Low-cost lipid production by an oleaginous yeast cultured in non-sterile conditions using model waste resources
- Fabio Santamauro†1, 2,
- Fraeya M Whiffin2,
- Rod J Scott†1Email author and
- Christopher J Chuck†2Email author
© Santamauro et al.; licensee BioMed Central Ltd. 2014
Received: 29 July 2013
Accepted: 17 February 2014
Published: 4 March 2014
The yeast Metschnikowia pulcherrima, previously utilised as a biological control agent, was evaluated for its potential to produce lipids for biofuel production.
Cultivation in low cost non-sterile conditions was achieved by exploiting its ability to grow at low temperature and pH and to produce natural antimicrobial compounds. Although not previously classified as oleaginous, a combination of low temperature and restricted nutrient availability triggered high levels of oil production in M. pulcherrima cultures. This regime was designed to trigger the sporulation process but prevent its completion to allow the accumulation of a subset of a normally transitional, but oil-rich, ‘pulcherrima’ cell type. This approach resulted in yields of up to 40% lipid, which compares favourably with other oleaginous microbes. We also demonstrate that M. pulcherrima metabolises glycerol and a diverse range of other sugars, suggesting that heterogeneous biomass could provide a suitable carbon source. M. pulcherrima also grows well in a minimal media containing no yeast extract. Finally, we demonstrate the potential of the yeast to produce lipids inexpensively on an industrial scale by culturing the yeast in a 500 L, open air, tank reactor without any significant contamination.
The production of antimicrobial compounds coupled to efficient growth at low temperature and pH enables culture of this oleaginous yeast in inexpensive, non-sterile conditions providing a potential route to economic biofuel production.
KeywordsBiodiesel Lipid Yeast Heterotrophic Waste Pilot scale Glycerol Lignocellulose
The rising cost of fossil fuels, coupled to concerns over security of supply and an increasing awareness of the environmental impact of associated CO2 emissions, have led to the development of alternative energy solutions becoming a global priority. Some of the most ambitious national targets have been set by the United Kingdom, where the government is committed to sourcing 15% of the total energy demand from renewable resources by 2020, and 80% by 2050 . Since a quarter of emissions are produced by the transport sector, meeting these reduction targets requires urgent development of biofuels compatible with the current infrastructure. Currently, the main source of biofuels is sugar-derived bioethanol or biodiesel produced from lipids, derived from oil palm, rapeseed or soybean . However, these crops compete with food crops for agricultural land and consequently have both a negative public image and environmental impact . Oleaginous microorganisms such as microalgae, bacteria, fungi and yeasts are an attractive alternative to higher plants for lipid production, since their production does not require agricultural land, avoiding displacement of food production. While species from these taxa have the potential to produce lipids, only microalgae have been extensively investigated to date . Although algae have great theoretical potential as a fuel source, substantial technical hurdles currently prevent their cost-effective exploitation. These include the scarcity of suitable land, given the current levels of productivity, the lack of adequate photosynthetically active radiation and the prohibitive costs of delivering supplementary light, fertiliser, controlling temperature and critically, protecting against invasive non-lipid-producing species [5, 6].
Heterotrophic organisms, such as yeasts, offer a credible alternative to microalgae for the production of biofuel feedstocks, especially in Northern Europe. Oleaginous yeast species are highly productive on a per-cell basis, with lipid yields of up to 65% dry weight and grow to high densities with biomass yields of 10 to 100 g⋅L-1 reported over 3 to 7 days . In comparison, microalgae achieve only 0.15 to 0.25 g⋅L-1 per day in open pond systems . The most common yeasts so far examined for biofuel production are Rhodotorula glutinis, Yarrowia lipolytica and Lipomyces starkeyi[8–11].
Recently, the economic production of C5 and C6 sugars from waste cellulosic materials has become plausible, making plant sugars derived from lignocellulose a feasible source of renewable feedstocks [12, 13]. Unlike microalgae, yeast cultivation does not require light, which both reduces input costs and enables production 24 h per day. Essential inputs such as phosphorous and nitrogen are also available from waste streams such as waste water, again reducing production costs.
One key issue with the cultivation of heterotrophic organisms is maintaining an axenic culture. Bacteria from the surrounding environment or contaminating input waste streams have the potential to outcompete most types of oleaginous yeast. Ensuring the strict sterile conditions required for production of triglycerides by yeast, therefore, requires stringent pre-treatment of the feedstock and careful enclosure of the fermentation broth. Both significantly increase production costs, potentially limiting industrial-scale lipid biofuel production . One potential solution is to use an extremophile yeast that thrives at an extreme pH, temperature or salinity that significantly reduces or prevents competing microbial growth [15, 16]. As bacteria and other contaminating organisms can adapt to such environments, an extremophile with more than one protective mechanism is attractive for open-tank reactors.
The non-saccharomyces yeast, Metschnikowia pulcherrima, is known to severely limit the growth of other microbes in the natural environment and is consequently used commercially as a post-harvest control agent . However, the ecological significance and exact mechanisms are still poorly understood but offer the potential for commercial exploitation. M. pulcherrima, also reported in literature as Candida pulcherrima (anamorph), Saccharomyces pulcherrimus, Rhodotorula pulcherrima, Torula pulcherrima, Cryptococcus castellanii and Torulopsis pulcherrima,  is commonly found in flower nectar and fruit surfaces, and colonises the grape must after pressing, contributing to the flavour and aroma of the wine . The yeast thrives in the must despite its high acidity (pH 3 to 4) and sugar concentration (>100 g/L), and resists competition from other microorganisms. Cultures of M. pulcherrima do not require iron or vitamins other than biotin [20, 21]. M. pulcherrima is well-known for its production of the red pigment, pulcherrimin formed by irreversible binding of pulcherrimic acid and environmental iron to create a metal-organic framework. This sequestration prevents competing microbes accessing essential supplies of iron, and is the assumed basis of the use of M. pulcherrima as a biofungicide in post-harvest disease control [17, 22, 23]. M. pulcherrima was therefore selected for study for its potential for low-cost culture in non-sterile conditions due a combination of acidophilia and secretion of the potent anti-microbial compounds, pulcherriminic acid and 2-phenylethanol .
During its growth on the surface of grapes M. pulcherrima is assumed to catabolise constituents of the plant cell-wall, as well as accessing sugars from within the grape . This accords with recent studies demonstrating that M. pulcherrima produces a wide range of cell-wall degrading enzymes; this property indicates a degree of metabolic plasticity that lends itself to industrial applications in oenology [19, 24].
Whilst the innate ability of M. pulcherrima to grow at low pH and temperature combined with pulcherrimin production would facilitate its culture under relatively inexpensive non-sterile conditions, previous studies have concluded that the species is not oleaginous . However, when subjected to environmental stress, for example, nitrogen starvation, a subset of cells within an M. pulcherrima culture begin sporulation via a transitional pulcherrima cell-type that has shown some evidence of lipid accumulation [27, 28]. We reasoned that increasing the proportion of pulcherrima cells formed in response to a suitable stimulus, coupled with a means to prevent their development into oil-poor spores, could elevate overall lipid production by an M. pulcherrima culture into the oleaginous range.
Here we show that manipulating the culture regime successfully increased the proportion of oil-rich pulcherrima cells and achieved a 40% overall lipid yield. We also show that the yeast metabolises glycerol and a diverse range of other sugars, suggesting that heterogeneous biomass could provide a suitable carbon source. Finally, we illustrate the potential to produce lipids inexpensively on an industrial scale by culturing M. pulcherrima in a 500-L, open air, tank reactor.
Results and discussion
As a non-saccharomyces yeast involved in the first stages of wine fermentation, M. pulcherrima endures particularly stressful conditions in the grape must, such as high acidity due to the high levels of tartaric and malic acid, and high osmotic pressure due to the high sugar concentrations. The ability to thrive under such harsh conditions makes M. pulcherrima a potentially attractive organism for industrial applications, particularly where culture in non-sterile conditions would reduce production costs. Whilst the robustness of M. pulcherrima has been established, its status as an oleaginous yeast has not. We set out to determine whether M. pulcherrima is capable of high lipid-production in conditions that exploit its potential for low cost culture.
Triggering oil production in M. pulcherrima
Lipid accumulation in microbial cells normally requires a starvation step to promote energy storage at the expense of further cell division . To achieve this, the starting concentration of nitrogen in the culture medium is deliberately limited relative to the carbon source. A recent study evaluating the potential of yeasts to produce biofuels from glycerol found that M. pulcherrima did not accumulate lipids to any significant degree at 28°C, over 5 days, even at high C/N ratios . However, other types of stress induce M. pulcherrima to undergo sporulation via a transitory pulcherrima cell-type that accumulates high lipid levels, particularly in dilute media or at low temperatures, though the effectiveness of these treatments has not been quantified [27, 28]. Although pulcherrima cells have not been observed as the predominant cell-type in a culture, it seemed likely that conditions of nutrient starvation and below-ambient temperatures would act as a trigger for this stage. Such low temperatures would potentially halt sporulation, resulting in increased levels of lipid-containing cells. To determine whether temperature could trigger oil production in M. pulcherrima, the culture was grown at 20°C for 3 days, the temperature was then modified (increased or decreased) for the remaining length of the culture period. The lipid content of the biomass was measured for the cultures after 15 days.
The effect of restricted nutrients on the growth
Several other Candida species (Candida curvata, Rhodotorula graminis, Cryptococcus curvatus and Yarrowia lipolytica) reportedly produce between 28 and 53% lipid when grown on glycerol as a carbon source. This level is comparable to that produced by the M. pulcherrima strain presented in this work [32, 33]. M. pulcherrima grew at high glycerol concentrations, and although this has been reported for some oleaginous species such as Y. lipolytica, it is far from common. For example, other similar oleaginous species, such as C. curvatus are unable to grow on concentrations higher than 60 g/L, presumably due to the elevated osmotic pressure [31–33].
Sulphur is another key nutrient that can affect growth rates. Generally sulphur is present as sulphates, especially in waste water. To assess the flexibility of M. pulcherrima, a range of cultures were examined with reduced levels of sulphate (Figure 4b). There was no difference, in terms of dry biomass; on culturing M. pulcherrima with concentrations of sulphur between 0.321 g/L and 0.045 g/L, the same level of biomass were obtained.
A range of micronutrients is added to most yeast culture media, including salts of manganese, zinc and iron. Although these micronutrients are potentially present in waste water, the type and amount of the micronutrient will vary substantially . To determine the tolerance of M. pulcherrima to fluctuations in micronutrient availability, the yeast was grown in SMP (10% w/v glycerol) lacking Fe, Zn, or Mn, either singly or in combination. The biomass production after 15 days was then compared to a culture grown on SMP with all the nutrients present (Figure 4c). Surprisingly, biomass productivity was reduced by only 15% in SMP not supplemented with iron, zinc and manganese.
M. pulcherrima demonstrates excellent flexibility on being cultured with different types and amounts of nitrogen, sulphur and micronutrients. This demonstrates that although there is an optimal culture configuration that produces the highest levels of biomass from any given system, the effect of removing key nutrients does not dramatically reduce biomass yields. M. pulcherrima, therefore, has excellent adaptability to any changes in the waste water or alternative nutrient streams that could be used to culture the system.
Lipid accumulation in M. pulcherrima continues up to 15 days, after which the lipid content decreases, presumably due to cellular lipid turnover (Figure 5). This phenomenon has been reported for the yeast Y. lipolytica and other oleaginous fungi, for example, Cunninghamella echinulata. The authors propose that the lipid degradation occurs in C. echinulata by selective use of triglyceride to produce glycolipids, sphingolipids and phospholipids through the substitution of one or more fatty acid (FA) chains. The triglyceride can also be completely hydrolysed to glycerol and free fatty acids, which can then be funnelled into glycolysis and β-oxidation.
Lipid profile of fed-batch cultures of M. pulcherrima
Run 1 (with yeast extract)
Run 2 (without yeast extract)
Run 3 (with biotin)
Run 4 (without biotin)
Final biomass yield (g⋅L -1 )
Final lipid content (% dry weight)
M. pulcherrima was successfully grown in a fed-batch culture, with over 17 g⋅L-1 biomass being produced over 30 days. Unsurprisingly, the highest biomass yields were achieved with yeast extract, though this is not essential to growth as comparable productivity was observed for all cultures irrespective of the supplementary nutrients. The highest lipid content of 39%, was produced by the culture without yeast extract but with a supplementation of biotin and nitrogen. Surprisingly, dry biomass productivity around 13 g/L and lipid productivity around 33% were still observed in the cultures containing no yeast extract, biotin or additional nitrogen. The ability to produce lipids from a complex sugar feed, with no additional yeast extract or vitamin supplementation, has significant ramifications for the viability of any future industrial process.
Open-air stirred tank production
Biomass characteristics and lipid profile of the oil produced from M. pulcherrima grown in the raceway ponds
Although total biomass yield was lower than obtained at laboratory scale, these suboptimal conditions nevertheless outperformed similar systems used to culture microalgae . Alongside sterilisation, agitation is a major cost in running an industrial bioprocess . Though further work is needed to optimise cultures of M. pulcherrima on this scale, the preliminary findings demonstrate that the yeast can be cultured under non-sterile conditions, with minimal agitation. This has the potential to substantially reduce the costs of a process. The level of oil extracted from the harvested biomass was 34% of the dry weight.
The biological control yeast, M. pulcherrima, which was previously classified as non-oleaginous, is shown here capable of producing up to 40% lipid by manipulating pH and temperature. The yeast also uses a variety of sugars potentially sourced from waste lignocellulose. Fed-batch culture using these sugars achieved a total biomass yield of 17 g⋅L-1. High biomass productivity was also achieved using simplified media lacking expensive inputs such as yeast extract and biotin. The unique combination of acidophilia and production of antimicrobials enable culture in lost-cost non-sterile conditions providing a potentially economically viable biofuel production platform.
All chemicals were purchased from Sigma Aldrich unless otherwise stated and used without purification.
Culture conditions on a laboratory scale
Cultures were grown from a single colony taken from a yeast extract, malt extract and dextrose (YMD) agar plate suspended in 10 ml YMD (yeast extract 10 g⋅L-1; malt extract 20 g⋅L-1; glucose 20 g⋅L-1) and inoculated into liquid standard M. pulcherrima (SMP) medium adapted from Chatzifragkou et al. (KH2PO4 7 g⋅L-1; Na2HPO4 2.5 g⋅L-1; MgSO4. 7H2O 1.5 g⋅L-1; CaCl2. 2 H2O 0.15 g⋅L-1; ZnSO4. 7H2O 0.02 g⋅L-1; MnSO4 . H2O 0.06 g⋅L-1; FeCl3 0.15 g⋅L-1; (NH4)2SO4 0.5 g⋅L-1 and yeast extract 1 g⋅L-1) . The medium was autoclaved for 20 minutes at 120°C prior to use. Each experiment was conducted in triplicate using 10 ml SMP medium in 50-ml Falcon tubes.
The optimum glycerol concentration was determined using SMP medium containing between 1% and 25% (w/v) glycerol. To examine the effect of temperature on lipid production, cultures were grown for 3 days at 25°C then switched to either 15°C or 20°C for the remaining period. Alternative nitrogen sources were evaluated by replacing (NH4)2SO4 in SMP medium (10% w/v glycerol) and maintaining the total nitrogen concentration at 0.106 g⋅L-1. Micronutrient deficiency was investigated using SMP medium (10% w/v glycerol) lacking the specified micronutrient. Sulphur content was varied using modified SMP medium (10% w/v glycerol) containing 50%, 25% and 12.5% of the concentration in SMP media. pH was adjusted using dilute HCl or KOH to a achieve a range between pH 3 and pH 6. Each set of experiments were conducted over 15 days. Optical density (O.D.)600nm, cell number and lipid fluorescence were measured at 72-hour intervals.
Alternative sugar sources
The performance of M. pulcherrima on different sugar sources (glucose, glycerol, xylose, arabinose, cellobiose, lactose, sucrose and glycerol) was examined using a 96-well microtitre plate system (Versamax, Molecular devices, UK). Briefly, 200 μl of culture were made up in 96-well plates in SMP medium containing a 30 g⋅L-1 of the sugar. After 15 days at 25°C, 180 rpm, the O.D.600nm was measured using a plate reader. All the possible combinations of two sugars were tested using six repeats for each combination. An OMP medium was developed for low-cost culture of M. pulcherrima. OMP consisted of: KH2PO4 7 g⋅L-1; Na2HPO4 2.5 g⋅L-1; MgSO4. 7H2O 0.188 g⋅L-1; MgCl2. 6H2O 1.083 g⋅L-1; CaCl2. 2 H2O 0.15 g⋅L-1; ZnSO4. 7H2O 0.02 g⋅L-1; (NH4)2SO4 0.063 g⋅L-1; NH4Cl 0.405 g⋅L-1: yeast extract 1 g⋅L-1 and glycerol 30 g⋅L-1. OMP was adjusted to pH 5 using dilute HCl and utilised without sterilisation.
M. pulcherrima was cultured in a fed-batch regime. The media were comprised of final effluent waste water; all nutrients other than yeast extract, biotin and the carbon source were supplied according to the OMP media. The media were not sterilised prior to use. Each culture was fed every 3 days up to 24 days (25°C, 180 rpm) and then allowed to rest at 15°C for 12 days without agitation, except phosphate, which was supplemented only at 15 days.
Raceway pond cultivation
OMP with a reduced glycerol content of 30 g⋅L-1 was used to establish cultures in two 500-L-capacity raceway ponds situated in a climate-controlled glasshouse. The ponds were inoculated with 500 ml of M. pulcherrima culture grown for 48 hours at 25°C with agitation (180 rpm) in yeast extract, mannitol and sorbose (YMS) medium consisting of 30 g L-1 yeast extract, 5 g⋅L-1 mannitol and 5 g⋅L-1 sorbose.
The pond cultures were agitated using a close-fitting paddle wheel driven at 10 rpm and aerated through two spargers situated at opposite sides of the ponds. Culture temperature, pH and O.D.600nm were measured daily until the onset of the stationary phase, then every 4 days together with the addition of lipid fluorescence up to 28 days. With the onset of the stationary phase the temperature in the glasshouse was reduced from 25°C to 20°C, the aeration was stopped and the paddle wheels were set at the minimum rotating rate (4 rpm).
Lipid extraction and analysis
Lipid was extracted by an adaptation of a method first reported by Bligh and Dyer . The samples were stirred in a CHCl3 and MeOH mixture (2:1 w/v) over 48 hours, and the biomass was filtered off and washed with additional CHCl3. This was repeated three times. The volatiles were removed under reduced pressure and the lipid was analysed.
The lipid and sterol content were calculated by proton nuclear magnetic resonance (1H NMR) by comparison of the glyceride protons (δ 4.1 ppm) to the α-protons of the sterol alcohol group (δ 3.7 to 3.9 ppm). NMR spectroscopic measurements were carried out at 298 K using a Bruker AV500 spectrometer (Bruker GmbH, Germany), operating at 300 MHz. 1H spectra were typically acquired using a 30-degree excitation pulse and a repetition time of 4.2 sec; 0.3-Hz line broadening was applied before Fourier transform, and spectra were referenced to the residual CHCl3 peak from the solvent (δ 7.26 ppm).
The glyceride proportion of the oil was transesterified using typical reported techniques . The fatty acid profile was then calculated by gas chromatography-mass spectroscopy (GC-MS) with the resulting fatty acid methyl esters (FAME) component compared to known FAME standards. GC-MS analysis was carried out using an Agilent 7890A gas chromatograph equipped with a capillary column (60 m × 0.250 mm internal diameter) coated with DB-23 ((50%-cyanpropyl)-methylpolysiloxane) stationary phase (0.25-μm film thickness) and an He mobile phase (flow rate: 1.2 ml/minute) coupled with an Agilent 5975C inert MSD with triple axis detector (Agilent Technologies, CA, USA). FAME samples were initially dissolved in 2 ml of dioxane and 1 μl of this solution was loaded onto the column, pre-heated to 150°C. This temperature was held for 5 minutes and then heated to 250°C at a rate of 4°C/minute and then held for 2 minutes.
fatty acid methyl esters
gas chromatography-mass spectroscopy
nuclear magnetic resonance
optimised M. pulcherrima
standard M. pulcherrima
yeast extract, malt extract and dextrose.
The authors would like to extend their thanks to Graham Fisher and Miles Davis for their aid in securing funding, the Engineering and Physical Sciences Research Council (EPSRC) for partially funding this work through the Doctoral Training Centre at the Centre for Sustainable Chemical Technologies (CSCT), to Dr Marianne Harkins and Rhodri Jenkins for their help in harvesting the stirred tank reactors and to Roger and Sue Whorrod for their kind endowment to the University resulting in the Whorrod Fellowship in Sustainable Chemical Technologies held by Dr C Chuck.
- Department of Energy and Climate Change: UK renewable energy roadmap. 2011, London: Department of Energy and Climate Change, 107-Google Scholar
- Escobar JC, Lora ES, Venturini OJ, Yanez EE, Castillo EF, Almazan O: Biofuels: environment, technology and food security. Renew Sust Energ Rev. 2009, 13: 1275-1287. 10.1016/j.rser.2008.08.014.View ArticleGoogle Scholar
- Fargione J, Hill J, Tilman D, Polasky S, Hawthorne P: Land clearing and the biofuel carbon debt. Science. 2008, 319: 1235-1238. 10.1126/science.1152747.View ArticleGoogle Scholar
- Chisti Y: Biodiesel from microalgae beats bioethanol. Trends Biotechnol. 2008, 26: 126-131. 10.1016/j.tibtech.2007.12.002.View ArticleGoogle Scholar
- Sharma YC, Singh B, Korstad J: A critical review on recent methods used for economically viable and eco-friendly development of microalgae as a potential feedstock for synthesis of biodiesel. Green Chem. 2011, 13: 2993-3006. 10.1039/c1gc15535k.View ArticleGoogle Scholar
- Suali E, Sarbatly R: Conversion of microalgae to biofuel. Renew Sust Energ Rev. 2012, 16: 4316-4342. 10.1016/j.rser.2012.03.047.View ArticleGoogle Scholar
- Li Q, Du W, Liu D: Perspectives of microbial oils for biodiesel production. Appl Microbiol Biotechnol. 2008, 80: 749-756. 10.1007/s00253-008-1625-9.View ArticleGoogle Scholar
- Yen H-W, Yang Y-C: The effects of irradiation and microfiltration on the cells growing and total lipids production in the cultivation of Rhodotorula glutinis. Biores Technol. 2012, 107: 539-541.View ArticleGoogle Scholar
- Tsigie YA, Lien Huong H, Ahmed IN, Ju Y-H: Maximizing biodiesel production from Yarrowia lipolytica Po1g biomass using subcritical water pretreatment. Biores Technol. 2012, 111: 201-207.View ArticleGoogle Scholar
- Huang C, Wu H, Liu Z-J, Cai J, Lou W-Y, Zong M-H: Effect of organic acids on the growth and lipid accumulation of oleaginous yeast Trichosporon fermentans. Biotechnol Biofuel. 2012, 5: 4-10.1186/1754-6834-5-4.View ArticleGoogle Scholar
- Angerbauer C, Siebenhofer M, Mittelbach M, Guebitz GM: Conversion of sewage sludge into lipids by Lipomyces starkeyi for biodiesel production. Biores Technol. 2008, 99: 3051-3056. 10.1016/j.biortech.2007.06.045.View ArticleGoogle Scholar
- Sun Y, Cheng JY: Hydrolysis of lignocellulosic materials for ethanol production: a review. Biores Technol. 2002, 83: 1-11. 10.1016/S0960-8524(01)00212-7.View ArticleGoogle Scholar
- Larsen J, Petersen MO, Thirup L, Li HW, Iversen FK: The IBUS process - lignocellulosic bioethanol close to a commercial reality. Chem Eng Technol. 2008, 31: 765-772. 10.1002/ceat.200800048.View ArticleGoogle Scholar
- Koutinas AA, Chatzifragkou A, Kopsahelis N, Papanikolaou S, Kookos IK: Design and techno-economic evaluation of microbial oil production as a renewable resource for biodiesel and oleochemical production. Fuel. 2014, 116: 566-577.View ArticleGoogle Scholar
- Pulz O, Gross W: Valuable products from biotechnology of microalgae. Appl Microbiol Biotechnol. 2004, 65: 635-648. 10.1007/s00253-004-1647-x.View ArticleGoogle Scholar
- Cripps RE, Eley K, Leak DJ, Rudd B, Taylor M, Todd M, Boakes S, Martin S, Atkinson T: Metabolic engineering of Geobacillus thermoglucosidasius for high yield ethanol production. Metab Eng. 2009, 11: 398-408. 10.1016/j.ymben.2009.08.005.View ArticleGoogle Scholar
- Spadaro D, Vola R, Piano S, Gullino ML: Mechanisms of action and efficacy of four isolates of the yeast Metschnikowia pulcherrima active against postharvest pathogens on apples. Postharvest Biol Tech. 2002, 24: 123-134. 10.1016/S0925-5214(01)00172-7.View ArticleGoogle Scholar
- Kurtzman CP: The yeasts: A taxonomic study. 2011, Elsevier, 2081-5Google Scholar
- Fernandez M, Ubeda JF, Briones AI: Typing of non-saccharomyces yeasts with enzymatic activities of interest in wine-making. Int J Food Microbiol. 2000, 59: 29-36. 10.1016/S0168-1605(00)00283-X.View ArticleGoogle Scholar
- Kluyver AJ, Vanderwalt JP, Vantriet AJ: Pulcherrimin, the pigment of candida pulcherrima. Proc Natl Acad Sci U S A. 1953, 39: 583-593. 10.1073/pnas.39.7.583.View ArticleGoogle Scholar
- Roberts C: The effect of iron and other factors on the production of pigment by the yeast Torulopsis pulcherrima. Am J Bot. 1946, 33: 237-244. 10.2307/2437427.View ArticleGoogle Scholar
- Spadaro D, Ciavorella AA, Lopez-Reyes JG, Garibaldi A, Gullino ML: Effect of culture age, protectants, and initial cell concentration on viability of freeze-dried cells of Metschnikowia pulcherrima. Can J Microbiol. 2010, 56: 809-815. 10.1139/W10-068.View ArticleGoogle Scholar
- Fugelsang KL, Edwards CG: Wine Microbiology - Practical Applications and Procedures. 2006, Springer, 393-2Google Scholar
- Jolly NP, Augustyn OPH, Pretorius IS: The role and use of non-saccharomyces yeasts in wine production. S Afr J Enol Vitic. 2006, 27: 15-39.Google Scholar
- Rosini G, Federici F, Martini A: Yeast flora of grape berries during ripening. Microb Ecol. 1982, 8: 83-89. 10.1007/BF02011464.View ArticleGoogle Scholar
- Chatzifragkou A, Makri A, Belka A, Bellou S, Mavrou M, Mastoridou M, Mystrioti P, Onjaro G, Aggelis G, Papanikolaou S: Biotechnological conversions of biodiesel derived waste glycerol by yeast and fungal species. Energy. 2011, 36: 1097-1108. 10.1016/j.energy.2010.11.040.View ArticleGoogle Scholar
- Pitt JI, Miller MW: Sporulation in candida pulcherrima, candid reukaufii and chlamydozyma species – their relationships with Metschnikowia. Mycologia. 1968, 60: 663-685. 10.2307/3757434.View ArticleGoogle Scholar
- Pan LX, Yang DF, Shao L, Li W, Chen GG, Liang ZQ: Isolation of the oleaginous yeasts from the soil and studies of their lipid-producing capacities. Food Technol Biotechnol. 2009, 47: 215-220.Google Scholar
- Ratledge C, Wynn JP: The biochemistry and molecular biology of lipid accumulation in oleaginous microorganisms. Advances in Applied Microbiology. 2002, Academic Press, 1-51. 51Google Scholar
- Charoenchai C, Fleet GH, Henschke PA: Effects of temperature, pH, and sugar concentration on the growth rates and cell biomass of wine yeasts. Am J Enol Viticult. 1998, 49: 283-288.Google Scholar
- Meesters P, Huijberts GNM, Eggink G: High cell density cultivation of the lipid accumulating yeast Cryptococcus curvatus using glycerol as a carbon source. Appl Microbiol Biotechnol. 1996, 45: 575-579. 10.1007/s002530050731.View ArticleGoogle Scholar
- Liang Y, Cui Y, Trushenski J, Blackburn JW: Converting crude glycerol derived from yellow grease to lipids through yeast fermentation. Bioresour Technol. 2010, 101: 7581-7586. 10.1016/j.biortech.2010.04.061.View ArticleGoogle Scholar
- Papanikolaou S, Fakas S, Fick M, Chevalot I, Galiotou-Panayotou M, Komaitis M, Marc I, Aggelis G: Biotechnological valorisation of raw glycerol discharged after bio-diesel (fatty acid methyl esters) manufacturing process: production of 1,3-propanediol, citric acid and single cell oil. Biomass Bioenergy. 2008, 32: 60-71. 10.1016/j.biombioe.2007.06.007.View ArticleGoogle Scholar
- Burgess JE, Quarmby J, Stephenson T: Role of micronutrients in activated sludge-based biotreatment of industrial effluents. Biotechnol Adv. 1999, 17: 49-70. 10.1016/S0734-9750(98)00016-0.View ArticleGoogle Scholar
- Fakas S, Galiotou-Panayotou M, Papanikolaou S, Komaitis M, Aggelis G: Compositional shifts in lipid fractions during lipid turnover in Cunninghamella echinulata. Enz Microb Technol. 2007, 40: 1321-1327. 10.1016/j.enzmictec.2006.10.005.View ArticleGoogle Scholar
- Viikari L, Vehmaanpera J, Koivula A: Lignocellulosic ethanol: from science to industry. Biomass Bioenerg. 2012, 46: 13-24.View ArticleGoogle Scholar
- McMillan JD, Jennings EW, Mohagheghi A, Zuccarello M: Comparative performance of precommercial cellulases hydrolyzing pretreated corn stover. Biotechnol Biofuel. 2011, 4: 29-10.1186/1754-6834-4-29.View ArticleGoogle Scholar
- Strauss MLA, Jolly NP, Lambrechts MG, Van Rensburg P: Screening for the production of extracellular hydrolytic enzymes by non-saccharomyces wine yeasts. J Appl Microbiol. 2001, 91: 182-190. 10.1046/j.1365-2672.2001.01379.x.View ArticleGoogle Scholar
- Chisti Y: Biodiesel from microalgae. Biotech Adv. 2007, 25: 294-306. 10.1016/j.biotechadv.2007.02.001.View ArticleGoogle Scholar
- Bligh EG, Dyer WJ: A rapid method of total lipid extraction and purification. Can J Biochem Phys. 1959, 37: 911-917.View ArticleGoogle Scholar
- Freedman B, Butterfield R, Pryde E: Transesterification kinetics of soybean oil. J Am Oil Chem Soc. 1986, 63: 1375-1380. 10.1007/BF02679606.View ArticleGoogle Scholar
This article is published under license to BioMed Central Ltd. This is an Open Access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/2.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly credited.