ARTICLE HIGHLIGHTS
Oleaginous yeasts were isolated from the megadiverse canopies of the Mount Makiling Forest Reserve.
The lipid production of the isolates was tested in a nitrogen-limiting medium with glucose as the sole carbon source.
The capabilities of NFR6 and BUB8 for lipid production from glycerol were tested.
BUB8 belongs to the Rhodotorula genera with R. paludigena as its closest relative.
1. INTRODUCTION
Alternative sustainable energy resources are actively being explored due to the imminent exhaustion of fossil fuel reserves and global calls to address climate change. Countries have been crafting policies to decrease dependence on petroleum-based fuels and to lessen emissions of greenhouse gases. Biofuels, unlike fossil fuels, are sustainable and renewable [1]. Biodiesel, a first-generation biofuel is produced mostly from the conversion of plant oils into fatty acid methyl esters (FAME) through transesterification (or transmethylation). Oils from crops such as soy, rapeseed, or palm tend to have a high oleic acid content, which provides ideal ignition properties and oxidative stability [2,3]. Biodiesel has a similar combustion property to its traditional counterpart and is considered environmentally sustainable and eco-friendly, due to the reduced carbon dioxide emission and absence of sulfur and aromatic content associated with its production and use [4,5].
The Philippines has begun to explore domestically produced biodiesel to minimize dependency on imported petroleum and to promote energy security. The country has been using coconut methyl ester-blended biodiesel in compliance with Republic Act 9367 also known as Biofuels Act of 2006. However, the utilization of vegetable oils for biodiesel production compromises food security as it directly competes with the utilization of agricultural lands that would otherwise be dedicated to food production. Although the Philippines has targeted higher eventual biodiesel blends, the mandated blends have remained at B2 (2%) and have stalled in recent years [6]. Many countries have difficulty meeting their target blends due to several production bottlenecks in expanding production. Industrial-scale production cannot be realized without diverting the land intended for food production. This shift will be triggered by the high demand for biofuels, which could ultimately lead to high food commodity prices and deforestation [7]. Because of the massive demand in the food sector, large-scale biodiesel production using plant oils will be unsustainable in the long haul [8]. Thus, new bioenergy sources should be explored to promote sustainability.
Microbial lipids or single-cell oils are now being explored as alternative feedstock for biodiesel production because they are quite similar to plant-based oils [9]. The use of microbial oils has numerous advantages: Short microbial life cycles; high productivities; low labor requirement; reduced dependency on site, season, and climate; amenability to genetic modifications for the specific products; and ease of scale-up [2,10,11]. These lipids can be obtained from oleaginous yeasts, bacteria, and microalga that are able to produce lipids more than 20% of their cell weight in the form of triacylglycerols (TAGs), which is the starting material for biodiesel production [12,13]. A number of these microbes can accumulate ~70–80% w/w lipids per dry cell weight under set cultivation conditions [14]. Among these groups of oleaginous microbes, oleaginous yeasts are perceived to be the most versatile because of their capability to utilize a wide range of substrates including agricultural residues and industrial wastes [15]. They can consume both hexoses and pentoses from various sources, along with non-sugar carbon substrates [16]. Lipid production starts when the yeast proliferates in a medium lacking key non-carbon nutrients such as nitrogen. Nitrogen-limited conditions prevent cell proliferation by inhibiting protein and nucleic acid synthesis, forcing the yeast to alternatively amass intracellular lipids which are then stored in cytoplasmic inclusion bodies in the form of triglycerides [2]. Hydrophilic substrates can be readily metabolized by oleaginous yeasts into lipids through the de novo lipid biosynthetic route whereas only a few strains can accumulate lipids from hydrophobic substrates through the ex novo route [Figure 1] [8].
 | Figure 1: Lipid biosynthesis via the de novo and ex novo pathways. Dotted lines indicate conversion of the substrates through the ex novo route. [Click here to view] |
Oleaginous yeasts have been discovered from different environments such as soil and plant parts [17]. Forest canopies are home to a diversity of yeast species, making them a suitable location for the discovery of novel microorganisms. For instance, the Mount Makiling Forest Reserve (MMFR) has a rich biological diversity consisting of endemic indigenous and introduced exotic species. In fact, a novel microorganism that produces extracellular lipase was isolated from a fern thriving in a tree canopy in the MMFR [18]. The rich yeast diversity in forest canopies increases the probability to isolate new yeast strains with high-value yeast oils. In this study, the MMFR canopies were explored to isolate and identify oleaginous yeasts which can be potentially used for biodiesel production using nitrogen-limited medium. Succeeding analyses were focused on the selection of the best oleaginous yeast in terms of biomass and lipid production, FAME profile and Fourier transform infrared spectroscopy (FTIR) analyses of the extracted and transmethylated lipids were carried out to establish the suitability of the strain as a biodiesel feedstock. The best-performing isolate was identified through molecular analyses.
2. MATERIALS AND METHODS
2.1. Sample Collection
Samples were collected along the two-hectare long-term ecological plot in the Molawin-Dampalit sub-watershed in MMFR. The forest reserve is geographically located in the southern part of Metro Manila, Luzon Island, Philippines at 14°08’14” N and 121°11’33” E [19]. Soil and various tree materials such as bark, canopy leaves, and epiphytic plants were collected from the identified dominant canopy trees that are distributed across the whole plot area. The set of trees included species of Pterocarpus indicus (Narra), Meliosma pinnata (Balilang Uak), Parashorea malaanonan (Bagtikan), Ficus benjamina (Salisi), Ficus balete (Balete), and Syzygium decipiens (Malaruhat Pula). Samples were aseptically collected and placed in sterile plastics, contained in a jar cooler for transport, and stored in ice until further processing.
After collection, samples were serially diluted in 0.85% sodium chloride solution and 10-3 to 10-5 dilutions were spread-plated on Dichloran Rose Bengal Chloramphenicol agar (per liter of distilled water: 10.0 glucose; 5.0 g peptone; 1.0 KH2PO4; 0.5 g MgSO4•6H2O; 0.5 mL 5% w/v rose bengal solution; 1.0 mL 0.2% w/v dichloran in ethanol solution; 0.1 g chloramphenicol; and 15.0 g agar) [20]. Single unique colonies were selected and streaked onto yeast extract peptone dextrose (YEPD) agar (per liter of distilled water: 10.0 g yeast extract; 20.0 g peptone; 20.0 g glucose; and 16.0 agar) and incubated at 28°C for 24 h. Streaking for isolation was performed thrice to ensure the purity of the isolates.
2.2. Screening for Oleaginous Yeasts
The purified yeasts were grown on YEPD agar plates for 96 h at 30°C. Replica plates were made by transferring the colonies from the initial YEPDA to a ~90-mm round filter paper. The filter paper was dried at 60°C for 15 min and then stained with 0.08% Sudan Black B (in 96% ethanol) with mild shaking (KS 260 Basic Orbital Shaker, IKA® Works, Selangor, Malaysia) for 20 min at room temperature. The filter was then washed 2x with ethanol for 5 min. Blue colonies were selected as potentially oleaginous yeasts [21,22].
2.3. Cultivation of Yeasts for Microbial Oil Production
Selected oleaginous yeasts were cultured in flasks with 30 mL of YEPD broth. Cultures were incubated at 150 rpm at 30°C for 24 h in a Thomas Shaking Incubator (Thomas Kagaku Co., Japan). A volume of the seed culture equivalent to 1.0 absorbance unit at 600 nm was harvested by centrifugation at 2000 × g for 15 min (CD-50SR, Tomy Seiko, Japan). The resulting cell pellet was resuspended in 5 mL of nitrogen-limited medium (per liter: 2.0 g (NH4)2SO4; 7.0 g KH2PO4; 2.0 g NaH2PO4; 1.50 g MgSO4•6H2O; and 1.0 g yeast extract) either with glucose or glycerol as C source (40 g/L) and finally transferred to 45 mL of the same medium [14,23]. The flasks were incubated in a rotary shaker at 30°C for 120 h at 150 rpm agitation. 10-mL samples were then taken to determine biomass and lipid concentrations.
To measure biomass concentration, samples were centrifuged at 10,000 × g for 5 min. The resulting cell pellets were washed twice with distilled water and resuspended in water. Cell suspensions were dried at 105°C to constant weight.
2.4. Lipid Extraction
Samples were centrifuged for 5 min at 10,000 × g and washed 2× with distilled H2O. The pellets were resuspended in 2 mL of 4 M hydrochloric acid and incubated for 2 h at 60°C with mild shaking to hydrolyze the biomass. The acid-hydrolyzed masses were transferred to 20 mL of chloroform-methanol (1:1 v/v) mixture and mildly agitated at RT for 3 h. The acid mixtures were centrifuged at 2000 × g for 5 min to separate the organic and aqueous layers [24]. The lower organic layers containing the lipids were recovered using a Pasteur pipette and evaporated in a low-temperature vacuum oven and dried at 70°C and 12 psi until constant weight. All solvents used for lipid extraction, including hydrochloric acid (RCI Labscan Limited, Thailand), chloroform (RCI Labscan Limited, Thailand), and methanol (J.T. Baker®, Mumbai, India) were prepared from analytical grade reagents. All experiments were carried out in triplicates. The total lipid was expressed as the amount of lipids (g/L) extracted from yeast biomass whereas lipid content (YL/X) was expressed as the total lipids per dry biomass. The lipid content was calculated using Equation 1 wherein Xi and Li are the final dry cell weight and lipid concentration, respectively. X0 and L0 are the initial dry cell weight and lipid concentration, respectively.
 |
The one-way analysis of variance module of IBM® SPSS® Statistics 25 (IBM Corp. Armonk, New York) was used for statistical analysis. Tukey’s honestly significant difference (HSD) test for post hoc comparison was applied to find means that are significantly different from each other (P o find; followed by the compact letter display (CLD) method to identify variables that have statistically different means from the ones that do not have statistically different means. Variables were labeled with letters and were only considered as distinguishable if and only if they do not share at least one letter.
2.5. Lipid Transmethylation
The oleaginous strains which exhibited maximum lipid accumulation were selected for transmethylation and fatty acid profile analysis. Fatty acids were converted to FAME according to the AOAC Official Method 969.33 [25]. Briefly, 5 mL 1.0 N methanolic sodium hydroxide and 5 mL methanol were added to 0.5 g of the extracted lipid sample. The mixture was refluxed for 10 min. 10 mL of 14% boron trifluoride in methanol (Sigma-Aldrich, Sigma-Aldrich Pte Ltd, Singapore) was added to the mixture and refluxed again for 2 min. About 5 mL of n-heptane (Mallinckrodt, Ireland) was then added, and the mixture was refluxed further for 1 min. After heating, the mixture was allowed to cool for a few minutes. 15 mL of saturated NaCl solution was added to the mixture and shaken vigorously. After standing, the upper heptane layer was collected and added with 0.36 g anhydrous sodium sulfate to remove any excess moisture. The resulting product was subjected to gas chromatography.
2.6. Lipid Characterization
Qualitative and quantitative compositions of the fatty acids were analyzed using a Shimadzu GC-14B gas chromatograph (Shimadzu Corporation, Japan) equipped with a flame ionization detector and an Ulbon HR-SS-10 capillary column (50 m × 0.25 mm) (Shinwa Chemical Industries Ltd., Japan). 1 μL sample was injected into the machine and was carried by ultra-high purity N2 at a constant flow of 40 mL/min. Detector and injector temperatures were set at 260°C. The temperature program used for the separation of FAME is as: 160–220°C ramped at 1.5°C/min and held for 10 min once 220°C was reached. The retention times and peak areas of the samples were compared with FAME standards (Supelco 37 FAME Mix, Sigma-Aldrich Pte Ltd, Singapore). The FTIR spectra between the microbial oils before and after transmethylation were recorded on a Shimadzu IR Prestige-21 (Japan) using the standard attenuated total reflection method.
2.7. Morphological and Phylogenetic Characterization of Oleaginous Yeast Strain
The morphological attributes of the selected yeast strain were observed. Cell shape, dimension, and reproductive features of the strain grown in a liquid YM medium were also observed microscopically (Supplementary Method).
Yeast strains were identified by sequencing the 5.8S-ITS region [26]. Isolates cultured on YEPD medium for 24 h at 30 °C were subjected to DNA extraction using Quick DNA™ Fungal/Bacterial DNA Extraction Kit following the manufacturer’s specifications (Zymo Research, California). The fungus-specific universal primers ITS-1 (5′◊3’: TCCGTAGGTGAACCT GCGG) and ITS-4 (5’◊3’: TCCTCCGCTTATTGATATGC) were used to amplify the 5.8S-ITS region by polymerase chain reaction. The PCR mixture contained 1× PCR buffer, 1.5 mM MgCl2, 0.2 mM dNTP mix, 1.0 μM each of ITS-1 and ITS-4, 1.25 U Taq pol (Invitrogen), 200 ng DNA template, and to 50 μL, nuclease-free water. The PCR steps were as follows: (1) 95°C for 3 min; (2) 94°C for 1 min; (3) 60°C for 1 min; (4) 72°C for 2 min; (5) 72°C for 5 min; and (6) 4°C. Steps 2–4 were repeated for 30 cycles [27]. The amplicons were sent for sequencing to Macrogen (South Korea). The basic local alignment search tool of the National Center for Biotechnology Information (NCBI) (https://blast.ncbi.nlm.nih.gov/Blast.cgi) was used to search for related sequences and to identify the yeast isolates. A phylogram was generated using the QIAGEN CLC Sequence Viewer ver. 8.0 (https://digitalinsights.qiagen.com/). The ITS-containing region of BUB8 was deposited to NCBI with accession number OP808033.1.
3. RESULTS AND DISCUSSION
3.1. Isolation and Screening of Oleaginous Yeasts
A total of 258 colonies that appeared morphologically to be yeasts were isolated from the epiphytic ferns, moss, roots, orchids, bark, and leaves of several forest trees in the MMFR. Twenty-two colonies that were blue after staining with Sudan Black B were selected for further studies. The presence of lipids is usually demonstrated using fat stains such as Sudan Black B, Sudan III, or Nile red staining [28,29]. This staining technique, in combination with replica plating, serves as a qualitative screening method to select potential oleaginous yeasts isolated from different sources such as sludge, soil, and other environmental samples [30]. The putative oleaginous yeast isolates from this study were obtained from Narra, Bagtikan, and Malaruhat Pula trees. Oleaginous yeasts have been obtained from various natural habitats such as soils, plant and flower surfaces, mangroves, bark-beetles, tree exudates, and water, among others [31,32].
3.2. Biomass and Lipid Production of Selected Yeast Isolates
Biomass production, lipid accumulation, and lipid contents of the isolates were evaluated in a nitrogen-limited medium supplemented with glucose [Figure 2]. A nitrogen-limited medium was used since lipid accumulation is induced by nitrogen exhaustion. To be considered oleaginous, lipid content must be more than 20% of cell biomass [33]. BTB8, MPB8, MPB7, NFR2, and NF15 are not oleaginous yeasts due to their low lipid content (10.83 ± 0.52, 11.56 ± 0.70, 17.07 ± 6.32, 17.46 ± 2.16, and 19.52 ± 3.94%, respectively, all of which are statistically indistinguishable since they share the same CLD a). NFR9 was the best lipid-yielding strain (55.10 ± 1.70%) however, its biomass accumulation was low (1.68 ± 0.07 g/L). Ideally, oleaginous yeasts should be able to grow at high densities since lipids are extracted from the cells [34]. Among the oleaginous yeast isolates, NFR6 and BUB8 accumulated the most biomass (10.09 ± 0.12 and 8.87 ± 0.17 g/l, respectively) and had good lipid content. The two strains were thus selected for the succeeding study based on Tukey’s test and CLD.
 | Figure 2: Biomass and lipid production and lipid content of the isolates. Values that share the same italicized letters are not statistically different. [Click here to view] |
The biomass and lipid production of NFR6 and BUB8 were evaluated in the same medium but with glycerol as substrate [Figure 3]. BUB8 had significantly higher biomass (8.08 ± 0.19 g/l) and lipid (2.91 ± 0.12 g/l) production, with a lipid content of 36.03 ± 0.75%. The cultivation studies demonstrated that BUB8 is capable of both de novo and ex novo lipid accumulation. This is a remarkable attribute since it suggests that the isolated strain can utilize various materials including wastes of agricultural and industrial origins. Other microbes capable of utilizing both pathways include Yarrowia, Cryptococcus, Rhodospiridium, Geotrichum, and Trichosporon [35,36]. The utilization of glycerol as substrate will encourage circular bioeconomy for biodiesel processing. Crude glycerol is the major waste product of biodiesel processing using plant oils and biomass-based substrates with at least 10% for every batch of biodiesel [37]. This by-product has been used as an alternative feedstock for single-cell oil production by the oleaginous yeasts, Yarrowia lipolytica, Cryptococcus curvatus, Trichosporonoides spathulate, and Candida viswanathii [38-41]. Crude glycerol mixed with hemicellulosic hydrolysate was also evaluated for lipid production by R. toruloides [42].
 | Figure 3: Biomass and lipid production and lipid content with glycerol as sole carbon source. [Click here to view] |
Most oleaginous yeasts can utilize various sugar carbon sources via the de novo lipid biosynthesis pathway but only a few strains can accumulate lipids through the ex novo route [8]. There are major differences between the two pathways in terms of biochemical mechanisms. In the de novo pathway, nitrogen depletion in the culture medium causes cellular AMP to rapidly decrease. This leads to the accumulation of citric acid, which is then cleaved by ATP-citrate lyase to acetyl-COA. Acetyl-CoA is further converted to TAGs. It should be noted that ATP-citrate lyase is unique to oleaginous yeasts [43,44]. The de novo pathway is a non-growth-associated process. The ex novo pathway, on the other hand, is growth-associated and is entirely independent of nitrogen levels [9,14]. Hydrophobic substrates such as fatty acids, oils, and TAGs from the culture medium would result to the unmodified form of the lipids [44].
3.3. Characterization of Microbial Oil
The lipid content and fatty acid profiles differ among species; however, the main fatty acids made by oleaginous yeast are acknowledged to be similar to those of vegetable oils [33,45-47]. They consist of myristic (C14:0), palmitic (C16:0), palmitoleic (C16:1), stearic (C18:0), oleic (C18:1), linoleic (C18:2), and α-linolenic acid (C18:3). Notably, this is the oil composition (and the lipid classes of C16 to C18) that has been prescribed for biodiesel production [48,49]. In particular, the typical relative composition of lipids in oleaginous yeasts consists of oleic acid > palmitic acid > linoleic acid = stearic acid [50]. The fatty acid profile of strain BUB8 exhibited oleic acid as the most abundant fatty acid present, followed by palmitic acid, linoleic acid, and stearic acid [Table 1]. This profile closely resembles the TAGs produced from glycerol by R. toruloides [51].
Table 1: Fatty acid methyl ester profile of transmethylated lipid extracted from BUB8.
| Number of carbon atoms | Common name | Composition (% w/w) |
|---|---|---|
| C12 | Lauric acid | 0.07±0.01 |
| C14 | Myristic acid | 1.23±0.01 |
| C15 | Pentadecylic acid | 0.06±0.00 |
| C16:0 | Palmitic acid | 23.54±0.90 |
| C16:1 | Palmitoleic acid | 1.29±0.04 |
| C17:0 | Margaric acid | 0.14±0.01 |
| C17:1 | Heptadecenoic acid | 0.39±0.02 |
| C18:0 | Stearic acid | 4.68±0.10 |
| C18:1 | Oleic acid | 52.59±0.37 |
| C18:2 | Linoleic acid | 12.33±0.06 |
| C20:0 | Arachidic acid | 1.04±0.01 |
| C20:1 | Paullinic acid | 0.23±0.01 |
| C20:2 | Eicosadienoic acid | 0.23±0.01 |
| C20:3 | Mead acid | 0.39±0.01 |
| C20:4 | Arachidonic acid | 0.56±0.01 |
| C22 | Behenic acid | 0.46±0.01 |
| C24 | Lignoceric acid | 0.83±0.01 |
The transmethylation of the fatty acids was further confirmed by ATR-FTIR. The FTIR spectra (Appendix B) of pure and transmethylated microbial oil extracted from BUB8 showed the following peaks at 2850 and 2920 cm-1 corresponding to the symmetric and asymmetric stretching around the –CH of methylene groups; 1740 cm–1 (stretching of the ester carbonyl group); 1160 cm-1 (ester stretching (–CO)); and 720 cm-1 (unsaturated alkene) [52,53]. A band is present at 1430 cm-1 in the FAME spectra, which are attributed to asymmetric –CH3 bending vibration. The stretching of O=C–CH3 bonds, represented by peaks at around 1190 cm-1, typical in FAME but not in microbial oils, was also observed [53]. Only subtle differences can be observed between the spectra since the FAME sample is chemically similar to its precursor.
The fatty acid composition is a major determinant of the quality of biodiesel [54,55]. Fuel properties such as cetane number, heat of combustion, kinematic viscosity, melting point, and oxidative stability are assessed through fatty acid compositional analysis [56-58]. The presence of high levels of monounsaturated fatty acids is desirable for a biodiesel substrate because of the favorable balance they conferred on these parameters [5,59]. In assessing biodiesel feedstocks, a mixture of high amounts of monounsaturated fatty acids and a balanced level of saturated and polyunsaturated fatty acids is preferable for biodiesel production [60]. Studies have demonstrated that biodiesel production from oleic acid has the property ideal to be a diesel substitute and thus BUB8 would be a good contender for biodiesel production [61,62].
3.4. Characterization and Molecular Identification of BUB8
Due to its significant lipid productivities both in glucose and glycerol and FAME and FTIR profiles, BUB8 was considered for further identification and characterization. The colonies of BUB8 are pink in color, circular, smooth, shiny, opaque, convex with an entire margin, and measuring 1.0–2.0 mm in diameter. SEM imaging showed that BUB8 cells appear to be ellipsoidal to subcylindrical [Figure 4]. The cells are arranged in singles, pairs, and clusters, exhibiting unipolar budding. Furthermore, the cells measure 3.0–5.0 × 2.0–3.0 μm (length × width) [Figure 4].
 | Figure 4: Appearance of BUB8 cells in petri plate (a) and under SEM (b). [Click here to view] |
ITS-PCR is widely used for the rapid identification and taxonomic study of yeasts from various sources [27,63-65]. To identify strain BUB8, the 5.8S-ITS rDNA was amplified, sequenced, and compared against available sequences. Phylogenetic analysis showed that BUB8 shares 99% identity with Rhodotorula paludigena [Figure 5]. The evolutionary tree showed the relationship of Rhodotorula spp. with related genera such as Sporidiobolus and Sporobolomyces, all of which belong to the Order Sporidiobolales, subphylum Pucciniomycotina. These genera are referred to as the “red yeasts,” producing pigmented colonies [66,67]. Rhodosporidiobolus is more closely related to Rhodotorula than to Sporobolomyces as shown in the tree [68]. Rhodotorula species are lipid-accumulating yeasts that have been considered to produce microbial oils as biodiesel feedstock [69-72].
 | Figure 5: Phylogeny of BUB8. The phylogram was constructed using the neighbor-joining method. GenBank accession numbers for the ITS regions are shown in brackets. The evolutionary distances were computed using the Kimura 2-parameter method. Bar, 0.110 substitutions per nucleotide position. Distance calculations were done based on 1000 replicates. [Click here to view] |
R. paludigena was only characterized as oleaginous in recent years, and thus, few publications on its lipid productivity were available [58]. In a recent study, a novel deep-sea strain, R. paludigena P4R5, can produce high levels of intracellular lipids (16.9 g/L) and extracellular mixture (48.5 g/L) of mannitol esters of 3-OH C14, C16 and C18 fatty acids simultaneously [73]. The intracellular lipids were composed mostly of C16 and C18 fatty acids, which could serve as substrates for biodiesel production. Likewise, the lipid production potential of another R. paludigena strain (CM33) was also evaluated using molasses and crude glycerol found to be suitable substrates for the microorganism [74]. When grown in molasses, it produced remarkable biomass (16.5 g/L) and lipid (6.1 g/L) concentrations with 37% lipid content. These results, together with the information provided by this study, highlight the potential of R. paludigena of contributing to a high-value carbon chain through the transformation of various substrates into lipids for biodiesel production.
4. CONCLUSION
Microbial oils are explored as substitute feedstock for the synthesis of biodiesel because they are comparable to vegetable oils in terms of fatty acid profiles. Oleaginous yeasts are promising microorganisms characterized by their significant intracellular accumulation of fatty acid in the form of triglycerides. The selection of potential oleaginous yeasts isolated from the MMFR was based on primary screening using Sudan Black stain, biomass, and lipid productivities. This study has identified a promising oleaginous strain isolated from the upper bark of a Bagtikan tree. It has presented both de novo and ex novo lipid biosynthesis, as it exhibited high lipid production using glucose and glycerol, respectively as substrates. FAME and FT-IR analyses have shown the presence of fatty acids, the most abundant of which is oleic acid, that are applicable for biodiesel production. Molecular identification of BUB8 using ITS-rDNA rDNA sequencing revealed 99% sequence homology with R. paludigena. This oleaginous yeast could be used for further optimization to harness its potential for biodiesel production and make it a better alternative to petroleum-based diesel. Likewise, this strain could kickstart developments of local microbial-based technology for sustainable biodiesel production in the Philippines to augment the production of coco-methyl ester biodiesel.
5. ACKNOWLEDGMENT
The authors would like to thank the Forest Canopy Observation Positioning and Investigation (FOREST CANOPI) Program: Developing Canopy Science in the Philippines funded by the Department of Science and Technology (DOST) for providing the biological samples.
6. AUTHOR CONTRIBUTIONS
IG Pajares and PJ Requiso performed microbiology, fermentation, and molecular biology experiments whereas LM Fabro conducted the chemical analyses. IG Pajares and KM Ramos prepared the manuscript. AK Raymundo is the first author’s dissertation adviser. All authors have seen and approved the manuscript and all its contents.
7. FUNDING
This study is funded by UPLB-BIOTECH CORE Project ID 16214.
8. CONFLICTS OF INTEREST
The authors report no financial or any other conflicts of interest in this work.
9. ETHICAL APPROVALS
This study does not involve experiments on animals or human subjects.
10. DATA AVAILABILITY
All the data obtained in the study are represented as table or figures available in the main text and appendices.
11. PUBLISHER’S NOTE
This journal remains neutral with regard to jurisdictional claims in published institutional affiliation.
REFERENCES
1. Mahlia TM, Syazmi ZA, Mofijur M, Abas AE, Bilad MR, Ong HC, et al. Patent landscape review on biodiesel production:Technology updates. Renew Sustain Energy Rev 2020;118:109526. [https://doi.org/10.1016/j.rser.2019.109526]
2. Sitepu IR, Garay LA, Sestric R, Levin D, Block DE, German JB, et al. Oleaginous yeasts for biodiesel:Current and future trends in biology and production. Biotechnol Adv 2014;32:1336-60. [https://doi.org/10.1016/j.biotechadv.2014.08.003]
3. Brandenburg J, Blomqvist J, Shapaval V, Kohler A, Sampels S, Sandgren M, et al. Oleaginous yeasts respond differently to carbon sources present in lignocellulose hydrolysate. Biotechnol Biofuels 2021;14:124. [https://doi.org/10.1186/s13068-021-01974-2]
4. Hill J, Nelson E, Tilman D, Polasky S, Tiffany D. Environmental, economic, and energetic costs and benefits of biodiesel and ethanol biofuels. Proc Natl Acad Sci U S A 2006;103:11206-10. [https://doi.org/10.1073/pnas.0604600103]
5. Knothe G. Dependence of biodiesel fuel properties on the structure of fatty acid alkyl esters. Fuel Process Technol 2005;86:1059-70. [https://doi.org/10.1016/j.fuproc.2004.11.002]
6. Mojica-Sevilla F. Biofuels Annual Country Report. US Department of Agriculture Foreign Agricultural Service Report No:RP2021-0075;2021. Available from:https://www.fas.usda.gov/data/philippines-biofuels-annual-6 [Last accessed on 2022 Sep 09].
7. Elder M, Hayashi S. A regional perspective on biofuels in Asia. In:Takeuchi K, Shiroyama H, Saito O, Matsuura M, editors. Biofuels and Sustainability. Science for Sustainable Societies. Tokyo:Springer;2018. 223-46. [https://doi.org/10.1007/978-4-431-54895-9_14]
8. Patel A, Matsakas L. A comparative study on de novo and ex novo lipid fermentation by oleaginous yeast using glucose and sonicated waste cooking oil. Ultrason Sonochem 2019;52:364-74. [https://doi.org/10.1016/j.ultsonch.2018.12.010]
9. Papanikolaou S, Aggelis G. Lipids of oleaginous yeasts. Part II:Technology and potential applications. Eur J Lipid Sci Technol 2011;113:1052-73. [https://doi.org/10.1002/ejlt.201100015]
10. Dourou M, Aggeli D, Papanikolaou S, Aggelis G. Critical steps in carbon metabolism affecting lipid accumulation and their regulation in oleaginous microorganisms. Appl Microbiol Biotechnol 2018;102:2509-23. [https://doi.org/10.1007/s00253-018-8813-z]
11. Mukhtar H, Suliman SM, Shabbir A, Mumtaz MW, Rashid U, Rahimuddin SA. Evaluating the potential of oleaginous yeasts as feedstock for biodiesel production. Protein Pept Lett 2018;25:195-201. [https://doi.org/10.2174/0929866525666180122112805]
12. Li Q, Du W, Liu D. Perspectives of microbial oils for biodiesel production. Appl Microbial Biotech 2008;80:749-56. [https://doi.org/10.1007/s00253-008-1625-9]
13. Kongruang S, Roytrakul S, Sriariyanun M. Renewable biodiesel production from oleaginous yeast biomass using industrial wastes. E3S Web Conf 2020;141:03010. [https://doi.org/10.1051/e3sconf/202014103010]
14. Papanikolaou S, Aggelis G. Lipids of oleaginous yeasts. Part I:Biochemistry of single cell oil production. Eur J Lipid Sci Technol 2011;113:1031-51. [https://doi.org/10.1002/ejlt.201100014]
15. Di Fidio N, Minonne F, Antonetti C, Galletti AM. Cutaneotrichosporon oleaginosus:A versatile whole-cell biocatalyst for the production of single-cell oil from agro-industrial wastes. Catalysts 2021;11:1291. [https://doi.org/10.3390/catal11111291]
16. Maina S, Pateraki C, Kopsahelis N, Paramithiotis S, Drosinos EH, Papanikolaou S, et al. Microbial oil production from various carbon sources by newly isolated oleaginous yeasts. Eng Life Sci 2017;17:333-44. [https://doi.org/10.1002/elsc.201500153]
17. Hoondee P, Wattanagonniyom T, Weeraphan T, Tanasupawat S, Savarajara A. Occurrence of oleaginous yeast from mangrove forest in Thailand. World J Microbiol Biotechnol 2019;35:108. [https://doi.org/10.1007/s11274-019-2680-3]
18. Elegado F, Legaspi CL, Paet JM, Querubin F, Tolentino JE, Vilela J, et al. Screening, identification and optimization of extracellular lipase production of yeast (Cryptococcus flavescens) isolated from a tree canopy fern in the Mount Makiling Forest Reserve, Philippines. AIP Conf Proc 2019;2155:020029. [https://doi.org/10.1063/1.5125533]
19. Gonzalez JC, de Guia AP, Dimalibot JC, Pantua KV, Gustilo WO, Bantayan NC. Understorey to canopy vertebrate fauna of a lowland evergreen forest in Mt. Makiling Forest Reserve, Philippines. Biodivers Data J 2020;8:e56999. [https://doi.org/10.3897/BDJ.8.e56999]
20. Dichloran Rose Bengal Chloramphenicol (DRBC) Agar. In:Corry JEL, Curtis GD, Baird RM, editors. Progress in Industrial Microbiology. Netherlands:Elsevier;1995. 303-5. [https://doi.org/10.1016/S0079-6352(05)80036-0]
21. Evans CT, Ratledge C, Gilbert SC. A rapid screening method for lipid-accumulating yeast using a replica-printing technique. J Microbiol Methods 1985;4:203-10. [https://doi.org/10.1016/0167-7012(85)90038-7]
22. Thancharoen K, Malasri A, Leamsingkorn W, Boonyalit P. Selection of oleaginous yeasts with lipid accumulation by the measurement of Sudan black B for benefits of biodiesel. Int J Pharm Med Biol Sci 2017;6:53-7. [https://doi.org/10.18178/ijpmbs.6.2.53-57]
23. 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;47:215-20.
24. Neema PM, Kumari A. Isolation of lipid producing yeast and fungi from secondary sewage sludge and soil. Aust J Basic Appl Sci 2013;7:283-8.
25. Association of Official Analytical Chemists. AOAC Official Method 969.33. Fatty Acids in Oils and Fats:Preparation of Methyl Esters-Boron Trifluoride Method. Official Methods of Analysis of AOAC International. 19th ed. Gaithersburg, Maryland, USA:AOAC International;2012.
26. Jiru TM, Abate D, Kiggundu N, Pohl C, Groenewald M. Oleaginous yeasts from Ethiopia. AMB Express 2016;6:78. [https://doi.org/10.1186/s13568-016-0242-8]
27. Heslam AE, Wambui V, Ogola JO, Maina JM. Phylogenetic analysis of isolated biofuel yeasts based on 5.8S-ITS rDNA and D1/D2 26S rDNA sequences. J Genet Eng Biotechnol 2014;12:37-43. [https://doi.org/10.1016/j.jgeb.2014.01.001]
28. Nouri H, Moghimi H, Rad MN, Ostovar M, Mehr SS, Ghanaatian F, et al. Enhanced growth and lipid production in oleaginous fungus, Sarocladium kiliense ADH17:Study on fatty acid profiling and prediction of biodiesel properties. Renew Energy 2019;135:10-20. [https://doi.org/10.1016/j.renene.2018.11.104]
29. Ayadi I, Belghith H, Gargouri A, Guerfali M. Screening of new oleaginous yeasts for single cell oil production, hydrolytic potential exploitation and agro-industrial by-products valorization. Process Saf Environ Prot 2018;119:104-14. [https://doi.org/10.1016/j.psep.2018.07.012]
30. Jape A, Harsulkar A, Sapre VR. Modified Sudan black B staining method for rapid screening of oleaginous marine yeasts. Int J Curr Microbiol Appl Sci 2014;3:41-6.
31. Paserakung A, Pattarajinda V, Vichitphan K, Froetschel MA. Selection and identification of oleaginous yeast isolated from soil, animal feed and ruminal fluid for use as feed supplement in dairy cattle. Lett Appl Microbiol 2015;61:325-32. [https://doi.org/10.1111/lam.12475]
32. Caporusso A, Capece A, De Bari I. Oleaginous yeasts as cell factories for the sustainable production of microbial lipids by the valorization of agri-food wastes. Fermentation 2021;7:50. [https://doi.org/10.3390/fermentation7020050]
33. Beopoulos A, Nicaud JM. Yeast:A new oil producer?OCL 2012;19:22-8. [https://doi.org/10.1051/ocl.2012.0426]
34. Lamers D, van Biezen N, Martens D, Peters L, van de Zilver E, Jacobs-van Dreumel N, et al. Selection of oleaginous yeasts for fatty acid production. BMC Biotechnol 2016;16:45. [https://doi.org/10.1186/s12896-016-0276-7]
35. Beopoulos A, Chardot T, Nicaud JM. Yarrowia lipolytica:A model and a tool to understand the mechanisms implicated in lipid accumulation. Biochimie 2009;9:692-6. [https://doi.org/10.1016/j.biochi.2009.02.004]
36. Papanikolaou S, Aggelis G. Yarrowia lipolytica:A model microorganism used for the production of tailor-made lipids. Eur J Lipid Sci Technol 2010;112:639-54. [https://doi.org/10.1002/ejlt.200900197]
37. Molis MH, Cheng S, Cross JS. Current and future trends for crude glycerol upgrading to high value-added products. Sustainability 2023;15:2979. [https://doi.org/10.3390/su15042979]
38. Papanikolaou S, Aggelis G. Lipid production by Yarrowia lipolytica growing on industrial glycerol in a single-stage continuous culture. Bioresour Technol 2002;82:43-9. [https://doi.org/10.1016/S0960-8524(01)00149-3]
39. Liang Y, Cui Y, Trushenski J, Blackburn JW. Converting crude glycerol derived from yellow grease to lipids through yeast fermentation. Bioresour Technol 2010;101:7581-6. [https://doi.org/10.1016/j.biortech.2010.03.087 https://doi.org/10.1016/j.biortech.2010.04.061]
40. Kitcha S, Cheirsilp B. Enhancing lipid production from crude glycerol by newly isolated oleaginous yeasts:Strain selection, process optimization, and fed-batch strategy. BioEnergy Res 2013;6:300-10. [https://doi.org/10.1007/s12155-012-9257-4]
41. Guerfali M, Ayadi I, Sassi HE, Belhassen A, Gargouri A, Belghith H. Biodiesel-derived crude glycerol as alternative feedstock for single cell oil production by the oleaginous yeast Candida viswanathii Y-E4. Ind Crops Prod 2020;145:112103. [https://doi.org/10.1016/j.indcrop.2020.112103]
42. Chmielarz M, Blomqvist J, Sampels S, Sandgren M, Passoth V. Microbial lipid production from crude glycerol and hemicellulosic hydrolysate with oleaginous yeasts. Biotechnol Biofuels 2021;14:65. [https://doi.org/10.1186/s13068-021-01916-y]
43. Adrio JL. Oleaginous yeasts:Promising platforms for the production of oleochemicals and biofuels. Biotechnol Bioeng 2017;114:1915-20. [https://doi.org/10.1002/bit.26337]
44. Ratledge C, Wynn JP. The biochemistry and molecular biology of lipid accumulation in oleaginous microorganisms. Adv Appl Microbiol 2002;51:1-51. [https://doi.org/10.1016/S0065-2164(02)51000-5]
45. Gientka I, Gadaszewska M, Blazejak S, Kieliszek M, Bzducha-Wrobel A, Stasiak-Rozanska L, et al. Evaluation of lipid biosynthesis ability by Rhodotorula and Sporobolomyces strains in medium with glycerol. Eur Food Res Technol 2017;243:275-86. [https://doi.org/10.1007/s00217-016-2742-9]
46. Johnravindar D, Karthikeyan OP, Selvam A, Murugesan K, Wong JW. Lipid Accumulation potential of oleaginous yeasts:A comparative evaluation using food waste leachate as a substrate. Bioresour Technol 2018;248:221-8. [https://doi.org/10.1016/j.biortech.2017.06.151]
47. Behera AR, Dutta K, Verma P, Daverey A, Sahoo DK. High lipid accumulating bacteria isolated from dairy effluent scum grown on dairy wastewater as potential biodiesel feedstock. J Environ Manag 2019;252:109686. [https://doi.org/10.1016/j.jenvman.2019.109686]
48. Steen EJ, Kang Y, Bokinsky G, Hu Z, Schirmer A, McClure A, et al. Microbial production of fatty-acid-derived fuels and chemicals from plant biomass. Nature 2010;463:559-62. [https://doi.org/10.1038/nature08721]
49. Singh S, Pandey D, Saravanabhypathy S, Daverey A, Dutta K, Arunachalam K. Liquid wastes as a renewable feedstock for yeast biodiesel production:Opportunities and challenges. Environ Res 2022;207:112100. [https://doi.org/10.1016/j.envres.2021.112100]
50. Ageitos JM, Vallejo JA, Veiga-Crespo P, Villa TG. Oily yeasts as oleaginous cell factories. Appl Microbiol Biotechnol 2011;90:1219-27. [https://doi.org/10.1007/s00253-011-3200-z]
51. Xu J, Zhao X, Wang W, Du W, Liu D. Microbial conversion of biodiesel byproduct glycerol to triacylglycerols by oleaginous yeast Rhodosporidium toruloides and the individual effect of some impurities on lipid production. Biochem Eng J 2012;65:30-6. [https://doi.org/10.1016/j.bej.2012.04.003]
52. Wade LG. Organic Chemistry. India:Pearson Education;2008.
53. Rabelo SN, Ferraz VP, Oliveira LS, Franca AS. FTIR analysis for quantification of fatty acid methyl esters in biodiesel produced by microwave-assisted transesterification. Int J Environ Sci Dev 2015;6:964-9. [https://doi.org/10.7763/IJESD.2015.V6.730]
54. Tiwari AK, Kumar A, Raheman H. Biodiesel production from Jatropha oil (Jatropha curcas) with high free fatty acids:An optimized process. Biomass Bioenergy 2007;31:569-75. [https://doi.org/10.1016/j.biombioe.2007.03.003]
55. Wang R, Song B, Zhou W, Zhang Y, Hu D, Bhadury PS, et al. A facile and feasible method to evaluate and control the quality of Jatropha curcas L. seed oil for biodiesel feedstock:Gas chromatographic fingerprint. Appl Energy 2011;88:2064-70. [https://doi.org/10.1016/j.apenergy.2010.12.078]
56. Demirbus A. Relationships derived from physical properties of vegetable oil and biodiesel fuels. Fuel 2008;87:1743-8. [https://doi.org/10.1016/j.fuel.2007.08.007]
57. Knothe G. Improving biodiesel fuel properties by modifying fatty ester composition. Energy Environ Sci 2009;2:759-66. [https://doi.org/10.1039/b903941d]
58. Ramos MJ, Fernandez CM, Casas A, Rodriguez L, Perez A. Influence of fatty acid composition of raw materials on biodiesel properties. Bioresour Technol 2009;100:261-8. [https://doi.org/10.1016/j.biortech.2008.06.039]
59. Knothe G. Designer biodiesel:Optimizing fatty acid ester composition to improve fuel properties. Energy Fuels 2008;22:1358-64. [https://doi.org/10.1021/ef700639e]
60. Garay LA, Sitepu IR, Cajka T, Chandra I, Shi S, Lin T, et al. Eighteen new oleaginous yeast species. J Ind Microbiol Biotechnol 2016;43:887-900. [https://doi.org/10.1007/s10295-016-1765-3]
61. Kusmiyati K, Agung S. Production of biodiesel from oleic acid and methanol by reactive distillation. Bull Chem React Eng Catal 2010;5:1-6. [https://doi.org/10.9767/bcrec.5.1.37.1-6 https://doi.org/10.9767/bcrec.5.1.7103.1-6]
62. Nakkash NB, Al-Karkhi SR. Production of biodiesel fuel from oleic acid and comparison of its properties with petroleum diesel. Iraqi J Chem Pet Eng 2012;13:13-25. [https://doi.org/10.31699/IJCPE.2012.4.2]
63. Heras-Vazquez FJ, Mingorance-Cazorla L, Clemente-Jimenez JM, Rodriguez-Vico F. Identification of yeast species from orange fruit and juice by RFLP and sequence analysis of the 5.8S rRNA gene and the two internal transcribed spacers. FEMS Yeast Res 2003;3:3-9. [https://doi.org/10.1111/j.1567-1364.2003.tb00132.x]
64. Nisiotou AA, Nychas GJ. Yeast populations residing on healthy or Botrytis-infected grapes from a vineyard in Attica, Greece. Appl Environ Microbiol 2007;73:2765-8. [https://doi.org/10.1128/AEM.01864-06]
65. El-Sharoud WM, Belloch C, Peris D, Querol A. Molecular identification of yeasts associated with traditional Egyptian dairy products. J Food Sci 2009;74:M341-6. [https://doi.org/10.1111/j.1750-3841.2009.01258.x]
66. Mannazzu I, Landolfo S, da Silva TL, Buzzini P. Red yeasts and carotenoid production:Outlining a future for non-conventional yeasts of biotechnological interest. World J Microbiol Biotechnol 2015;31:1665-73. [https://doi.org/10.1007/s11274-015-1927-x]
67. Kot AM, Kieliszek M, Piwowarek K, Blazejak S, Mussagy CU. Sporobolomyces and Sporidiobolus-non-conventional yeasts for use in industries. Fungal Biol Rev 2021;37:41-58. [https://doi.org/10.1016/j.fbr.2021.06.001]
68. Li CJ, Zhao D, Li BX, Zhang N, Yan JY, Zou HT. Whole genome sequencing and comparative genomic analysis of oleaginous red yeast Sporobolomyces pararoseus NGR identifies candidate genes for biotechnological potential and ballistospores-shooting. BMC Genomics 2020;21:181. [https://doi.org/10.1186/s12864-020-6593-1]
69. Amaretti A, Raimondi S, Sala M, Roncaglia L, De Lucia M, Leonardi A, et al. Single cell oils of the cold-adapted oleaginous yeast Rhodotorula glacialis DBVPG 4785. Microb Cell Fact 2010;9:73. [https://doi.org/10.1186/1475-2859-9-73]
70. Enshaeieh M, Abdoli A, Madani M. Single cell oil (SCO) production by Rhodotorula mucilaginosa and its environmental benefits. J Agric Sci Technol 2015;17:387-400.
71. Jiru TM, Steyn L, Pohl C, Abate D. Production of single cell oil from cane molasses by Rhodotorula kratochvilovae (syn, Rhodosporidium kratochvilovae) SY89 as a biodiesel feedstock. Chem Cent J 2018;12:91. [https://doi.org/10.1186/s13065-018-0457-7]
72. Maza DD, Viñarta SC, García-Ríos E, Guillamón JM, Aybar MJ. Rhodotorula glutinis T13 as a potential source of microbial lipids for biodiesel generation. J Biotechnol 2021;331:14-8. [https://doi.org/10.1016/j.jbiotec.2021.03.002]
73. Wang M, Mao W, Wang X, Li F, Wang J, Chi Z, et al. Efficient simultaneous production of extracellular polyol esters of fatty acids and intracellular lipids from inulin by a deep-sea yeast Rhodotorula paludigena P4R5. Microb Cell Fact 2019;18:149. [https://doi.org/10.1186/s12934-019-1200-3]
74. Gosalawit C, Imsoonthornruksa S, Gilroyed BH, Mcnea L, Boontawan A, Ketudat-Cairns M. The potential of the oleaginous yeast Rhodotorula paludigena CM33 to produce biolipids. J Biotechnol 2021;329:56-64. [https://doi.org/10.1016/j.jbiotec.2021.01.021]