Abstract
We tested different alga-bacterium-archaeon consortia to investigate the production of oil-like mixtures, expecting that n-alkane-rich biofuels might be synthesized after pyrolysis. Thermosipho globiformans and Methanocaldococcus jannaschii were cocultured at 68°C with microalgae for 9 days under two anaerobic conditions, followed by pyrolysis at 300°C for 4 days. Arthrospira platensis (Cyanobacteria), Dunaliella tertiolecta (Chlorophyta), Emiliania huxleyi (Haptophyta), and Euglena gracilis (Euglenophyta) served as microalgal raw materials. D. tertiolecta, E. huxleyi, and E. gracilis cocultured with the bacterium and archaeon inhibited their growth and CH4 production. E. huxleyi had the strongest inhibitory effect. Biofuel generation was enhanced by reducing impurities containing alkanenitriles during pyrolysis. The composition and amounts of n-alkanes produced by pyrolysis were closely related to the lipid contents and composition of the microalgae. Pyrolysis of A. platensis and D. tertiolecta containing mainly phospholipids and glycolipids generated short-carbon-chain n-alkanes (n-tridecane to n-nonadecane) and considerable amounts of isoprenoids. E. gracilis also produced mainly short n-alkanes. In contrast, E. huxleyi containing long-chain (31 and 33 carbon atoms) alkenes and very long-chain (37 to 39 carbon atoms) alkenones, in addition to phospholipids and glycolipids, generated a high yield of n-alkanes of various lengths (n-tridecane to n-pentatriacontane). The gas chromatography-mass spectrometry (GC-MS) profiles of these n-alkanes were similar to those of native petroleum crude oils despite containing a considerable amount of n-hentriacontane. The ratio of phytane to n-octadecane was also similar to that of native crude oils.
INTRODUCTION
Crude petroleum oil reservoirs are generally located in deep geological formations with environments characterized by anaerobic, high-temperature, and high-pressure conditions that harbor various microorganisms (1, 2). Petroleum fluids are produced by biological modification and thermal cracking of fossil organic materials in source rocks during geologically long-term subsurface storage, and expelled fluids migrate toward traps that are eventually formed by reservoir and cap rocks (3). The major detectable hydrocarbons in crude oils are n-alkanes and related compounds (4). Understanding the activity of such microorganisms is essential to search for potential associations with crude oil generation.
We previously sampled mixtures of crude oils and production water directly from subsurface oil deposits to search for subsurface microorganism activity during oil generation. Analysis of 16S rRNA gene sequences led to the identification of a Thermacetogenium-like bacterium (92% identity in 16S rRNA gene sequence with Thermacetogenium phaeum) and a Methanothermobacter-like archaeon (96% identity with Methanothermobacter thermautotrophicus) as the major indigenous populations in one oil deposit at 74°C (5). Cocultured T. phaeum and M. thermautotrophicus syntrophically degraded acetate to CH4 at 55°C under anaerobic conditions in a methanogenic reactor for wastewater (6). Based on these findings, we hypothesized that syntrophic pairs of bacteria and archaea would be the major indigenous microorganisms in oil reservoirs and that thermophilic pairs might have some involvement in the subsurface generation of petroleum from organic materials. We then supposed that the oil generation process could be analyzed in subsurface microalgae during a short period under anaerobic, high-temperature, and atmospheric conditions using syntrophic pairs that have rapid-growth ability. However, we could not isolate either the bacterium or the archaeon from either production water or oils from our samples.
On the other hand, Kuwabara et al. (7) isolated a thermostable, anaerobic bacterium (Thermosipho globiformans) from a hydrothermal vent at Suiyou Seamount in the Western Pacific Ocean. The growth of this bacterium at 68°C under anaerobic conditions was enhanced 5- to 7-fold in coculture with the thermostable archaeon Methanocaldococcus jannaschii (8, 9) compared with monoculture. Furthermore, cocultures of the two microorganisms rapidly proliferated and reached the stationary phase of growth within 16 h (K. Igarashi and T. Kuwabara, unpublished data).
This growth was similar to that of cocultures of Thermotoga maritima and M. jannaschii at 80°C (10, 11), whereas it was 30- to 50-fold faster than that of pairs of T. phaeum and M. thermautotrophicus at 55°C (6) and of Pelotomaculum thermopropionicum and M. thermautotrophicus at 55°C (12). Therefore, we cocultured T. globiformans and M. jannaschii because they comprised a bacterial and archaeal pair similar to those found in the subsurface oil deposit. We then tested various alga-bacterium-archaeon consortia to determine the generation of oil-like mixtures within a short period under anaerobic, thermophilic, and atmospheric conditions. The processes involved in oil generation, such as n-alkane production, can be duplicated by laboratory pyrolysis. Higher temperatures are needed to drive these reactions within a few hours or days rather than the millions of years in nature. Pyrolysis is an established and accepted research technology that has been applied to generate liquid fuels from higher plants, including woody materials (13). The production of biofuels from marine microalgae has also been studied in detail because microalgae are primarily renewable producers in oceans and they constitute the largest biomass in nature. Furthermore, they do not compete with the food industry for starch stocks and arable land (14–17).
The present study aimed to compress the time required to generate oil from microalgae and to mimic the subsurface reaction using rapidly proliferating model cocultures of T. globiformans and M. jannaschii with pyrolysis at 300°C for 4 days. This strategy generated high yields of n-alkane-rich oils from microalgae. We also identified a close relationship between the n-alkane composition in hexane subfractions after pyrolysis and the lipid composition of Arthrospira platensis (Cyanobacteria), Dunaliella tertiolecta (Chlorophyta), Emiliania huxleyi (Haptophyta), and Euglena gracilis (Euglenophyta) microalgae as biofuel sources. A considerable amount of n-alkanes with compositions similar to those of native petroleum crude oils was generated from E. huxleyi, which contains more long-chain alkenes and very long-chain alkenones (18).
E. huxleyi is a unicellular, calcifying coccolithophorid alga of the phylum Haptophyta that is widely distributed in oceans, where it frequently forms massive blooms that can cover >100,000 km2 of the ocean surface. During such blooms, most of the algae tend to sink and sediment on the ocean floor (19, 20). Alkenones are less labile and have been identified in sea sediments from the Eocene age and in Cretaceous black shale (21).
MATERIALS AND METHODS
Bacterial and archaeal strains.
The bacterial strain Thermosipho globiformans MN14T was isolated from a hydrothermal vent at Suiyo Seamount, Izu-Bonin Arc, western Pacific Ocean (28°34′N, 140°38′E), at a depth of 1,384 m and a temperature of 230°C and was identified in our laboratory (7). The archaeal strain Methanocaldococcus jannaschii DSM 2661 was isolated from black sediments sampled at the East Pacific Rise hydrothermal vents at 21oN (8, 9). The strain was purchased from Deutsche Sammlung von Mikroorganismen und Zell Kulturen (DSMZ) GmbH, Braunschweig, Germany. Monocultures of T. globiformans and M. jannaschii grow poorly in Tc medium without sulfur powder (Tc-So [see below]) at 68°C under N2-H2-CO2 (80:10:10) and H2-CO2 (80:20) gas phases, respectively. The cell densities were 7 × 107 and 5 × 106 cells/ml, respectively, after 16 h of cultivation. In contrast, cocultures of the two strains at 68°C for 16 h in Tc-So medium under a gas phase of N2-H2-CO2 (80:10:10) stimulated their growth 5- to 7-fold (T. globiformans, 4 × 108 cells/ml; M. jannaschii, 3 × 107 cells/ml).
The Tc-So medium contained (liter−1) 25 g NaCl, 0.33 g KCl, 2.8 g MgCl2 · 6H2O, 1.66 g MgSO4, 0.3 g K2HPO4, 0.25 g NH4Cl, 10 mg NaBr, 25 mg Fe2SO4 · 7H2O, 10 ml each of trace mineral and vitamin solutions (22), 3 g of yeast extract (Difco), 3 g of tryptone (Difco), and 1 mg of resazurin. Sulfur powder was omitted from the Tc medium (7). After adjusting the pH to 6.5, the medium was sterilized by autoclaving and stored at room temperature.
Before anaerobic cultivation, the medium was anoxidized by adding 1/20 volume of 10% (wt/wt) Na2S · 9H2O and 10% (wt/wt) l-cysteine HCl in an anaerobic workstation (Ruskinn Technology Ltd., Leeds, United Kingdom) under an N2-H2-CO2 (80:10:10) gas phase. Separate T. globiformans and M. jannaschii monocultures were simultaneously inoculated into serum bottles (final concentrations, 1 × 106 and 1 × 105 cells/ml, respectively) with an initial N2-H2-CO2 (80:10:10) gas phase in the workstation.
Microalgae.
Dry, powdered freshwater algae, A. platensis (Cyanobacteria) and E. gracilis (Euglenophyta), were provided by DIC Life Tec Co. Ltd., Tokyo, Japan, and by Euglena H1 Co. Ltd., Tokyo, Japan, respectively. Saltwater algae, D. tertiolecta NIES 2258 (Chlorophyta) and E. huxleyi NIES 837 (Haptophyta), were cultured in artificial seawater (Marine Art SF-1; Osaka Yakken, Osaka, Japan) enriched with Erm-Schreiber medium (ESM) and 10 nM sodium selenite. The ESM, containing (liter−1) 12 g NaNO3, 0.5g K2HPO4, 25.9 mg FeEDTA, 33.2 mg MnEDTA, 10 mg thiamine-HCl, 0.1 mg biotin, 0.1 mg cyanocobalamin, and 100 g Tris at pH 8.0, was mixed (10 ml) with 990 ml of artificial seawater immediately before use (23). The cells were constantly illuminated at 100 μmol of photons m−2 s−1 by fluorescent lights in a 2-liter flat, oblong glass vessel aerated by air bubbling at 20°C for 12 days. The cells were harvested by centrifugation at 5,000 × g for 10 min, lyophilized, and stored at 5°C.
Microscopy.
Bacterial and archaeal monocultures and cocultures were routinely sampled for cell counting using a Nikon model Eclipse E600 microscope with phase-contrast and epifluorescence capabilities.
Pyrolysis.
Lyophilized powdered algae (100 mg each) were placed in Pyrex glass tubes (12 by 120 mm) to make ampoules. The air was completely replaced with nitrogen gas, and then the ampoules (12 by 70 to 75 mm) were evacuated using an oil vacuum pump, sealed, and heated in a muffle furnace at 300°C for 4 days. Thereafter, the ampoules were cooled and connected to a silicon tube, one side of which was sealed using a syringe sampler with a silicon septum. The air in the capped silicon tubes was evacuated, the tip of the ampoule was broken, 0.5 ml of the gas phase of the pyrolysis product was withdrawn, and hydrocarbons were analyzed by gas chromatography (GC). The remaining organic materials in the ampoules were Vortex mixed and then extracted twice with 0.5 ml of chloroform to analyze n-alkanes and other hydrocarbons.
Lipid analysis.
Microbial cells were sedimented by centrifugation, and then total lipids were extracted using chloroform-methanol as described by Ames (24). The total lipids in the chloroform fraction were dried by an evaporator and nitrogen blow down, and then they were dissolved in 1.0 ml of 5% hydrogen chloride-methanol (Wako Pure Chemicals Industries Ltd., Osaka, Japan) and heated at 100°C for 60 min for methanolysis. The samples were dried by an evaporator and nitrogen blow down, dissolved in 1 ml of chloroform, and fractionated by silica gel column chromatography.
Silica gel column chromatography.
A 150- by 5-mm column was plugged at the bottom with glass wool, rinsed with methanol, left to dry, and then dry packed with 0.5 g of activated Wako Gel C200 silica gel (Wako Pure Chemical Industries Ltd.). The gel was conditioned with 5 ml of hexane, which was discarded. Dried samples were suspended in hexane, applied to the column, and washed with an additional 0.5 ml of hexane. Samples were then eluted stepwise with 5 ml of hexane, 5 ml of 5% (vol/vol) diethyl ether in hexane (5% EH), 5 ml of 30% (vol/vol) diethyl ether in hexane (30% EH), and 5 ml of diethyl ether. The eluates were collected in calibrated glass centrifuge tubes, concentrated by evaporation, and dried under nitrogen gas. Dried samples were weighed, dissolved in 0.5 ml of chloroform, and stored at −20°C.
GC-mass spectrometry (MS).
The lipid and oil composition was analyzed using a 6890N gas chromatograph (Agilent Technology Inc., Santa Clara, CA) equipped with an MS-600H mass spectrometer (JEOL, Tokyo, Japan) and a 30-m by 0.25-mm (inside diameter [i.d.]) (0.25-μm film) DB-5MS fused-silica capillary column (Agilent Technology). The carrier gas was helium (1 ml/min). The injection and detector temperatures were set at 300°C and 320°C, respectively. The temperature program comprised 1 min at 50°C, ramping to 320°C in 10°C/min increments, and holding for 12 min.
Coculture of T. globiformans and M. jannaschii with sonicated microalgae.
The microalgae A. platensis, D. tertiolecta, E. huxleyi, and E. gracilis (0.5 g each) were suspended in 5 ml of Tc-So medium, frozen at −20°C, thawed, and sonically disrupted for 5 min at 5 kc with glass beads (except for E. huxleyi). The pH of the suspensions was adjusted to 6.5, and the volume was brought up to 100 ml using Tc-So medium. Media in 500-ml bottles were anoxidized using Na2S · 9H2O and l-cysteine HCl in the anaerobic workstation and then inoculated with T. globiformans and M. jannaschii cells (which had been separately cultured). We monitored T. globiformans and M. jannaschii populations, as well as CH4 production in the gas phases of the media. We then established practically optimal culture conditions as follows: microalgae were cocultured during the first stage at 68°C for 48 h under an N2-H2-CO2 (80:10:10) gas phase and in the second phase under H2-CO2 (80:20) at 68°C for 7 days. The gas phases were replaced at room temperature.
Insoluble materials in the media were collected after the second stage by centrifugation at 5,000 × g for 20 min, lyophilized, and stored at 5°C before being sealed in Pyrex test tubes for pyrolysis.
Flowchart of the study.
Figure 1 summarizes possible relationships between lipid composition and the generation of n-alkane-rich biofuels from cocultured microalgae in a flowchart.
Fig 1.
Flowchart of the study. A to D represent experiments.
Gas chromatography.
The methods for gas chromatography are described in the supplemental material.
RESULTS
Characterization of T. globiformans and M. jannaschii cocultures to determine optimal culture conditions for media containing microalgae.
We assessed the growth of T. globiformans and M. jannaschii cells and CH4 production in various culture media to determine the optimal coculture conditions for these organisms in modified Tc-So medium containing sonically oscillated microalgae. Basal Tc-So medium comprised 0.3% tryptone and 0.3% yeast extract. We prepared three additional media that included 0.5, 0.75, and 1.0% (each) tryptone and yeast extract. Both microorganisms were cultured under an N2-H2-CO2 (80:10:10) gas phase at 68°C under atmospheric conditions for 48 h during the first stage of coculture. The gas phase was then replaced with H2-CO2 (80:20), and the second stage of culture continued at 68°C for 7 days under atmospheric conditions. Figure 2 indicates the amounts of CH4 produced in the gas phases. The populations of T. globiformans and M. jannaschii reached 2 × 109 and 7.2 × 107 cells/ml, respectively, and the CH4 content in the N2-H2-CO2 (80:10:10) gas phase reached 12% after 48 h of the first stage of culture in medium enriched with 1% tryptone and 1% yeast extract. In contrast, the bacterial cell density was reduced to 1 × 109 cells/ml, while that of the archaeal cells increased to 1.1 × 108 cells/ml. In addition, the gas phase comprised almost 60% CH4 after 7 days of second-stage cultivation under an H2-CO2 (80:20) gas phase. At the third stage, when the gas phase was replaced with H2-CO2 (80:20), M. jannaschii maintained CH4 production. Both T. globiformans and M. jannaschii rapidly grew in a syntrophic fashion during the first stage (Fig. 2) by utilizing H2 produced via the bacterial metabolism of organic materials. However, only M. jannaschii proliferated during the second stage under a high H2 concentration and produced more CH4. The second stage of the coculture in basal Tc-So medium resulted in 4 × 108 cells/ml of T. globiformans, 3 × 107 cells/ml of M. jannaschii, and high (35%) levels of CH4 release into the gas phase. We also added organic acids, carbohydrates, and amino acids to the Tc-So medium and measured CH4 production and the growth of the two microorganisms. However, none of these compounds exerted any significant effect.
Fig 2.
Production of CH4 in three gas phases by cocultured T. globiformans and M. jannaschii in Tc-So medium containing various concentrations of tryptone and yeast extract at 68°C under an N2-H2-CO2 (80:10:10) gas phase in stage I and H2-CO2 (80:20) in stages II and III. □, Basal Tc-So medium containing 0.3% tryptone and 0.3% yeast extract; ●, Tc-So medium containing 0.5% tryptone and 0.5% yeast extract; ▲, Tc-So medium containing 0.75% tryptone and 0.75% yeast extract; ○, Tc-So medium containing 1% tryptone and 1% yeast extract.
Analysis of cell growth and CH4 production in the coculture of T. globiformans and M. jannaschii in the presence of microalgae.
The cell populations and CH4 production in medium containing A. platensis were almost identical to those in Tc-So basal medium during the first and second stages. In contrast, CH4 production and cell growth were reduced to 70% of those in Tc-So basal medium when media contained D. tertiolecta and E. gracilis after 7 days of cultivation at the second stage. Growth and CH4 production were obviously repressed during the first and second stages in medium containing E. huxleyi. The CH4 concentration in the gas phase was <0.5%, and the densities of T. globiformans and M. jannaschii were <5 × 107 and <1 × 106 cells/ml, respectively, at both stages. The growth of T. globiformans and M. jannaschii reached the stationary phase at 24 h of cultivation in the first stage of Tc-So and in enriched medium and needed 48 h for growth to the stationary phase in cocultivation with microalgae. These results indicated that the microalgae interacted with T. globiformans and M. jannaschii to repress their growth and CH4 formation. Regardless, such repression enhanced the production of n-alkane-rich oils because of a decrease in the number of impurities during pyrolysis. Based on these findings, we established optimal coculture conditions in basal Tc-So basal medium containing microalgae.
Lipid composition of microalgae cocultured with T. globiformans and M. jannaschii and n-alkanes produced by pyrolysis.
We analyzed biofuel production from the four microalgae by comparing the lipid composition of the cocultured algae with the content and composition of n-alkanes produced by pyrolysis. Lipids extracted with chloroform-methanol from the algae were dried and heated with 5% HCl-methanol at 100°C for 60 min for methanolysis, and then the lipid composition was analyzed by GC-MS (Table 1). The major lipid components of A. platensis and D. tertiolecta were methyl esters of 16- and 18-carbon-chain fatty acids that were derived from glycolipids and phospholipids and small amounts of isoprenoids. The lipid contents of the two microalgae accounted for 15 to 20% (wt/wt). In addition to these methyl esters, considerable amounts of docosahexaenoic acid (DHA) methyl ester, long-chain (31 and 33 carbon atoms) alkenes, and very long-chain (37, 38, and 39 carbon atoms) hydrocarbon alkenones (18) were detected in lipid fractions from the cocultured E. huxleyi sample. The lipid content in E. huxleyi was 40 to 45% (wt/wt), and the total alkene and alkenone content reached over 55% of the total lipid.
Table 1.
Main components of fatty alcohols, fatty acids, wax esters, alkenes, and alkenones in the total lipids extracted from microalgae cocultured with T. globiformans and M. jannaschii
| Lipid component | Contenta |
|||
|---|---|---|---|---|
| A. platensis | D. tertiolecta | E. huxleyi | E. gracilis | |
| Fatty alcohols | ||||
| C12 | — | — | — | 0.9 |
| C13 | — | — | — | 1.4 |
| C14 | — | — | — | 13.9 |
| C15 | — | — | — | 1.1 |
| C16 | — | — | — | 3.2 |
| Isoprenoid hydrocarbons (5 components) | 4.3 | 4.5 | 2.2 | 3 |
| Fatty acid methyl esters | ||||
| C12:0 | — | — | — | 1.9 |
| C13:0 | — | — | — | 4.1 |
| C14:0 | — | 0.7 | 7.7 | 18.8 |
| C16:0 | 68.3 | 24.5 | 5.1 | 14.9 |
| C16:1 | 3.8 | 0.8 | 0.8 | — |
| C16:2 | — | 1.6 | — | — |
| C16:4 | — | 11.2 | — | — |
| C18:0 | 1.3 | 0.4 | 0.5 | 11.4 |
| C18:1 | — | 4.8 | 1 | 10.5 |
| C18:2 | 11.1 | 2.4 | 0.2 | 0.2 |
| C18:3 | 11.2 | 46.3 | 8.5 | — |
| C18:4 | — | 2.8 | 2.8 | — |
| C18:5 | — | — | 6.7 | — |
| C20:2 | — | — | — | 1.5 |
| C20:3 | — | — | — | 2.9 |
| C20:4 | — | — | — | 3.9 |
| C22:0 | — | — | 1.6 | — |
| C22:4 | — | — | — | 0.2 |
| C22:6 | — | — | 7.7 | — |
| Wax esters | ||||
| C24 | — | — | — | 0.6 |
| C26 | — | — | — | 1.1 |
| C28 | — | — | — | 2.4 |
| C30 | — | — | — | 0.7 |
| C32 | — | — | — | 1.2 |
| C34 | — | — | — | 0.2 |
| Long-chain alkenes | ||||
| C31:1 | — | — | 2.5 | — |
| C31:2 | — | — | 6.9 | — |
| C33:3 | — | — | 5.6 | — |
| Very long-chain alkenones | ||||
| C37 | — | — | 23.2 | — |
| C38 | — | — | 16.1 | — |
| C39 | — | — | 0.9 | — |
Components were calculated as ratios (%) of each total lipid sample solubilized in chloroform from peak areas of GC-MS profiles after extracted samples were methanolized and analyzed. —, Undetectable or very small peaks. The values are averages of triplicate experiments.
Many fatty alcohols, wax esters, and fatty acid methyl esters were detected in the lipid fraction of E. gracilis (Table 1), as described by Koritala (25). The lipid content in E. gracilis was 20 to 25% of the dry weight.
C16:0 fatty acid methyl ester accounted for 80% of the lipid composition of cocultured T. globiformans and M. jannaschii cells, and GC-MS showed that the methanolysis products of lipids extracted from microalgae with and without cocultures of the two microorganisms essentially did not differ.
Table 2 shows the main hydrocarbons and their relative amounts in hexane subfractions separated by Wako Gel C200 column chromatography after pyrolysis and GC-MS analysis of the four cocultured algae. Samples of A. platensis and D. tertiolecta, in which the major lipids comprised fatty acids, both contained short-chain n-alkanes (nC13 to nC19). In addition, we detected large amounts of phytane (iC20) in the A. platensis sample, as well as pristane (iC19) and phytane in the D. tertiolecta sample. Short-chain n-alkanes (nC13 to nC19) and iC20 were also major components of the E. gracilis sample, which contained nC20 to nC25 alkanes as minor components. In contrast, the E. huxleyi sample of major lipids, comprising long-chain alkenes and very long-chain alkenones, contained n-alkanes of various chain lengths (nC13 to nC35) and small amounts of pristane and phytane. These n-alkane and isoprenoid profiles were similar to those of native crude oils (4, 26) except for the considerable amount of nC31 alkane. Two peaks indicating biomarkers (steranes) were detected in the profiles of an extracted ion (m/z 217). However, the ion (m/z 191) for hopanes was very small. These results indicated that the n-alkane profiles determined by GC-MS in hexane subfractions from the pyrolysis products of microalgae samples are closely related to the lipid composition of the microalgae. Long-chain lipid components, in addition to basic fatty acid methyl esters, increased the amounts of generated oils and the appearance of long-chain n-alkanes in each alga. The final concentrated samples of hexane subfractions from A. platensis, D. tertiolecta, and E. gracilis were liquid, whereas that from E. huxleyi was solid.
Table 2.
Main hydrocarbons in hexane subfractions from Wako Gel C200 column chromatography after pyrolysis of the four cocultured algae
| Hydrocarbon | Abbreviation | Contenta |
|||
|---|---|---|---|---|---|
| A. platensis | D. tertiolecta | E. huxleyi | E. gracilis | ||
| n-Dodecane | nC12 | 0.5 | 0.5 | 2.1 | 1.8 |
| n-Tridecane | nC13 | 2.2 | 1.4 | 4.5 | 12.5 |
| n-Tetradecane | nC14 | 4.8 | 3 | 4.7 | 22.6 |
| n-Pentadecane | nC15 | 16.5 | 9.5 | 5.2 | 15.3 |
| n-Hexadecane | nC16 | 5.1 | 3.7 | 3.1 | 21.5 |
| n-Heptadecane | nC17 | 31.1 | 3.1 | 5.1 | 12.6 |
| Pristane | iC19 | 3.1 | 44.1 | 0.2 | 0.6 |
| n-Octadecane | nC18 | 2.6 | 2.3 | 2.7 | 1.9 |
| Phytane | iC20 | 30.9 | 30.1 | 3.6 | 7.7 |
| n-Nonadecane | nC19 | 3.2 | 1.5 | 6.5 | 1.3 |
| n-Eicosane | nC20 | — | 0.8 | 2.7 | 0.5 |
| n-Heneicosane | nC21 | — | — | 3.9 | 0.4 |
| n-Docosane | nC22 | — | — | 3.4 | 0.4 |
| n-Tricosane | nC23 | — | — | 3.1 | 0.7 |
| n-Tetracosane | nC24 | — | — | 2.1 | 0.1 |
| n-Pentacosane | nC25 | — | — | 1.1 | 0.1 |
| n-Hexacosane | nC26 | — | — | 0.8 | — |
| n-Heptacosane | nC27 | — | — | 1.4 | — |
| n-Octacosane | nC28 | — | — | 1.9 | — |
| n-Nonacosane | nC29 | — | — | 0.9 | — |
| n-Triacontane | nC30 | — | — | 5.5 | — |
| n-Hentriacontane | nC31 | — | — | 17.4 | — |
| n-Dotriacontane | nC32 | — | — | 1.8 | — |
| n-Tritriacontane | nC33 | — | — | 5.3 | — |
| n-Tetratriacontane | nC34 | — | — | 3.9 | — |
| n-Pentatriacontane | nC35 | — | — | 7.1 | — |
Hydrocarbon contents were calculated as percentages (%) in total amounts of each subfraction from peak areas of GC-MS profiles. —, undetectable. The values are averages of at least two experiments.
Effect of coculture with T. globiformans and M. jannaschii on biofuel production.
To demonstrate the possible effects of cocultures on the production of n-alkane-rich oils, we analyzed the n-alkane compositions of pyrolysis products from the four algae that were not cocultured. Table 3 compares the compositions and amounts of n-alkanes and nitriles detected in the GC-MS profiles of chloroform-soluble pyrolysis fractions of E. gracilis samples with and without coculture. All peaks detected by total ion scan chromatography in the cocultured samples coincided with those of the extracted ion profile of the m/z 71 fragment, which is one characteristic fragment of n-alkanes. In contrast, many noncoincident peaks were detected in samples without coculture. The major noncoincident peaks were detected as tetradecanenitrile (C14N), hexadecanenitrile (C16N), and octadecanenitrile (C18N), eluted in the 5% EH subfraction on Wako Gel C200. Those alkanenitriles were also detected in the other three microalgal samples without coculture. Therefore, we speculated that reducing the amounts of impurities, such as nitriles, during pyrolysis enhanced the production of n-alkane-rich oil.
Table 3.
Comparison of main hydrocarbon and nitrile contents in chloroform-soluble fractions from pyrolysis products of cocultured and noncocultured E. gracilis
| Hydrocarbon or nitrile | Contenta |
|
|---|---|---|
| Cocultured alga | Noncocultured alga | |
| nC12 | 8.7 | 6.4 |
| nC13 | 16.1 | 12.2 |
| nC14 | 24.1 | 19.7 |
| nC15 | 12.6 | 9.1 |
| nC16 | 14.6 | 12.4 |
| Tetradecanenitrile (C14N) | 0.3 | 17.6 |
| nC17 | 14.1 | 3.9 |
| iC19 | 0.1 | 0.4 |
| nC18 | 1.5 | 0.6 |
| iC20 | 6.6 | 3.1 |
| Hexadecanenitrile (C16N) | 0 | 9.2 |
| nC19 | 1.3 | 0.3 |
| Octadecanenitrile (C18N) | 0 | 5.1 |
Components were calculated as ratios (%) of total amounts of chloroform- soluble fractions based on peak areas of GC-MS profiles. All values are averages of two experiments.
Table 4 summarizes the total dry weight (mg) of crude oil production in the hexane, 5% EH, 30% EH, and diethyl ether subfractions (from Wako Gel C200) of the chloroform-soluble fractions and the oil recovered (mg) in the hexane subfraction of the four microalgae (100 mg each) with and without coculture after pyrolysis. Table 4 also summarizes the ratios (%) of isoprenoids, fatty acid methyl esters, fatty alcohols, wax esters, alkenes, and alkenones that were calculated from each GC-MS peak area of lipid samples. More crude oil was produced by pyrolysis, and almost twice as much of the n-alkane-rich biofuels was recovered from microalgae that had been cocultured with the two microorganisms. About 30% of the dry weight of cocultured E. huxleyi was recovered as crude oil, and 8 to 10% was recovered as n-alkane-rich oil in the hexane subfraction after pyrolysis.
Table 4.
Lipid compositions of cocultured algae and comparison of oil production from pyrolyzed microalgae with and without coculture of T. globiformans and M. jannaschii
| Parameter | Value in microalgaed |
|||
|---|---|---|---|---|
| A. platensis (Cyanobacteria) | D. tertiolecta (Chlorophyta) | E. huxleyi (Haptophyta) | E. gracilis (Euglenophyta) | |
| Lipid composition (%)a | ||||
| Isoprenoids | 4.3 | 4.5 | 2.2 | 3.0 |
| Fatty acid methyl esters | 95.7 | 95.5 | 42.6 | 70.3 |
| Fatty alcohols | ND | ND | ND | 20.5 |
| Wax esters | ND | ND | ND | 6.2 |
| Alkenes | ND | ND | 15.0 | ND |
| Alkenones | ND | ND | 40.2 | ND |
| Crude oil productionb (%) in chloroform fraction | 17.5 | 13.2 | 29.4 | 20.4 |
| 14.6 | 8.7 | 23.2 | 17.2 | |
| Oil recoveryc (%) in hexane subfractions | 4.2 | 2.8 | 8.9 | 5.8 |
| 1.2 | 1.5 | 5.7 | 3.1 | |
Contents of lipid compositions were calculated as ratios (%) of total lipid composition in samples treated with 5% HCl-methanol determined from peak GC-MS areas.
Total dry weight of Wako Gel C200 subfractions (hexane, 5% EH, 30% EH, and diethyl ether subfractions) of chloroform fractions obtained from pyrolysis products of algae (100 mg) with and without (boldface) coculture.
Dry weight of Wako Gel C200 hexane subfractions of chloroform fractions obtained from pyrolysis products of 100 mg of algal samples with and without (boldface) coculture.
All values are averages of at least three experiments, and the standard deviation of each is ±10 to 20%. ND, not detected.
Characterization of hydrocarbon gases resulting from algal pyrolysis.
We used gas chromatography to analyze the composition of hydrocarbons in 0.5 ml of the 6 ml of gases resulting from the pyrolysis of microalgal samples (100 mg each). Figure S1 in the supplemental material shows that at least 20 peaks were detected in E. huxleyi-cocultured samples. The same peaks were detected in all four microalgal samples with and without coculture. Among these peaks, the contents of methane, CO2, ethylene, ethane, propylene, propane, i-butane, n-butane, and n-pentane were measured using a calibrated chromatograph (see Table S1 in the supplemental material). The CO2 content was very high, and the gases were rich in methane and ethane. The compositions and ratios of the hydrocarbon gases produced by pyrolysis were similar among the algae and with and without cocultivation. The n-hexane content was very low, and a peak for n-hexane could not be quantified.
Cocultured T. globiformans and M. jannaschii cells produced essentially no n-alkane-rich biofuel.
We prepared two batches of lyophilized cocultured bacterial and archaeal cells to determine whether they can produce biofuels. One batch was cocultured for 2 days in Tc-So medium under a N2-H2-CO2 (80:10:10) gas phase, followed by 7 days under an H2-CO2 (80:20) gas phase at 68°C for comparison with cocultured microalgae (9-day cocultures). The other was cocultured in Tc-So medium enriched with 1% yeast extract and 1% tryptone at 68°C for 24 h under an N2-H2-CO2 (80:10:10) gas phase. After pyrolysis at 300°C for 4 days (100 mg per sample), hydrocarbons in the chloroform-soluble fractions and the hexane, 5% EH, 30% EH, and diethyl ether subfractions from the two samples separated by column chromatography on Wako Gel C200 were analyzed by GC-MS. Figure S2A to E in the supplemental material shows the results for cells that were cocultured for 9 days. The major components of the two chloroform-soluble fractions were tetradecanenitrile (C14N), pentadecanenitrile (C15N), and hexadecanenitrile (C16N) and a minimal amount of n-alkanes (nC14, nC15, and nC16) (see Fig. S2A in the supplemental material). The n-alkanes eluted in the hexane subfractions (see Fig. S2B in the supplemental material), and nitriles eluted in the 5% EH subfraction (see Fig. S2C in the supplemental material). Branched alkanes were detected only in the hexane subfraction (see Fig. S2B in the supplemental material) from the cells cocultured for 9 days. The 30% EH subfractions (see Fig. SD in the supplemental material) contained phenol and naphthalene, as well as their derivatives, which were detected by analyzing the ion profiles of m/z 94, 108, 122, 128, 142, and 156 extracted from the GC-MS values (see Fig. S3 in the supplemental material). The dry weight of each subfraction was 0.1, 2.5, 2.5, and 3.4 mg of the 9-day cocultures and 0.2, 6.1, 5.4, and 4.8 mg of the cells cocultured in enriched medium. The total weight (mg) of the four subfractions was assumed to be the dry weight of the chloroform-soluble fraction, and the total values corresponded to 8.5 and 16.5% of the lyophilized starter cells, respectively. Since the ratios of the hexane subfractions to the starter cells were only 0.1% and 0.2% (wt/wt), we assumed that the cocultured cells essentially did not generate n-alkane-rich biofuels. All values are averages of triplicate experiments.
DISCUSSION
We tested different alga-bacterium-archaeon consortia to investigate the production of oil-like mixtures, suggesting that they could potentially be used for the synthesis of n-alkane-rich biofuels for a very short period. A model coculture of T. globiformans and M. jannaschii under anaerobic and atmospheric conditions at 68°C and subsequent pyrolysis at 300°C for 4 days were introduced in the process. The yield of crude and n-alkane-rich oil was highest when the marine microalga E. huxleyi served as the raw material. The GC-MS profiles of n-alkanes and isoprenoids in the pyrolysis products were similar to those of natural crude oils, except for the higher nC31 alkane levels. Petroleum crude oils contain many compounds that are undetectable by GC-MS. Therefore, analysis of hydrocarbons in pyrolysis products is limited. However, n-alkanes and related hydrocarbons can be quantitatively recovered using gas and silica gel column chromatography (27).
Adding tryptone and yeast extract to cocultures in the enriched media enhanced the growth of the two microorganisms and CH4 production (Fig. 2), whereas adding sonicated microalgae (in particular E. huxleyi) repressed or did not affect either cell growth or CH4 production. Introducing the coculture step increased the production of crude and n-alkane-rich oils generated from the microalgae by pyrolysis. We suppose that one reason for the enhanced production of biofuels from microalgae is a reduction in the formation of impurities containing alkanenitriles during pyrolysis. Alkanenitriles comprised the major pyrolysis products from cocultured T. globiformans and M. jannaschii cells (see Fig. S2 in the supplemental material) and from algal samples without coculture. However, small amounts of alkanenitriles were detected in the pyrolysis products of cocultured algal samples. These results suggest that cocultured cells themselves and/or the culture environment, such as anaerobic conditions, prevented alkanenitrile production and enhanced n-alkane production during pyrolysis. However, the actual reasons for the elevated production remain unresolved.
Phospholipids and glycolipids are major lipids in the cell membrane and photosynthetic apparatus of microalgae, and they are generated from 14- to 22-carbon chain fatty acids. These lipids were common and essential to the four microalgae. We considered that most of the methyl esters in the four microalgae detected by GC-MS were derived from these 14- to 22-carbon chain fatty acids by methanolysis. Analysis of A. platensis and D. tertiolecta lipids by GC-MS indicated mainly methyl esters and small amounts of isoprenoids. Short-chain (C13 to C19) n-alkanes and more isoprenoids (iC19 and iC20) were produced from the microalgae by pyrolysis. The ratios of iC20 to nC18 in A. platensis and D. tertiolecta were 11.9 and 13.1, respectively. In contrast, fatty alcohols and wax esters, in addition to these methyl esters, were detected in E. gracilis. The fatty alcohol and wax ester contents were still low (20.5% and 6.2%, respectively). Therefore, short-chain n-alkanes (C13 to C19) were also rich in pyrolyzed E. gracilis. However, this sample contained some nC20 to nC25 alkanes that reduced the iC20/nC18 ratio to 4.1. In contrast, more long-chain (C31 and C33) alkenes (15%) and very long-chain (C37 to C39) alkenones (40.2%) were detected in E. huxleyi lipids. Concomitant with these findings, short-chain (nC13) to very long-chain (nC35) n-alkanes were detected in pyrolysis products of the E. huxleyi sample, and the iC20/nC18 ratio was also reduced to 1.3. The reported ratio of iC20 to nC18 in natural crude oils is <1.0 (28). Thus, the GC-MS profile of the hexane subfraction of E. huxleyi was similar to those of natural crude oils despite the presence of nC31 alkane.
We found that oil generation from E. huxleyi can be accelerated by introducing T. globiformans and M. jannaschii cocultures and pyrolysis and that essentially no n-alkane-rich biofuel was produced from the cocultured bacterial and archaeal cells themselves after pyrolysis. We suppose that alkenones of coccolithophorid algae like E. huxleyi might serve as key raw materials for subsurface crude oil generation.
Supplementary Material
ACKNOWLEDGMENTS
This study was supported in part by a grant-in-aid for Science Research from the Ministry of Education, Culture, Sports, Science and Technology of Japan.
We are grateful to N. Foster for critical reading of the manuscript.
We declare no conflict of interest.
Footnotes
Published ahead of print 26 November 2012
Supplemental material for this article may be found at http://dx.doi.org/10.1128/AEM.01685-12.
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