources reveals the extent of carbon incorporation into the individual fatty acids and polar headgroup, based on mass shift and reversal of the 13C isotopic distribution.

Experimental Procedures Sample Preparation. Rhodococcus rhodochrous (ATCC# 53968) cultures (5 mL) in Ledbetter-Foster basal salts were grown at 28 °C for 48 h on both natural abundance and 99% 536

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FIGURE 2. Full-range microelectrospray negative-ion FT-ICR mass spectrum (bottom) of a raw bacterial lipid extract from Rhodococcus rhodochrous (ATCC# 53968) grown on natural isotopic abundance media. The SWIFT-isolated 851 Da isotopic distribution, prior to SORI fragmentation, is shown in the top right inset. enriched 13C 1:1 (vol/vol) mixtures of n-hexadecane and n-octadecane (because 13C-labeled n-alkanes were commercially available as a hexadecane/octadecane mixture). The cells were then isolated and subjected to a modified Bligh-Dyer lipid extraction (54). All solvents were HPLC grade and used as received. The extraction was initiated by the addition of 7.6 mL of 2:1:0.8 methanol:chloroform:water to produce a single-phase extraction system. After 3 h, 2 mL each of water and chloroform were added to bring the final volume ratio to 2:2:1.8 and disrupt the single-phase extraction system to a two-phase system. The two phases were allowed to separate for 24 h before the organic phase, containing the cellular neutral lipids, glycolipids, and phospholipids, was collected, and the solvents were removed with (blown to dryness) in a SpeedVac (Savant, Holbrook, NY). The dried lipid was stored at -20 °C; for analysis, the lipid was redissolved in 2:1 methanol:chloroform and vortexed for 1 min before negative ion microelectrospray into an FT-ICR mass spectrometer. Mass Analysis. All mass analyses were carried out on a home-built FT-ICR mass spectrometer (55) equipped with a 22 cm horizontal room-temperature bore 9.4 T magnet (Oxford Corp., Oxney Mead, England) and an Odyssey data station (ThermoQuest, Bremen, Germany)). Negative ions were generated from a microelectrospray source (52) at a flow rate of 300 nL/min and a source potential of ∼3 kV before external accumulation (5-15 s) in a 67 cm long octopole (56). Ions were then transferred through a 200 cm long rf-only octopole into a open cylindrical ICR cell (10 cm i.d. × 30 cm long) prior to broadband frequency chirp excitation (50 kHz-1.44 MHz at a sweep rate of 150 Hz/µs and amplitude, Vp-p ) 210 V) and direct mode image current detection to yield 1 Mword time-domain data. The data were processed by an in-house FT-ICR MS analysis package (57). The time-domain data was Hanning-apodized, followed by a single zero-fill before FFT and magnitude calculation. Frequency-to-mass conversion was from a formula based on assumed three-dimensional axial quadrupolar trapping potential (58), to generate the spectra shown in Figures 2-8. Collisionally induced dissociation (CID) was achieved by isolating the ions of interest by stored waveform inverse Fourier transform (SWIFT) radial dipolar mass-selective ejection (59, 60) followed by sustained off-resonance irradiation (SORI) (61, 62) fragmentation. The collision gas was argon, pulsed to 1 × 10-6 Torr 100 ms prior to a 500 ms single frequency SORI excitation 2 kHz below the base frequency of the most abundant peak in the SWIFT-isolated isotopic envelope. Fragmentation FT-ICR mass spectra were

FIGURE 3. Full-range SORI fragmentation FT-ICR mass spectrum (bottom) of the SWIFT-isolated unenriched parent ions shown in Figure 2 (top, right). Neutral losses of both 256 and 298 Da tentatively identify the fatty acids as C16:0 and C19:0. A subsequent neutral loss of 162 Da indicates that the headgroup is a hexose (in this case, inositol [Brugger, 1997 #42)). A mass scale-expanded segment (top) verifies both the fatty acids that comprise the lipid (C16:0 at 255 Da and a (branched (64, 65)) C19:0 fatty acid at 297 Da) and the inositol headgroup (inositol phosphate anion minus H2O at 241 Da and inositol phosphate anion minus two H2O at 223 Da).

FIGURE 4. Full-range FT-ICR mass spectrum (bottom) of the raw bacterial lipid extract from Rhodococcus rhodochrous (ATCC# 53968) grown on 13C enriched media resulting from negative-ion microelectrospray ionization. The SWIFT-isolated 895 Da isotopic distribution, prior to SORI fragmentation, is shown in the top right inset. produced from 20 coadded time-domain data sets and were processed as previously described.

Results and Discussion Composition of Lipids from Bacteria Grown in Natural Abundance 13C Media. A 100 µL aliquot of the raw lipid extract was microelectrosprayed into the ESI FT-ICR mass spectrometer at a rate of 300 nL/min to yield a rather complex negative-ion mass spectrum consisting of ∼650 peaks, representing ions of 300 < m/z < 1500 at a mass resolving power (m/∆m50% ≈ 30 000 (Figure 2), in which m is ion mass in Da, z is ion charge in multiples of the elementary charge, and ∆m50% is the full magnitude-mode peak width at halfmaximum peak height. The five most abundant ions were isolated and fragmented. Because all five yielded similar information about bioremediation, we shall describe only one. The upper inset in Figure 2 illustrates SWIFT isolation of singly charged ions of 851 Da at a S/N of ∼1000:1. After

FIGURE 5. Full-range SORI fragmentation FT-ICR mass spectrum (bottom) of the SWIFT-isolated 13C-enriched parent ions shown in Figure 4 (top, right). Again, neutral losses of both 272 and 317 Da tentatively identify the fatty acids as C16:0 and C19:0. A subsequent loss of 168 Da identifies the headgroup as a hexose (inositol). A mass scale-expanded segment (top) enables the assignment of both fatty acids that comprise the lipid (C16:0 at 271 m/z and a branched (64, 65) C19:0 fatty acid at 316 Da) and the inositol headgroup (inositol phosphate minus H2O at 247 Da and inositol phosphate minus two H2O at 230 Da). Note the increase in mass and reversal of the phospholipid isotopic distribution as a result of complete 13C incorporation.

FIGURE 6. Mass scale-expanded segments for both unenriched and 99% 13C enriched lipid species isolated in Figure 2 (top, right) and Figure 4 (top, right), respectively. Notice the 44 Da mass shift (equal to the total number of carbons in the phosphatidylinositol) and reversal of the isotopic pattern due to 13C incorporation. isolation, the parent ions were fragmented by SORI to yield structurally significant product ions (see Figure 3 (bottom)). Neutral losses from the parent ion of both 256 and 298 Da are characteristic of C16:0 and C19:0 fatty acids. Moreover, as expected from prior MS/MS examples (38, 53), the (C19) fatty acid on the central C-2 carbon of glycerol appears at higher relative abundance than the (C16) fatty acid at the C-3 glycerol carbon. Thus, MS/MS can determine which fatty acid is R1 and which is R2 in the schematic phospholipid shown in Figure 1. A subsequent neutral loss of 162 Da points to a hexose (C6H10O5) from either species, corresponds to elimination of a hexose substituent, and is consistent with the presence of an inositol phosphate group (51). A zoom mass inset (Figure 3, top) of the lower-mass fragments shows both C16:0 and C19:0 fatty acid anions (m/z 255 and 297) and hexose phosphate headgroup ions (m/z 223 and 241). The fatty acids and polar headgroup of the phospholipid are therefore identified unambiguously (except for positional and branching isomers) consistent with a (35:0) phosphatidylinositol parent ion (m/z 851). VOL. 34, NO. 3, 2000 / ENVIRONMENTAL SCIENCE & TECHNOLOGY

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FIGURE 7. A series of mass scale-expanded segments clearly shows a shift to higher mass (equal to the number of carbons in each fatty acid) and reversal of the isotopic envelope in each of the fatty acids that comprise the C35:0 phosphatidylinositol.

FIGURE 8. A series of mass scale-expanded segments shows a shift to higher mass (equal to the number of carbons in the characteristic inositol headgroup fragments) and reversal of the isotopic envelope for inositol phosphate minus one (left) or two (right) water molecules. The odd number of carbon atoms in the C19:0 fatty acid suggests the possibility of a branched hydrocarbon chain. In fact, previous experiments for similar experimental growth conditions, prior mass analysis, and permanganate-sulfuric acid oxidation (63) identified a C19:0 fatty acid as C18:10Me (i.e., a methyl substituent at carbon 10 (64, 65)). Although such a branch is likely the correct structure for our C19:0 fatty acid, it is not possible to assign the location and carbon length of the branched segment fatty acids with low-energy CID in our instrument, due to the high energy required to produce structurally significant alkane fragmentation products (66-70). However, our analysis of the SWIFT-isolated lipids was simply to determine the degree of incorporation and confirm that all structural lipid elements showed a similar percent 13C incorporation, rather than to determine the complete lipid structure. Therefore, no further experiments were conducted to determine the branching structure of the C19:0 fatty acid. Composition of Lipids from Bacteria Grown in 13CEnriched Media. Based on the number of carbons in the m/z 851 phosphatidylinositol determined through CID fragmentation, the magnitude of the mass shift upon uptake and metabolism of 99% 13C hexadecane and octadecane is predicted to be 44 Da. Thus, the chemically identical (35:0) phosphatidylinositol characterized in the unenriched lipid 538

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should shift to 851 + 44 ) 895 Da due to the incorporation of 44 13C heavy carbons. Negative-ion electrospray FT-ICR MS of the raw lipid extract from Rhodococcus rhodochrous grown from 13C enriched media was performed under identical conditions as for lipids isolated from the unenriched media. The broadband mass spectrum (Figure 4, bottom) reveals a lipid profile similar to that for natural abundant growth medium. The expected species observed at m/z 895 was SWIFT-isolated (Figure 4, top) and fragmented (same SORI conditions as for natural abundance lipid). Figure 5 (bottom) shows the broadband CID fragmentation spectra along with a zoom mass inset (top) that reveals the same fatty acid and headgroup composition as for natural abundance lipid. Specifically, neutral losses of 272 and 317 Da again indicate C16:0 and C19:0 fatty acids but now heavier (by 16 and 19 Da) due to 13C incorporation. Likewise, a subsequent loss of 168 Da characteristic of a hexose minus H2O was observed for both peaks. At lower m/z (Figure 5, top, zoom inset) C16:0 and C19:0 fatty acid ions are seen, along with two characteristic fragment ions of a hexose phosphate. Finally, Figure 6 shows SWIFT-isolated parent species from bacteria grown from both unenriched (top) and enriched (bottom) media. The shift to 44 Da higher mass (predicted from the CID fragmentation of the unenriched parent) and reversal of the carbon isotope distribution provide direct evidence for contaminant mineralization. The total shift of 44 Da is now seen to arise from one additional Da (13C vs 12C) from each of the carbons in the C16 and C19 fatty acids, a C6 hexose, and C3 from the glycerol backbone. Unambiguous evidence for mineralization is evident from closer inspection (Figure 7) of the zoom mass insets for the C16:0 (left) and C19:0 (right) fatty acids from both the unenriched (top) and enriched (bottom) CID fragmentation spectra. In both examples, we see a shift to higher mass equal to the number of carbons in the fatty acid (16 or 19) as well as reversal of the isotope pattern is observed. Further proof of contaminant mineralization is evident from inspection of characteristic headgroup CID spectra (Figure 8), exhibiting a similar shift to higher mass (by 6 Da, corresponding to six carbon atoms in the hexose) and reversal of the isotope pattern), for both the singly (m/z 241) and doubly (m/z 223) dehydrated inositol phosphate anions. In summary, observation of complete 13C incorporation into all carbon atoms of bacterial lipids isolated from bacteria whose only hydrocarbon food source was isotopically enriched to 99% 13C constitutes definitive evidence for bioremediation. By microelectrospray FT-ICR negative-ion mass spectrometry (and tandem mass spectrometry by use of collisionally induced dissociation (CID)) 13C incorporation has been proved in two ways for each of four species for each phospholipid: (a) increase in mass by the same number of Da as the number of carbons in the parent phospholipid ion, CID fragment ions corresponding to each of its collisionally induced fatty acid and hexose phosphate headgroup fragments, and CID fragment ions corresponding to neutral loss of each of those components from the parent phospholipid ion; and (b) reversal of the 13C isotopic distribution for enriched (99% 13C) vs natural-abundance (99% 13C) species. A second important advantage of the present approach, independent of bioincorporation, is that determining the number of carbon atoms in a parent or product ion (based on mass shift for all-13C vs all-12C species) makes it possible to determine the elemental composition unequivocally, even though our mass accuracy would not have been good enough to determine elemental composition directly from the natural-abundance mass. Thus, being able to compare all13C and all-12C versions of the same molecule effectively extend by a factor of ∼3 the upper mass limit below which elemental composition can be determined.

Specifically, mineralization of hexadecane and octadecane hydrocarbon contaminant by Rhodococcus rhodochrous (ATCC# 53968) was confirmed through the use of 13C-enriched hexadecane and octadecane growth substrates. Interestingly, although the food source was C16 and C18 hydrocarbons, we observed bacterial synthesis of a phospholipid with C16 and C19 fatty acids. The present method could obviously be used to test for mineralization of hydrocarbons of other lengths, with or without branching, with or without double bonds. The method is definitive, rapid, and simple, requiring only a simple extraction. We therefore propose that ESI FTICR MS and MS/MS provide an attractive method for rapid characterization of bacterial phospholipids (to the level of elemental composition of headgroup and fatty acid side chains), as a test for the extent of bioremediation of hydrocarbons (e.g., gasoline, diesel fuel, kerosene, etc.) by a candidate microorganism. In fact, we have already performed similar experiments with Rhodococcus rhodochrous (ATCC#19067), with results very similar to those obtained here. Finally, the present isotopic-labeling approach could also be pursued with other types of mass analyzers with collisioninduced fragmentation. In particular, high-energy collisioninduced dissociation is well suited for fragmenting fatty acid chains to locate double bonds and branches (66-70). However, FT-ICR mass resolving power is typically 10 times higher than that of the best double-focusing mass analyzers, and only FT-ICR offers ultrahigh resolving power in both MS1 and MS2, so that FT-ICR is best-suited to biological analyses involving complex mixtures of many components.

Acknowledgments The authors thank Daniel McIntosh for machining all of the custom parts required for the 9.4 T instrument construction. The authors also thank Christopher L. Hendrickson and John P. Quinn for many helpful discussions. This work was supported by NSF (CHE-94-13008), NIH (GM-31683), the American Chemical Society Division of Analytical Chemistry graduate Fellowship (R.P.R.), and the National High Magnetic Field Laboratory in Tallahassee, Florida.

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Received for review August 2, 1999. Revised manuscript received November 12, 1999. Accepted November 18, 1999. ES990889N