Conversion of membrane lipid acyl groups to triacylglycerol and formation of lipid bodies upon nitrogen starvation in biofuel green algae Chlorella UTEX29
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- Goncalves, E.C., Johnson, J.V. & Rathinasabapathi, B. Planta (2013) 238: 895. doi:10.1007/s00425-013-1946-5
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Algal lipids are ideal biofuel sources. Our objective was to determine the contributors to triacylglycerol (TAG) accumulation and lipid body formation in Chlorella UTEX29 under nitrogen (N) deprivation. A fivefold increase in intracellular lipids following N starvation for 24 h confirmed the oleaginous characteristics of UTEX29. Ultrastructural studies revealed increased number of lipid bodies and decreased starch granules in N-starved cells compared to N-replete cells. Lipid bodies were observed as early as 3 h after N removal and plastids collapsed after 48 h of stress. Moreover, the identification of intracellular pyrenoids and differences in the expected nutritional requirements for Chlorella protothecoides (as UTEX29 is currently classified) led us to conduct a phylogenetic study using 18S and actin cDNA sequences. This indicated UTEX29 to be more phylogenetically related to Chlorella vulgaris. To investigate the fate of different lipids after N starvation, radiolabeling using 14C-acetate was used. A significant decrease in 14C-galactolipids and phospholipids matched the increase in 14C-TAG starting at 3 h of N starvation, consistent with acyl groups from structural lipids as sources for TAG under N starvation. These results have important implications for the identification of key steps controlling oil accumulation in N-starved biofuel algae and demonstrate membrane recycling during lipid body formation.
The development of alternative sources of renewable energy is of utmost importance as world energy consumption is projected to grow by 53 % from 2008 to 2035 (Hu et al. 2008). Although renewable fuels are the fastest growing form of energy, its contribution to total energy production is projected to increase a mere 4 % by 2035 (Hu et al. 2008). In addition, the fact that US transportation fuels must contain 36 billion gallons of renewable fuels by 2022 (Chisti 2007) highlights the importance of all efforts put into developing renewable fuels to an affordable, sustainable and scalable level. Algal lipid from microalgae is one of the best sources for biofuels due to the following reasons: impressively high algal growth rates (commonly doubling its biomass within 24 h) (Chisti 2007), tolerance to extreme conditions (desert and arid lands) (Hu et al. 2008), their abilities to recycle wastewater (agricultural run-off, industrial and municipal wastewater) and CO2 from flue gases emitted from power plants (Hu et al. 2008; Wilkie et al. 2011), reduced competition with food crops (Griffiths and Harrison 2009) and, finally, the production of multiple value-added co-products (e.g., polymers, surfactants, proteins and pigments) (Foley et al. 2011; Wilkie et al. 2011) in parallel to the accumulation of large quantities of neutral lipids (up to 80 % by weight of dry biomass) (Chisti 2007).
Green microalgae, especially within the genus Chlorella, are known for their great potential for high-quality biofuel production as they can grow in a variety of environmental conditions, use different carbon sources, achieve high biomass and lipid content (up to 55 % of cellular dwt) with adequate fatty acid composition (Chen and Walker 2011; Guarnieri et al. 2011; Heredia-Arroyo et al. 2010; Miao and Wu 2004; O’Grady and Morgan 2011; Xu et al. 2006). Several studies have investigated the effect of nitrogen starvation as an inducer for triacylglycerol (TAG) accumulation in green algae, with most of the studies conducted in the model green algae Chlamydomonas reinhardtii (Goodson et al. 2011; James et al. 2011; Miller et al. 2010; Mujtaba et al. 2012; Nguyen et al. 2011; Yeh and Chang 2011). However, many aspects of TAG accumulation in response to nitrogen starvation remain unresolved. For instance, most studies on TAG accumulation examined cells after 24 h or later time points of N starvation, not clarifying the early events of the process. Moreover, it is not known whether what is observed in Chlamydomonas is conserved in other green algae with greater potential for outdoor, large-scale production of biofuels.
A study on Chlamydomonas suggested that the de novo synthesis of fatty acids in the chloroplast is a limiting factor for TAG production, although the recycling of membrane lipids could account for up to 30 % of the TAG produced under N starvation (Fan et al. 2011). Moreover, a tracer experiment with 13C-labeled glucose in Chlorella protothecoides indicated overall stability in global carbon flux during N starvation but did not assess the contribution of recycling of membrane lipids into TAG (Xiong et al. 2010). In this study, a series of radiolabeling experiments with 14C-acetate was performed to track membrane lipids’ remodeling into TAG during N starvation in a green algal strain with potential for outdoor cultivation, with further identification of lipids through HPLC/UV/(+)ESI–MS2. Molecular phylogeny of cDNAs was used to confirm the taxonomic status of the strain. A time-course transmission electron microscopy (TEM) study revealed major ultrastructural changes occurring during the early and late events of lipid droplet formation.
Materials and methods
Algal cultures and nitrogen starvation
UTEX29 (as of Jan 2012, named as C. protothecoides by the stock center) was purchased from the algae culture collection at the University of Texas, Austin, and grown in liquid medium under a light:dark regime of 18:6 h (80 μmol photons/m2/s), under continuous agitation at 130 rpm. A nitrogen-replete (+N) mineral basal algal medium (BAM) (Sorokin and Krauss 1958) was prepared with the following composition (in g/L): KNO3, 1.25; KH2PO4, 1.25; MgSO4·7H20, 1.0; CaCl2, 0.0835; H3BO3, 0.1142; FeSO4·7H20, 0.0498; ZnSO4·7H20, 0.0882; MnCl2·4H2O, 0.0144; MoO3, 0.0071; CuSO4·5H2O, 0.0157; Co(NO3)2·6H20, 0.0049; EDTA, 0.5; and filter-sterilized glucose, 9. The pH was adjusted to 6.8 before autoclaving. For the nitrogen-free (−N) medium, KNO3 was replaced by KCl for the same final potassium concentration. To induce nitrogen deficiency, algal cells previously grown in N-replete medium were washed once in −N medium, suspended in fresh −N medium and cultured for 3–48 h. Cell counting was done using a hemocytometer counting chamber (Fisher Scientific, USA) and turbidity measurements were done using a spectrophotometer (DU730, Beckman Coulter, Pasadena, CA). The relationship between OD600 and cell density was determined as 1 OD600 = 8.3 × 108 cells/mL within a maximum deviation of 14 % of that relationship in both nitrogen-replete and nitrogen-deficient cultures. For calculations, we therefore employed OD600 for normalizing for cell numbers.
Lipid staining using Nile red
Detection of intracellular lipid bodies was conducted as previously described (Chen et al. 2009). Corresponding amounts of cells were treated with a solution of the vital fluorescent stain Nile Red (Greenspan et al. 1985) (50 μg/L) and DMSO (20 % v/v) to increase membrane permeability. After 10 min of incubation, the relative fluorescence (excitation at 530 nm and emission at 570 nm) was measured in a Synergy HT microplate reader (Biotek, USA).
Gravimetric method to quantify lipid productivity
Cells from nitrogen-replete or nitrogen-depleted cultures were collected by centrifugation at 3,500 rpm at 10 °C for 15 min in a Heraeus Labofuge 400R centrifuge in 50-mL tubes. The cell pellets were extracted for total lipids as described elsewhere. The solvent in the lipid extract was evaporated in a nitrogen evaporator (Organomation, MA) and the total lipid fraction was dried in an oven set at 50 °C until two consecutive weighings were constant. The lipid yield was expressed as g/100 g dry weight of cells.
Transmission electron microscopy
Algal cells were collected by centrifugation (5 min, 3,500 rpm in swing-out rotors of a Heraeus Labofuge 400R centrifuge) fixed in 3 % (v/v) glutaraldehyde in BAM medium overnight and washed three times with 0.1 M sodium cacodylate, 2 mM MgCl2, 1 mM CaCl2, 0.25 % (wt/v) NaCl pH 7.23. Fixed cells were processed with the aid of a Pelco BioWave laboratory microwave (Ted Pella, Redding, CA, USA). The samples were buffer washed, post-fixed with 2 % (wt/v) OsO4, water washed, dehydrated in a graded ethanol series 25, 30, 40, 50, 55, 60, 65, 80, 90, 100 % (v/v), infiltrated with LRWhite acrylic resin (Electron Microscopy Sciences, Hatfield, PA) and cured at 60 °C for 2 days. Cured resin blocks were trimmed, thin sectioned and collected on formvar copper 200 mesh grids, post-stained with 2 % aqueous uranyl acetate and Reynold’s lead citrate. Sections were examined with a Hitachi H-7000 TEM (Hitachi High Technologies America, Inc., Schaumburg, IL) and digital images were acquired with a Veleta 2k × 2k camera and iTEM software (Olympus Soft-Imaging Solutions Corp., Lakewood, CO). The TEM experiments were done in triplicates and repeated twice. Three slides were prepared from each replicate and 30 cells were analyzed from each slide.
Radiotracer labeling using 14C-acetate
[2-14C]Acetic acid (Na salt in water) was purchased from American Radiolabeled Chemicals Inc (St. Louis, MO). For labeling studies, the cells were initially grown in 250 mL flasks containing 100 mL of +N medium until the mid-logarithmic phase, transferred to 4.5 mL glass vials for a 24-h incubation with 3 mL of +N medium containing [14C]-acetate (50 nci/μL), followed by centrifugation for 10 min at 3,500 rpm in a swing-out rotor of a Heraeus Labofuge 400R centrifuge, removal of supernatant, and finally addition of +N or −N, label-free medium, for a 48-h growth period.
In the pulse-chase experiment, cells were incubated for 1 h in +N medium containing [14C]-acetate (50 nci/μL), centrifuged for 10 min at 3,500 rpm in a swing-out rotor of a Heraeus Labofuge 400R centrifuge, supernatant was removed, cell pellets were washed with −N medium and suspended in −N medium containing 0.5 mM unlabeled acetate. The experiments were conducted in triplicate.
Extraction of total lipids
Equal number of cells were harvested from different treatments by centrifugation (10 min, 8,000 rpm, in an Eppendorf 5418 centrifuge) and frozen in liquid nitrogen. The lipid extraction method reported by Fan et al. (2011) was used with minor modifications. Total lipids were extracted with methanol–chloroform–formic acid (2:1:0.1, v:v:v) in the presence of glass beads under maximum agitation for 5 min using a Vortex Genie 2 vortex (Fisher Scientific). Maximum extraction efficiency was achieved by repeating this step 7×, followed by phase separation using 1 M KCl, 0.2 M H3PO4 solution. This procedure was sufficient to extract all lipids detectable by iodine staining in thin layer chromatography (TLC).
Analysis of lipids using thin layer chromatography
Total lipid extracts were separated in baked ammonium sulfate-impregnated silica plates (J.T. Baker, USA) by thin layer chromatography (TLC) using a double-development solvent system (2/3 in acetone–toluene–water (91:30:3 by volume) followed by full development in hexane–diethyl ether–acetic acid (70:30:1 by volume) (Fan et al. 2011). TLC plates were briefly stained with iodine vapor for quick visualization of lipid bands. Bands were identified based on their relative mobilities (Rf) compared to standard lipids. For the quantification of radioactivity, the bands were scrapped off the plate, mixed with 2 mL distilled water and 2 mL Ready Gel liquid scintillation cocktail, homogenized and radioactivity counted in a LS6500 liquid scintillation counter (Beckman, USA).
HPLC/UV/(+)ESI–mass spectrometry–mass spectrometry
Lipid bands isolated from the TLC plates were dissolved in isopropanol and separated using an HPLC (1100 series model G1312A binary pump, Agilent, Palo Alto, CA) equipped with a C8 column (Hypurity 5 μm; 2.1 × 100 mm with a C8 guard column; Thermo Scientific) and gradients utilizing a binary mobile phase system of water and isopropanol (both were HPLC-grade; Honeywell Burdick & Jackson, Muskegon, MI) with a flow rate of 0.15 mL/min. For the data presented here, the gradient was set to 30 % isopropanol at time 0 and then increased linearly to 60 % isopropanol in 15 min and then to 95 % isopropanol in 50 min and then held at 95 % isopropanol for 20 min. All mass spectrometric data were obtained with a ThermoFinnigan (San Jose, CA) LCQ fitted with the conventional electrospray ionization source. The sheath and auxiliary gases were nitrogen (65 and 5, respectively, instrumental unitless parameter) and the heated capillary temperature was set at 250 °C. For (+)ESI–MSn, the spray voltage was set at 3.3 kV, the heated capillary voltage at +12.5 V and the tube lens offset at 0 V. Collision-induced dissociation (CID) tandem mass spectrometry (MSn) of [M + Na]+ ions was performed with 3–7 u precursor ion isolation, 30 ms activation time and either 37.5 % normalized CID energy at 0.25 qCID or 40 % normalized CID energy at 0.30 qCID. Ultraviolet/visible (UV) detection at 210 or 220 nm was obtained with an Agilent 1100 series G1314A UV/Vis detector.
Cloning 18s rRNA and actin mRNA partial sequences from UTEX29
RNA extraction was done using NucleoSpin RNA Plant Kit (Macherey–Nagel, Germany). Synthesis of cDNA and RT-PCR were performed using Superscript One-Step RT-PCR with Platinum Taq (Invitrogen, USA) according to the manufacturer’s instructions. The primers for 18s rRNA were 5′-WACCTGGTTGATCCTGCCAGT-3′ and 5′-GATCCTTCYGCAGGTTCACCTAC-3′. The primers for actin were 5′-GTGACCAACTGGGACGAC-3′ and 5′-CGGGCAGCTCGTAIGTCTT-3′. The gel-extracted PCR products were cloned into pCR 2.1-TOPO vector and sequenced (Sambrook 2001).
Molecular phylogeny of UTEX29 partial sequences
Database searches were performed using UTEX29 18s rRNA and actin cDNA sequences as queries. Homologous sequences within the taxa “green algae” were obtained by a Basic Local Alignment Search Tool (BLAST) search at the National Center for Biotechnology Information. The sequences with the best e value, coverage and supporting literature were chosen for further analysis (Table S1). Multiple sequence alignment was conducted using ClustalW software in MEGA5 (Tamura et al. 2011). Phylogenetic trees were inferred from the aligned sequence data using the neighbor-joining method (Saitou and Nei 1987) in MEGA5, with the tree being tested by bootstrapping with 1,000 replicates.
Quantitative data were analyzed using JMP software, Version 7 (SAS Institute Inc., Cary, NC). Means were compared using Student’s t test, α = 0.05. All experiments were conducted in triplicate.
Ultrastructure of nitrogen-starved cells
Taxonomic characterization of Chlorella sp. UTEX29
14C-acetate labeling of fatty acids
Identification of TLC lipid fractions
Pulse-chase of 14C-acetate-labeled fatty acids
The accumulation of LBs by microalgae is a metabolic response that may enable the cells to efficiently store energy, especially under unfavorable environmental conditions, and to rapidly utilize the acyl groups from TAG to synthesize new membranes or other metabolites once favorable growth conditions resume (Murphy 2012; Waltermann and Steinbuchel 2005). In this study, ultrastructural observations clearly showed the effect of N removal in the collapse of membrane systems with concomitant accumulation of LBs after 48 h (Fig. 1). While the pulse-chase experiment suggested TAG accumulation as early as 3 h of −N (Fig. 7), no significant difference (α = 0.05) in TAG was detected using the Nile Red fluorescence assay (data not shown). However, the early event of LB formation (after 3 h of −N) was confirmed by the transmission electron microscopic observations (Fig. 2), demonstrating the importance of this approach to overcome the limitations in detection levels of indirect methods for measuring intracellular TAG, such as the Nile Red fluorescence assay. Although some studies have shown ultrastructural changes in response to N deprivation in different algal species (García-Ferris and de los Ríos 1996; Goodson et al. 2011; Pyliotis and Goodchild 1975), this is the first study in commercially important biofuel green algae to investigate the ultrastructural changes occurring in the early events (3 h) of N deprivation. Another interesting ultrastructural observation was that the starch granules observed in abundance and filling large areas of the single chloroplast in the control cells (Figs. 1a, 2a, b) were not observed in the cells under N deficiency (Figs. 1b, 2c, d). This result is consistent with previous observations (Wang et al. 2009), in which it was demonstrated that starchless mutants of Chlamydomonas accumulate up to 30-fold the amount of TAG (approximately 400 mg/109 cells) compared to the wild-type strain, after 48 h of N starvation. This indicates that starch and TAG are competing carbon sinks and that degradation of starch during −N may increase the carbon flux into acetyl-CoA and TAG de novo synthesis. Since the presence of plastoglobuli was observed in one of the examined cells (Fig. 3) as also observed by Orus and Martinez (1991), it is possible that a chloroplastic pathway for TAG synthesis may also be present in Chlorella as it was described in Chlamydomonas (Fan et al. 2011; Goodson et al. 2011). However, the fact that it was observed only once indicates it may not be the preferential route for TAG synthesis under the experimental conditions employed. Interestingly, N deficiency in the higher plant Arabidopsis thaliana induces the partial mobilization of acyl groups from galactolipids to plastoglobules in the chloroplast, where fatty acid phytyl esters accumulate (Gaude et al. 2007), but the enzyme systems involved have not been identified.
The correct identification of green microalgae, especially within the Chlorella genus, which comprises more than 100 species described (Krienitz et al. 2004), is a complex task that may be overlooked in some studies and that is key for drawing correct conclusions when studying axenic cultures. The identification of pyrenoids, intracellular structures found in C. vulgaris, in our TEM study (Figs. 1, 2) prompted us to examine if Chlorella sp. UTEX29 is correctly assigned as C. protothecoides. The nutritional requirements experiment (Supplemental Fig. 2) showed that UTEX29 does not require thiamine and can utilize ammonium as N source for normal growth. The phylogenetic studies with UTEX29 18S rRNA and actin cDNA partial sequences (Fig. 4) indicate that UTEX29 strain is more structurally, nutritionally and phylogenetically related to C. vulgaris than to C. protothecoides. This result raises the question of how reliable is the species-level identification and the importance of using complementary approaches when investigating the identity of algae used in biofuel industry.
This study demonstrates that the oil-accumulating characteristic of the green algae Chlorella sp. UTEX29 is reproducibly triggered by N removal from the medium, with a ~fivefold increase in detectable lipid content within 24 h of −N (Supplemental Fig. 1). Several studies have investigated the effect of nitrogen starvation as an inducer for TAG accumulation in both the genus Chlorella (Hortensteiner et al. 2000; Pyliotis and Goodchild 1975; Xiong et al. 2010; Yeh and Chang 2011) and a related species, the model green algae C. reinhardtii (Fan et al. 2011; Goodson et al. 2011; James et al. 2011; Miller et al. 2010; Nguyen et al. 2011; Yeh and Chang 2011). However, these studies focus at 24 h or later time points of N starvation, without clarifying the early events on lipid body formation. Moreover, information regarding the contribution of membrane lipids toward TAG synthesis is also very limited to date. A study on Chlamydomonas indicated that the de novo synthesis of fatty acids in the chloroplast is a limiting factor for TAG production, and assuming that the decreases in polar lipid amounts during −N are entirely mobilized into TAG, Fan et al. (2011) suggested that recycling of membrane lipids could account for up to 30 % of the TAG synthesis after 48 h of N deprivation. By labeling the fatty acids with 14C-acetate and tracking the radioactivity shifts after 24 h of N removal (Fig. 5), Fan et al. (2011)’s assumption was confirmed, as a 34 % decrease in polar lipids radioactivity was detected, which accounted for the same percentage increase in TAG radioactivity (Fig. 5c). The 14C-acetate-labeling experiment in this study provides evidence in vivo to demonstrate a substrate–product relationship between TAG and structural polar lipids upon N deficiency, providing a key support for the hypothesis of membrane recycling into TAG. While our study was in progress, Li et al. (2012) reported that the activity of a galactoglycerolipid lipase (PGD1) was critical in TAG accumulation under N starvation in C. reinhardtii (Li et al. 2012), consistent with our findings in Chlorella.
The TLC lipid zones P3 and P5, which showed a significant decrease in radioactivity under −N conditions, were identified by ESI–MS/MS as the galactolipids MGDG and DGDG, which account for up to 80 % of membrane lipids in green plant tissues (Guella et al. 2003). This result suggests that galactolipids may be a major contributing source of acyl groups for the synthesis of TAG under −N. Although GC–MS is routinely used to identify fatty acid composition of lipid compounds, the LC(+)ESI–MS/MS approach performed in this study has the advantage of generating high-resolution lipid profiles without the fatty acid methyl esterification (FAME) step, and still obtain information about the acyl chains. The fatty acid composition of the lipids was possible by analyzing the CID products of the [M + Na]+ ions (Tables S1, S2), and the putative regiochemical distribution was deduced as previously described (Guella et al. 2003). The possible identification of odd-chain fatty acids C19:1, and C19:2 at the sn-1 position of galactolipids needs further study.
The significant increase in radioactivity (α = 0.05) in TAG observed as early as 3 h of N starvation with concomitant decreased radioactivity in both MGDG and phospholipids observed with the pulse-chase experiment (Fig. 7) suggests that lipases and acyltransferases may be involved in the response to this nutrient stress at an early time point following N starvation. In fact, two recent transcriptomic studies in Chlamydomonas under −N detected upregulated genes encoding putative lipases that could play a role in releasing fatty acids from membrane lipids (Boyle et al. 2012; Miller et al. 2010). Moreover, a recent proteomics study in Chlamydomonas identified lipase-like oil body associated proteins, which may be localized at contact sites between LBs and chloroplasts, and thus, be involved in the degradation of plastidial membranes (Nguyen et al. 2011). In Chlamydomonas, one of the enzymes involved in this process is phospholipid:diacylglycerol acyltransferase (PDAT), which was recently shown to have both acyltransferase and lipase activities in a variety of substrates, such as phospholipids, galactolipids, and TAG (Yoon et al. 2012). While PDAT and PGD1 homologs would be expected to play a similar role in Chlorella, other enzymes which have not been fully characterized may also contribute to this phenomenon of membrane recycling of fatty acids into TAG. In fact, the replacement of the membrane lipid phosphatidylcholine present in Chlorella by diacylglycerol-N, N, N-trimethylhomoserine (Fan et al. 2011) in Chlamydomonas, confirmed in this study, illustrates the evolutionary divergences between the two genera.
The present study shows that the recycling of structural membrane lipids significantly contributes to the increased flux of acyl moieties into TAG starting as early as 3 h of N starvation in Chlorella. It also provides key ultrastructural information that will serve as a basis for subsequent high-throughput proteomic and transcriptomic studies to investigate the initial and late metabolic changes, leading to the trigger of LB formation and TAG accumulation in oleaginous green algae.
We thank Dr. Byung-Ho Kang, Karen Kelley, and Kim Backer-Kelley (University of Florida, Interdisciplinary Center for Biotechnology Research, Electron Microscopy and Bio-Imaging lab) for help with transmission electron microscopy. The HPLC–MS-MS analyses of lipid fractions were done at Chemistry Department, University of Florida. EG thanks the Plant Molecular and Cellular Biology program, the College of Agriculture and Life Sciences and the Horticultural Sciences Department, University of Florida for a graduate research assistantship.