Introduction
Arundo donax L. a nonfood lignocellulose biomass does not compete with
other feed and food crops considered as potential alternate renewable crop
yielding high proportions of transportation fuel by employing cost effective
and operative conversion methods. This
species is considered as a
valuable source of biomass feedstock due to persistent yield, malleability to
marginal environments with low input requirements among other lignocellulosic
biomass (Amaducci and Perego 2015). The
survivability to drought resistance and salt tolerance make it suitable energy
crop which can be cultivated in low quality irrigation waters (Accardi et al.
2015).
A. donax a member of Poaceae family represents numerous remarkable
features as potential dedicated biomass energy crop (Corno et al. 2014; Lemões et al. 2018) and acquainted by
number of common names including, bamboo reed, bamboo, false bamboo, Arundo grass, reed grass, giant reed,
giant reed grass, giant Danube reed, bamboo cane, giant cane, Canne-de Provence,
wild cane, Spainsh cane and A. donax cane. In Pakistan it is commonly
known by “Nurr”, “Nurro” or “Nurru,” (Maria et al. 2013) and saroot. This
plant flourishes abundantly and spontaneously in southern Europe and several
subtropical temperate regions. A. donax was
habituated from Asia through Middle East to whole Mediterranean basin during
prehistory (Corno et al. 2014). Widely divergent archeological and historical
evidences witness the origin of A. donax from
Asia, North Africa, North and South America, South Europe, Middle East moreover
in Australia (Saikia et al. 2015). In its native range, A. donax is abundant
in Pakistan and India ascending to 2500 m elevations
in Himalayas and spreads throughout China and South-East Asia.
A. donax is an erect, tall, sterile rhizomatous (Scordia et al.
2011) perennial C3 grass grows well up to 9 m
(Saikia et
al. 2015) in dense stands (Krička et al. 2017). Due to fast growth
of A. donax in water, it usually
falls in category of emergent aquatic plant (Angelini et al. 2009). The potential
yield of A. donax dry biomass is
29–46 tons ha-1 year-1 that
entirely depends on geographical positions and climatic conditions (Pari et al.
2015) besides moisture content, density and time of cultivation (Krička et
al. 2017).
Various approaches are also evolving to utilize crop
lands that are not fit for traditional food crops better for growing energy crops
to avoid competition for land between nonfood and food crops (Giacobbe et al.
2016). Keeping in view, the cost effectiveness, lignocellulosic
feedstocks have several advantages over other agricultural feedstocks like;
potatoes, cornstarch, sugarcane juice in addition to easy production with low
cost unlike other food crops. A. donax
can also be considered a good candidate to supplement or replace maize, sorghum
and other energy food crops in particular to produce green energy (Pilu et al.
2013).
The lignocellulosic biomass structure is highly complex
polymer, primarily composed of cellulose, hemicelluloses and lignin which is
not directly accessible for microbial or enzymatic degradation (Ghorbani et al.
2015). This makes it a major limitation for efficient bioethanol
production. Prior to the conversion from biomass to bioethanol, effective
pretreatment approach is a prerequisite (Scordia et al. 2011) in order to break
lignin and hemicellulose making cellulose available to hydrolyzing enzymes for
the release of sugar monomers that can finally be converted into ethanol or any
other valuable products (Huang et al. 2015).
The pretreatment processes are used to increase
carbohydrate degradability and porosity, remove lignin and preserve
hemicellulosic constituents (Gupta and Lee 2010;
Chiaramonti et al. 2012). The
pretreatment and conversion of lignocellulosic biomass into hydrolysable
constituents by alkali, acid, microbes and commercial enzymes is very
challenging. Alkaline (NaOH) pretreatment studies
reported for Miscanthus, wheat straw
and cotton stalk show effects on delignification and ultimately on enzymatic
hydrolysis yield. Some contents of hemicelluloses and cellulose were also
degraded and removed from biomass feedstocks by the action of hydroxide ions in
addition to delignification during alkaline pretreatment (Cheng et al.
2010).
Ultrasound, a sound wave, through agitation and
cavitation in liquid can produce energy and has great potential to damage the
surface structure of biomass. Ultrasound mainly applied to supplement
pretreatment of various lignocellulosic biomass with different reaction
solutions (Wang et al. 2016). However, for both bioethanol and biogas
production, a higher susceptibility of biomass to pretreatments will allow the
use of more environmentally friendly processes, by lowering pollution and
energy costs.
Biological pretreatments are based on using
microorganisms capable of degrading cellulose, lignin and hemicellulose.
Cellulose fraction is perhaps the most resistant component to biological
attack. White, soft and brown rot fungi are mainly used to pretreat
lignocellulosic feedstocks and enhance the enzymatic hydrolysis yield (Anwar et al.
2019). Brown rots fungi mostly degrade cellulose, whereas soft and white
rots mainly involved in both lignin and cellulose degradation. White rot fungi
are considered as among the most effective basidiomycetes for biodegradation of
lignocellulosic biomass (Sun and Cheng 2002).
The low energy consumption, ecofriendly, cost effective, absence of chemicals
and inhibitory compounds, simple and lesser requirements are imperative aspects
of microbial pretreatment which attracts attention of scientists and
researchers (Chiaramonti et al. 2012). Fungi can efficiently produce ligninolytic
enzymes, which play a key part in biological pretreatments. Fungi by
biodegrading lignin improve availability of enzymes to the cellulose in
lignocellulosic biomass structure. Consequently, modified biomass is more
vulnerable to enzymatic degradation and digestion (Ghorbani et al. 2015).
Moisture content, particle size, pretreatment time and temperature
significantly affect degradation of lignin and enzymatic saccharification yield
(Kumari and Singh 2018).
Although an enormous amount of literature is available
with respect to second generation biomass, but no work has been reported so far
on the potential of Arundo biomass
after being pretreated comparatively with fungus and sonication. The current
study was designed with the goal to investigate effective approaches amongst different
fungal and sonication pretreatments in order to improve bio-delignification,
comparative importance of each pretreatment on rate of fermentable sugars and
efficacy of conversion by enzymatic hydrolysis of A. donax. Comparative evaluation of different pretreatment
approaches and their impact on saccharification yield from A. donax was the main objective of current study.
Collection and preparation of A. donax
The
sampling of A. donax was carried out
in October from Kallar Kahar Lake, a brackish lake with geographical
coordinates, Latitude: 32° 47' 0" North; Longitude: 72° 42' 0" East (Ahmad and Erum 2012) situated in Jhelum
respectively, Punjab, Pakistan. In current research plant culms of A. donax were obtained for estimation of
reducing sugar and bioethanol production. Physiological parameters were
measured then finely cut from internode 2 with sharp sterile plant cutter.
Three biological repeats were collected. A.
donax biomass was brought to lab, cleaned, weighed and the leaves were
removed. The Arundo biomass was
initially dried at 45°C for 72 h (Zakir et al.
2016) and stored in airtight bags.
Fungal strains and growth conditions
The fungal
strains; Trichoderma koningii and Aspergillus niger were provided by the
Department of Microbiology, Quaid-e-Azam University, Islamabad, Pakistan. The
strains were aseptically cultured for 7 days at 30oC on potato dextrose agar (PDA)
plates. Fungal strains were maintained on sterile PDA plates and preserved at 4oC
(Ghorbani
et al. 2015). The freshly grown mycelium were further inoculated
into 250 mL Erlenmeyer flasks in 30 mL of PD broth growth medium at pH
5.6 and incubated at 30oC with 180 rpm
for 7 days for fungal pretreatments.
Reagents and enzymes
The
reagents used in experiments were sodium acetate, glucose, citrate buffer,
glacial acetic acid, antibiotics i.e., tetracycline hydrochloride and
cyclohexamide and all the chemicals used throughout current experimental study
were procured from Sigma-Aldrich (Beijing, China), cellulase of T. reesei from Shanghai Boao Biotech.
Corp., Shanghai, China were of highest purity and analytical grade. Water was
purified using (Master-D series) high
performance ultra-pure water system.
Pretreatment of Arundo biomass
In order to
investigate higher yield of reducing sugar and release efficiency; physical,
fungal and sonication pretreatments were carried out. Untreated biomass was
taken as control. All experiments were carried out in triplicates.
Physical fragmentation: The internodes 2–5 of each collected plant were combined
for making a composite biomass then chipped and pulverized in micro soil plant disintegrator crusher pulverizer
grinding mill (FT102). For
achieving uniform particle size, the ground biomass was screened through 20
mesh sieves to attain homogenous particle size (850 µm)
for efficient pretreatment of Arundo biomass.
The ground biomass was stored in sterile airtight polybags at room temperature
under dry conditions until use for further analysis and pretreatments (Giacobbe et al.
2016; Silverstein et al. 2007).
Bio-delignification: For bio-delignification experiments the respective
sterile aqueous culture medium containing 5 g L-1 yeast extract, 15 g L-1 glucose and 15 g
L-1 peptone were prepared aseptically. The aqueous
solution was then enriched by copper (CuSO4·H2O),
manganese (MnSO4·H2O) and zinc
(ZnSO4·7H2O) ions with final
concentrations equal to 2.5 µM, 0.1 mM and 5 µM, respectively pH of the prepared solution
adjusted at 4 ± 0.05. Fungal pretreatment carried
out separately by addition of 100 mL prepared culture medium to 4 g of Arundo dry biomass in 500 mL Erlenmeyer
flasks. Following sterilization, each flask was then inoculated by two plugs of
10 mm diameter with 5 days fresh grown PDA fungal
culture medium (T. koningii and A. niger), capped with hydrophobic
sterile cotton plugs and incubated in an orbital shaker at 30oC, 160
rpm (Ghorbani
et al. 2015) up to 14 days. The biopretreated and untreated A. donax samples were withdrawn from
triplicate flasks of each fungal culture; T.
koningii and A. niger after 7 and
14 days. Pretreated and untreated samples were washed with double distilled
water (50 mL) at 28oC at 180 rpm for 1 h, then vacuum filtrated
through ceramic Buchner funnel (SHB III, TOPTION)
with filter paper lining to separate liquid and solid and remove most of
water-soluble components. Solid fraction was extensively washed with distilled
water until neutral pH attained followed by last washing with 50 mM citrate
buffer (pH; 5) that subsequently used in enzymatic hydrolysis (Carvalho et al.
2013; Amezcua-Allieri et al. 2017),
samples were vacuum filtered through Buchner funnel and oven dried at 60°C for
24 h to a constant weight (Mishra et al. 2014; Ghorbani et al. 2015; Wang et al. 2016; Zakir et al.
2016). After cooling down, pretreated dried residues were kept in
desiccator, collected and stored in zip-lock bags at room temperature for
enzymatic hydrolysis. Untreated (non-inoculated) biomass samples were taken as
control, incubated and further treated under same conditions.
Sonication: A. donax (1 g DM)
ground biomass was mixed in 100 mL of distilled water and subjected to
sonication at frequency of 50 Hz for 15, 30, 45 and 60 min. The slurry was
vacuumed filtered, washed with distilled water (H2O)
and dried at 70°C for 24 h (Mishra et al. 2014). The solid fraction
of Arundo biomass was then proceeded
for enzymatic hydrolysis.
Enzymatic hydrolysis (EH)
Enzymatic
saccharification of fungal and sonication pretreated biomass as well as their
respective control samples of A. donax was
carried out using commercial cellulase derived from T. reesei (≥ 700 units) containing
60 FPU g-1 (Filter
Paper Unit per gram) enzyme activity. The 30 FPU g-1 of enzyme
was added to dried biomass (Wu et al. 2016). Enzymatic
hydrolysis was performed in triplicate using 250 mL sterile glass reactors,
each containing 50 mL of sodium acetate buffer: 50 mM L-1; pH 5 at
room temperature which was prepared in autoclaved distilled water. The residual
pretreated dry biomass 1g was mixed with acetate buffer resulting concentration
of substrate 2% (w/v). Enzymatic reaction proved more effective when diluted
with buffer as compared to distilled water (Zakir et al. 2016). Each enzymatic
hydrolysis mixture containing 40 µg mL-1
tetracycline (Cheng et al. 2010) and 30 µg mL-1 cyclohexamide was
incubated at 48oC for 72 h with 120 rpm in an orbital incubator
shaker (MaxQ 8000, Thermo Fisher). Adding cyclohexamide inhibits DNA
translation of the eukaryotic cells to inhibit cell growth which ultimately
leads to death of cell. The main target of using cyclohexamide and tetracycline
hydrochloride was the inhibition of microbial growth that affects pH during
enzymatic hydrolysis process and enzymatic activity (Silverstein et al. 2007; Wang et al. 2018).
Samples of 2 mL were withdrawn after 24, 48 and 72 h of
enzymatic saccharification to evaluate glucose concentration. Hydrolysates were
heated for 10 min in boiling water to stop enzymatic reaction ( Martin-Sampedro et al. 2017), cooled at room temperature and separated by
centrifugation at 10,000 rpm for 10 min. Then collected the supernatant and
filtered through 0.2 µm nylon syringe filters (Wang
et al. 2018) and stored in
refrigerator at -20oC for further analysis. Consequently, quantity
of reducing sugars was measured by using glucose calibration curve (Amezcua-Allieri
et al. 2017) using (Beckman DU640 UV/Vis) spectrophotometer at 540 nm. Reducing sugar (glucose) concentration was measured
by 3, 5-dinitrosalicylic acid (DNS) by Miller
(1959). The values presented in the results were means of triplicates with the
values of standard deviation and calculated in mg of reducing sugar per g dry
weight of A. donax biomass by
following equation:
Reducing sugar yield (mg g-1 dry biomass) = (r xn) /m
where “r” is reducing sugars concentration (mg mL-1) from
hydrolysate, “n” total volume hydrolyzed (mL), “m” initial dry weight (g) of Arundo biomass. The values were expressed in mg g-1 basis.
Physico-chemical characteristics of Arundo biomass
The moisture, volatile matter and
ash of well dried raw A. donax of
October were analyzed according to standard protocols;
ASTM D 3174, ASTM D 3173 and ASTM D 3175, respectively on the dry weight
basis. Fixed carbon content of biomass was calculated by the difference. The
fixed carbon content is the value of difference from 100% biomass to ash,
moisture and volatile matter percent on dry weight basis (Saikia et al.
2015). While Cellulose, hemicelluloses and lignin content of raw Arundo biomass were determined by acid detergent fiber (ADF), neutral detergent fiber
(NDF) and acid detergent lignin (ADL) methods (Omar et al. 2011; Saikia et al. 2015) with some
modifications. The elemental analysis carbon, hydrogen, nitrogen and sulphur of
well dried raw Arundo biomass was
analyzed by automatic CHNS analyzer (Vario EL cube). The
content of oxygen was measured by calculating the difference of O (%) = 100 (%)
– C (%) – H (%) – N (%) – S (%) (Licursi et al. 2015). The values
reported in results are the mean ± standard deviation of three replicates.
Scanning electron microscopy (SEM)
The
characteristics of surface morphology of A.
donax biomass were scanned by using SEM (HITACHI-S-3400N), current; 30 mA, voltage; 15 kV and
distance; 14.3 mm. Untreated and pretreated A.
donax biomass samples were dried in oven at 50oC for 24 h (Wang et al.
2018) for removing moisture content. Dried Arundo biomass (1 mg) was examined and photographed by using SEM to
investigate surface morphology of each sample both in degraded (pretreated) and
intact (untreated) biomass.
Statistical analysis
To verify
differences regarding untreated and pretreated Arundo biomass by an analysis of variance; univariate
one-way analysis of variance (ANOVA), following
post hoc multiple comparison Tukey and Duncan’s test by using Statistical Product and Services Solutions (SPSS) version 23 at statistical significance level of
0.05. A multifactorial design was employed comprising dependent variables;
fungal strains T. ressei and A. niger, duration of pretreatment
7, 14 days and harvest time, sonication time and independent variables was
glucose yield.
Results
Characterization of Arundo biomass
The
proximate analysis reveals fixed carbon (20.8%), volatile matter (71% ± 0.76),
ash (4% ± 0.01) and moisture content (4.2% ± 0.01%) whereas compositional
analysis showed cellulose (24%), hemicellulose (30%) lignin (26.8%) and other
components (16.9%) and elemental analysis demonstrated nitrogen (0.31 ± 0.02%),
sulfur (0.09% ± 0.004), carbon (45.1% ± 0.01), hydrogen (5.9% ± 0.03) and
oxygen (48.69%) were found in Arundo biomass
on percent dry weight basis are presented in Table 1. Based on the results of
elemental analysis, H:C ratio computed is 0.13 and O:C ratio is 1.07.
SEM analysis
The SEM
images of degraded and non-degraded biomass indicated that pretreated biomass
exposed many discernible surface morphologies with porous and rough surfaces
might be due to removal of lignin and hemicelluloses after pretreatments
compared to untreated biomass (Fig. 1). Untreated Arundo biomass (Fig. 1a) had intact, compact and smooth surfaces.
Fig. 1b revealed layering, scaling and visible abrasions of fibers due to
degradation of lignin and hemicelluloses after T. koningii pretreatment. A.
niger pretreated biomass (Fig. 1c) resulted in mild scaling and layering as
compared to Fig. 1b which may possibly be owing to partial decomposition of
hemicelluloses. However, significant cavitation and shock effect of sonication
enhanced removal of lignin beside considerably increased degradation of
hemicelluloses (Fig. 1d–g). The surface of
Arundo biomass revealed few sunken areas beside layering and scaling (Fig.
1d), also with erosion troughs and clear cracks (Fig. 1e). Also, there was
disintegration of upper layer (Fig. 1f) as well as scaling and cavitation in A.
donax biomass was noted (Fig. 1g). Among all pretreated biomass highest
disintegration of surface morphology could be observed in Trichoderma pretreated biomass rendering it advantageous for enzyme
accessibility and processivity. Trichoderma
pretreated biomass was more accessible for enzyme among all other
pretreated biomass. Among all pretreated biomass samples, morphological
structure of T. koningii pretreated
biomass was highly destroyed and disintegrated, making it more advantageous for
saccharification. This observation was in accordance with our results of T. koningii having maximum yield of
glucose after enzymatic hydrolysis (Fig. 2).
Table 1: Characteristics of raw A. donax biomass% dry weight
|
Characteristics |
%age |
|
Proximate |
|
|
Moisture content |
4.20 ± 0.01 |
|
Ash |
4.00 ± 0.01 |
|
Volatile matter |
71.00 ± 0.76 |
|
Fixed carbona |
20.80 |
|
Ultimate |
|
|
Carbon |
45.10 ± 0.01 |
|
Hydrogen |
5.90 ± 0.03 |
|
Nitrogen |
0.31 ± 0.02 |
|
Sulfur |
0.09 ± 0.004 |
|
Oxygenb |
48.69 |
|
Compositional |
|
|
Cellulose |
24.00 |
|
Hemicellulose |
30.00 |
|
Lignin |
26.80 |
|
Othersc |
16.90 |
Results are presented as means of three repeats with the values of
standard deviations. (a) Remaining
organic matter after biomass volatile matter and moisture have been driven off;
(b) Oxygen content was calculated by
difference. O (%) = 100 (%) – C (%) – H (%) – N (%); (c) Calculated as the difference between 100% and the sum of the
composition of four components i.e. ash, cellulose, hemicellulose and lignin
%201449-1456_files/image002.jpg)
Fig. 1: Scanning electron microscopy (SEM) images of untreated
and pretreated A. donax biomass at 3k x magnification with 10 µm scale
bar. a: Untreated, b: T. koningii, c:
A. niger, d: Sonication 15 min (S15),
e: Sonication 30 min (S30), f: Sonication 45 min (S45), g: Sonication 60 min
(S60)
Effect of fungal and sonication pretreatments on enzymatic
saccharification
Fig. 2a showed the effect of incubation for 7 and 14 days after
pretreatment with two fungal strains T. koningii and A. niger along with untreated Arundo
biomass and the production of reducing sugars (glucose) at 24, 48 and 72 h of
enzymatic hydrolysis process. The cellulose digestibility was substantially
enhanced by T. koningii resulting in
maximum fermentable sugars release. The optimum glucose 237.7 ± 0.7 mg g-1
dry solids was released after 14 days of T. koningii pretreatment at 72 h of enzymatic hydrolysis and significantly higher than untreated Arundo
biomass 106.6 ± 0.3 mg g-1 solid dry biomass (p value
<0.05) while A. niger pretreated biomass liberated 214.8 ± 0.8 mg g-1 solid dry biomass of glucose with same conditions. All pretreated
biomass exhibited similar trend of increased glucose yield by increasing
duration of enzymatic hydrolysis. The prolonged duration of pretreatment and
enzymatic hydrolysis favored efficient generation of glucose. Among enzymatic saccharification of each
pretreatment 72 h released maximum glucose yield followed by 48 and 24 of the
reaction which is statistically significantly different (Table 2). Data
showed that with sonication time ranging from 15–60 min at 15 min interval on
glucose harvest the digestibility of
cellulose content and glucose yield was significantly enhanced by increased
duration of sonication (Fig. 2b). It was found that sonication for 60 min
liberated more reducing sugars (glucose) followed by 45, 30 and 15 min (Table
3).
Discussion
%201449-1456_files/image004.png)
Fig. 2: (a) Effect of 7
and 14 days pretreatment on glucose yield (mg g-1) of A. donax biomass
by T. koningii and A. niger at 24, 48 and 72 h of enzymatic
hydrolysis. Untreated biomass was taken as control. (b) Effect of
untreated, sonication 15 min (S15), sonication 30 min
(S30), sonication 45 min (S45) and sonication 60 min (S60) of A.
donax biomass at 24, 48 and 72 h of enzymatic hydrolysis. Untreated
biomass was taken as control
Table 2: Estimating effect of untreated, T. koningii and A. niger pretreated A.
donax biomass on glucose yield (mg g-1) during enzymatic
hydrolysis (h)
|
Time (h) |
Pretreatment
|
Glucose yield (mg g-1) |
|
|
|
|
7 days |
14 days |
|
24 |
Untreated |
35.7 ± 0.4c |
58.3 ± 0.2c |
|
A. niger |
91.7 ± 0.3b |
110.7 ± 0.2b |
|
|
T. koningii |
115.6 ± 0.4a |
132.4 ±0.5a |
|
|
48 |
Untreated |
49.6 ± 0.2c |
73.5 ± 0.1c |
|
A. niger |
124.1 ± 0.2b |
168.2 ± 0.2b |
|
|
T. koningii |
159.2 ± 0.5a |
192.3 ± 0.5a |
|
|
72 |
Untreated |
65.9 ± 0.0c |
106.6 ± 0.3c |
|
A. niger |
168.6 ± 0.2b |
214.8 ± 0.8b |
|
|
T. koningii |
204.1 ± 0.2a
|
237.7 ± 0.7a
|
|
Results are presented as means of three repeats with the values of
standard error (SE). Values
followed by different letter within each column are significantly different (P < 0.05) by Univariate Analysis of
Variance “UNIANOVA” following Tukey’s test analysis using Statistical Product
and Service Solutions (SPSS) version 23
Table 3: Estimating effect of untreated and sonication pretreated
A. donax biomass on glucose yield (mg g-1) during enzymatic
hydrolysis (h)
|
Time (h) |
Treatment |
Glucose
yield (mg g-1) |
|
24 |
Untreated |
24.6±0.4 |
|
S15 |
47.9±0.1 |
|
|
S30 |
80.7±0.3 |
|
|
S45 |
92.3±0.3 |
|
|
S60 |
119.9±0.6 |
|
|
48 |
Untreated |
44.5±0.2 |
|
S15 |
63.1±0.0 |
|
|
S30 |
112.5±0.5 |
|
|
S45 |
125.4±0.6 |
|
|
S60 |
164.9±0.5 |
|
|
72 |
Untreated |
67.1±0.4 |
|
S15 |
76.1±0.2 |
|
|
S30 |
152.6±0.4 |
|
|
S45 |
161.4±0.2 |
|
|
S60 |
204.5±0.4 |
*Pretreatments:
Untreated, S15; Sonication 15 min, S30; Sonication 30 min, S45; Sonication 45 min,
S60; Sonication 60 min
Proximate,
ultimate and compositional analyses reveal the optimal values of their
respective compositions in A. donax (Table 1). The moisture content (4.2
± 0.01%) of A. donax in current study
is in accordance to the values of moisture content and dry matter for A. donax reported in previous studies (Mejdi et al.
2010; Natalia and Adam 2011). Moisture content of the crop depends on
many factors and remains relatively low, rendering the dry biomass high. Moisture content is of paramount importance since lower
the moisture content of biomass would decrease the tendency of decomposition
and save energy (Sánchez et al. 2019), in addition, less energy would be
required for size reduction (Ani 2015) which would reduce pretreatment time and
decrease the cost of pretreatment (Quintero et al. 2011); Low moisture
content would be more favorable to stop anaerobic microbial degradation thus
also permit safe long term storage of the biomass (Rentizelas 2016). Lower ash
content (4 ± 0.01%) in current study
is a favorable characteristic regarding biofuel production and is in line with
the previous reports (Vernersson et al. 2002; Krička et al. 2017). The ash content is one of the most remarkable
characteristics of LB Biomass, as it comprises inorganic matter and minerals,
being an integral part of biomass, it affects the combustion rate. Lower ash
content is optimally most conducive to a favorable outcome with respect to
biofuel production (Singh 2019). A high volatile matter (71 ± 0.76%) in A. donax
biomass found in current study is another good attribute for biofuel production
(McKendry 2002), which is also consistent with
the results (71.3%) reported by Vernersson et al. (2002). The fixed carbon
content and volatile matter affects the biological conversion mechanisms of the
fuel (Vassilev et al. 2010). In current study, fixed carbon content was
20.8% slightly different from previously reported values. It is the valuable
characteristic of the biomass as it represents the potential of biomass to
release sugars and be used as biofuel source (García et al. 2012; Vanja et al. 2017). As given in Table 1, the elemental analysis
indicated carbon, hydrogen, nitrogen, sulfur and oxygen were found with
insignificant difference (Licursi et al. 2015; Saikia et al. 2015). Upper surface of biomass generally contains
hemicellulose, lignin and ash enclosing interior cellulose fiber (Wang et al.
2016). Fuel efficacy of biomass depends on the atomic ratio of H/C and
O/C. Lower ratio (0.13) found in our results reveals the higher energy content (Singh 2019). Current study
shows that A. donax biomass comprises
cellulose, hemicelluloses and lignin (Table 1), which is sufficient holocellulose
to be converted into monomeric sugars and ultimately to bioethanol (Saikia et al.
2015; Lemões et al. 2018).
It was found
that both fungal strains T. koningii and A. niger produced
maximum sugars after 14 days of pretreatment. T. viride resulted in
conversion of cellulose to glucose as 56% of the theoretical yield after
enzymatic saccharification of rice straw pretreated biomass (Ghorbani et al. 2015). However, reducing sugar yield
obtained by fungal-pretreated cotton stalk (10.91–55.6 mg g-1) reported
by Wang et al. (2016) was still lower than the current study. Pretreatment
of rice straw by Pleurotus florida showed total reducing sugars at 353
mg g-1 of dry biomass with 75% efficacy at 72 h of saccharification (Naresh Kumar et
al. 2018). The lignocellulosic waste sawdust at 72 h of
saccharification with cellulase enzyme (derived from T. estonicum) produced 78.56% glucose (Saravanakumar and Kathiresan 2014).
The enzymatic digestibility
following fungal pretreatments, was reported higher as a result of significant
lignin degradation (Wan and Li 2012). Sonication pretreatment of Arundo biomass for
60 min was found more efficient in producing reducing sugar yield. This is in
accordance with the findings reported for maximum cellulose release at 60 min
from A. donax biomass (Mishra et al.
2014). Alkali assisted microwave
pretreatment by using cotton stalk biomass, collected 0.495 g g-1 reducing
sugars (Vani
et al. 2012). The higher levels of sugar yield can be attributed to
lower lignin portion (Wang et al. 2016). The SEM images of pretreated biomass
illustrated disintegration and abrasions as compared to untreated biomass. The
disruption of the integrated structure after various pretreatments, increased
the accessibility of cellulose, which may enhance the effective absorption of
enzyme in the interior of cellulose, as is obvious too by the increase in more
glucose release from cellulose (Fig. 2). T.
koningii pretreated biomass was highly susceptible to enzyme degradation as
compared to other pretreated biomass and hence, produced highest glucose yield
among all the pretreatments. The destruction and disintegration after different
pretreatment approaches enhance the accessibility of cellulose, which in return
increase efficient absorption of enzyme resulting in maximum fermentable
sugars. The SEM images of 60 min
sonication pretreatment, exhibited greater
disruption of Arundo biomass, which also validated our findings of
higher glucose yield. These outcomes evidently implied advantage of T. koningii pretreatment over A. niger and sonication for effective conversion of A.
donax to bioethanol.
Conclusion
A. donax is rich in
cellulose but difficult to degrade into fermentable sugars. In untreated Arundo biomass, 72 h hydrolysis glucose
yield was found significantly lower than pretreated biomass. SEM observations
evidenced advantageous intensive structural changes at varying degrees after
pretreatments of Arundo biomass. T. koningii pretreated Arundo biomass had maximum surface
disintegration and degradation of fibers, which led to higher fermentable sugar
yield as compared to A. niger.
Enzymatic hydrolysis showed that Arundo biomass
incubated with T. koningii also
yielded higher glucose content. T.
koningii pretreatment proved to be a feasible and eco-friendly alternative
for higher amount of fermentable sugars and second-generation ethanol.
Sonication is not viable alternate for Arundo
biomass pretreatment. Prolonged pretreatment is a main barrier for
application of fungal pretreatments. Results further showed the effectiveness
of fungal pretreatment for improved glucose yield from Arundo biomass. Further research studies based on combined
pretreatments, characterization of new enzymes, fungal consortiums, cost
effectiveness, environmental sustainability, genetic engineering, enzymatic hydrolysis optimization,
delignification and enhancement of fermentation are needed to explore
more efficient strategies to obtain highest yield of fermentable sugars from
lignocellulosic biomass.
Acknowledgments
First
author (MS) acknowledges the financial support from Higher Education Commission
Pakistan; under “International Research Support Initiative Program” (Project
no: 1-8/HEC/HRD/2017/8371). MS also acknowledges COMSATS University, Islamabad, Pakistan for
providing scientific facilities; Director Zheng Peng for laboratory and
equipment’s facilitations at Joint Laboratory of Advanced Applied Technology of
Water Treatment, Gansu Academy of Membrane Science and Technology Lanzhou;
Abdul Wahab, Habib Ullah, Falak Shair and Zafar Hayat; PhD scholars of Applied
Microbiology and Biotechnology Laboratory COMSATS University Islamabad for
assistance in handling analytical data and preparing illustrations.
Author Contributions
MU
conceived of the presented idea and supervised the research work. SM designed
the study, conducted experiments, collected data, performed analytical methods,
drafted manuscript. ESS, XKL, FYH and MU supervised the research work. XKL and
ESS worked on almost all technical details of this research study and improved
write-up. KM and JJ performed analytical computations and took lead for
structural analysis. All authors discussed the results provided critical
feedback and helped final shape the research, analysis and manuscript.
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