Boosting accumulation of neutral lipids in Rhodosporidium kratochvilovae HIMPA1 grown on hemp (Cannabis sativa Linn) seed aqueous extract as feedstock for biodiesel production

Bioresource Technology xxx (2014) xxx–xxx

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Boosting accumulation of neutral lipids in Rhodosporidium kratochvilovae HIMPA1 grown on hemp (Cannabis sativa Linn) seed aqueous extract as feedstock for biodiesel production Alok Patel, Mohammad Pravez, Farha Deeba, Vikas Pruthi, Rajesh P. Singh, Parul A. Pruthi ⇑ Molecular Microbiology Laboratory, Biotechnology Department, Indian Institute of Technology Roorkee (IIT R), Roorkee, Uttarakhand 247667, India

h i g h l i g h t s

g r a p h i c a l a b s t r a c t

 Novel oleaginous yeast isolated from

Himalayan permafrost soil.  Hemp seed extract used as cheap

triglyceride feedstock for biodiesel production.  Sonicated HSAE had 10–15% higher lipid yield than GSM and lacks toxic components.  TAG accumulated supersized lipid droplet of 5.95 ± 1.05 lm visualized by BODIPY stain.  Unique FAME profile of this yeast showed C20:0, C22:0, C27:0 similar to algal oils.

a r t i c l e

i n f o

Article history: Received 3 January 2014 Received in revised form 25 March 2014 Accepted 26 March 2014 Available online xxxx Keywords: Rhodosporidium kratochvilovae HIMPA1 Triacylglyceride Sonicated hemp seed aqueous extract BODIPY Biodiesel

a b s t r a c t Hemp seeds aqueous extract (HSAE) was used as cheap renewable feedstocks to grow novel oleaginous yeast Rhodosporidium kratochvilovae HIMPA1 isolated from Himalayan permafrost soil. The yeast showed boosted triglyceride (TAG) accumulation in the lipid droplets (LDs) which were transesterified to biodiesel. The sonicated HSAE prepared lacked toxic inhibitors and showed enhanced total lipid content and lipid yield 55.56%, 8.39 ± 0.57 g/l in comparison to 41.92%, 6.2 ± 0.8 g/l from industrially used glucose synthetic medium, respectively. Supersized LDs (5.95 ± 1.02 lm) accumulated maximum TAG in sonicated HSAE grown cells were visualized by fluorescent BODIPY (505/515 nm) stain. GC–MS analysis revealed unique longer carbon chain FAME profile containing Arachidic acid (C20:0) 5%, Behenic acid (C22:0) 9.7%, Heptacosanoic acid (C27:0) 14.98%, for the first time in this yeast when grown on industrially competent sonicated HSAE, showing more similarity to algal oils. Crown Copyright Ó 2014 Published by Elsevier Ltd. All rights reserved.

1. Introduction Worldwide concern on the obtainability of fossil fuel, depleting oil reserves and skyrocketing petroleum oil prices has ignited widespread interest in unconventional renewable energy ⇑ Corresponding author. Tel.: +91 1332 285530 (O), +91 1332 285110 (R), +91 9760214585 (Mobile); fax: +91 1332273560. E-mail address: [email protected] (P.A. Pruthi).

resources. To combat this challenge, diverse raw material sources are viewed as feedstocks. Depending on the source of the raw material used biofuels obtained from biomass feedstocks can be classified into triglyceride-based biomass, starch or sugar derived biomass, and lignocellulosic biomass (Sawangkeaw and Ngamprasertsith, 2013; Atabani et al., 2013). Among them, triglyceridesbased biomass utilizes raw material obtained from sources such as vegetable oils, animal fat, and microbial oil are considered as prominent feedstock for its conversion into biodiesel (Yan et al.,

http://dx.doi.org/10.1016/j.biortech.2014.03.142 0960-8524/Crown Copyright Ó 2014 Published by Elsevier Ltd. All rights reserved.

Please cite this article in press as: Patel, A., et al. Boosting accumulation of neutral lipids in Rhodosporidium kratochvilovae HIMPA1 grown on hemp (Cannabis sativa Linn) seed aqueous extract as feedstock for biodiesel production. Bioresour. Technol. (2014), http://dx.doi.org/10.1016/ j.biortech.2014.03.142

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A. Patel et al. / Bioresource Technology xxx (2014) xxx–xxx

2014; Singh et al., 2014; Russo et al., 2012). Compared to vegetable oils and animal fats, oleaginous microbes possessing triglyceride (TAG) accumulating ability have an edge over them. They can be easily cultivated under controlled environment conditions and easier to scale up for higher productivity and above all they do not compete ethically or economically with food crops (Liang and Jiang, 2013). TAGs are the major form of storage neutral lipids present in intracellular lipid bodies of the organism having similar fatty acid composition from yeast to man and are reported to be upcoming biodiesel feedstock for industrial acclimatization (Reue, 2011; Fei et al., 2008). The major advantage of using oleaginous yeast for the production of TAG feedstock is that they can grow on various kind of simple to complex sugars like lignocellulose hydrolysates as substrates and convert them into TAG at optimized conditions (Chen et al., 2009; Meng et al., 2009). Abundance of lignocellulosic raw materials, their sustainability and availability at cheaper cost makes them unique for its usage as TAG feedstock (Papanikolaou and Aggelis, 2011; Zhu et al., 2008, 2012). However, to obtain fermentable sugars from lignocelluloses, a harsh physical or chemical process like steam explosion or dilute acid pretreatment is applied. These pretreatment processes produce degradation products including acetic acid, furfural and 5-hydroxymethylfurfural (HMF) which strongly inhibit microbial fermentation (Dam et al., 1986; Liu et al., 2012). Recently, oleaginous yeast that can accumulate neutral lipids more than 70% of cell dry weight and also tolerate toxic inhibitors belongs to the genera of Rhodosporidium sp., Rhodotorula sp. and Lipomyces sp. (65%) including strains of Rhodotorula glutinis (72%), Rhodosporidium toruloides (48–67.5%) have been reported (Sitepu et al., 2013; Zhu et al., 2012; Chen et al., 2009; Yan et al., 2014). Industrial production of biodiesel from them depends on the availability of feedstock in both large quantities and at low cost. So chasing for a cheaper raw material resource to boost commercial biodiesel production is of current research interest. Industrial hemp (Cannabis sativa Linn.) seeds are considered as an inexpensive triacylglyceride feedstock as they possesses required features like high biomass content and high oil yield (0.14 and 0.70 t/ha) out of seed output 0.5–2 t/ha (Rehman et al., 2013). This crop effectively grows on arable land under diverse climatic conditions with seed oil content varying from 28% to 35% making it as an upcoming bioenergy crop for the production of bioethanol, biogas, biohydrogen, solid fuel, and biodiesel (Rehman et al., 2013; Kreuger et al., 2011; Latif and Anwar, 2009). Hemp seeds reported to have a concentrated balance of essential amino acids, essential fatty acids, along with its major vitamin and mineral content (Callaway, 2004). Earlier, researchers have shown the feasibility of converting hemp oil to produce hemp oil biodiesel (HOB) having comparable traits to meet ASTM 6751-09 standards (Li et al., 2010). Besides this, HOB had high content of PUFA and better transesterification efficiency for biodiesel production (Su et al., 2013; Da Porto et al., 2012). In this investigation we are reporting for the first time the usage of nontoxic HSAE prepared by sonication for accumulation of neutral lipids in the intracellular bodies of novel oleagenic yeast, Rhodosporidium kratochvilovae HIMPA1 (GenBank accession No. KF772881) isolated from Himalayan permafrost soil to produce biodiesel. The effect of ultrasonication as a mild pretreatment method was applied for efficient extraction of lipid, sugar and protein components in HSAE used for cultivation of the isolated oleaginous yeast. The accumulation of TAG content was visually monitored by intracellular labeling of neutral lipid with fluorescent BODIPY (505– 515 nm) stain at different time intervals imaged with digital inverted epiflourescent microscope (EVOS FL, AMG Group USA). In-vivo live cell imaging showed that accumulated TAG directly correlates with the lipid body size and can be studied without distorting the cellular integrity. Growth studies and nutritional requirements conditions producing maximum TAG in sonicated

HSAE were optimized and compared with industrially used glucose synthetic media (GSM). FAME profiles of biodiesel obtained were verified by TLC, FTIR and GCMS analysis, and its transesterification was done using BF3-Methanol method. This study opens up feasibility for preparation of FAME profile having longer hydrocarbon chain for biodiesel production via microbial interventions. 2. Methods 2.1. Material The raw material used in this study was hemp seeds obtained from local market of Roorkee, India. All solvents and reagents used in this study were HPLC grade. Standards for TLC (Triolein), Internal standard (Nonadecanoic acid) and FAME standard (AOCS low erucic rape seed oil O7756-1AMP) for GC–MS analysis were acquired from Sigma Aldrich (St. Louis, MO, USA). BODIPY 505/515 (4,4-difluro-1,3,5,7-tetramethyl-4-bora-3a, 4a-diaza-s-indacene) was purchased from Invitrogen (Life Technology, USA). For the cultivation of yeast strains growth media (YEPD), yeast nitrogen base (YNB) and its supplements were obtained from Himedia, India. 2.2. Isolation and screening of oleaginous yeast strain The permafrost soil samples collected from Tungnath Hill area (coordinates: 30.290 22°N 79.55°E), in the Himalayan Garhwal ranges, Uttarakhand (India) were subjected to serial dilutions in 0.9% sterilized saline water and cultured at 30 °C for 48 h on YEPD agar plates (g/l): glucose, 10; peptone, 5; yeast extract, 3; malt extract, 3; agar, 15. The YEPD plates were supplemented with ampicillin 5–10 lg/ml to avoid bacterial growth. Yeast colonies appeared on agar plates were picked and patched on the nitrogen limiting medium (NLM) agar plates containing Salt solution (g/l): KH2PO4, 1; MgSO4, 0.5; (NH4)2SO4, 0.1; CaCl2, 0.1; Trace elements (mg/l): boric acid, 0.5, CuSO4, 0.04, KI, 0.1; FeCl3, 0.2; MnSO4, 0.4; NaMO3, 0.2; ZnSO4 0.4; Vitamins: D-biotin, 0.002; calcium pantothenate, 0.4; folic acid, 0.002; inositol, 2; niacin, 0.4; PABA, 0.2; pyridoxine HCl, 0.4; riboflavin, 0.2; thiamine, 0.4 supplemented with glucose 20 g/l, pH was adjusted to 6.0 and the plates were incubated at 30 °C. The selected oleaginous yeast colonies possessing high amount of TAG accumulating ability were then grown on NLM broth to monitor cell and lipid droplet size (diameter) using BODIPY staining protocol as describe in Section 2.6.7 below. Images of their early stationary growth phase obtained after fluorescence microscopy was directly analyzed by ImageJ software. The media composition of glucose synthetic media (GSM) used in g/l: YNB (w/o amino acids), 6.7; Glucose, 20; KH2PO4, 1; (NH4)2SO4, 5.0. Amino acid mix (0.2 g/l) added to GSM had the composition L-Tryptophan, 0.02; L-Histidine, 0.02; L-Arginine, 0.02; L-Methionine, 0.02; L-Leucine, 0.03; L-Isoleucine, 0.03; L-Phenylalanine, 0.05; L-Valine, 0.15; L-Serine, 0.4; L-Threonine, 0.2; L-Glutamic Acids, 0.1; L-Aspartic acids, 0.1; L-Tyrosine, 0.1. 2.3. Identification and characterization of selected oleaginous yeast strain The selected oleaginous yeast strains were identified by BIOLOG Microbial Identification System, USA. Genotypic characterization was performed by sequencing the D1/D2 domains of the gene encoding subunit 26S of ribosomal RNA. The universal primers NL1F (50 GCATATCAATAAGCGGAGGAAAAG30 ) and NL4R (50 GGTC CGTGTTTCAAGACGG30 ) were used for D1/D2 amplification (Kurtzman and Robnett, 1998). The genomic DNA was extracted, amplified and sequenced. The sequences obtained were compared with the sequences available in the NCBI (http://www.ncbi.nlm.

Please cite this article in press as: Patel, A., et al. Boosting accumulation of neutral lipids in Rhodosporidium kratochvilovae HIMPA1 grown on hemp (Cannabis sativa Linn) seed aqueous extract as feedstock for biodiesel production. Bioresour. Technol. (2014), http://dx.doi.org/10.1016/ j.biortech.2014.03.142

A. Patel et al. / Bioresource Technology xxx (2014) xxx–xxx

nih.gov/), using BLAST programme. Gene specific primers Forward: 50 GCATATCAATAAGCGGAGGAAAAGA30 and Reverse: 50 CAAGACG GGTCGTTTAAAGTCA30 were used for confirming the PCR product of length 609 bp. To determine the taxonomic position and relationships phylogenetic tree of R. kratochvilovae HIMPA1was constructed using MEGA 6 software (Tamura et al., 2013). 2.4. Preparation of the hemp seeds aqueous extract (HSAE) Hemp seeds (5 g) were thoroughly washed twice with sterile distilled water and dried in oven at 60 °C overnight. The seeds were then finally grinded to powdered form in an electric grinder (Philips, India) for 2 min. The grinded samples were boiled in a 500 ml Erlenmeyer flask with 100 ml of sterile distilled water for 1 h and then filtered. The filtrate obtained was subjected to sonication at 20 kHz for 5 min to be used as lipid production medium for oleaginous yeast cultivation. 2.5. Batch cultivation of R. kratochvilovae HIMPA1for lipid production R. kratochvilovae HIMPA1 was cultivated at 30 °C on a rotary shaker at 200 rpm for 48 h using YEPD broth. To obtain the seed culture, cells were harvested by centrifugation, washed twice with sterilized distilled water and resuspended in 0.9% sterilized saline water to attain cell density of 6.5–7.8  108 cells/ml. Batch cultivation of R. kratochvilovae HIMPA1 for lipid production was performed in a 250 ml conical flask separately containing 100 ml each of sonicated HSAE and unsonicated HSAE, respectively inoculated with 2% of seed culture for 216 h at 30 °C with 200 rpm. In control experiment glucose synthetic medium (GSM) containing 30 g/l glucose in yeast nitrogen base (YNB) was taken. After fermentation, the cell dry weight (g), lipid yield (g/l) and lipid content (%) were determined. All experiments of lipid production were done in triplicate and results were averaged. The lipid yield is the amount of lipid extracted from the cells per litre of the fermentation medium (g/l) while the lipid content Y (%), was calculated using the following equation: Y = WL/DCW  100. Where WL is the weight of the total lipid obtained gravimetrically and DCW is the weight of the cell biomass. 2.6. Biochemical analysis of HSAE 2.6.1. Estimation of sugars, nitrogen, phosphorus and inhibitors concentration Total sugars and reducing sugars contents in HSAE were estimated using phenol sulphuric acid method and DNS method respectively. Total nitrogen was determined by the Kjeldahl method. Standard AOAC Official Methods of Analysis were used for estimation of total fiber content (soluble and insoluble contents) and other components. The concentration of inhibitors such as acetic acid, HMF and furfural in the sonicated HSAE were determined by HPLC (Waters Corp., USA) equipped with Waters 600 E pump, UV-PDA detector (Waters 996) and LiChroCART 250-4 C-18 column (5 lm particle size, 250  4.6 mm, Merck, Germany). 2.6.2. Dry cell weight determination The growth of yeast cells were observed by measuring cell optical density (OD) at 600 nm with the spectrophotometer. Cell biomass, expressed as dry cell weight (DCW) was harvested by centrifuging it at 3000 rpm for 5 min and its weight was determined gravimetrically after drying the cells in an oven at 105 °C for 12 h. 2.6.3. Total lipid extraction The total cellular lipid was extracted using modified protocol of Bligh and Dyer (1959). Briefly, 50 ml of culture broth was trans-

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ferred to Teflon-capped Pyrex tubes (16  100 mm), centrifuged at 3000 rpm for 5 min and the supernatant was removed. The pellet was washed twice with distilled water. The yeast cells were then sonicated at 20 kHz for 5 min followed by addition of 10 ml of chloroform:methanol (2:1; v/v) and stirred for 30 min. The extract so obtained was filtered with sintered glass funnel to which 5 ml of 0.034% MgCl2 was then added and centrifuged at 3000 rpm for 5 min. Upper aqueous layer was aspirated and the organic phase was washed twice with 1 ml of 2 N KCl/methanol (4:1, v/v) followed by addition of 5 ml of artificial upper phase (chloroform/methanol/water; 3:48:47, v/v/v) until the phase boundary becomes clear. Nonadecanoic acid (0.1 lg/ml) was used as internal standard. The bottom chloroform layer was transferred to a new screw cap test tube, and the lipid yield was determined gravimetrically.

2.6.4. TLC & FTIR analysis for neutral lipid determination TAG analysis of the extracted total lipid was carried out by using TLC (0.25-mm-thick silica gel G-60, F254) plates (Merck, India) and chromatograms were developed in hexane:diethyl ether:acetic acid (85:15:1, v/v/v) with triolein as standard. For quantification of SEs and TAGs, plates were dipped into methanolic MnCl2 solution (0.63 g MnCl24H2O, 60 ml water, 60 ml methanol and 4 ml concentrated sulphuric acid), dried and heated at 120 °C for 15 min (Fei et al., 2009). Conversion of TAG into FAMEs was detected by dual solvent system. The TLC plate was firstly developed to 2.5 cm from the origin with hexane:tert-butyl methyl ether:acetic acid (50:50:0.5, v/v/v), and after air dried, it was redeveloped to 8 cm from the origin with hexane:tert-butyl methyl ether:acetic acid (97:3:0.5, v/v/v) as a developing agent. The samples were visualized by spraying sulphuric acid 50% (w/w) and then heating at 135 °C and compared with triolein, used as standard (Ichihara and Fukubayashi, 1996). The lipid were also analyzed by FTIR spectrometer (Thermo Nicolet NEXUS, Maryland, USA). The range of spectrum was set from 400 to 4000 cm1 and triolein was used as standard.

2.6.5. Transesterification of fatty acids by methanolic BF3 Transesterification of extracted yeast lipid samples was done using the method of Morrison and Smith (1964). Briefly, fatty acid samples dried with anhydrous sodium sulphate were evaporated to dryness under nitrogen in a Teflon coated screw cap tube. Boron fluoride–methanol reagent was added under nitrogen, in the proportions 1 ml reagent per 4–16 mg of lipid. The samples were then heated at 80 °C for 20 min in a boiling water bath. The esters were extracted by adding 2 volumes of hexane and then 1 volume of water followed by centrifugation at 3500g until both layers were clearly visible.

2.6.6. Determination of FAMEs composition by GC–MS The products of transesterification were analyzed by GC–MS (Clarus 500, Perkin Elmer) using protocol of Härtig (2008). GC– MS analysis was done by injecting 1 lL of sample at 250 °C in splitless injection mode, and helium was used a carrier gas (1 ml/min). The column temperature was initially set at 50 °C and held for 1.5 min thereafter, the temperature was ramped to 180 °C (25 °C/ min) for 1 min, followed by a further increased to 220 °C (10 °C/ min), and held for 1 min. Finally, the temperature was increased to 250 °C (15 °C/min) and held for 3 min. The mass transfer line and ion source were set at 250 and 200 °C, respectively. The FAMEs were detected with electron ionization (70 eV) in scan mode (50–600 m/z). The identification of unknown FAMEs from yeast cells was achieved by comparing their retention times and mass spectrum profiles with known standards of rape seed oil FAMEs.

Please cite this article in press as: Patel, A., et al. Boosting accumulation of neutral lipids in Rhodosporidium kratochvilovae HIMPA1 grown on hemp (Cannabis sativa Linn) seed aqueous extract as feedstock for biodiesel production. Bioresour. Technol. (2014), http://dx.doi.org/10.1016/ j.biortech.2014.03.142

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2.6.7. BODIPY (505/515) staining and fluorescence microscopy For fluorescence microscopy BODIPY stock solution was prepared by adding 0.1 mg/ml with DMSO (Govender et al., 2012). Initially, 100 ll of the culture broth of R. kratochvilovae grown at different time intervals was pelleted, washed and suspended in 1 ml of sterile PBS (pH 6.0) to obtain appropriate cell concentration (5  109 cells/ml). 2 ll of BODIPY stock solution was then added to it and mixed well. After 5 min of incubation in dark the cell suspension was washed with 0.9% saline water before viewing. Fluorescence imaging was performed on a digital inverted fluorescence microscope equipped with GFP light cube (EVOS-FL, Advance Microscopy Group, AMG, USA). 2.6.8. Using ImageJ to estimate the cell size, LDs size and LDs volume The size of cells and intracellular LDs were analyzed by ImageJ 1.48a software (Schneider et al., 2012). Images obtained after fluorescence microscopy containing the fluorescence channel were converted to a Z-projection in ImageJ. Using straight tool in ImageJ two lines were manually drawn which bisect the cell and each fluorescent LD followed by adding the ROI manager. Each line represents the diameter of the cell and LD. The radius of each LD (calculated by taking half of the length of the line) was used to calculate the volume of a sphere. The length and width of LDs were measured for droplets that show a non-spherical geometry and the volume was estimated using the ellipsoid volume. 100 cells were counted to calculate the average size of cells and intracellular LDs, which was presented as mean ± S.D. 3. Results and discussion 3.1. Screening and identification of oleaginous yeast Permafrost soil samples collected from Tungnath Hill areas of the Himalayan Garhwal Ranges, Uttarakhand, India at a sea level of 4000 m height and temperatures ranging from 0 to 25 °C were screened for oleaginous yeast. Initially 479 microbial colonies were obtained on YEPD supplemented with ampicillin. These colonies were then picked and patched on nitrogen limiting media (NLM) agar plates which resulted in selection of 79 yeast colonies. The selection criteria was based on the fact that maximum TAG accumulation takes place in NLM by oleaginous yeasts having maximum lipid droplet size. Among these 79 screened yeast colonies, 08 oleaginous yeast isolates (Rhodosporidium spp., Trichosporan spp., Lipomyces spp., Candida spp., R. glutinis, Cryptococcus spp., Rhodotorula arauceriae, Rhodotorula minuta) having lipid droplet sizes (4.25 ± 0.53, 3.89 ± 0.46, 3.46 ± 0.65, 1.79 ± 0.32, 3.48 ± 0.34, 3.43 ± 0.68, 4.17 ± 0.97, 4.22 ± 0.35 lm) respectively were identified using BIOLOG microbial identification system. The identified colonies were characterized using live in vivo fluorescence imaging technique based on the size of LDs stained with BODIPY fluorescent dye (505/515 nm) which specifically stain TAG stored in the LDs. These LDs were then measured by ImageJ software via digital inverted epifluorescent microscope (EVOS FL). The identity of the selected strain Rhodosporidium spps having maximum LD size (4.25 ± 0.53 lm) was further confirmed by genotypic characterization performed by sequencing the D1/D2 domains of the gene encoding subunit of 26S of ribosomal RNA gene. The partial sequence of 609 bp using universal primers for D1/D2, and 606 bp with gene specific primers were obtained. These nucleotide sequences were searched for homology with other sequences by doing BLAST search tool (http://.ncbi.nlm.nih.gov/Blast.cgi) from NCBI Database which revealed its 100% identity, e-value score of zero with complete homology to following strains: R. kratochvilovae strain CECT 11973 isolate Y2 (Acc. No. AY296049 of 602 bp); R. kratochvilovae (Acc. No. DQ778989 of 602 bp); R. kratochvilovae

strain CECT 11956 (Acc. No. AY167603 of 593 bp); R. kratochvilovae strain PYCC 5580 (Acc. No. AF444778 of 593 bp). Data indicated that the partial sequence of 26S ribosomal RNA gene (609 bp) of our yeast strain HIMPA1 belongs to R. kratochvilovae and was assigned GenBank Accession No. KF772881. To infer the genetic diversities within and among groups, and to identify the taxonomic position and relationships, the phylogenetic trees based on the D1/ D2 domain of 26S rRNA gene sequence of R. kratochvilovae HIMPA1 (GenBank Accession No. KF772881) and the other related strains were constructed by MEGA 6 software (Tamura et al., 2013). The evolutionary history of the taxa was inferred using the NeighborJoining method. The optimal tree with the sum of branch length = 0.37297148 was shown in Fig. 1. The evolutionary distances were computed under minimum evolution and maximum parsimony criteria. The tree was drawn to scale, with branch lengths in the same units as those of the number of base substitutions per site. The percentage of replicate trees in which the associated taxa clustered together in the bootstrap test (1000 replicates) was shown next to the branches. The analysis involved 37 nucleotide sequences. There were a total of 508 positions in the final dataset. Dendrograms drawn further revealed that the isolate HIMPA1 was affiliated to the pedigree of Rhodosporidium spps. The evolutionary hierarchy of this strain was Basidiomycota > Pucciniomycotina > Microbotryomycetes > Sporidiobolales > R. kratochvilovae. 3.2. Preparation of the hemp seeds aqueous extract (HSAE) Hemp seeds have been used from prehistoric times as a source of fiber, oil, lubricants and fuel. They represent perfect and natural blend of the major essential amino acids, essential fatty acids, vitamins and mineral content (Callaway, 2004; Latif and Anwar, 2009). Hemp seed oil contains 75%, essential fatty acids (EFAs) content as triglycerides especially linoleic (LA) and linolenic (LNA) acids in a 3:1 ratio as its major omega-6 and omega-3 fatty acids, respectively. This leads to selection of hemp seed rich in TAGs as a good source for commercial biodiesel production. Hence, in this investigation hemp seeds aqueous extract were used as an alternate to commercially available expensive fermentation ingredients such as yeast extract, peptone, vitamins, minerals. The feedstock preparation from lignocellulosic material usually had to pass the technical barrier of a harsh physical or chemical process in the form of steam explosion or dilute acid pretreatment, which is applied to break the lignin or hemicellulose shell into simpler sugars. These cumbersome steps of hydrolysis by dilute acid and neutralization leads to production of toxic components (formic acid, acetic acid and furfural) which hinders growth of microorganisms during the down streaming process. Formation of HMF occurs in acidic solutions due to substantial reversible dimerization and dehydration of fructose at elevated temperatures in high-boiling molecular solvents or by inorganic salt in alcohol (Dam et al., 1986). Interestingly, Liu et al. (2012) reported that aqueous extraction at 100– 130 °C do not produce HMF since water is an inefficient solvent for dehydration of fructose but is required to solubilize proteins, vitamins and minerals. As the aqueous extraction method is not suitable for extracting the highly hydrophobic (30–35%) lipid content present in hemp seeds which agglomerate together as large water insoluble lipid complexes. We have used sonication treatment for efficient solubilization and homogenization of oil content into very fine emulsions. Recently, Jianguo et al. (2013) revealed that ultrasound pretreatment boosted the production of volatile fatty acids and significantly increase soluble proteins and reducing sugars in food waste fermentations. Thus, HSAE prepared through sonication, in this study uniquely paved out the way as simple option for pretreatment process. We observed both the sonicated and unsonicated HSAE lacks toxic components of 5-HMF and furfural

Please cite this article in press as: Patel, A., et al. Boosting accumulation of neutral lipids in Rhodosporidium kratochvilovae HIMPA1 grown on hemp (Cannabis sativa Linn) seed aqueous extract as feedstock for biodiesel production. Bioresour. Technol. (2014), http://dx.doi.org/10.1016/ j.biortech.2014.03.142

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Fig. 1. The phylogenetic tree of Rhodosporidium kratochvilovae HIMPA1 Acc. No. KF772881 showing evolutionary history was inferred using the Neighbor-Joining method. The optimal tree with the sum of branch length = 0.37297148 is shown. Evolutionary analyses were conducted using MEGA6 software. GenBank sequence and strain names are indicated after species designation.

when checked using HPLC. To reveal the effect of sonication on HSAE, its composition analysis of various components present in these growth media was done (Table 1). We observed that the sonication enhances the availability of soluble and monomeric contents like; soluble proteins (15.78%), DAG (25.18%), MAG (187.35%), free fatty acids (228.08%), soluble fiber (282.35%), total sugars (350.69%) and reducing sugars (531.42%) in sonicated HSAE as compare to unsonicated HSAE. This has also increased the amount of total proteins (14.63%), and total carbohydrates (60.58%), tremendously. It was further supplemented by the fact that the decrease in polymeric and macromolecular structure leads to reduction of total fats (10.69%), insoluble fiber (15.6%),

triglycerides (33.4%) and lignin content (17.3%) in sonicated HSAE in comparison to unsonicated HSAE. To evaluate the effect of sonication on triglycerides compositions of unsonicated HSAE and sonicated HSAE fatty acid percentage were compared (Table 2). Data showed relative decrease in the percentage of unsaturated fatty acids and an increase in the percentage of saturated fatty acids when ultrasound-treatment was done. This could be due to the fact that unsaturated fatty acids are more susceptible to oxidation. The ratio of unsaturated to saturated fatty acids was decreased from 7.413 to 5.035 in unsonicated to sonicated HSAE. This ratio is an indicator of the extent of fat deterioration. The enhanced solubility and easier uptake of soluble particles along

Table 1 Percentage composition of dry cell weight of industrial hemp seeds, unsonicated and sonicated aqueous seed extracts. Composition

Hemp seeds (%)

Unsonicated HSAE (%)

Sonicated HSAE (%)

Total proteins Total nitrogen Soluble protein Total fats Saturated fats Polyunsaturated fats Monounsaturated fats Triacylglycerides (TAG) Diacylglycerides Monoacylglycerides Free fatty acid (FAs) Number of TAGs/FAs Carbohydrates Total dietry fibers Soluble fibers Insoluble fibers Lignin Total sugars Reducing sugars

22.5–24.8 32.4–34.5 22.4–28.6 30.0–35.5 3.3–5.2 23.0–27.2 2.7–3.85 25.2 18.4 6.85 6.39 25.2/6.39 34–35.8 28–32 3.0–5.4 22.8–29.0 4.04–5.76 2.28–2.44 2.22–2.54

20.5–21.8 32.4–33.5 22.8–26.6 23.0–25.5 2.3–2.5 16.4–17.2 1.7–1.85 19.32 13.54 4.35 4.45 19.32/4.45 34–35.8 14.05–16.45 3.4–5.2 14.8–15.2 1.04–1.76 2.86–2.94 2.10

23.5–28.8 36.4–37.5 26.4–28.6 20.54–21.85 2.63–2.68 14.57–15.85 2.27–3.35 12.85–12.25 16.95–18.65 12.5–14.7 14.6–15.5 10.85/14.5 54.6–58.88 12.65–13.45 13.0–15.4 12.48–13.93 0.86–0.94 12.89–13.26 13.26

Please cite this article in press as: Patel, A., et al. Boosting accumulation of neutral lipids in Rhodosporidium kratochvilovae HIMPA1 grown on hemp (Cannabis sativa Linn) seed aqueous extract as feedstock for biodiesel production. Bioresour. Technol. (2014), http://dx.doi.org/10.1016/ j.biortech.2014.03.142

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Table 2 Triglyceride composition (%) of unsonicated hsae and sonicated HSAE. Fatty acid content

Unsonicated HSAE (%)

Sonicated HSAE (%)

Saturated fatty acids Octanoic acid (C8:0) Decanoic acid (C10:0) Dodecanoic acid (C12:0) Tetradecanoic acid (C14:0) Palmitic acid (16:0) Stearic acid (18:0) Arachidic acid (20:0) Behenic acid (22:0) Tetracosanoic acid (C24:0) (A) Total saturated fatty acids (%)

– – – – 8.5 2.6 0.5–0.8 0.5 0.1 12.1

0.02 0.03 0.01 0.01 11.05 3.05 1.02 0.8 0.8 16.7

Unsaturated fatty acids Oleic acid (18:1 x-9) Linoleic acid (18:2 x-6) G-Linolenic acid (18:3 x-6) a-Linolenic acid (18:3 x-3) Eicosaenoic acid (20:1w-9) Eruic acid (C22:1) Tetracosenoic acid (C24:1) (B) Total unsaturated fatty acids (%) Ratio of (A)/(B)

11.5 54.2 20.2 3.3 0.3 0.02 0.02 89.7 7.413

10.2 52.2 18.5 3.02 0.02 0.01 0.04 84.1 5.035

with emulsified fat content makes sonicated HSAE a better media for cultivation of microbes. Another important aspect for boosting TAG accumulation is the C:N:P ratio of the media. Our data showed total Carbon:Nitrogen:Phosphate ratio (C:N:P) of sonicated HSAE to be 230:10:1 which indicated that the sonicated HSAE possess natural nutritional composition with high C:N ratio and can be exploited for enhanced TAG lipid production. Earlier, Zhu et al. (2008), showed high C/N ratio of 238 for maximum lipid accumulation in Trichosporan fermentans. 3.3. Batch cultivation of R. kratochvilovae HIMPA1 The cultivation conditions of R. kratochvilovae HIMPA1 when grown on sonicated HSAE, unsonicated HSAE and GSM were optimized for maximum lipid production. The maximum cell dry weight (CDW), lipid content and lipid yield produced by R. kratochvilovae HIMPA1 on different medium (100 ml) at 30 °C were monitored till 216 h and their time courses were recorded (Fig. 2A–C). When grown on sonicated HSAE maximum cell dry weight (15.10 ± 0.98 g/l) were attained after 168 h while maximum total lipid yield (8.39 ± 0.57 g/l) and lipid content (55.56% of CDW) was observed during early stationary phase (Fig. 2B). The phase between (122–168 h) showed maximum lipid accumulation and was defined as Lipid Accumulating Phase (LAP) However, in comparison to sonicated HSAE, unsonicated HSAE showed drastic reduction in maximum total lipid yield to 5.10 ± 0.37 g/l and lipid content to 37.92% of CDW, while cell dry weight was reduced marginally to 13.45 ± 0.56 g (Fig. 2A). This reduction of lipid content (17.64%) and lipid yield (3.29 g/l) in unsonicated hemp seeds extract could be due to emulsification of lipids attained by sonication. In GSM, the maximum cell dry weight achieved was 14.46 ± 0.12 g/ l and total lipid yield produced was 6.2 ± 0.8 g/l, while maximum total lipid content (41.92% of CDW) was observed at 168 h (Fig. 2C). The LAP in GSM was recorded to be from 96 to 168 h. Data indicates that sonicated HSAE produces 14–15% higher lipid content than GSM, making it a better alternative TAG feedstock for lipid production. The uptake of reducing sugars and total nitrogen during cultivation of R. kratochvilovae HIMPA1 on these media were plotted at different time intervals during cultivation of HIMPA1 (Fig. 2D–F). The results obtained showed the substrate utilization capability of oleaginous yeasts. The new oleaginous yeast species belonging to Rhodosporidium, Rhodotorula representing basidiomyceta reported to have highest extractable lipid levels

ranging from 8.81–65.32% of CDW obtained in cells grown under nitrogen starvation (Sitepu et al., 2013). These genera can convert both C6 and C5 sugars present in lignocellulosic hydrolysate to lipids. Various yeast genera Rhodosporidium, Rhodotorula, Lipomyces, Trichosporon and fungal Mortierella, Cunninghamella cultivated under nutrient limiting conditions (Papanikolaou and Aggelis, 2011) demonstrated to grow on both hexoses and pentoses making utilization of waste hydrolysates from cheaper lignocellulosic material as a feasible possibility for biodiesel production. 3.4. Quantifying cell size and lipid droplet size in R. kratochvilovae HIMPA1 Lipophilic bright green fluorescent dye, BODIPY 505/515 which has resistance to photobleaching, and can maintain its fluorescence for longer than 30 min was explored for monitoring intracellular neutral lipids stored in LDs of R. kratochvilovae HIMPA1 via live cell imaging. We demonstrated that cells grown on sonicated HSAE have LD sizes (0.56 ± 0.076, 2.3 ± 0.97, 4.65 ± 0.73, 5.95 ± 1.02 lm) with cell size (2.43 ± 0.37, 4.67 ± 0.43, 6.28 ± 0.34, 7.02 ± 0.91 lm) after 48, 96, 144, 192 h respectively while data obtained from the TAG accumulating ability in GSM having (0.22 ± 0.014, 0.89 ± 0.023, 2.06 ± 0.43, 2.38 ± 0.52 lm) with cell size (2.23 ± 0.21, 3.01 ± 0.14, 4.12 ± 0.24, 5.53 ± 0.31 lm) after similar time intervals. Results showed that the intensity of fluorescence dye when visualised by fluorescence microscopy directly correlate with accumulated TAG in LDs. Maximum LD size of the cells grown on sonicated HSAE (192 h) have 2.5 times more TAG accumulation ability than in GSM. The exponential increase in accumulation of TAGs was observed between 144 and 192 h and was defined as TAG accumulating phase. The size of the LDs was measured by ImageJ software which is used for projecting volume of the accumulated TAG in them. Data showed increase in average LDs volume 110.29 lm3 and 7.06 lm3 in sonicated HSAE and GSM respectively, accounting for 15.7 folds increase in sonicated HSAE. The volume of LDs calculated gave direct correlation to biodiesel produced per unit cell. We observed that the storage lipids as TAG was maximum in the stationary phase (192 h) and harvesting of cellular lipids should be done up to 216 h, to obtain maximum lipid yield. Supersized LDs (SLDs) that are up to 50 times the normal volume in cells were formed to be reported in yeast mutants lacking FLD1 and seipins (Feil et al., 2011; Fei et al., 2008). Recently researchers have highlighted the role of lipid droplet structure and its function in mammalian physiology and pathophysiology. They have also shown that the mechanism of storage and metabolism of lipids in the LDs are conserved from yeast to man (Reue, 2011). 3.5. FTIR spectroscopy analysis The FTIR spectra of lipid extracted from sonicated HSAE grown R. kratochvilovae HIMPA1 showed transmittance spectral similar to triolein used as standard. No bands were observed in the spectral region 4000–3450 cm1 indicating thereby the absence of hydroxyl and amine groups. Both triolein and extracted lipid showed infrared absorption peaks in the second spectral region, 3100– 2850 cm1. The bands obtained in this region were close to the wave numbers 2923 and 2856 cm1 respectively and assigned to the symmetrical and asymmetrical C–H stretching vibration of the CH2 and CH3 aliphatic groups, which are found in large quantities in vegetable oils. The peak obtained at 1743 cm1, confirms the presence of carbonyl group. 3.6. Transesterification of total lipids into biodiesel Transesterification of total lipids obtained from batch cultivation of R. kratochvilovae HIMPA1 grown on sonicated HSAE and

Please cite this article in press as: Patel, A., et al. Boosting accumulation of neutral lipids in Rhodosporidium kratochvilovae HIMPA1 grown on hemp (Cannabis sativa Linn) seed aqueous extract as feedstock for biodiesel production. Bioresour. Technol. (2014), http://dx.doi.org/10.1016/ j.biortech.2014.03.142

A. Patel et al. / Bioresource Technology xxx (2014) xxx–xxx

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Fig. 2. Panel I: Showing dry cell weight (g/l), Total lipid yield (g/l) and lipid content (%) of Rhodosporidium kratochvilovae HIMPA1 grown at different time intervals on (A) unsonicated HSAE (l = 0.117 h1); (B) sonicated HSAE (l = 0.249 h1); (C) glucose synthetic medium (l = 0.158 h1). Panel II: Showing substrate utilization: Reducing sugar (g/l) and total nitrogen (g/l) by Rhodosporidium kratochvilovae HIMPA1 grown at different time intervals on (D) unsonicated HSAE; (E) sonicated HSAE; (F) glucose synthetic medium.

GSM was done using BF3-Methanol method yielded conversion efficiency of 98.5%, 95.5% and biodiesel yield 95%, 93% respectively. The total lipids extracted from both the mediums used for growing R. kratochvilovae HIMPA1 were compared for TAG accumulated as detected by MnCl2 charring method. It was observed that TAG band present in the HSAE lane was almost twice of GSM while, MAG content was higher in GSM. FAMEs obtained after transesterification of total lipids were detected by TLC using dual solvent system, first on hexane:tert-butyl methyl ether:acetic acid (50:50:0.5, v/v/v) and then by second development with hexane:tert-butyl methyl ether:acetic acid (97:3:0.5, v/v/v) verified conversion of TAG with 98.5% (HSAE) and 95.5% (GSM) conversion efficiency, respectively. Earlier, investigation on the viability of hemp seed oil to produce hemp oil biodiesel (HOB) via basecatalyzed transesterification process gives conversion efficiency of oil more than 99.5% and biodiesel yield about 97%, which showed HOB to have comparable traits to meet ASTM 6751-09 standards (Li et al., 2010; Rehman et al., 2013; Su et al., 2013). 3.7. Fatty acid methyl ester composition The FAMEs profile of lipids obtained from GSM and sonicated HSAE medium were analyzed by GC–MS (Fig. 3). When GSM was

Fig. 3. FAMEs profiles – lane 1: Rape seed oil FAMEs standard (AOCS-07756-1AMP); lane 2: R. kratochvilovae HIMPA1 grown on glucose synthetic media; lane 3: Hemp seed oil; lane 4: R. kratochvilovae HIMPA1 grown on sonicated HSAE.

Please cite this article in press as: Patel, A., et al. Boosting accumulation of neutral lipids in Rhodosporidium kratochvilovae HIMPA1 grown on hemp (Cannabis sativa Linn) seed aqueous extract as feedstock for biodiesel production. Bioresour. Technol. (2014), http://dx.doi.org/10.1016/ j.biortech.2014.03.142

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A. Patel et al. / Bioresource Technology xxx (2014) xxx–xxx

Table 3 FAME composition (%) of Rape seed oil Standard AOCS-07756-1AMP; hemp seed oil, and biodiesel produced by R. kratochvilovae HIMPA1 grown on GSM and sonicated HSAE. FAME composition (%)

Myristic acid (C14:0)

Palmitic acid (C16:0)

Stearic acid (C18:0)

Oleic acid (C18:1)

Linoleic acid (C18:2)

Linolenic acid (LA) (C18:3)

Arachidic acid (C20:0)

Behenic acid (C22:0)

Erucic acid (C22:1)

Tetracosanoic acid (C24:0)

Heptacosanoic acid (C27:0)

Rape seed oil Hemp seed oil

1 ND

4 8.5

3 3

60 15

12 52.2

5

3 ND

3 ND

5 ND

3 ND

ND ND

GSM Sonicated HSAE

ND 3

5.9 11.15

25.1 2

37.3 45.55

ND 8.47

22 5

6.5 9.7

ND ND

ND ND

3.00 14.98

used as medium for lipid production by R. kratochvilovae HIMPA1, the FAMEs profile contains mainly methyl esters of palmitic acid (C16:0) 5.90%, stearic acid (C18:0) 25.10%, oleic acid (C18:1) 37.5%, along with arachidic acid (C20:0) 22%, behenic acid (C22:0) 6.5%, and an unusual fatty acid heptacosylic acid (C27:0) 3% which were reported for the first time by this oleaginous yeast (Table 3). Since, GSM medium is prepared without additions of any other lipid source clearly indicates that FAME profile is specific for the yeast strain grown. The profile revealed similarity to other genera specific oleaginous yeast of Rhodosporidium spp. having characteristics shown by presence of palmitic acid (C16:0) 5%, stearic acid (C18:0) 25.10%, oleic acid (C18:1) 37.30%. The three additional fatty acids were strain specific, arachidic acid (C20:0) 22%, behenic acid (C22:0) 6.5%, and heptacosylic acid (C27:0) 3%, as these were also present in the FAME profile of the lipids extracted from medium having glycerol as sole carbon source using this strain (unpublished data). FAME profile attained when sonicated HSAE was used as growth medium for R. kratochvilovae HIMPA1 showed methyl esters of myristic acid (C14:0) 3%, palmitic acid (C16:0) 11.15%, stearic acid (C18:0) 2%, oleic acid (C18:1) 45.55%, linoleic acid (C18:2) 8.47%, arachidic acid (C20:0) 5%, behenic acid (C22:0) 9.7% and heptacosanoic acid (C27:0) 14.98% (Fig. 3). The longer carbon chain esters of fatty acid showed increase in percentage content of behenic acid (C22:0) 9.7% and heptacosanoic acid (C27:0) 14.98% in sonicated HSAE as compared to 6.5% and 3% present in GSM respectively (Table 3). Thus, combination of sonicated HSAE and oleaginous yeast R. kratochvilovae HIMPA1 together gave the unusual FAME profile containing longer carbon chains C20:0, C22:0, C27:0 reported for the first time in this study along with usual C16:0, C18:0, C18:1, C18:1, C18:2 fatty acids. Interestingly, FAME profile of hemp seed oil lacked C27:0 molecules and diminuted amount of C20:0, C22:0, fatty acids but contains C16:0 (6–5%), C18:0 (2–5%), C18:1 (12–15%), C18:2 (52–56%), a-C18:3 (15–18%), c-C18:3 (3–4%), respectively (Table 3). The data showed that the combo (sonicated HSAE and oleaginous yeast R. kratochvilovae HIMPA1) can enhance biodiesel fuel characteristics by enhancing production of higher hydrocarbon chains C20:0 C22:0 C27:0 (Da Porto et al., 2012; Li et al., 2010) similar to algal oils (Amini et al., 2009). Thus, this opens up new avenues for the preparation of combination of desired FAME profile for biodiesel production using microbial interventions.

4. Conclusions Hemp seeds extracts are the economical triacylglyceride feedstock containing all essential nutrients components used for growing novel oleaginous yeast R. kratochvilovae HIMPA1 to boost maximum TAG accumulation in LDs of yeast cells. Pretreatment by sonicating HSAE makes it devoid of toxic inhibitors and produces lipid yield 15% more than GSM. Supersized lipid droplets of 5.95 ± 1.05 lm were visualized using BODIPY stain in HSAE grown yeast. Unique FAME profile having Arachidic acid (C20:0) 5%, Behenic acid (C22:0) 9.7%, Heptacosanoic acid (C27:0) 14.98%

aLA

cLA

18 0.20 0.15

3.3

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Please cite this article in press as: Patel, A., et al. Boosting accumulation of neutral lipids in Rhodosporidium kratochvilovae HIMPA1 grown on hemp (Cannabis sativa Linn) seed aqueous extract as feedstock for biodiesel production. Bioresour. Technol. (2014), http://dx.doi.org/10.1016/ j.biortech.2014.03.142