Significance
A number of organisms synthesize hydrocarbons, but the scale at which this occurs in the environment is unknown. Here, we provide the first global estimates of hydrocarbon production by the two most abundant cyanobacteria on Earth, Prochlorococcus and Synechococcus. We suggest that these organisms represent a significant and widespread source of hydrocarbons to the world’s oceans, which in turn may sustain populations of obligate hydrocarbon-degrading bacteria known to be important in consuming anthropogenic oil spills. Our study demonstrates the role cyanobacteria play in the ocean ‟hydrocarbon cycle” and reveals the massive scale of this process. The widespread distribution of cyanobacteria and hydrocarbon-degrading bacteria in freshwater, marine, and terrestrial environments suggests the hydrocarbon cycle is pervasive in many natural ecosystems.
Keywords: cyanobacteria, hydrocarbons, hydrocarbon cycle, hydrocarbon-degrading bacteria, oil remediation
Abstract
Hydrocarbons are ubiquitous in the ocean, where alkanes such as pentadecane and heptadecane can be found even in waters minimally polluted with crude oil. Populations of hydrocarbon-degrading bacteria, which are responsible for the turnover of these compounds, are also found throughout marine systems, including in unpolluted waters. These observations suggest the existence of an unknown and widespread source of hydrocarbons in the oceans. Here, we report that strains of the two most abundant marine cyanobacteria, Prochlorococcus and Synechococcus, produce and accumulate hydrocarbons, predominantly C15 and C17 alkanes, between 0.022 and 0.368% of dry cell weight. Based on global population sizes and turnover rates, we estimate that these species have the capacity to produce 2–540 pg alkanes per mL per day, which translates into a global ocean yield of ∼308–771 million tons of hydrocarbons annually. We also demonstrate that both obligate and facultative marine hydrocarbon-degrading bacteria can consume cyanobacterial alkanes, which likely prevents these hydrocarbons from accumulating in the environment. Our findings implicate cyanobacteria and hydrocarbon degraders as key players in a notable internal hydrocarbon cycle within the upper ocean, where alkanes are continually produced and subsequently consumed within days. Furthermore we show that cyanobacterial alkane production is likely sufficient to sustain populations of hydrocarbon-degrading bacteria, whose abundances can rapidly expand upon localized release of crude oil from natural seepage and human activities.
Hydrocarbons are ubiquitous in the oceans, where natural seepage and human activities are estimated to release between 0.4 and 4.0 million tons of crude oil into the ocean ecosystem annually (1). Even in minimally polluted marine surface waters, alkanes such as pentadecane and heptadecane have been found at concentrations ranging from 2 to 130 pg/mL (2, 3), although their sources remain unclear. A small proportion of alkanes, from 1 to 60 fg/mL, is associated with particulate matter >0.7 µm in diameter (4). Larger amounts may be associated with particulate matter <0.7 µm in diameter, because ocean concentrations are higher than the solubility of pentadecane and heptadecane, which is ∼10 pg/mL and 1 pg/mL, respectively (2). Populations of hydrocarbon-degrading bacteria, referred to as hydrocarbonoclastic bacteria, including many species that cannot use other carbon sources, are present in marine systems and play an important role in turnover of these compounds (5–9). Because obligate hydrocarbon-degrading bacteria are found in waters without significant levels of crude oil pollution, these organisms must use an alternate hydrocarbon source (9–11).
Here, we investigate the extent to which cyanobacteria may contribute to these marine hydrocarbon pools. Cyanobacteria (oxygenic photosynthetic bacteria) can synthesize C15 to C19 hydrocarbons via two separate pathways. The first produces alkanes, predominantly pentadecane, heptadecane, and methyl-heptadecane, in addition to smaller amounts of alkenes, via acyl-ACP reductase (FAR) and aldehyde deformylating oxygenase (FAD) enzymes (12). The second pathway generates alkenes, primarily nonadecene and 1,14-nonadecadiene, via a polyketide synthase enzyme (Ols) (13). The abundance and ubiquity of cyanobacteria in the marine environment suggests hydrocarbon production in the oceans could be considerable and broadly distributed geographically (14, 15).
We focused our studies on the two most abundant marine cyanobacteria, Prochlorococcus and Synechococcus (16). These genera have estimated global population sizes of 2.9 ± 0.1 × 1027 and 7.0 ± 0.3 × 1026 cells, respectively (14), and are together responsible for approximately a quarter of marine net primary production (14). These are also the only cyanobacterial genera for which global population size estimates have been compiled (14). Although the distribution patterns of both genera overlap (14, 17), Prochlorococcus cells dominate low-nutrient open-ocean areas between 40°N and 40°S and can be found at depths of up to 200 m (16, 18). Synechococcus are more numerous in coastal and temperate regions where conditions and nutrient levels are more variable (14, 16) but are still widely distributed in high abundance.
Results and Discussion
Marine Cyanobacteria Encode Alkane Biosynthetic Pathways.
We first examined the genetic capability for hydrocarbon production in marine cyanobacterial genomes. Previous studies have demonstrated that hydrocarbon biosynthetic pathways are conserved in sequenced cyanobacteria, encompassing isolates from marine, freshwater, and terrestrial environments (19). This suggests that hydrocarbons have a key, as-yet-unidentified, function in cyanobacteria that is independent of the ecosystem these organisms inhabit. We extended this work by including newly sequenced Prochlorococcus strains, isolated from diverse regions of the ocean (20). All 36 Prochlorococcus and 15 marine Synechococcus strains examined have the predicted capacity to synthesize alkanes via FAR/FAD enzymes (Table S1). With the exception of Leptolyngbya sp. PCC7376 and Moorea producens 3L, which encode Ols homologs, all of the other marine cyanobacteria including Cyanobium, Acaryochloris, Crocosphaera, Trichodesmium, Lyngbya, Oscillatoria, Nodularia, and Microcoleus species encode FAR/FAD homologs (15, 19). Thus, it is likely that alkanes are the predominant hydrocarbons released by cyanobacteria into the marine environment. Homologs of FAR/FAD and Ols were not identified in any other bacterial, plant, or algal species, suggesting that these pathways for hydrocarbon production are unique to cyanobacteria.
Table S1.
Hydrocarbon pathways in sequenced cyanobacterial strains
| Strain | fad | far | ols |
| Acaryochloris marina MBIC11017 | 292/337(86%) | 191/230(83%) | |
| Anabaena cylindrica PCC7122 | 301/338(89%) | 197/223(88%) | |
| Anabaena sp. 90 | 297/338(87%) | 196/223(87%) | |
| Anabaena variabilis ATCC29413 | 305/338(90%) | 204/230(88%) | |
| Arthrospira maxima CS-328 | 295/338(87%) | 206/231(89%) | |
| Arthrospira platensis NIES-39 | 295/338(87%) | 206/231(89%) | |
| Calothrix sp. PCC6303 | 298/338(88%) | 202/228(88%) | |
| Calothrix sp. PCC7507 | 299/338(88%) | 202/228(88%) | |
| Candidatus atelocyanobacterium thalassa | 296/338(87%) | 198/228(86%) | |
| Chamaesiphon minutus PCC6605 | 299/340(87%) | 193/230(83%) | |
| Chroococcidiopsis thermalis PCC7203 | 301/339(88%) | 200/228(87%) | |
| Coleofasciculus chthonoplastes PCC7420 | 298/338(88%) | 199/230(86%) | |
| Crinalium epipsammum PCC9333 | 299/339(88%) | 190/225(84%) | |
| Crocosphaera watsonii WH8501 | 291/339(85%) | 206/231(89%) | |
| Cyanobacterium aponinum PCC10605 | 290/340(85%) | 193/225(85%) | |
| Cyanobacterium stanieri PCC7202 | 1,472/2,442(60%) | ||
| Cyanobium gracile PCC6307 | 256/337(75%) | 181/220(82%) | |
| Cyanobium sp. PCC7001 | 261/334(78%) | 182/220(82%) | |
| Cyanothece sp. ATCC51142 | 294/339(86%) | 206/228(90%) | |
| Cyanothece sp. CCY0110 | 301/338(89%) | 207/230(90%) | |
| Cyanothece sp. PCC7424 | 1,862/2,796(66%) | ||
| Cyanothece sp. PCC7425 | 290/338(85%) | 196/230(85%) | |
| Cyanothece sp. PCC7822 | 1,871/2,796(66%) | ||
| Cyanothece sp. PCC8801 | 306/339(90%) | 205/230(89%) | |
| Cylindrospermopsis raciborskii CS-505 | 301/338(89%) | 191/222(86%) | |
| Cylindrospermum stagnale PCC7417 | 299/338(88%) | 199/231(86%) | |
| Dactylococcopsis salina PCC8305 | 282/338(83%) | 199/231(86%) | |
| Geitlerinema sp. PCC7407 | 292/339(86%) | 204/231(88%) | |
| Gloeobacter violaceus PCC7421 | 266/338(78%) | 183/221(82%) | |
| Gloeocapsa sp. PCC7428 | 299/338(88%) | 193/226(85%) | |
| Halothece sp. PCC7418 | 287/338(84%) | 202/228(88%) | |
| Leptolyngbya sp. PCC7376 | 2,151/2,724(78%) | ||
| Lyngbya sp. PCC8106 | 296/338(87%) | 196/229(85%) | |
| Microcoleus sp. PCC7113 | 301/340(88%) | 201/230(87%) | |
| Microcoleus vaginatus FGP-2 | 297/338(87%) | 194/230(84%) | |
| Microcystis aeruginosa NIES-843 | 299/338(88%) | 207/231(89%) | |
| Moorea producens 3L | 1,300/2,202(59%) | ||
| Nodularia spumigena CCY9414 | 302/338(89%) | 199/230(86%) | |
| Nostoc azollae' 0708 | 299/338(88%) | 197/223(88%) | |
| Nostoc punctiforme PCC73102 | 298/338(88%) | 196/222(88%) | |
| Nostoc sp. PCC7107 | 305/338(90%) | 195/230(84%) | |
| Nostoc sp. PCC7120 | 305/338(90%) | 204/230(88%) | |
| Nostoc sp. PCC7524 | 303/338(89%) | 202/230(87%) | |
| Oscillatoria acuminata PCC6304 | 294/338(86%) | 202/231(87%) | |
| Oscillatoria nigro-viridis PCC7112 | 297/338(87%) | 194/230(84%) | |
| Oscillatoria sp. PCC6506 | 301/338(89%) | 196/230(85%) | |
| Pleurocapsa sp. PCC7327 | 1,890/2,861(66%) | ||
| Prochlorococcus marinus str. AS9601 | 262/337(77%) | 173/214(80%) | |
| Prochlorococcus marinus str. CCMP1375 | 263/337(78%) | 169/218(77%) | |
| Prochlorococcus marinus str. CCMP1986 | 261/337(77%) | 173/214(80%) | |
| Prochlorococcus EQPAC1 | 261/337(77%) | 173/214(81%) | |
| Prochlorococcus GP2 | 262/337(78%) | 173/214(81%) | |
| Prochlorococcus LG | 263/337(78%) | 169/218(78%) | |
| Prochlorococcus marinus str. MIT9107 | 258/337(77%) | 171/214(80%) | |
| Prochlorococcus marinus str. MIT9116 | 258/337(77%) | 171/214(80%) | |
| Prochlorococcus marinus str. MIT9123 | 258/337(77%) | 171/214(80%) | |
| Prochlorococcus marinus str. MIT9201 | 261/337(77%) | 172/214(80%) | |
| Prochlorococcus marinus str. MIT9202 | 262/337(77%) | 148/186(80%) | |
| Prochlorococcus marinus str. MIT9211 | 257/334(76%) | 171/219(78%) | |
| Prochlorococcus marinus str. MIT9215 | 262/337(77%) | 171/214(79%) | |
| Prochlorococcus marinus str. MIT9301 | 262/337(77%) | 173/214(80%) | |
| Prochlorococcus marinus str. MIT9302 | 262/337(78%) | 173/214(81%) | |
| Prochlorococcus marinus str. MIT9303 | 237/303(78%) | 183/236(77%) | |
| Prochlorococcus marinus str. MIT9311 | 260/337(77%) | 174/214(81%) | |
| Prochlorococcus marinus str. MIT9312 | 260/337(77%) | 174/214(81%) | |
| Prochlorococcus marinus str. MIT9313 | 265/337(78%) | 178/218(81%) | |
| Prochlorococcus marinus str. MIT9314 | 261/337(77%) | 172/214(80%) | |
| Prochlorococcus marinus str. MIT9515 | 263/337(78%) | 169/214(78%) | |
| Prochlorococcus marinus str. MIT9321 | 262/337(78%) | 173/214(81%) | |
| Prochlorococcus marinus str. MIT9322 | 262/337(78%) | 173/214(81%) | |
| Prochlorococcus marinus str. MIT9401 | 262/337(78%) | 173/214(81%) | |
| Prochlorococcus marinus str. MIT9515 | 263/337(78%) | 169/214(79%) | |
| Prochlorococcus marinus str. MIT0601 | 260/337(77%) | 172/214(80%) | |
| Prochlorococcus marinus str. MIT0602 | 259/338(77%) | 171/218(78%) | |
| Prochlorococcus marinus str. MIT0603 | 259/338(77%) | 171/218(78%) | |
| Prochlorococcus marinus str. MIT0604 | 262/337(78%) | 172/214(80%) | |
| Prochlorococcus marinus str. MIT0701 | 267/337(80%) | 177/220(80%) | |
| Prochlorococcus marinus str. MIT0702 | 268/337(80%) | 177/220(80%) | |
| Prochlorococcus marinus str. MIT0703 | 268/337(80%) | 177/220(80%) | |
| Prochlorococcus marinus str. MIT0801 | 259/334(78%) | 174/222(78%) | |
| Prochlorococcus marinus str. NATL1A | 258/334(77%) | 174/222(78%) | |
| Prochlorococcus marinus str. NATL2A | 257/334(76%) | 174/222(78%) | |
| Prochlorococcus marinus str. PAC1 | 257/334(77%) | 174/222(78%) | |
| Prochlorococcus marinus str. SB | 260/337(77%) | 173/214(81%) | |
| Prochlorococcus marinus str. SS2 | 263/337(78%) | 169/218(78%) | |
| Prochlorococcus marinus str. SS35 | 263/337(78%) | 169/218(78%) | |
| Prochlorococcus marinus str. SS51 | 263/337(78%) | 169/218(78%) | |
| Prochlorococcus marinus str. SS52 | 263/337(78%) | 169/218(78%) | |
| Prochlorococcus marinus str. UH18301 | 260/337(77%) | 172/214(80%) | |
| Pseudanabaena sp. PCC 7367 | 285/339(84%) | 201/226(88%) | |
| Raphidiopsis brookii D9 | 299/338(88%) | 194/221(87%) | |
| Rivularia sp. PCC 7116 | 294/338(86%) | 194/228(85%) | |
| Stanieria cyanosphaera PCC7437 | 1,897/2,788(68%) | ||
| Synechococcus elongatus | 279/337(82%) | 195/231(84%) | |
| Synechococcus sp. BL107 | 259/337(76%) | 172/210(81%) | |
| Synechococcus sp. CC9311 | 265/339(78%) | 181/219(82%) | |
| Synechococcus sp. CC9605 | 266/337(78%) | 171/210(81%) | |
| Synechococcus sp. CC9902 | 259/337(76%) | 172/210(81%) | |
| Synechococcus sp. JA-2–3B'a (2–13) | 268/338(79%) | 179/221(80%) | |
| Synechococcus sp. JA-3–3Ab | 268/338(79%) | 180/221(81%) | |
| Synechococcus sp. PCC6312 | 294/338(86%) | 188/221(85%) | |
| Synechococcus sp. PCC7002 | 2,720/2,720(100%) | ||
| Synechococcus sp. PCC7335 | 285/339(84%) | 196/231(84%) | |
| Synechococcus sp. PCC7502 | 283/339(83%) | 188/222(84%) | |
| Synechococcus sp. RCC307 | 265/337(78%) | 182/220(82%) | |
| Synechococcus sp. RS9916 | 263/337(78%) | 173/210(82%) | |
| Synechococcus sp. RS9917 | 262/337(77%) | 176/210(83%) | |
| Synechococcus sp. WH5701 | 264/334(79%) | 182/220(82%) | |
| Synechococcus sp. WH7803 | 264/337(78%) | 180/219(82%) | |
| Synechococcus sp. WH7805 | 265/337(78%) | 175/210(83%) | |
| Synechococcus sp. WH8102 | 263/337(78%) | 174/210(82%) | |
| Synechococcus sp. WH8109 | 265/337(78%) | 174/210(82%) | |
| Synechocystis sp. PCC6803 | 340/340(100%) | 231/231(100%) | |
| Thermosynechococcus elongatus BP-1 | 290/338(85%) | 186/221(84%) | |
| Trichodesmium erythraeum IMS101 | 297/338(87%) | 192/220(87%) |
The Synechocystis sp. PCC 6803 Fad/Far and Synechococcus sp. PCC 7002 Ols genes were subjected to BLAST analysis against 115 sequenced cyanobacterial genomes. The positive values of these BLAST results are listed. Only matches greater than 30% identity over the length of the query sequence are shown.
Table S2.
Hydrocarbon amounts from GC-MS analysis, dry cell weights, and cell counts for each sample
| Species | Pentadecane per pellet, µg | Heptadecane per pellet, µg | 8-heptadecene per pellet, µg | Total hydrocarbons per pellet, µg | Dry cell weight per pellet, mg | Cells per pellet (×109) |
| Prochlorococcus marinus str. CCMP1986 | 29.042 ± 13.354 | 1.147 ± 0.522 | n.d. | 30.187 ± 13.876 | 7.933 ± 1.745 | 65.383 ± 13.415 |
| Prochlorococcus marinus str. MIT9312 | 15.509 ± 14.795 | 0.595 ± 0.508 | n.d. | 16.104 ± 15.303 | 9.452 ± 1.899 | 48.57 ± 4.045 |
| Prochlorococcus marinus str. MIT9313 | 10.339 ± 6.404 | 0.403 ± 0.162 | n.d. | 10.742 ± 6.566 | 7.082 ± 0.811 | 16.51 ± 2.923 |
| Synechococcus sp. WH5701 | 0.152 ± 0.036 | 1.846 ± 0.381 | 2.073 ± 0.409 | 4.071 ± 0.826 | 2.967 ± 0.681 | 1.573 ± 0.084 |
| Synechococcus sp. WH7803 | 0.434 ± 0.148 | n.d. | 0.039 ± 0.002 | 0.473 ± 0.150 | 1.433 ± 0.115 | 1.098 ± 0.032 |
| Synechococcus sp. WH7805 | 0.394 ± 0.094 | n.d. | 0.104 ± 0.015 | 0.498 ± 0.109 | 2.1 ± 0.1 | 1.598 ± 0.042 |
| Synechococcus sp. WH8102 | 0.293 ± 0.052 | n.d. | 0.048 ± 0.004 | 0.341 ± 0.056 | 1.533 ± 0.153 | 1.123 ± 0.038 |
Values are from three biological replicates. n.d., not detected.
Prochlorococcus and Synechococcus Accumulate Predominantly Heptadecane and Pentadecane.
Next, we measured the hydrocarbon contents of cultured Prochlorococcus and marine Synechococcus cells via gas chromatography-mass spectrometry (GC-MS). Hydrocarbon content has been quantified in a wide range of freshwater and terrestrial cyanobacteria, all of which produced either alkanes or alkenes, ranging between 0.024 and 0.262% of dry cell weight (19). However, the hydrocarbon content of Prochlorococcus and marine Synechococcus species has not been quantified. Our analysis included axenic cultures of three Prochlorococcus strains: CCMP1986 (MED4) and MIT9312—both high-light-adapted strains, representative of the most numerically abundant Prochlorococcus ecotypes (21)—and one low-light-adapted strain, Prochlorococcus MIT9313. Axenic cultures of three diverse marine Synechococcus strains, WH7803, WH7805, and WH8102 (16), and one estuarine strain, WH5701, were also examined.
In all Prochlorococcus strains examined, pentadecane was the dominant hydrocarbon, ∼96% of the total, with the remainder consisting of heptadecane (Table 1 and Fig. S1). Pentadecane was previously identified as the dominant hydrocarbon in Prochlorococcus CCMP1986, although the presence of heptadecane was not observed (12). Total hydrocarbon contents were between 0.350 and 0.711 fg per cell or 0.149–0.368% of dry cell weight (Table 1). Pentadecane was also the dominant hydrocarbon in Synechococcus sp. WH7803, WH7805, and WH8102, ranging between 79 and 92% of total hydrocarbons (Table 1 and Fig. S1). The remainder consisted of 8-heptadecene. The hydrocarbon composition of Prochlorococcus and marine Synechococcus species differs from that of other alkane-producing cyanobacteria, where heptadecane was the dominant hydrocarbon (19). The hydrocarbon composition of Synechococcus sp. WH5701 was significantly different from the marine strains, consisting of 3.7% pentadecane, 45.3% heptadecane, and 50.9% 8-heptadecene (Fig. S1), which may reflect its evolutionary distance from the other Synechococcus strains examined here (22). Total hydrocarbons in Synechococcus were between 0.304 and 2.580 fg per cell or 0.022–0.138% of dry cell weight (Table 1). With the exception of Prochlorococcus CCMP1986, which has the highest hydrocarbon yields so far observed in a cyanobacterium, hydrocarbon contents were within the range previously observed for other cyanobacteria (19).
Table 1.
Hydrocarbon production by Prochlorococcus and Synechococcus
| Strain | Pentadecane, fg per cell | Heptadecane, fg per cell | 8-heptadecene, fg per cell | Total hydrocarbons, fg per cell | Total hydrocarbons, % dry cell weight | Hydrocarbons in global cellular population, million tons | Hydrocarbon production, million tons per year |
| Prochlorococcus str. CCMP1986 | 0.448 ± 0.223 | 0.018 ± 0.009 | n.d. | 0.466 ± 0.232 | 0.368 ± 0.126 | ||
| Prochlorococcus str. MIT9312 | 0.337 ± 0.343 | 0.013 ± 0.012 | n.d. | 0.350 ± 0.355 | 0.181 ± 0.187 | ||
| Prochlorococcus str. MIT9313 | 0.685 ± 0.548 | 0.026 ± 0.014 | n.d. | 0.711 ± 0.562 | 0.149 ± 0.080 | ||
| Mean Prochlorococcus | 0.509 ± 0.383 | 1.48 | 269–539 | ||||
| Synechococcus sp. WH5701 | 0.096 ± 0.018 | 1.170 ± 0.211 | 1.314 ± 0.225 | 2.580 ± 0.454 | 0.138 ± 0.009 | ||
| Synechococcus sp. WH7803 | 0.396 ± 0.136 | n.d. | 0.035 ± 0.001 | 0.431 ± 0.137 | 0.033 ± 0.008 | ||
| Synechococcus sp. WH7805 | 0.248 ± 0.065 | n.d. | 0.065 ± 0.009 | 0.313 ± 0.074 | 0.024 ± 0.005 | ||
| Synechococcus sp. WH8102 | 0.261 ± 0.047 | n.d. | 0.043 ± 0.005 | 0.304 ± 0.052 | 0.022 ± 0.002 | ||
| Mean Synechococcus | 0.907 ± 0.179 | 0.635 | 39–232 |
Cellular hydrocarbon amounts were quantified by GC-MS. Measurements are from three biological replicates. SD is indicated. Total hydrocarbons per cell were quantified by dividing the total hydrocarbon mass by the total number of cells in the pellet (Table S2). The amount of hydrocarbons as a percentage of dry cell weight was quantified by dividing the total hydrocarbon mass by the pellet cell mass (Table S2). The average mass of hydrocarbons per cell was used to quantify hydrocarbon amounts in global ocean cell populations given global population size estimates of 2.9 ± 0.1 × 1027 Prochlorococcus and 7.0 ± 0.3 × 1026 Synechococcus cells (14). This total and the turnover rate were used to quantify the annual production amount. n.d., not detected.
Fig. S1.
Chromatograms showing separation of hydrocarbons extracted from Synechococcus species (WH8103, WH7803, WH7805, and WH5701) and Prochlorococcus species (CCMP1986, MIT9312, and MIT9313) by GC-MS (Thermo Scientific Trace GC 1310 – ISQ LT Single Quadruple EI MS, A1-1310 Autosampler) using a Thermo TG-SQC GC column (15 m × 0.25 mm, 0.25-µm film thickness). Peaks were identified as A, (10.25 min) pentadecane; B, (12.16 min) 8-heptadecene; C, (12.26 min) 8-heptadecene; and D, (12.45 min) heptadecane. Samples are shown from a single replicate. The chemical identity of the peak at 11.22 min in the Synechococcus species could not be resolved by MS so was excluded from further analysis.
Significant Potential for Hydrocarbon Production by Prochlorococcus and Synechococcus in the Oceans.
Given the average measured abundances of hydrocarbons in Prochlorococcus and Synechococcus cells and their population sizes, we estimate that, at any given point in time, these cyanobacteria represent a pool of 2.12 million tons of alkanes in the oceans (Table 1). Although total population sizes of Prochlorococcus and Synechococcus remain largely stable on an annual timescale, their turnover rates are high. Prochlorococcus divide once every 1–2 d (23–25), with cellular losses balancing division in a quasi-steady-state manner. Cyanobacterial mortality can be mediated by a variety of factors, including predation by grazers or viruses, UV-induced lysis, or spontaneous cell death (18), resulting in release of organic carbon compounds, including alkanes, into the environment. At these cellular turnover rates, production of alkanes by Prochlorococcus globally is estimated to be between 269 and 539 million tons per annum (Table 1). Synechococcus maximum growth rates are comparable to or slighter faster than those of Prochlorococcus (25, 26), with similar mechanisms of mortality (27, 28). However, estimates of in situ specific growth rates are broader, between 1–6 d (29, 30), due to the varying environments Synechococcus occupies. Therefore, total annual production of hydrocarbons by Synechococcus could vary between 39 and 232 million tons per annum (Table 1).
Natural oil seepage from within marine sediments is estimated to be between 0.2 and 2 million tons per annum and is geographically concentrated at the continental margins (1). An equivalent release of hydrocarbons into the ocean is thought to result from human activities such as oil leakage from drilling rigs and shipping vessels (1). Localized events such as the Deepwater Horizon oil spill, which released ∼0.435 million tons of oil (7), can further increase anthropogenic hydrocarbon inputs into the marine environment. In contrast, the 308–771 million tons of hydrocarbons produced annually by Prochlorococcus and Synechococcus significantly exceeds the inputs from these other natural and anthropogenic sources and, due to the broad distribution of cyanobacteria across the world’s surface oceans, is more widespread (14). As mentioned above, measurements of pentadecane and heptadecane in marine surface waters indicate that they are present only at femtogram- to picogram-per-mL levels (2, 4, 31). From our analyses, we estimate that the amounts of hydrocarbons produced by marine cyanobacteria to be on the order of 2–540 pg⋅mL−1⋅d−1 (Materials and Methods), indicating that the majority of pentadecane and heptadecane in surface waters may derive from cyanobacteria and that these compounds do not accumulate significantly; hence, the continuous production of cyanobacterial hydrocarbons must be balanced by degradation or loss. Some hydrocarbon exchange occurs between the ocean and the atmosphere (2, 32), but not at rates sufficient to balance estimated cyanobacterial production rates. Given that the temperature-dependent half-life of pentadecane and heptadecane is between 0.8 and 5 d at surface ocean temperatures, a significant fraction of the hydrocarbons will likely escape these abiotic loss pathways (31). Therefore, biological degradation is likely to be responsible for the majority of turnover (31).
Both Obligate and Facultative Hydrocarbon-Degrading Bacteria Can Metabolize Cyanobacterial Alkanes.
Alkane degradation pathways have not been identified in grazers of cyanobacteria (33). Therefore, the majority of cyanobacterially produced hydrocarbons are likely to be released into the environment and subsequently degraded by bacteria (34). Obligate hydrocarbon-degrading bacteria, including Cycloclasticus, Thalassolituus, Oleiphilus, Oleispira, and Alcanivorax species, have been isolated from geographically diverse coastal and open-ocean regions in all seas, in sediments and both surface and deep waters, including areas with minimal oil pollution (9–11, 35). Alcanivorax species are typically among the dominant bacteria found metabolizing crude oil during large spill events (34–38). Other bacteria, including Marinobacter, Pseudomonas, and Acinetobacter species, which can use hydrocarbons in addition to a broader range of carbon sources, have also been detected in oil-polluted ocean samples (10, 34).
To determine whether cyanobacterial alkanes can support the growth of both obligate as well as nonobligate hydrocarbon-degrading bacteria, we tested the ability of three axenic and well-characterized oil degraders (5, 8) to grow on either heptadecane or crude oil: Alcanivorax borkumensis SK2, an obligate hydrocarbon-degrading bacterium (39), and Acinetobacter baylyi ADP1 and Marinobacter aquaolei VT8, two strains that can facultatively grow on crude oil. We observed that all three strains could grow in media containing either oil or heptadecane as the only source of reduced carbon (Fig. 1). These findings suggest that diverse groups of marine hydrocarbon degraders may contribute to the degradation of the hydrocarbons produced by Prochlorococcus and Synechococcus.
Fig. 1.
Hydrocarbon-degrading bacteria can grow on both oil and heptadecane. Growth of (A) Alcanivorax borkumensis SK2, (B) Acinetobacter baylyi ADP1, and (C) Marinobacter aquaolei VT8 in ASW medium (triangles), ASW medium with heptadecane (7.77 mg/mL) (squares), and ASW medium with oil (circles). Values represent the mean and SD from three biological replicates. The growth rate constants (µ) on oil and heptadecane were, respectively, 0.12 and 0.12 h−1 for Alcanivorax, 0.019 and 0.006 h−1 for Acinetobacter, and 0.010 and 0.004 h−1 for Marinobacter.
Cyanobacterial Hydrocarbon Production Can Support Populations of A. borkumensis SK2.
A. borkumensis SK2 has previously been demonstrated to use a broad range of alkanes, with similar growth rates observed when either pentadecane or heptadecane was added as the sole carbon source (40). However, in both this study and the previous report (40) an excess of alkanes, similar to concentrations in oil spills, was used (41). Whether cyanobacterial alkane production is sufficient to support hydrocarbon-degrading bacterial populations in minimally polluted seawater has not been determined.
Information on population sizes of specific hydrocarbon degraders in the environment is limited, with estimates of Alcanivorax varying between 101 and 5 × 103 cells/mL (35, 41). However, in most studies Alcanivorax cell numbers are below the detection limit (34), as assayed via quantitative PCR (5, 6) or in situ hybridization (36), and their population dynamics in the oligotrophic ocean are unknown. To determine the yield of A. borkumensis SK2 on cyanobacterial alkanes, cells were grown in artificial sea water (ASW) medium either without any organic carbon or containing only the minimum amount of heptadecane that could be added to the culture. A small increase in cell number was observed in cultures containing heptadecane, before declining to the original cell number by day 3 (Fig. 2). The addition of heptadecane to this culture sustained cell numbers for a further 3 d, although a smaller increase in population size was observed compared with the initial growth period, suggesting that some other component of the medium became limiting. Based on these data, we find that the yield of A. borkumensis SK2 is 0.1 pg dry cell weight per pg heptadecane (Materials and Methods)—lower than other yields previously reported for other hydrocarbon-degrading bacteria, which range from 0.5 to 1.77 pg/pg hydrocarbon (42). Thus, we expect that a marine cyanobacterial hydrocarbon production rate of 2–540 pg−1⋅mL⋅d−1 could support a population of 1.5 × 101 to 4.1 × 103 A. borkumensis SK2 cells per mL if used as the only source of reduced carbon, comparable to the population of obligate hydrocarbon-degrading bacteria observed in natural environments.
Fig. 2.
Marine heptadecane concentrations support the growth of A. borkumensis SK2. Growth of A. borkumensis SK2 was quantified in ASW medium (triangles) or ASW medium supplemented with 40 µg/mL heptadecane (squares and circles). The cultures containing heptadecane were split and an additional 40 µg/mL heptadecane added to one set at day 3 (squares). Results are from three biological replicates. SD is indicated. Control cultures containing only ASW medium or ASW medium and heptadecane (i.e., without added bacteria) showed no evidence of cell contamination.
In the cultures to which no heptadecane was added, cell numbers decreased significantly over the course of the experiment. This suggests that populations of Alcanivorax, and possibly other obligate hydrocarbon-degrading bacteria in the oceans, may be unable to persist for long in the absence of a constant hydrocarbon supply. Given the widespread distribution of both Prochlorococcus and marine Synechococcus, it is highly likely that hydrocarbon production is continuous and therefore hydrocarbon-degrading bacteria are constantly supplied with a fresh source of alkanes. Although pyruvate is present in the oceans (43, 44), and A. borkumensis SK2 can use it as an energy and carbon source (40), it likely faces greater competition from other heterotrophs for pyruvate than for alkanes (43, 44). Other compounds, predominantly hydrocarbons but possibly other unidentified substrates, may also contribute to the pool available to sustain obligate hydrocarbon-degrading species. These include pristane, a C19 saturated terpenoid alkane produced in significant quantities by some algal species (45), which A. borkumensis SK2 can use (10), and a branched C21 alkane produced by some algae (45).
The Ocean Hydrocarbon Cycle.
Catabolic degradation of hydrocarbons by marine bacteria produces significant amounts of CO2, which can be incorporated back into alkanes within cyanobacteria (33). Based on our estimates of cyanobacterial alkane production rates and observed concentrations in the ocean, we expect that this “short-term hydrocarbon cycle” (46) occurs on the order of days (Fig. 3). Given the significant contribution of Prochlorococcus and Synechococcus to marine carbon flux, the levels of hydrocarbon production we have measured indicate that there must be a notable biogeochemical cycle of these compounds in the world’s upper oceans. In terms of annual hydrocarbon fluxes within the ocean, the short-term hydrocarbon cycle occurs at rates several orders of magnitude greater than the “long-term hydrocarbon cycle,” whereby organic matter is converted to oil in sediments over a period of thousands to millions of years before release into the marine environment by natural seepage or human activity and subsequent breakdown.
Fig. 3.
The hydrocarbon cycle. A schematic representation of the short-term hydrocarbon cycle, which occurs over days, and the long-term hydrocarbon cycle, which takes place over thousands to millions of years. A simplified process showing the conversion of CO2 to alkanes using energy derived from photosynthesis is detailed in the enlarged cyanobacterial cell on the left. A simplified process showing the metabolism of alkanes is detailed in the enlarged hydrocarbon degrading bacterial cell on the right.
The short- and long-term hydrocarbon cycles are linked in that hydrocarbon-degrading bacteria are responsible for removing both cyanobacterial alkanes and crude oil from the environment. Upon release of crude oil, a significant increase in hydrocarbon-degrading bacteria is observed (5–7, 41). Crude oil is a complex mixture of ∼20,000 compounds broadly grouped into four categories: saturated hydrocarbons, predominantly C5–C40 alkanes (40–60%), aromatic hydrocarbons (20–40%), resins (5–20%), and asphaltenes (1–10%) (8, 34). It is notable that after release of crude oil into the environment, saturated hydrocarbons are the first to be degraded by bacteria (7, 9, 34). This suggests that populations of bacteria, which normally degrade cyanobacterial alkanes and other hydrocarbons derived from biological sources, quickly acclimate to metabolize crude oil hydrocarbons, followed by rapid population expansion to use this new energy source.
Although cyanobacterial hydrocarbons are only a small proportion (0.00032%) of the estimated 662 billion tons of dissolved organic carbon (DOC) present in the ocean at any point in time (47), only a relatively small fraction of this bulk DOC (∼0.2 billion tons) is turned over within days (48). Cyanobacterial hydrocarbons, with an estimated pool of 2.12 million tons, likely belong to the labile subset of rapidly cycled DOC and constitute a notable proportion (∼1%) of that bioavailable fraction. The short-term hydrocarbon cycle should therefore be considered a component of the “microbial loop” within marine food webs (49, 50).
Conclusions
This study sets a minimum estimate for the total amount of hydrocarbons produced by cyanobacteria in marine environments. Other abundant marine cyanobacteria, notably Trichodesmium species (51), may add significantly to hydrocarbon output, as might some algal species. Although population sizes of freshwater and terrestrial cyanobacteria are not well constrained, given that these organisms are also capable of producing hydrocarbons (19), alkane and alkene production is likely to be considerable here as well. The widespread distribution of hydrocarbon-degrading bacteria in freshwater, marine, and terrestrial environments suggests that the short-term hydrocarbon cycle is pervasive in many natural ecosystems (9, 11).
Materials and Methods
Bioinformatics.
FASTA BLAST comparisons (52) were performed using inferred protein sequences for Synechocystis sp. PCC6803 sll0209 (FAR) and sll0208 (FAD) and Synechococcus sp. PCC7002 Syn7002_A1173 (Ols) (WP_012306795) with the 115 completed cyanobacterial genomes listed in the NCBI database (www.ncbi.nlm.nih.gov/genome/browse/) and Biller et al. (20). Synechococcus sp. PCC7002 was not classified as a marine species because it was isolated from mud flats in Puerto Rico, is evolutionarily distant from other Synechococcus species (16, 22), and has not been detected in the open ocean.
Bacterial Strains, Media, and Growth Conditions.
Axenic cultures of Prochlorococcus CCMP1986 (MED4), MIT9312, and MIT9313 were used. Cultures were routinely assessed for purity by confirming a lack of turbidity after inoculation into three different purity test broths (53). Triplicate 2-L cultures of each strain were grown in Pro99 medium (54) prepared with 0.2 µm filtered, autoclaved seawater (collected from Vineyard Sound, MA) and supplemented with 10 mM filter-sterilized sodium bicarbonate upon inoculation. Cells were grown under constant light flux (30–40 µmol photons⋅m−2⋅s−1 for CCMP1986 and MIT9312; 10–20 µmol photons⋅m−2⋅s−1 for MIT9313) at 24 °C, in acid-washed polycarbonate containers. Growth was monitored by measuring bulk culture fluorescence using a 10-AU fluorometer (Turner Designs).
All Synechococcus species were grown in ASW medium (55). Synechococcus sp. WH5701 was cultured in conical flasks at 30 °C and 40 µmol photons⋅m−2⋅s−1 with shaking at 160 rpm. Synechococcus spp. WH7803, WH7805, and WH8102 were cultured in culture flasks at 24 °C and 40 µmol photons⋅m−2⋅s−1 without shaking. Cultures were routinely assessed for purity by plating aliquots on ASW solid medium containing 0.8% wt/vol yeast extract and 1.5% wt/vol agar.
All hydrocarbon degrading bacteria were cultured at 30 °C with shaking at 160 rpm. Acinetobacter baylyi ADP1 (56) was routinely grown in BHI medium. Alcanivorax borkumensis SK2 (39) was routinely grown in A. borkumensis medium 809 (DSMZ). Marinobacter aquaolei VT8 (57) was routinely grown in Marinobacter medium 970 (DSMZ). To test growth on hydrocarbons cultures were grown to stationary phase and washed twice with ASW medium before being cultured in 10 mL ASW medium with no carbon source or with either 1% (vol/vol) crude oil (Nigerian bonny light crude oil) or filter-sterilized, analytical standard-grade (≥99.5%) heptadecane (100 µL equivalent to 77.7 mg) (Sigma). Samples of ASW medium, ASW medium and crude oil, and ASW medium and heptadecane were also set up as negative controls. Growth was determined by measuring the optical density at 600 nm, and specific growth rate constants (µ) were calculated during exponential phase (21–45 h for A. borkumensis, 21–71 h for A. baylyi, and 21–138 h for M. aquaolei). Three biological replicates of each sample were measured. Cultures were routinely assessed for purity by plating aliquots on ASW solid medium containing 0.8% wt/vol yeast extract and 1.5% wt/vol agar and on either BHI medium, A. borkumensis medium 809, or Marinobacter medium 970 with 1.5% wt/vol agar for A. baylyi ADP1, A. borkumensis SK2, and M. aquaolei VT8 cultures, respectively.
For growth of A. borkumensis SK2 in 40 µg/mL heptadecane, 4 × 107 cells per mL were inoculated so that hydrocarbon amounts were equivalent to 1 pg of heptadecane per cell at the time of inoculation. This was necessary because a heptadecane concentration of 40 µg/mL, equivalent to 0.5 µL per 10 mL, was the minimum amount that could be used in this experiment, without diluting heptadecane in another similar solvent. These could be either used by A. borkumensis SK2 as an energy and carbon source or demonstrate high cellular toxicity or growth inhibition, which has been shown with alkanols (40). The higher inoculum population size was also essential to accurately count cell numbers using a Beckman Coulter 2Z particle counter. Three biological replicates of each sample were measured.
The dry cell weight of A. borkumensis SK2 was determined by culturing three biological replicates in ASW medium and heptadecane (100 µL in 10 mL) to late exponential phase. Samples were washed three times with water and cell counts per milliliter determined using a Beckman Coulter 2Z particle counter. The dry cell weight was determined by first washing 10 mL of culture three times with water, filtering the solution via Whatman glass microfiber filters (GE Healthcare), and drying the samples before measurement. Dry cell weights were recorded using an Adam PW 214 analytical balance. A mass of 0.39 ± 0.042 pg per A. borkumensis SK2 cell was calculated via dividing the dry cell weight by the cell count. To calculate the growth yield the increase in cell population of 1.01 × 107 observed in the first 2 d in media containing 40 µg/mL of heptadecane was multiplied by the average mass of a cell. This amount was then divided by the mass of heptadecane per mL, resulting in a growth yield of 0.1 pg dry cell weight per pg of heptadecane.
Cell Counting and Sample Preparation for GC-MS Analysis.
Prochlorococcus cultures were harvested in exponential phase by centrifugation at 15,000 × g for 15 min in a JLA-8.1000 rotor (Beckman Coulter) at 4 °C. Pellets were resuspended in fresh Pro99 medium and transferred into tared glass vials and lyophilized for 48 h. Total Prochlorococcus cells in each sample were enumerated by flow cytometry using an Influx Cell Sorter (BD Biosciences) as previously described (58, 59).
Synechococcus samples were harvested in exponential phase (Table S1). An aliquot was removed and cells counted by flow cytometry using a BD FACScan. Cell counting was performed by running 12 µL of sample per minute through the FACS device until 100,000 particles were counted. The background particle count from the media (∼200 counts per second) was subtracted from the total count. Cell counts were performed in triplicate for each sample. Fifteen milliliters of each sample was centrifuged at 2,800 × g, washed twice with water, and freeze-dried using a ScanVac CoolSafe Freeze Dryer. Dry cell weights were recorded using an Adam PW 214 analytical balance.
Extraction and Analysis of Total Hydrocarbons.
All chemicals were purchased from Sigma chemicals. To extract total hydrocarbons, dichloromethane (1.5 mL for Prochlorococcus and 1 mL for Synechococcus) was added to pelleted dried cells in glass vials. Samples were placed in a sonicator bath for 30 min then centrifuged (GeneVac EZ-2; SP Scientific) for 10 min (1,800 × g) to pellet any remaining material. The supernatant was transferred to a glass GC sample vial and stored at −80 °C. The pellets were extracted twice more in dichloromethane as above to ensure complete extraction of hydrocarbons (80–90% of hydrocarbons were extracted in the first extraction). For negative controls, extraction blanks were carried out on growth media without cyanobacteria and positive controls consisted of adding 1 mg/mL standard alkane mix (C8-C20 alkane mix; Sigma) to the extraction procedure. Three biological replicates of each sample were analyzed.
Identification and Quantification of Hydrocarbons.
Hydrocarbons were identified by GC-MS (Thermo Scientific Trace GC 1310 – ISQ LT Single Quadruple EI MS, A1-1310 Auto sampler) with a Thermo TG-SQC GC column (15 m × 0.25 mm, 0.25-µm film thickness). The injection volume was 1 µL with a 10:1 split ratio with an injector temperature of 230 °C, using helium as a carrier gas at a constant flow of 1.2 mL⋅min−1. The following gradient was used: initial oven temperature 30 °C, 2 min; 150 °C at 15 °C⋅min−1; 230 °C at 3.4 °C⋅min−1. A transfer line temperature of 240 °C was used. The MS conditions in the positive mode were as follows: ion source, 250 °C; mass range 45–650 Da; scan time of 0.35 s. Pentadecane and heptadecane were identified by co-retention with standards and NIST mass spectral search libraries (National Institute of Standards and Technology NIST v2.0); two peaks were identified as 8-heptadecene using the NIST library alone. Pentadecane and heptadecane were quantified using standard curves derived from peak areas of pentadecane and heptadecane alkane standards (0.06–31 µg/mL) and 8-heptadecene was quantified using peak areas derived from heptadecane standards (0.06–31 µg/mL). Any background signal in the extraction blank was subtracted when determining hydrocarbon amounts.
Calculations.
Amounts of hydrocarbons produced by Prochlorococcus and Synechococcus per mL of seawater were calculated by multiplying the numbers in the ocean, which range from 3 × 103 to 1 × 106 cells per mL and 3 × 103 to 5 × 104 cells per mL, respectively (14), at the sea surface, by the average amount of hydrocarbons per cell (Table 1). This equates to 1.5–509 pg/mL of hydrocarbons produced every 1–2 d by Prochlorococcus and 1.9–31.7 pg/mL of hydrocarbons produced every 1–6 d by Synechococcus.
Acknowledgments
We thank Katherine Helliwell and Alexandra Jamieson of the A.G.S. laboratory for assistance with culturing of Synechococcus and hydrocarbon analysis, respectively; Nigel Millar (Department of Pathology, University of Cambridge) for flow cytometry assistance; and Paul Berube and Jamie Becker (Massachusetts Institute of Technology) for helpful discussions. This work was supported by the Environmental Services Association Educational Trust (D.J.L-S.), European Research Council (Starting grant to A.V.T.), National Science Foundation Grant OCE-1356460 and Gordon and Betty Moore Foundation Grant GMBF495 (to S.W.C.), and the Simons Foundation SCOPE project (S.W.C.). This work was also funded in part by a University of Warwick International Chancellors Scholarship (to B.P.S.) and by a Natural Environment Research Council Grant NE/I00985X/1(to D.J.S.).
Footnotes
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
See Commentary on page 13434.
This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1507274112/-/DCSupplemental.
References
- 1.Kvenvolden KA, Cooper CK. Natural seepage of crude oil into the marine environment. Geo-Mar Lett. 2003;23(3-4):140–146. [Google Scholar]
- 2.Schwarzenbach RP, Bromund RH, Gschwend PM, Zafiriou OC. Volatile organic compounds in coastal seawater. Org Geochem. 1978;1:93–107. [Google Scholar]
- 3.Gschwend P, Zafiriou OC, Gagosian RB. Volatile organic-compounds in seawater from the Peru upwelling region. Limnol Oceanogr. 1980;25(6):1044–1053. [Google Scholar]
- 4.Martí S, Bayona JM, Albaigés J. A potential source of organic pollutants into the northeastern Atlantic: The outflow of the Mediterranean deep-lying waters through the Gibraltar Strait. Environ Sci Technol. 2001;35(13):2682–2689. doi: 10.1021/es000258p. [DOI] [PubMed] [Google Scholar]
- 5.Kostka JE, et al. Hydrocarbon-degrading bacteria and the bacterial community response in Gulf of Mexico beach sands impacted by the Deepwater Horizon oil spill. Appl Environ Microbiol. 2011;77(22):7962–7974. doi: 10.1128/AEM.05402-11. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 6.Gutierrez T, et al. Hydrocarbon-degrading bacteria enriched by the Deepwater Horizon oil spill identified by cultivation and DNA-SIP. ISME J. 2013;7(11):2091–2104. doi: 10.1038/ismej.2013.98. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 7.Dubinsky EA, et al. Succession of hydrocarbon-degrading bacteria in the aftermath of the Deepwater Horizon oil spill in the Gulf of Mexico. Environ Sci Technol. 2013;47(19):10860–10867. doi: 10.1021/es401676y. [DOI] [PubMed] [Google Scholar]
- 8.McGenity TJ, Folwell BD, McKew BA, Sanni GO. Marine crude-oil biodegradation: A central role for interspecies interactions. Aquat Biosyst. 2012;8(1):10. doi: 10.1186/2046-9063-8-10. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 9.Leahy JG, Colwell RR. Microbial degradation of hydrocarbons in the environment. Microbiol Rev. 1990;54(3):305–315. doi: 10.1128/mr.54.3.305-315.1990. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 10.Yakimov MM, Timmis KN, Golyshin PN. Obligate oil-degrading marine bacteria. Curr Opin Biotechnol. 2007;18(3):257–266. doi: 10.1016/j.copbio.2007.04.006. [DOI] [PubMed] [Google Scholar]
- 11.Nie Y, et al. Diverse alkane hydroxylase genes in microorganisms and environments. Sci Rep. 2014;4:4968. doi: 10.1038/srep04968. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 12.Schirmer A, Rude MA, Li X, Popova E, del Cardayre SB. Microbial biosynthesis of alkanes. Science. 2010;329(5991):559–562. doi: 10.1126/science.1187936. [DOI] [PubMed] [Google Scholar]
- 13.Mendez-Perez D, Begemann MB, Pfleger BF. Modular synthase-encoding gene involved in α-olefin biosynthesis in Synechococcus sp. strain PCC 7002. Appl Environ Microbiol. 2011;77(12):4264–4267. doi: 10.1128/AEM.00467-11. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 14.Flombaum P, et al. Present and future global distributions of the marine Cyanobacteria Prochlorococcus and Synechococcus. Proc Natl Acad Sci USA. 2013;110(24):9824–9829. doi: 10.1073/pnas.1307701110. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 15.Whitton BA, Potts M. The Ecology of Cyanobacteria: Their Diversity in Time and Space. Kluwer; Dordrecht, The Netherlands: 2000. [Google Scholar]
- 16.Scanlan DJ, et al. Ecological genomics of marine picocyanobacteria. Microbiol Mol Biol Rev. 2009;73(2):249–299. doi: 10.1128/MMBR.00035-08. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 17.Vincent WF. Cyanobacterial dominance in the polar regions. In: Whitton BA, Potts M, editors. The Ecology of Cyanobacteria. Kluwer; Dordrecht, The Netherlands: 2000. pp. 321–340. [Google Scholar]
- 18.Biller SJ, Berube PM, Lindell D, Chisholm SW. Prochlorococcus: The structure and function of collective diversity. Nat Rev Microbiol. 2015;13(1):13–27. doi: 10.1038/nrmicro3378. [DOI] [PubMed] [Google Scholar]
- 19.Coates RC, et al. Characterization of cyanobacterial hydrocarbon composition and distribution of biosynthetic pathways. PLoS One. 2014;9(1):e85140. doi: 10.1371/journal.pone.0085140. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 20.Biller SJ, et al. 2014. Genomes of diverse isolates of the marine cyanobacterium Prochlorococcus. Scientific Data 1:140034.
- 21.Johnson ZI, et al. Niche partitioning among Prochlorococcus ecotypes along ocean-scale environmental gradients. Science. 2006;311(5768):1737–1740. doi: 10.1126/science.1118052. [DOI] [PubMed] [Google Scholar]
- 22.Shih PM, et al. Improving the coverage of the cyanobacterial phylum using diversity-driven genome sequencing. Proc Natl Acad Sci USA. 2013;110(3):1053–1058. doi: 10.1073/pnas.1217107110. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 23.Vaulot D, Marie D, Olson RJ, Chisholm SW. Growth of Prochlorococcus, a photosynthetic prokaryote, in the equatorial Pacific Ocean. Science. 1995;268(5216):1480–1482. doi: 10.1126/science.268.5216.1480. [DOI] [PubMed] [Google Scholar]
- 24.Mann EL, Chisholm SW. Iron limits the cell division rate of Prochlorococcus in the eastern equatorial Pacific. Limnol Oceanogr. 2000;45(5):1067–1076. [Google Scholar]
- 25.Zubkov MV. Faster growth of the major prokaryotic versus eukaryotic CO2 fixers in the oligotrophic ocean. Nat Commun. 2014;5:3776. doi: 10.1038/ncomms4776. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 26.Moore LR, Goericke R, Chisholm SW. Comparative physiology of Synechococcus and Prochlorococcus - Influence of light and temperature on growth, pigments, fluorescence and absorptive properties. Mar Ecol Prog Ser. 1995;116(1–3):259–275. [Google Scholar]
- 27.Wang K, Wommack KE, Chen F. Abundance and distribution of Synechococcus spp. and cyanophages in the Chesapeake Bay. Appl Environ Microbiol. 2011;77(21):7459–7468. doi: 10.1128/AEM.00267-11. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 28.Llabres M, Agusti S, Alonso-Laita P, Herndl GJ. Synechococcus and Prochlorococcus cell death induced by UV radiation and the penetration of lethal UVR in the Mediterranean Sea. Mar Ecol Prog Ser. 2010;399:27–37. [Google Scholar]
- 29.Field CB, Behrenfeld MJ, Randerson JT, Falkowski P. Primary production of the biosphere: Integrating terrestrial and oceanic components. Science. 1998;281(5374):237–240. doi: 10.1126/science.281.5374.237. [DOI] [PubMed] [Google Scholar]
- 30.Liu HB, Campbell L, Landry MR. Growth and mortality rates of Prochlorococcus and Synechococcus measured with a selective inhibitor technique. Mar Ecol Prog Ser. 1995;116(1–3):277–287. [Google Scholar]
- 31.Wakeham SG, Davis AC, Karas JA. Mesocosm experiments to determine the fate and persistence of volatile organic compounds in coastal seawater. Environ Sci Technol. 1983;17(10):611–617. doi: 10.1021/es00116a009. [DOI] [PubMed] [Google Scholar]
- 32.Duce RA, Gagosian RB. The input of atmospheric N-C10 to N-C30 alkanes to the ocean. J Geophys Res. 1982;87(Nc9):7192–7200. [Google Scholar]
- 33.Rojo F. Degradation of alkanes by bacteria. Environ Microbiol. 2009;11(10):2477–2490. doi: 10.1111/j.1462-2920.2009.01948.x. [DOI] [PubMed] [Google Scholar]
- 34.Head IM, Jones DM, Röling WFM. Marine microorganisms make a meal of oil. Nat Rev Microbiol. 2006;4(3):173–182. doi: 10.1038/nrmicro1348. [DOI] [PubMed] [Google Scholar]
- 35.Hara A, Syutsubo K, Harayama S. Alcanivorax which prevails in oil-contaminated seawater exhibits broad substrate specificity for alkane degradation. Environ Microbiol. 2003;5(9):746–753. doi: 10.1046/j.1468-2920.2003.00468.x. [DOI] [PubMed] [Google Scholar]
- 36.Syutsubo K, Kishira H, Harayama S. Development of specific oligonucleotide probes for the identification and in situ detection of hydrocarbon-degrading Alcanivorax strains. Environ Microbiol. 2001;3(6):371–379. doi: 10.1046/j.1462-2920.2001.00204.x. [DOI] [PubMed] [Google Scholar]
- 37.Harayama S, Kishira H, Kasai Y, Shutsubo K. Petroleum biodegradation in marine environments. J Mol Microbiol Biotechnol. 1999;1(1):63–70. [PubMed] [Google Scholar]
- 38.Kasai Y, et al. Predominant growth of Alcanivorax strains in oil-contaminated and nutrient-supplemented sea water. Environ Microbiol. 2002;4(3):141–147. doi: 10.1046/j.1462-2920.2002.00275.x. [DOI] [PubMed] [Google Scholar]
- 39.Schneiker S, et al. Genome sequence of the ubiquitous hydrocarbon-degrading marine bacterium Alcanivorax borkumensis. Nat Biotechnol. 2006;24(8):997–1004. doi: 10.1038/nbt1232. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 40.Naether DJ, et al. Adaptation of the hydrocarbonoclastic bacterium Alcanivorax borkumensis SK2 to alkanes and toxic organic compounds: A physiological and transcriptomic approach. Appl Environ Microbiol. 2013;79(14):4282–4293. doi: 10.1128/AEM.00694-13. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 41.Coulon F, McKew BA, Osborn AM, McGenity TJ, Timmis KN. Effects of temperature and biostimulation on oil-degrading microbial communities in temperate estuarine waters. Environ Microbiol. 2007;9(1):177–186. doi: 10.1111/j.1462-2920.2006.01126.x. [DOI] [PubMed] [Google Scholar]
- 42.Widdel F, Musat F. Energetic and other quantitative aspects of microbial hydrocarbon utilization. In: Timmis KN, editor. Handbook of Hydrocarbon and Lipid Microbiology. Springer; Berlin: 2010. pp. 732–764. [Google Scholar]
- 43.Obernosterer I, Kraay G, de Ranitz E, Herndl GJ. Concentrations of low molecular weight carboxylic acids and carbonyl compounds in the Aegean Sea (Eastern Mediterranean) and the turnover of pyruvate. Aquat Microb Ecol. 1999;20(2):147–156. [Google Scholar]
- 44.Kieber DJ, Mcdaniel J, Mopper K. Photochemical source of biological substrates in seawater: Implications for carbon cycling. Nature. 1989;341(6243):637–639. [Google Scholar]
- 45.Blumer M, Guillard RR, Chase T. Hydrocarbons of marine phytoplankton. Mar Biol. 1971;8(3):183–189. [Google Scholar]
- 46.Gibson DT. Biodegradation of aromatic petroleum hydrocarbons. In: Wolfe DA, editor. Fate and Effects of Petroleum Hydrocarbons in Marine Organisms and Ecosystems. Pergamon; Oxford: 1977. pp. 36–46. [Google Scholar]
- 47.Hansell DA, Carlson CA, Repeta DJ, Schlitzer R. Dissolved organic matter in the ocean a controversy stimulates new insights. Oceanography (Wash DC) 2009;22(4):202–211. [Google Scholar]
- 48.Hansell DA. Recalcitrant dissolved organic carbon fractions. Annu Rev Mar Sci. 2013;5(5):421–445. doi: 10.1146/annurev-marine-120710-100757. [DOI] [PubMed] [Google Scholar]
- 49.Pomeroy LR, Williams PJI, Azam F, Hobbie JE. The microbial loop. Oceanography (Wash DC) 2007;20(2):28–33. [Google Scholar]
- 50.Azam F, et al. The ecological role of water-column microbes in the sea. Mar Ecol Prog Ser. 1983;10(3):257–263. [Google Scholar]
- 51.Capone DG, Zehr JP, Paerl HW, Bergman B, Carpenter EJ. Trichodesmium, a globally significant marine cyanobacterium. Science. 1997;276(5316):1221–1229. [Google Scholar]
- 52.Altschul SF, Gish W, Miller W, Myers EW, Lipman DJ. Basic local alignment search tool. J Mol Biol. 1990;215(3):403–410. doi: 10.1016/S0022-2836(05)80360-2. [DOI] [PubMed] [Google Scholar]
- 53.Berube PM, et al. Physiology and evolution of nitrate acquisition in Prochlorococcus. ISME J. 2015;9(5):1195–1207. doi: 10.1038/ismej.2014.211. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 54.Moore LR, et al. Culturing the marine cyanobacterium Prochlorococcus. Limnol Oceanogr Methods. 2007;5:353–362. [Google Scholar]
- 55.Wilson WH, Carr NG, Mann NH. The effect of phosphate status on the kinetics of cyanophage infection in the oceanic cyanobacterium Synechococcus sp WH7803. J Phycol. 1996;32(4):506–516. [Google Scholar]
- 56.Barbe V, et al. Unique features revealed by the genome sequence of Acinetobacter sp. ADP1, a versatile and naturally transformation competent bacterium. Nucleic Acids Res. 2004;32(19):5766–5779. doi: 10.1093/nar/gkh910. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 57.Huu NB, Denner EB, Ha DT, Wanner G, Stan-Lotter H. Marinobacter aquaeolei sp. nov., a halophilic bacterium isolated from a Vietnamese oil-producing well. Int J Syst Bacteriol. 1999;49(Pt 2):367–375. doi: 10.1099/00207713-49-2-367. [DOI] [PubMed] [Google Scholar]
- 58.Olson RJ, Vaulot D, Chisholm SW. Marine-phytoplankton distributions measured using shipboard flow-cytometry. Deep-Sea Res, Part A. 1985;32(10):1273–1280. [Google Scholar]
- 59.Cavender-Bares KK, Mann EL, Chisholm SW, Ondrusek ME, Bidigare RR. Differential response of equatorial Pacific phytoplankton to iron fertilization. Limnol Oceanogr. 1999;44(2):237–246. [Google Scholar]