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.
-
- 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 MWi is the molecular weight of each of the fatty acid peaks obtained in
GC-MS and
%Mi 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.