Materials and Methods

Conventional biodiesel production protocol is given in Fig. 1 and Fig. 2 outlines the method followed in the present endeavor to optimize biodiesel production through novel biocatalyst as well as a conventional chemical catalyst.

• 1. Isolation, screening and identification of lipase producing fungi

Indigenous fungi that are efficient in extracellular lipase pro- duction, were explored in natural environments. Endophytic fungi (from root, shoot, bark and leaf of a mangrove plant) and free spores (from sediment and water column) were isolated from samples sourced from a backwater region (14 24058.14"N, 74 24030.30"E, images

Fig. 1. Process outline of conventional algal biodiesel production.

* - protocol explained in detail in Fig. 2.

Fig. 2. Flowchart describing the workflow to optimize biodiesel production through novel biocatalyst.

with a salinity of 35 ppt) in Uttara Kannada district along the west coast of India. The collected samples were plated in the Potato Dextrose (PD) agar under aseptic conditions. Twenty-four fungal strains (CS1 e CS24) belonging to different genera were isolated during initial plating which was further sub cultured to obtain pure fungal isolates. These fungal isolates were subjected to multiple screening tests on agar plates for lipase using tween 80 and olive oil as sole carbon sources substituted with CaCl₂.2H₂O and phenol red as extracellular lipase indicators respectively. Phenol red/olive oil agar plate was prepared using chemicals with composition (g/L) of: 0.01% (w/v) phenol red, 2% (w/v) agar, 1% (v/v) olive oil, 1% (w/v) CaCl₂ (anhydrous) with pH adjusted to 7.2 using 10 M NaOH [38]. Tween 80 plates were prepared as follows: 1% peptone, 0.5% NaCl, 0.01% CaCl₂.2H₂O, 2% agar and 1 ml (v/v) tween 80 [39]. The screening tests resulted in eight lipase positive strains (CS1 e CS8), and further one single strain (CS4) was screened based on the observation of the highest zone of clearance.

Morphological and molecular characterization of fungi

The morphological structure of the selected strain was studied after staining the fungus CS4 with lactophenol cotton blue and observing in the high-resolution phase-contrast optical microscope (Olympus BX51) under 40 magnification. The fungal strain was also viewed under a scanning electron microscope (JEOL JSM IT- 300) for better visualization of structural morphology. Further, the selected strain was subjected to 18S rRNA based molecular identification after extracting genomic DNA using DNeasy plant maxi extraction kit (Qiagen Inc., USA) and the extracted DNA were amplified using PCR with ITS1F (50-TCCGTAGGTGAACCTGCGG-30) and ITS 4R (50-TCCTCCGCTTATTGATATGC-30) desalted 20 nm oligonucleotide universal primers. A single discrete PCR amplicon band of 500 bp was observed when resolved on an agarose gel. The PCR amplicon was then purified to remove contaminants and subjected to forward and reverse DNA sequencing reactions along with forward and reverse primers using BDT v3.1 cycle sequencing kit on ABI 3730xl Genetic Analyzer. The consensus sequence of 18S rRNA genes generated using forward and reverse sequences were subjected to nucleotide BLAST in National Centre for Biotechnology Information (NCBI) GenBank database to identify the strain based on maximum identity score. The first ten sequences that showed higher statistical significance with the strain of interest were selected and aligned using CLUSTAL-W multiple alignment soft- ware. The phylogenetic tree was constructed by multiple sequence alignment with the neighbour-joining method using MEGA soft- ware version 7. The 18S rRNA based molecular identification revealed a maximum homology of the strain CS4 to Cladosporium tenuissimum identified hereafter as Cladosporium sp. CS4.

Inoculum and lipase enzyme production medium

The Cladosporium sp. CS4 was inoculated in PD broth liquid medium (potato (infused form) 20% (w/v) and dextrose 2% (w/v)) at 28 C for 7 days. After 7 days, the cell pellets formed a mat over the liquid layer were collected by centrifugation at 11,392xg for 20 min at 4 C. The cell pellets were collected, which was used as inoculum for lipase production medium consisting of ingredients (g/L): KH₂PO₄ - 2.0, bacto-peptone - 5.0, yeast extract - 1.0, NaNO₃ -

0.5, KCl - 0.5, MgSO₄$7H₂O - 0.5, olive oil - 10.0, pH adjusted to 5.5 [40]. Cultivated with a culture volume of 1 L for 5 days at 28 C. Then, the crude extract was separated from the mycelial mat by centrifugation at similar operating conditions for inoculum

preparation. The cell-free supernatant obtained after centrifugation with subsequent concentration and purification was used as a lipase enzyme source.

Enzyme concentration and purification

The cell-free supernatant was saturated with ammonium sul- phate at a final concentration of 80% saturation with constant and slow stirring at 4 C, followed by centrifugation at 18,900xg for 20 min at 4 C [41]. The pellet obtained after centrifugation was dissolved in 50 mM Tris HCl pH 7.0 and dialyzed against 50 mM Tris-Chloride buffer (pH 7.2) with 0.1 M NaCl [42] at4 C overnight

in a dialysis tubing having Molecular Weight Cut-Off (MCWO) of

3.5 KDa (Sigma-Aldrich). The concentrated enzyme after dialysis was subjected to a subsequent purification step that involved size exclusion-based gel filtration chromatography in a pre-equilibrated Superdex 200, 10/300 GL (GE Healthcare) with an elution rate of

0.15 mL/min. The eluted enzyme fractions (1 mL each) were collected in an AKTA pure 25 (GE Healthcare) FPLC collector. The collected enzyme fractions were subjected to protein estimation spectrophotometrically using Braford protein assay. The fractions that exhibited the highest protein content was subjected to lipase assay test and lipase active fractions were pooled and dialyzed extensively in 50% glycerol-based dialysis buffer containing 50 mM Tris-Chloride and 0.1 M NaCl with pH 7.2 and stored at 4 C until use. The dialyzed fractions were run on SDS- PAGE to evaluate lipase purity.

Lipase activity determination

The rate of release of p-nitrophenol (p-NP) was used as a proxy to measure lipase activity through a spectrophotometer, using p- nitrophenol palmitate (p-NPP) as a synthetic lipid substrate [43]. The substrate solution was prepared according to Ref. [44] with the required modifications. Briefly, the substrate solution was prepared using 30 mg of p-NPP (sigma, AR grade) dissolved in 10 mL 2- propanol and mixed with 90 mL emulsifying solution containing 0.1% acacia (Arabic gum) and 0.4% Triton X-100, which resulted in a

markers (10e200 KDa) (Thermo Scientific) was used as a reference to determine the molecular weight of purified protein [49]. Silver staining was used to visualize protein bands on SDS PAGE gel [50]. Protein concentration was measured using Bradford protein assay with Bovine Serum Albumin (BSA) of increasing concentrations as standard at a wavelength of 595 nm in VERSA max tunable microplate reader (CARE Biosystems, Mumbai, India). Bradford re- agent (5 ) was prepared as follows in (g L^-1) - 0.05% Coomassie

Brilliant Blue (CBB) G-250, 23.5% (v/v) of methanol and 50% (v/v) of

85% phosphoric acid which was diluted to 1 concentration before protein assay analysis.

Characterization of purified lipase

Effect of pH and temperature on lipase activity and stability the lipase enzyme activities at varying pH ranges were measured at standard assay conditions with varying pH buffers of 50 mM concentration ranging from pH 3.0 e pH 10.0 using p-NPP as synthetic lipid substrate. The buffers used for pH optimization were as follows: Citrate phosphate (pH 3.0e6.0), Tris-HCl (pH 7), Potassium-phosphate (pH 8.0), Glycine-NaOH (pH 9.0e10.0). pH stability was determined by preincubating the enzyme aliquots with respective buffers for a period of 24 h at 4 C [40]. Residual enzyme activity of the preincubated enzyme aliquots was measured by adjusting the pH to 7.0 and post-incubation of reac- tion mixture at 60 C for 30 min after introducing p-NPP substrate. Temperature optimization of the enzyme was carried out by eval- uating the enzyme activity at various incubation temperatures ranging from 30 C to 80 C (with increments of 10 C) at pH 6.0. For determining thermal stability, the enzyme aliquots were pre- incubated with pNPP substrate at pH 6.0 under different temper- atures for a period of 30 min and the residual enzyme activity was determined after post-incubation of respective enzymes at 60 C for 30 min and assayed following standard protocols. Relative and residual activity expressed as a percentage (%) is determined by the ratio of activity at each temperature and pH range to that of the maximum recorded lipase activity.

Relative activity (%) = Activity / Maximum Activity * 100 - (1)

final substrate concentration of 0.8 mM of p-NPP. A volume of 100 mL cell-free supernatant was added to 100 mL of 0.5 M citrate phosphate buffer, pH 6.0 and 900 mL of substrate solution. The re- action was carried out in Eppendorf tubes in an incubator shaker maintained at 37 C for 30 min under a rotation speed of 120 rpm.

Effect of surfactants on lipase activity

The effect of various surfactants on purified lipase was determined after incubating the enzyme aliquots with 0.125%, 0.25%, 0.5% and 0.75% (w/v) of SDS and similar concentrations in (v/v) of Triton-X, tween 20 and tween 80 at 37 C without substrate for 1 h. After 1 h, the residual activity was determined after adding pNPP substrate and incubating the reaction mixture at 60 C for 30 min with respect to control [40].

Isolation, cultivation & harvesting of microalgae

Indigenous diatom strains that are tolerant to higher salinities and capable of adapting to extreme environmental conditions were isolated and sediments samples were collected from a salt pan situated in vicinity to downstream of the Aghanashini estuary, Uttara Kannada, Karnataka, India (14 32048.84"N latitude and 74 20046.12"E longitude). The collected samples were subjected to serial dilution, agar plating and subsequent sub culturing of pure diatom isolate. The diatom isolates were later inoculated in F/2 medium [51] and sequentially scaled up in a 100 mL Erlenmeyer flask with 50 mL working volume under constant exposure of cells to a light intensity of 210 mmol m^-2 s^-1 with 12:12 h light-dark cycles. An aliquot of grown algal biomass was then subjected to acid digestion following standard protocols [52] and the acid-treated sample was subjected to morphological observation and imaging using a high-resolution phase contrast microscope Olympus BX51 as well as Scanning Electron Microscope (SEM) in JEOL JSM-IT-300 with dry silicon drift detector (EDAX) and accelerating voltage up to 30 KV under a scanning magnification of 1000 to facilitate observation of striae and accurate diatom valve measurements. The isolated diatom was identified as Nitzschia punctata (Nitzschia sp.) based on morphological characterization through high-resolution imaging and comparison with standard diatom taxonomical iden- tification keys [53,54] The stock cultures of Nitzschia sp. were maintained in F/2 medium with routine sub culturing in filtered and sterilized estuarine water of salinity 35 ppt. Growth experi- ments were conducted on the rooftop with natural light (sunlight), 12: 12 h light/dark period after adding the inoculum of pure Nitz- schia sp. having an approximate cell density of 1 10⁶ cells/ml in 10 L translucent plastic tub with 5 L working volume. The cultures were maintained at ambient temperature for nine days with peri- odic mixing. At the end of the growth period, the diatom cells were harvested by siphoning the spent culture medium as the diatom cells sink and settle out in stationary cultures. The harvested biomass was washed twice with double distilled water, centrifuged at 3500 g [55] for 10 min and oven-dried at 85 C overnight.
The biomass productivity (mg L^-1d^-1) = (C - C₀)/T
where C is the final biomass concentration of algae after harvest, C₀ is the initial biomass concentration at the time of inoculation. T is the culture period in days.

Microalgal oil extraction and biodiesel production
Oil extraction and quantification

The cultivated and harvested diatom (microalgal) biomass was oven-dried at 85 C overnight and pulverized to a fine powder using mortar and pestle. Oil (lipid) extraction was performed on dried and pulverized algal biomass by following the modified Folch method [56]. The algal biomass was ultrasonicated at 35 kHz fre- quency at 45 C for 15 min after adding chloroform and methanol in the ratio of 2:1 (v/v). The ultrasonicated sample was treated with 0.8% NaCl solution to enable a clear phase separation of aqueous and organic phases with cell debris at the interface between the two phases resulting in a solvent mixture ratio of (2:1:0.8) Chlo- roform/methanol/water. The lower organic phase consisting of lipids that are dissolved in chloroform was separated using sepa- rating funnel and the procedure was repeated twice to ensure maximum lipid recovery. The lipids thus extracted were evaporated under vacuum using a rotavap rotary vacuum evaporator (Model: PBV 7D) under reduced pressure and the oil obtained were weighed gravimetrically by dividing the lipid content obtained in mg with the biomass used for lipid extraction in terms of dry cell weight (dcw) to obtain lipid content (% dcw). Oil yield is calculated using equation (3).

Oil yield (%) = mass of oil extracted (mg) / mass of algal biomass used (mg) * 100 - (3)
Fatty acid characterization

Fatty acids of lipids extracted in a 250 mL Soxhlet extraction unit, fitted with a reflux condenser was characterized using GC-MS. A known quantity of algal biomass (~500 mg) was taken in a cel- lulose extraction thimble and refluxed using hexane as a solvent in Soxhlet extractor for 5 h at 50 C. The extracted fatty acids in hexane was concentrated using rotary vacuum evaporator under reduced pressure and characterized using GCMS: gas chromatog- raphy (Agilent Technologies 7890A GC, single quadruple analyser) e mass spectroscopy (Agilent 5975C inert MSD with triple-axis detector) model with helium as inert carrier gas and temperature at 35 C for 2 min, with a ramp input rate of 35 Ce300 C at 20 C per min with final hold time of 5 min at 300 C. The operating conditions were set with an initial solvent delay of 4 min [57]. The GC-MS generated peaks were interpreted through AMDIS data analysis software and the organic compounds eluted were matched with spectral mass spectroscopy database NIST V11.

Acid-catalyzed transesterification

Aliquots of a known quantity of diatom extracted lipid (~20 mg) were subjected to acid-catalyzed transesterification. The reaction was carried out in a water bath connected to a reflux condenser and the reaction temperature was set at 80 C until a reaction time of 2.5 h. A 2 wt% H₂SO₄ was used as a catalyst along with 2.5 ml of methanol as a co-reactant [58]. On reaction completion, the mixture was cooled to room temperature and phase separation was induced using hexane and distilled water in the ratio of 2:1. The upper phase containing Fatty Acid Methyl Ester (FAME) in hexane was collected in a sterile glass vial and the traces of water was removed by adding anhydrous sodium sulphate and stored in an Eppendorf tube for FAME composition analysis.

Lipase catalyzed transesterification

A known quantity of lipid (~5 mg) extracted from 10 mg of pulverized algal biomass was taken in a sterile screw-capped vial and was added with 50 mM Tris-HCl buffer pH 6.0 in the ratio of 0.5:1 and purified lipase was added to about 10% by volume of the lipid substrate taken for analysis [59]. The methanol and substrate concentrations were maintained in a ratio of 6:1 [20]. The reaction was carried out at 40 C for 48 h with constant shaking at 150 rpm. Methanol was added in parts at 0 h, 12 h and 24 h to avoid lipase inhibition to higher concentrations of methanol [60]. The FAME component analysis was carried out by collecting the upper organic phase containing biodiesel dissolved in hexane after visible phase separation on reaction completion.

Quantification of biodiesel and conversion efficiency estimation

Biodiesel (FAME) composition and the variation in FAME yield between acid and enzyme-catalyzed transesterification reaction was quantified using peaks observed in GC-MS. The mass spec- troscopy (ms) source and quadruple temperatures were 230 C and 150 C respectively. The program for oven operating conditions was set at 50 C initially for a hold time of 2 min, then increased at a ramp heating rate of 10 C/min until 280⁰C with a 4 min. Methyl laurate (C₁₃) was used as an internal standard. FAME composition was determined in terms of percentage of each FAMEs out of the total present in the sample. The peaks were analysed using AMDIS data processing tool and FAME compounds were identified and matched using NIST V11 Mass spectral search pro- gram. The percentage conversion of fatty acids into corresponding

Fatty acids of lipids extracted in a 250 mL Soxhlet extraction unit, fitted with a reflux condenser was characterized using GC-MS.

fatty acid methyl esters (FAME) was estimated using equation (3) [59,61].

where MW_i is the molecular weight of each of the fatty acid peaks obtained in GC-MS and %M_i corresponds to the percentage of fatty acids recorded in GC peaks.

Fourier transform infrared analysis (FTIR)

The IR spectrum of fatty acids and FAMEs of diatom oil was recorded in FTIR spectrometer (PerkinElmer GX FTIR) to identify the functional groups corresponding to lipids by scanning the liquid sample (containing fatty acids/FAMEs extracted in hexane) at mid infra-red region (4000e650 cm^-1) under transmission mode. The absorption spectra of fatty acids and FAMEs were plotted between % transmittance versus wave number.