Effect
of Pretreatments on Physicochemical Characteristics of Olive Pomace and on
Production of Cellulases from Trichoderma reesei RUT C30 under Solid-State
Fermentation
Malika Boutiche1,2*,
Fatma Sahir-Halouane1, Leila Meziant2, Fairouz Saci2,
Kahina Oudjedi3, Mouna Derdour2,
Karima Ouffroukh2, Ibtissem Maghboune2, Samah Fiala2
and Amel Bekrar2
1Université M'Hamed BOUGARA de Boumerdes, faculté des
Sciences, Laboratoire VALCORE, Avenue de l’indépendance –35000 Algeria
2National Centre for Biotechnology Research, BPE 73, Ali
Mendjeli, Nouvelle Ville, 25000-Constantine, Algeria
3Institut National de l’Alimentation, la Nutrition et des
Technologies Agro-Alimentaires (INATAA), 25000 Constantine, Algeria
*For correspondence:
m.boutiche@univ-boumerdes.dz
Received 31 March 2022;
Accepted 10 December 2022; Published 27 January 2023
Abstract
Olive pomace (OP) is a cheap and
abundant agricultural by-product that could be valorized by different
biotechnological processes. The present study was conducted to better
understand the effects of alkaline, milling and thermal pretreatments on OP for
obtaining high value-added products (cellulases). Trichoderma reesei RUT
C-30 fungus was used for cellulases production on OP substrate under
Solid-State Fermentation (SSF) process and cellulases activity was assessed by
the filter paper method (FPase). The effect of the three pretreatments
and their combinations on physico-chemical composition and cellulases
production was investigated. Results showed that untreated OP was a favorable
environment for the growth of T. reesei and a good fermentation
substrate that gave FPase activity of 0.83 IU/g DS. The chemical
composition (lipids, proteins, carbohydrates and ash) was significantly (P <
0.05) affected by the different pretreatments as well as their
combinations. Regarding the fiber fraction, alkaline and thermal pretreatments
did not affect the NDF content, while a remarkable decrease (29.88%) was
recorded after milling pretreatment. Alkaline pretreatment decreased
significantly the cellulose contents whereas milling increased it of 35%. ADL
fraction was only decreased by the milling treatment of 42%. No significant
effect of thermal pretreatment was noticed on ADL and cellulose. The alkaline
pretreatment with 1% NaOH improved the cellulase activity to a value of 1.28 IU/g
DS, while lower yields were obtained after milling (0.2 IU/g DS) and thermal
(0.15 IU/g DS) pretreatments. This study showed that only alkaline treatment
improved the production of cellulase from OP without being combined with
milling and thermal treatment. © 2023 Friends Science Publishers
Keywords:
Olive pomace; Trichoderma reesei;
Pretreatments; Cellulases; Solid-state Fermentation
Introduction
The world production of olive
oil is valued to be more than 18 million tones/year (Coimbra et al. 2010) and Algeria covers more
than 1.5% of this production (Stamatelatou et
al. 2012). 42% of the national production of olive oil is located in the
central region: Béjaia, Bouira, TiziOuzou and Jijel (Rives
2021). However, olive oil extraction process produces great amount of wastes.
According to Nefzaoui (1991), 100 kg of olives produced about 35 kg of crude
olive pomace and 100 liters of vegetation water. Depending on the extraction method, olive
pomace (OP) can reach up to 30–40% of olive oil production (Aliakbarian et al. 2011). Olive pomace by-product
contains fragments of skin, pulp, stones and oil (Mirabella et al. 2014). OP is composed of lignin
(31%), hemicellulose (24%), cellulose (14%), fat (11%), soluble sugars (6.5%),
protein (6%) and many mineral salts (Roig et
al. 2006). This physico-chemical composition of OP depends principally on
the type and origin of olives, environmental conditions and storage time
(Papaioannou et al. 2013). Several
studies have proven the negative effects of this solid by-product on the
microbial flora of the soil and even on the aerial environment (Aranda et al. 2007). Therefore, it is important
to manage these wastes in order to minimize their negative effects on
the environment. In fact, olive pomace contains valuable raw material such as a
great proportion of organic matter and a varied
range of nutrients, which could be used for energy generation, as an animal
feed or as a fermentation substrate in biotechnological means for bio-fuels biofuels or enzymes production (Roig et al. 2006; Roussos et al. 2009). Still, some obstacles are associated with effective
utilization of lignocellulosic residues for enzymes production. The main
constraint is the recalcitrance of the plant cell walls of the lignocellulosic
fractions (Kumar and Sharma 2017) to develop effective and low-cost pretreatments
as potential ways of altering the structure and improving the degradability of
lignocellulose biomass (Hendriks and Zeeman 2009). In literature, several
pretreatments were described taking into account several aspects: (1)
mechanisms concerned (2) advantages and disadvantages and (3) economic
valuation (Menon and Rao 2012). According to the National Research Council
(1999), an effective pretreatment must preserve the hemicellulose fraction,
minimize the production of growth inhibitors of fermentation microorganisms and
reduce energy costs.
Different pretreatments have been investigated on olive solid wastes,
including physical (Neifar et al. 2013; Leite et al. 2016), chemical (López-Linares et al. 2013; Pellera et al. 2016; Erdocia et al. 2017),
thermal (Fernández-Bolaños et al.
2001; Aliakbarian et al. 2011) and
biological (Haddadin et al. 2009)
methods and various combinations thereof (El-Ghonemy et al. 2014; Ouyang et al. 2018). Physical pretreatment means include mechanical deterioration and
irradiation that lead to structural disruption and reduction of the particle
size, degree of polymerization and crystallinity of the raw material (Cara et al. 2007; Ravindran and Jaiswal
2016), which increases the enzymatic digestibility of cellulose and
hemicelluloses in the lignocellulosic biomass (Mtui 2009). Chemical
pretreatments are performed using alkalis (NaOH, Ammonia), acids (H2SO4),
oxidants (Ozone, Oxygen, H2O2), organic solvents (Alcohols, Organic acids) or
ionic liquids (Organic salts) (Gandla et
al. 2018). Chemical pretreatments favors hydrolysis of lignocellulosic
biomass by eliminating hemicelluloses, disrupting lignin or reducing the
crystallinity of cellulose during processing (Mosier et al. 2005; Zheng et al. 2009).
Among these, alkali has been most extensively investigated. The use of alkali
causes the degradation of ester and glycosidic side chains causing in
structural modification of lignin fraction, separation of structural bonds
between lignin and carbohydrates, cellulose swelling and its partial
decrystallinization, in addition to a dissolution of hemicellulose (Zheng et al. 2009;
Brodeur et al. 2011). Thermal pretreatments are effectively used on an industrial scale for
lignocellulosic residue processing: hydrothermal, steam-explosion and
hydro-chemical pretreatments are attested to cause elimination of
hemicelluloses without being hydrolyzed, re-localization of lignin and
hydration of cellulose and at the same time, swelling the pore size of the
fibers which facilitate the enzyme accessibility (Gandla et al. 2018). In biological pretreatments, lignocellulosic
degrading fungi are used to reduce the lignin barrier from the biomass prior to
fermentation. Although, this pretreatment is only significant if combined with
other pretreatments (Vasco-Correa et al.
2016). Filamentous fungi species are recognized for their high aptitude to
secrete large amount of enzymes into their environment, making them very
interesting for industrial enzyme production (Gudynaite-Savitch and White 2016;
Srivastava et al. 2018). Trichoderma reesei is the most used
fungus in enzyme industry, particularly for cellulases (Jun et al. 2011; Hinterdobler et al. 2021). Trichoderma cellulases are presently used in many industries such
as textile, food, biofuel production, agriculture, animal feed, paper and pulp
industries (Linke et al. 2015;
Imran et al. 2016). Cellulases enzymes catalyze the bioconversion of cellulose into fermentable
sugars. Cellulases complex are formed of three types of enzymes: endo-1,4-β-D-glucanase, exo-1,4-β-D-glucanase and β-glucosidase (Paloheimo et al. 2016). The production of
cellulolytic enzymes by T. reesei has
been the subject of various studies using different substrates (Belal 2013; Pirota et al. 2014; Abdullah et al.
2016). Among the several mutants of T. reesei, T. reesei RUT-C30 is known to be one of the best producing
cellulolytic strain studied (Aftab
and Vermette 2008; Dhillon et al. 2011; Fonseca et al.
2020). Solid-state fermentation (SSF) process has been
used for the cultivation of filamentous fungi because it simulates their living
conditions in their natural habitat (Ugwuanyi et al. 2009; Ray and Behera 2017). This process includes absence or
near absence of free water. The SSF is an attractive way to produce cellulases
from microorganisms because of its lower capital cost investment, simpler
equipment and higher productivity (Ray and Behera 2017; Soccol et al. 2017).
The use of olive pomace as substrate in fermentation processes for
cellulases production requires pretreatments due to the heterogeneity and
complexity of this lignocellulosic biomass. Several studies have been
previously carried out but, to the best of our knowledge, no study has included
at the same time, three pretreatments with their combinations, and provides
elements of answer to their effects on the physicochemical parameters, the
fiber fraction and the kinetics of cellulases production. Consequently, the
main objective of this research was to valorize the OP biomass from Jijel
region (Northen-east of Algeria) as a naturel medium for cellulases production
using Trichoderma reesi RUT-C30 using
three major combined pretreatments namely: alkaline pretreatment with different
concentrations of NaOH (1, 3, 5 and 7%), mechanical milling and thermal
pretreatment.
Materials and Methods
Substrate
In this study, olive pomace, used as substrate
for the solid-state fermentation, was provided by a traditional oil mill
located in Jijel region (Northern Algeria). After the oil pressing operation,
fresh olive pomace was immediately collected, transported to the laboratory and
kept in sealed bags at -20°C for further analysis. Then, divided into four
batches, three of them have undergone different pretreatments, namely
mechanical milling, alkaline and thermal treatments, while the last batch has
been reserved as is for the comparison of the results.
Substrate pretreatments
Mechanical pretreatment
(milling): Olive pomace (OP) samples from the batch 1 were pretreated according to
the method of Haddadin et al. (2009). Samples were oven-dried at 65°C
for 48 h. After that, dried olive pomace samples were crushed into fine
particles using a mechanical grinder with three-phase motor and then separated
through a sieve with a porosity of 1.25 mm. The fine powder was recovered and
stored at -20°C in closed containers.
Alkali pretreatment: OP
samples of the batch 2 were chemically treated with alkaline solution of sodium
hydroxide (NaOH) according to Bansal et al. (2012). 20 g of substrate
was weighed in Erlenmeyer flasks, then 100 mL of NaOH solution prepared at
different concentrations (1, 3, 5 and 7% w/v) were added and left at room
temperature standing for 2 h. After soaking, the treated substrates were
filtered through a metal sieve and carefully washed with distillated water
until the pH of the washing water become neutral. The washed residues were
then oven-dried at 65°C during 48 h. Treated samples were then kept frozen
at -20°C prior to fermentation.
Thermal pretreatment: OP
samples of the batch 3 were treated thermally by using boiled water. Aliquots
of 20 g of each sample were boiled in 100 mL of distilled water for 2 h on
hotplates in thermo resistant flasks. The treated samples were then drained in
a metal sieve and left to cool. After that, the residues were oven-dried at
65°C prior to analysis.
Physicochemical
characterization of olive pomace
In order to evaluate the chemical composition of
all OP samples before and after treatment, physico-chemical analyses were
carried out. The pH of OP samples was measured by pH-meter (Hanna, pH 210,
Romania) following the method of Haddadin
et al. (2009), OP samples were extracted by mixing 5 g of OP with
50 mL of distilled water on magnetic stirrer for 30 min. Moisture content and
dry matter (DM) of OP samples were determined gravimetrically after drying at
105°C to constant weight (Moftah et
al. 2012). Moisture and Dry matter were expressed in percentage (%).
The ash content was assessed through incineration in a muffle furnace
(Nabertherm GmbH, Germany) at 550°C, from aliquots of 5 g of each OP sample
until obtaining white ashes of constant weight. Ash content is expressed in
percentage. Total nitrogen of OP samples was analyzed by the Kjeldahl method (AOAC method number 954.18-B, 1990)
using a semi-automatic Kjeltech apparatus (BUCHI, Digest Automat + Distillation
Unit, Germany). Nitrogen content was expressed in percentage on dry weight
basis. The crude protein content of OP samples was calculated by multiplying
the total nitrogen values by 6.25 to obtain percentages. Lipid content of OP samples was determined by solvent extraction
from 1g aliquots using hexane (25 mL) at 130°C during 45 min using the Soxhlet
Apparatus (FOSS, SoxtecTM 2043 Sweden). Lipid content was expressed
as percentage of dry matter. The fiber contents of OP samples namely, neutral
detergent fibers (NDF), acid detergent fibers (ADF) and acid detergent lignin
(ADL) were evaluated by the method of Soest
and Robertson (1979), using the semi-automated Fibertec apparatus (FOSS
Fibertec2010, Sweden). Fiber contents were expressed in percentage on dry
weight basis. The total carbohydrates content was determined by the
phenol-sulfuric acid method described by Dubois et al. (1956), after hot extraction of 1 g of OP
samples using 16 mL of ethanol 80% at 100°C during 30 min. Extracts containing
carbohydrates were recovered by centrifugation (5000 rpm/10 min) and pellets
were re-extracted twice in the same conditions, then the volume of the extracts
was made up to 100 mL with ethanol 80%. Reducing sugars content of OP extracts was
determined according to the method described by Miller (1959) using the colored dinitro-3, 5-salicylic acid (DNS)
reagent. Carbohydrates and sugars contents were expressed as percentage on dry
weight basis. All treatments and measurements were carried out in triplicate.
Fungal strain and spore
suspension preparation
The fungal strain Trichoderma reesei RUT
C30 was provided by the Industrial Microbiology Laboratory of the University of
Reims Champagne-Ardenne (France). Suspension of spore was prepared by
incubating the cultures of fungus on PDA plates at 30°C for about 5 or 7 days,
then spores were collected by washing with 10 mL of sterile water containing
0.1% (v/v) of Tween 80 and the prepared suspension was adjusted to a
concentration of approximately 3 × 107 spores/mL, using Malassez
counting chamber.
Enzymes production
under solid-state fermentation
The solid fermentation was carried out in 250 mL
Erlenmeyer containing 5 g of substrate (fresh or pretreated OP) and then
humidified by distilled water to a proportion of 1:1 (w/v) taking into account
the initial moisture of each substrate. The preparations were sterilized at
121°C for 20 min. Once cooled, the sterilized substrates were inoculated with
spore suspension previously prepared (3 × 107 spores/mL) then
incubated at a temperature of 30°C for 6, 12, 18, 24 and 30 days of static
fermentation. The operation was performed in triplicate.
Extraction of crude
enzyme
Cellulases enzymes were extracted by mixing the
fermented OP with 50 mL of distilled water and homogenized by UltraTurrax (IKA,
T25 digital, Germany) for 1 min. The
Fig. 1: Scheme of the principal steps of
the experimental work including substrate pretreatments and fermentation
process.
supernatants were recovered after cold
centrifugation at 8500 rpm for 20 min at a temperature of 4°C. Supernatants
will be used for the determination of the pH, soluble proteins and for
measuring the total cellulolytic activity of the extracts.
Determination of total
cellulase activity using filter paper (FPase
activity)
Cellulase activity against
Whatman No. 1 filter paper (W1FP) was measured as described by Silveira et al. (2014). FPase
activity was determined by mixing 0.5 mL of citrate-phosphate buffer
(0.05 M, pH 4.8) with 0.5 mL of the
enzyme extract. After 10 min at 50°C, the W1FP strips, each weighing
approximately 50 mg (1.0 × 6.0 cm), were added to the test tubes. The mixture
was incubated in a water bath at 50°C for 60 min. The reducing sugars released
after the enzymatic reaction were revealed by adding 1.5 mL of DNS reagent,
placing the tubes in a boiling water bath for 5 min and adding 10 mL of
distilled water and then measuring the absorbances at 540 nm by a UV/visible
spectrophotometer (Agilent Technologies Cary 60 UV-Vis, Germany). The
cellulolytic activity was expressed by the international unit, corresponding to
one micromole of glucose released per minute and per mL of enzymatic extract
under the assay conditions. The APFase is
calculated by converting sample absorbances to released glucose concentration
by linear interpolation from a standard curve of D-glucose used as reference.
Table 1: Chemical characterization of olive pomace substrates before and after
milling, alkaline, thermal pretreatments and their combinations
Pretreatment |
Moisture (%) |
DM (%) |
Ash (%) |
Lipid (%) |
TN (%) |
Proteins (%) |
TC (%) |
RS (%) |
Untreated OP |
37.64 ± 1.47a |
62.36 ± 1.47d |
0.70 ± 0.05g |
7.68 ± 0.44b |
0.43 ± 0.03b |
2.68 ± 0.18b |
0.86 ± 0.10b |
0.68 ± 0.08b |
1% NaOH |
2.51 ± 0.94c |
97.49 ± 0.94b |
1.11 ± 0.02e |
1.48 ± 0.15d |
0.25 ± 0.01d |
1.56 ± 0.01d |
0.21 ± 0.01cde |
ND |
3% NaOH |
0.69 ± 0.55d |
99.31 ± 0.55a |
1.25 ± 0.10de |
ND |
0.32 ± 0.07c |
2.00 ± 0.46c |
0.23 ± 0.01cd |
ND |
5% NaOH |
0.83 ± 0.72d |
99.17 ± 0.72a |
1.51 ± 0.07bc |
ND |
0.37 ± 0.04bc |
2.31 ± 0.27bc |
0.22 ± 0.03cd |
ND |
7% NaOH |
0.91 ± 0.72d |
99.09 ± 0.72a |
1.68 ± 0.10b |
ND |
0.32 ± 0.03c |
2.02 ± 0.19c |
0.18 ± 0.01de |
ND |
Milled OP |
4.62 ± 0.04b |
95.38 ± 0.04c |
2.27 ± 0.03a |
18.30 ± 0.26a |
0.81 ± 0.04a |
5.08 ± 0.23a |
2.01 ± 0.05a |
1.33 ± 0.14a |
Milling + 1% NaOH |
1.44 ± 0.01cd |
98.56 ± 0.01ab |
0.87 ± 0.07f |
0.63 ± 0.05e |
0.18 ± 0.02e |
1.14 ± 0.12e |
0.27 ± 0.02c |
0.22 ± 0.01c |
T° OP |
2.51 ± 0.15c |
97.49 ± 0.15b |
0.16 ± 0.06h |
3.48 ± 1.38c |
0.41 ± 0.03b |
2.58 ± 0.20b |
0.18 ± 0.01de |
ND |
T° OP + 1% NaOH |
1.48 ± 0.11cd |
98.52 ± 0.11ab |
1.36 ± 0.21cd |
ND |
0.13 ± 0.02e |
0.79 ± 0.15e |
0.15 ± 0.02e |
ND |
All values were expressed in
percentage (%) on dry substrate basis. Values with different letters in the
same column (a, b, c, d, e, f, g, h) are significantly different (P < 0.05).
DM, Dry matter; ND, Not detected; OP, Olive pomace substrate; T°, Thermal
pretreatment; TN, total nitrogen; TC, Total carbohydrates; RS, reducing sugars.
Fig. 2: Effect of Alkaline pretreatment (1, 3, 5 and 7% NaOH)
on fiber composition of olive pomace substrate
Vertical bars indicate
standard error of three replicates. Different letters for the same parameter
(a, b, c) indicate significant differences (P < 0.05). DM, Dry
matter; NDF, Neutral detergent fibers; ADF, Acid detergent fibers; ADL, Acid
detergent lignin
Soluble proteins
The amount of soluble proteins in the crude
extract was quantified according to the method of Bradford (1976), which is a colorimetric method based on the
reaction of proteins with Coomassie brilliant blue G250 reagent (Bradford
reagent). 100 µL of enzymatic extract
are mixed with 2 mL of Bradford reagent. After stabilization of the color for
10 min, the absorbance of the reaction mixture is determined at 595 nm using a
spectrophotometer (Agilent Technologies Cary 60 UV-Vis, Germany). Protein
concentration was determined by a calibration curve using bovine serum albumin
(BSA) as standard. The different pretreatment and fermentation steps performed
above are summarized in Fig. 1.
Statistical analysis
All data were obtained from at least three
independent assays. Results of the physico-chemical analysis are expressed in
the form of mean ± standard deviation. The significance of the effect of
pretreatments on each physico-chemical characteristic of OP samples was tested
by the analysis of variance (ANOVA) followed by a post-hoc multiple comparison
of means using LSD test using the software STATISTICA (version 5.5, Stat Soft
Inc., USA). Differences with P-values < 0.05 were considered as statistically significant.
Results
Effect
of different pretreatments on OP chemical composition
The effects of all pretreatment used in this study on
the composition of OP are shown in Table 1. All treatments showed a significant
decrease in the moisture content of OP substrates (P < 0.05). A significant increase (P < 0.05) in ash content is noticed for all pretreatments,
except for thermal pretreatment where a significant decrease was observed.
Regarding the lipid content, significant decrease was obtained after all
pretreatments except the milling pretreatment which caused a significant
increase (P < 0.05). Thermal
pretreatment did not affect the protein content of OP substrate, however,
milling increased significantly the
Fig. 3: Effect of milling pretreatment (combined or not
with 1% NaOH) on fiber composition of olive pomace substrate
Vertical bars indicate
standard error of three replicates. Different letters for the same parameter
(a, b, c) indicate significant differences (P < 0.05). DM, Dry
matter; NDF, Neutral detergent fibers; ADF, Acid detergent fibers; ADL, Acid
detergent lignin
Fig. 4: Effects of thermal
pretreatment (combined or not with 1% NaOH) on fiber composition of olive
pomace substrate
Vertical
bars indicate standard error of three replicates. Different letters for the
same parameter (a, b, c) indicate significant differences (P < 0.05).
DM, Dry matter; NDF, Neutral detergent fibers; ADF, Acid detergent fibers; ADL,
Acid detergent lignin
proteins in OP substrate, while the other
pretreatments caused its decrease (P <
0.05). Total carbohydrates and reducing sugar contents of OP substrates
were significantly increased by milling, while the other pretreatments caused
their decrease (P < 0.05), in
addition to a total loss of reducing sugars.
Effect
of different pretreatments on OP fiber composition
As shown in Fig. 2, NaOH pretreatment did not affect
the NDF content. In parallel, a significant decrease was observed on ADF and
cellulose contents, in contrary to hemicellulose fraction, where a significant
increase was observed after alkaline pretreatments. Milling pretreatment caused
significant decrease of NDF (29.88%), ADF (16.22%) and hemicellulose (74.60%)
fractions compared to untreated OP (Fig. 3), whereas, combined pretreatment
augmented significantly NDF (31.51%), ADF (21.01%) and hemicellulose (69%)
fractions compared with the milled OP. It is interesting to note that after
milling pretreatment, ADL fraction decreased of about 42% compared to the
untreated OP, and a slight decrease of 9.3% was observed after combined
pretreatment with 1% NaOH. Contrary to the trends noted for the previous
fractions, the milling has augmented significantly the cellulose fraction of OP
of almost 35%, the same effect was observed for the combined pretreatment.
Regarding the thermal pretreatment, no clear significant effect was observed in
the NDF and hemicellulose fractions after both pretreatments, while ADF
fraction decreased slightly but significantly after thermal process (Fig. 4).
Regarding ADL and cellulose fractions, no significant effect of
Fig. 5: Evolution of PFase activity (a), protein content (b) and pH values (c) of extracts obtained from T. reesei culture on alkaline
pretreated plive pomace during 30 days of fermentation
Vertical bars indicate
standard error of three replicates. OP, Olive pomace substrate; DS, Dry
substrate; FPase activity, Filter paper cellulase activity.
thermal pretreatment on olive pomace was noticed
whether combined or not with 1% NaOH compared to untreated one.
Effect
of alkaline pretreatment on the Fpase activity
As shown in Fig. 5a, the highest FPase
activity (0.83 IU/g DS) was recorded at the 24th day of fermentation
on untreated OP. This activity was improved by 35.15% after the alkaline
pretreatment with 1% NaOH. In contrast, the activity decreased after alkaline
pretreatment with 3, 5 and 7% NaOH by 38.55, 10.84 and 60.24%, respectively. It
is worth noting that olive pomace used in this study was only moistened with
distilled water, without nutrient medium addition for both untreated and
pretreated OP substrates. Despite this, rapid and remarkable growth of the
mycelium of T. reesei RUT C30 was observed from the third day of
fermentation for untreated and pretreated OP with 1% NaOH compared to those
pretreated with 3, 5 and 7% NaOH.
The evolution of soluble proteins (Fig. 5b) for all used substrates
follows the same progression as the cellulolytic activity. The maximum content
of soluble proteins (6.99 mg/g DS) corresponds to the optimum enzymatic
activity for 1% NaOH pretreated OP (24th day of fermentation). It
was also seen that extracts from the untreated substrate presented the highest
soluble protein content in comparison with the other pretreated samples.
The pH evolution (Fig. 5c) showed the same shape for all the samples
analyzed. From different initial values, the pH has decreased during the 12th
first days of fermentation to reach values between 5.8 and 6.8, which
represents the adequate pH for the production of cellulolytic enzymes by T.
reesei. Taking into account the results of the alkaline pretreatment, 1%
NaOH was the most effective pretreatment on OP substrate for cellulases
production (FPase activity). Therefore, it was chosen for all further
combinations with milling and thermal pretreatments.
Effect
of milling pretreatment on the Fpase activity
The first notable result is that
a slow fungal colonization was observed on milled OP substrate during the
fermentation assay compared to OP pretreated with 1% NaOH. It was clearly shown
in Fig. 6a, that the optimal cellulolytic activities obtained on milled and
combined pretreatments were lower than the activity obtained on 1% NaOH
pretreated OP. However, an improvement of the FPase activity was
recorded when milling was combined with 1% NaOH.
It can also be noted, that protein contents (Fig. 6b) of milled OP
combined or not with alkaline pretreatment during the fermentation period were
much lower than those recorded in OP pretreated with 1% NaOH, which were
proportional to the cellulolytic activity.
As shown in Fig. 6c, pH evolution of milled OP during fermentation
followed the same trend as soluble proteins, with a remarkable drop in pH value
at the 12th day.
Effect
of thermal pretreatment on Fpase activity
Thermal pretreatments either
alone or in combination with 1% NaOH have negative effects on the cellulolytic
enzyme production (Fig. 7a). A very slow mycelial growth was observed during
the fermentation period on OP samples
Fig. 6: Evolution of PFase activity (a), protein content (b) and pH values (c) of extracts obtained from T. reesei culture on milled olive
pomace substrate during 30 days of fermentation
Vertical
bars indicate standard error of three replicates. OP, Olive pomace substrate;
DS, Dry substrate; FPase activity, Filter paper cellulase activity
Fig. 7: Evolution of FPase
activity (a), protein content (b) and pH values (c) of extracts obtained from T. reesei culture on thermal
pretreated olive pomace substrate during 30 days of fermentation
Vertical bars indicate
standard error of three replicates. OP, Olive pomace substrate; DS, Dry
substrate; FPase activity, Filter paper cellulase activity
thermally pretreated, combined
or not with 1% NaOH. The results of FPase activities showed that the
highest values were obtained on the 6th day of fermentation on
thermal pretreated OP alone or combined with 1% NaOH, but these values were
widely lower than the obtained activity with alkali pretreated OP.
As
illustrated in Fig. 7b, the soluble protein contents during the fermentation
period followed the same evolution as FPase activities. Soluble protein
contents recorded for both thermal and combined pretreatments were much lower
than those noted for alkaline pretreatment.
The pH evolution (Fig. 7c) showed stability during the fermentation period
maintaining the initial pH values. As expected, the pH values obtained for
combined pretreatment (temperature + 1% NaOH) were close to that of alkaline
pretreatment, but higher than the thermal pretreatment alone.
Discussion
This study revealed that the
alkaline pretreatment at different concentrations caused a significant decrease
in lipid content of the OP (Table 1) which can be explained by the
saponification process. The loss of carbohydrates is usually due to peeling and
hydrolytic reactions (Hendriks and Zeeman 2009).
In order to better demonstrate the effect of alkaline pretreatment on
fiber contents, the NDF, ADF, ADL, cellulose and hemicellulose contents were
calculated taking into account the dry matter of untreated olive pomace (Fig.
2). It is important to note that NDF comprises lignin, cellulose and
hemicellulose, ADF includes lignin and cellulose, while ADL represents only the
lignin content. Alkaline pretreatment at different concentrations had no effect
on NDF content. Unlike our findings, Pellera et al. (2016) found a significant increase of NDF content of OP
after alkaline pretreatment with different concentrations of NaOH (1–16%). It
can also be seen that ADF and cellulose contents decreased slightly but
significantly The decrease of ADF is mainly the consequence
of cellulose diminution after pretreatment.
The different pretreatments used in this study were designed to improve
cellulose production by increasing the accessibility of T. reesei RUT C30 fungi to the cellulosic fraction of the
substrate. Alkaline pretreatment is frequently used as a mean to modify the
originally complex and recalcitrant chemical structure of lignocellulosic
biomass (Yoon et al. 2014).
Bali et al. (2015), in a
comparison of various alkaline pretreatments, demonstrated that the highest
increase in cellulose accessibility was found with dilute NaOH solution (2%).
Moreover, El-Ghonemy et al. (2014)
found that pretreated substrates by 1% NaOH were much more efficient on
enzymatic hydrolysis compared to that treated by 4% NaOH. The same trend was
reported also by Rodríguez-Zúñiga et al.
(2015). In another study, Sun et al. (2008)
reported that the highest FPase activity
was found on alkali-treated rice straw compared to the untreated one using T. reesei Rut C-30 fungi. The
improvement of cellulolytic activity after pretreatment with 1% NaOH (Fig. 3a),
may be due to the swelling of the biomass that becomes more accessible for
enzymes after solvation and saponification reactions caused by the alkaline
treatment (Galbe and Zacchi 2007; Hendriks and Zeeman 2009). In fact, cellulose
can be swelled or dissolved in NaOH solutions, leading to the decrease of
lignocellulosic biomass crystallinity (Sun et
al. 2016). Besides, the improvement of enzyme activity after pretreatment
may be due to the fact that alkaline treatment did not make changes in the
fibers composition but, it depleted the medium of available carbon source
readily accessible for the strain used, like the lipids and carbohydrates. This
situation promoted the induction of cellulolytic enzyme synthesis by the
cellulose of the fermentation medium. Several authors have already affirmed
this suggestion. Ballerini (2006) explained that the production of cellulases
is regulated by the processes of induction and catabolic repression: induction
linked to the presence of substrates (in this case cellulose) and repression by
the sources of carbon molecules (such as glucose via catabolic repression mechanisms). Kaur et al. (2006) reported that it is difficult to deduce the nature of
the inducers that will be valid for the known cellulolytic microorganisms but
it is evident that the insoluble native cellulosic material is certainly the
best substrate for the production of cellulases. Candace and Weimer (1991)
suggested that close physical contact between fungus and cellulose can
stimulate induction and assumed that there is an appropriate sites recognized
in cellulose. It is also possible that this improvement of FPase activity after pretreatment with 1% NaOH is due to the nature
of the cellulose used by T. reesei RUT
C30. Since, there are two types of cellulose in OP composition. The cellulose
located in pulp of OP easily accessible for T.
reesei compared to that located in the OP stones (woody endocarp) which
contains, in addition to cellulose, high levels of lignin. Knowing that the
pure pulp of OP, contains about 20% only of the total crude cellulose as
reported by Sansoucy et al. (1984).
Besides the high amount of lignin in biomass, the enzymatic hydrolysis is also
controlled by lignin location in biomass and its surface area which play a
significant role (Kim et al.
2016). In addition, it was shown that alkali pretreatments are more effective
on agricultural residues than on woody materials (Kumar et al. 2009). In the other hand, Oke et al. (2016) observed
that the untreated mixed lignocellulosic substrates (MS) supported the highest
cellulase production, in comparison to MS treated with 1% NaOH. Similar results
were also found by Brijwani and Vadlani (2011) when soybean hulls were used as
substrate.
The pH diminution showed in Fig. 3c can be explained by NH4 cations
assimilation in the form of NH3 which induce H+ ions accumulation
(Roussos et al. 1983). This
diminution can be also related to the production of acidic metabolites that
neutralize the NaOH to give lower pH. It was found in this study that a higher
concentration of NaOH leads to a higher initial pH, and consequently, a high
residual NaOH content in the substrate pores, since an important characteristic
of alkali pretreatment is that the lignocellulosic biomass on itself absorbed
some of the alkali: approximately 3 g NaOH/100 g of total substrate (Hendriks
and Zeeman 2009). This affects the good growth of T. reesei strain and resulted in a slow mycelial growth on the
solid residue and explained the low celluolytic activities obtained for OP
pretreated with concentrations higher than 1% NaOH. Deswal et al. (2011) observed that increasing the initial pH of the medium
from 5.5 to 10.0 leads to a significant decrease in cellulases
production.
It appears that the milling pretreatment was more efficient on ADL
fraction than the combined one (Fig. 4). This contradiction can be explained by
the possible recondensation and reorganization of soluble lignin compounds,
solubilized during the alkaline pretreatment (Hendriks and
Zeeman 2009; Pellera et al. 2016). Rodhe et al. (2011) observed
about 75% loss of lignin on milled sorghum straw pretreated with 0.2 M of NaOH. The same fact was also found
by Haddadin et al. (1999) when milled
OP pretreated at 3%NaOH was used. After milling, the NDF and ADF diminutions
are mainly the consequence of ADL decrease. Mtui (2009) reported that ADL
decrease may be due to reduction of the degree of lignin depolymerization via the cleavage of uncondensed-aryl
ether linkages. After combined pretreatment in reference to untreated OP, a
significant increase in ADF and cellulose fractions was recorded, whereas NDF
and hemicellulose remained unchanged, only ADL fraction was decreased. Our
results are in agreement with the findings of Oke et al. (2016) who found that cellulose content of lignocellulosic
substrates increased after milling pretreatment combined with alkali (1% NaOH).
In contrast, Haddadin et al. (1999)
observed a significant decrease in cellulose content of milled OP pretreated at
3% NaOH, these findings may be the result of the NaOH high concentration,
compared to the dosage used in our research. The slow growth of T. reesei strain observed with milled OP
during the fermentation period may be probably due to the heterogenic
granulometry of milled sample, which contains particles with different sizes
(below 1.25 mm diameter). In fact, the fine particles formed after milling of
olive pomace in presence of high lipid contents (18.30%) causes a clogging that
prevents an effective transfer of oxygen in the culture medium, unlike larger
particles that improve breathing and aeration efficiency by increasing the
inter-particle space. Moreover, Sanchez (2009) thought that the mycelial
development is associated to the effective degradation of the lignocellulosic
biomass that constitutes its carbon source. In fact, the best solid substrate
should contain all the essential nutrients to the growing microorganism for
optimal function (Bansal et al. 2012; Gordillo-Fuenzalida et al. 2019).
Although milling resulted in a decrease in lignin content, cellulase
activity was not improved (Fig. 5a). According to Bali et al. (2015), lignin removal has been shown to increase the yield
of enzymatic hydrolysis, however, the direct effect of lignin removal on
cellulose accessibility is still not fully clear because lignin is also
associated with cellulases inhibition, and the relative contributions of these
two roles of lignin have not yet been fully defined. In addition, Berlin et al. (2006) stated that lignin
depolymerization has been considered as an effective inhibitor of cellulases.
The negative effect of milling process on cellulolytic activity may be due to
the formation of inhibitor compounds after degradation of lignocellulose during
pretreatment such as soluble phenolic compounds as reported by Vancov and
McIntosh (2011) and Sun et al.
(2016). In addition, Pellera et al. (2016)
reported that phenolic compounds are produced by the degradation of
lignocellulosic materials, and generally by hemicellulose and lignin
solubilization. The low PFAse activity
obtained on milled OP may also be due to the particle size of this substrate.
In fact, several authors concluded that the pore size of the substrate is in
relation to the size of the enzymes which constitute an important limiting
factor in the enzymatic hydrolysis of lignocellulosic biomass (Chandra et al. 2007; Alvira et al. 2010). Therefore, Grethlein (1985) found linear correlations
between the initial hydrolysis rate of pretreated biomass and the pore size
accessible to a molecule with a diameter of 51 A° similar to the size of T. reesei cellulase components.
Consequently, cellulase can get captivated in the pores of substrate if the
internal area is much superior than the external area which is the case for
many lignocellulosic substrates (Zhang and Lynd 2004).
Several authors reported that when milling is combined with alkaline
pretreatment an improvement in cellulolytic activity was noticed. Bansal et al. (2012) found an enhancement of
the cellulase production after alkali pretreatment (1% NaOH) using Aspergillus niger fungi cultured on
milled agricultural and kitchen wastes. Belal (2013) have also observed a
positive effect of combined pretreatment (milling + 5% NaOH) on conversion of
polysaccharide into sugar by T. reesei cellulases
on milled rice straw substrate. In another study, Wu et al. (2011) reported that milled bagasse pretreated with 2.5 M NaOH has given an enzymatic hydrolysis
yield of 98.7%.
The remarkable decrease in soluble protein content of milled OP (Fig. 5b)
was noted on the 12th day; this may be due to the assimilation of
the medium proteins during mycelium growth (Roussos et al. 1983). The thermal pretreatment of olive pomace did not
affect the ADL and cellulose fractions in comparison to the untreated one (Fig.
6). Aiello et al. (1996), in a study
on sugarcane bagasse substrate, found unchanged lignin and cellulose fractions
after combined pretreatment (100°C + 5% NaOH). The same finding was reported by
Rodríguez-Zúñiga et al. (2015) when
hydrothermal pretreatment (190°C, 10 min) was applied on sugarcane bagasse
substrate, in term of lignin fraction. Moreover, Xiao et al. (2017) reported that cellulose and lignin are unaffected by
the hydrothermal pretreatment. It seems that, this pretreatment has no effect
on lignin fraction, this can be due to the spatial re-localization or
reorganization of this later, which can occur with hydrothermal pretreatment
(Kristensen et al. 2008). In
addition, Agbor et al. (2011)
explained that not all pretreatments result in substantial
delignification: the structure of lignin may be altered without
extraction due to changes in the chemical properties of the lignin. Thermal
pretreatment combined with 1% NaOH caused a slight increase in hemicellulose
fraction compared to the untreated OP, whereas, theoretically, thermal
pretreatment causes a removal of hemicellulose of the solid fraction. In fact,
this increase of hemicellulose was the result of alkaline pretreatment only,
and as a consequence, thermal pretreatment had no effect on fiber fractions
because of the hardness of olive pomace and the mild temperature applied during
pretreatment. The study of Wu et al. (2011)
revealed that low thermal pretreatment (50°C) combined with 2.5 M NaOH significantly accelerated
the removal of hemicellulose and lignin on sorghum bagasse substrate. Further,
Hendriks and Zeeman (2009) reported that after thermal process, a certain part
of the hemicellulose is hydrolyzed and produce acids. This leads to conclude
that the improvement of the digestibility of lignocellulosic biomass depends on
the nature of the substrate and the operating conditions (pretreatments
combination). The low FPase activities
obtained after thermal pretreatment represented in Fig. 7a curve may possibly
be the result of changes in the chemical structure of olive pomace after
thermal pretreatment because the enzymatic hydrolysis of lignocellulosic
substrate could be influenced not only by the efficiency of the enzymes, but
also by the physical, chemical and morphological characteristics of these
biomass, as reported by Sun et al. (2016).
It could also be explained by the removal of certain nutrients from the
substrate after pretreatments (Abdullah et
al. 2016).
In our study, a slight production of cellulase was noted, while Aiello et al. (1996) found no detectable
activity during the fermentation using Trichoderma
reesei QM 9414 on sugarcane bagasse treated with alkali (5% NaOH at 100°C).
The authors suggested that the loss of activity could be the result of
absorption of cellulase on cellulose and lignin or the inhibition of enzymes by
glucose and cellobiose of the fermentation medium. This is an indication that
in some cases, pretreatment of substrate prior to cellulose production might
not be necessarily efficient, it could make a substrate less accessible and
less suitable for microbial growth and fermentation when compared with the
untreated one (Yoon et al. 2014).
Several studies confirmed this trend, when higher cellulolytic enzyme
production was obtained with untreated sugarcane bagasse (Rodríguez-Zúñiga et al. 2014; Ducom et al. 2019),
wheat straw (Sharma et al. 2015),
municipal solid wastes (Abdullah et al. 2016)
and mixed lignocellulosic substrates (Oke et
al. 2016) among others.
Gordillo-Fuenzalida et al. (2019),
confirm the ability of the Trichoderma
spp. to grow and produce cellulolytic enzymes in the presence of FMWs could
enable the onsite production of cellulase enzymes in lignocellulosic bioethanol
biorefineries. This would improve biorefinery economics by avoiding the
purchase of commercial enzymes, while improving their environmental
sustainability by utilizing waste materials, which goes in the line with our
finding.
Conclusion
In the present study, local
olive pomace (OP) was investigated as a substrate for solid-fermentation to
produce cellulases enzymes from Trichoderma reesei fungi. Three
different pretreatments and their combinations were applied on OP for the first
time to improve the cellulases production. Results showed that alkaline
pretreatment with 1% NaOH improved significantly the enzyme production from OP
substrate. The other pretreatments (milling, thermal and their combinations)
showed a negative effect on the improvement of cellulases production, besides
being expensive and require high energy consumption. Therefore, olive pomace
could be considered as a valuable substrate for fermentations process,
especially for cellulases production given its chemical composition and fiber
content.
Acknowledgments
The authors are grateful to
the Algerian Ministry of Higher Education and Scientific Research and the
National Centre for Biotechnology Research for their respective financial and
material supports. The authors wish also to thank Mr Ali Boutiche, from England
and Dr. Djouher Gaad for providing English language help. Our sincere thanks to
Miss Bennacer amel. Her relevant and effective contribution in the revision of
the English language and the structuring of the publication.
Author Contributions
Malika Boutiche: Conceptualization, Experimental work (enzymatic
activity), Writing original draft, Writing review and editing. Fatma Sahir-Halouane:
Supervision, Writing review and editing. Leila Meziant: Experimental work (enzymatic
activity), Statistical analysis, Writing original draft. Fairouz Saci: Writing
original draft. Kahina Oudjedi: Experimental work (pretreatments + enzymatic
activity). Mouna Derdour: Experimental work (microbiology + physicochemical analysis).
Karima Ouffroukh: Experimentral work (physicochemical analysis). Ibtissem
Maghboune: Experimental work (physicochemical analysis). Samah Fiala:
Experimental work (physicochemical analysis). Amel Bekrar: Experimental work
(physicochemical analysis).
Conflicts of Interest
The authors decrare that they have no conflicting interests.
Data Availability
Data related to this article are not confidential and could be supplied on
request to the authors.
Ethics Approval
Not applicable in this study.
References
Abdullah JJ, D Greetham, N
Pensupa, A Gregory, GA Tucker, C Du (2016). Optimizing cellulase production
from municipal solid waste (MSW) using solid state fermentation (SSF). J Fund Renew Ener Appl 6:3
Aftab A, P Vermette (2008).
Culture-based strategies to enhance cellulose enzyme production from Trichoderma reesei RUT C30 in bioreactor
culture conditions. Biochem Eng J
40:399‒407
Agbor VB, N Cicek, R Sparling, A
Berlin, DB Levin (2011). Biomass pretreatment: Fundamentals toward application.
Biotechnol Adv 29:675‒685
Aiello C, A Ferrer, A Ledesma
(1996). Effect of alkaline treatments at various temperatures on cellulase and
biomass production using submerged sugarcane bagasse fermentation with Trichoderma reesei QM 9414. Bioresour Technol 57:13‒18
Aliakbarian B, AA Casazza, P
Perego (2011). Valorization of olive oil solid waste using high pressure–high
temperature reactor. Food Chem
128:704‒710
Alvira P, E Tomás-Pejó, M
Ballesteros, MJ Negro (2010). Pretreatment technologies for an efficient
bioethanol production process based on enzymatic hydrolysis. Bioresour Technol 101:4851‒4861
AOAC Official Methods of
Analysis (1990). Method 945, 18‒B. In: Association of Official
Analytical Chemists, 15th edn. Arlington, Virginia, USA.
Aranda E, I Garcia-Romera, JA
Ocampo, V Carbone, A Mari, A Malorni, F Sannino, A De Martino, R Capasso
(2007). Chemical characterization and effects on Lepidium sativum of the native and bioremediated components of dry
olive mill residue. Chemosphere 69:229‒239
Bali G, X Meng, JI Deneff, Q
Sun, AJ Ragauskas (2015). The effect of alkaline pretreatment methods on
cellulose structure and accessibility. Chem Sustain Chem 8:275‒279
Ballerini D (2006). Les Biocarburants: Etat des Lieux,
Perspectives Etenjeux du Développement, Editions, Technip, Paris, France
Bansal N, R Tewari, R Soni, SK Soni (2012). Production
of cellulases from Aspergillus niger NS-2
in solid-state fermentation on agricultural and kitchen waste residues. Waste Manage 32:1341‒1346
Belal EB (2013). Bioethanol
production from rice straw residues. Braz
J Microbiol 44:225‒234
Berlin A, M Balakshin, N Gilkes,
J Kadla, V Maximenko, S Kubo, J Saddler (2006). Inhibition of cellulase,
xylanase and β-glucosidase
activities by soft wood lignin preparations. J Biotechnol 125:198‒209
Bradford MM (1976). A rapid
and sensitive method for the quantification of microgram quantities of protein
utilizing the principle of protein-dye binding. Anal Biochem 72:248‒254
Brijwani K, PV Vadlani (2011).
Cellulolytic enzymes production via
solid-state fermentation: Effect of pretreatment methods on physicochemical
characteristics of substrate. Enz Res
2011:1‒10
Brodeur G, E Yau, K Badal, J
Collier, KB Ramachandran, S Ramakrishnan (2011). Chemical and physicochemical
pretreatments of lignocellulosic biomass: A review. Enz Res 2011:1-17
Candace HH, PJ Weimer (1991). Biosynthesis and Biodegradation of Cellulose,
Edition. Marcel Dekker, New York, USA
Cara C, I Romero, JM Oliva, F
Sáez, E Castro (2007). Liquid Hot Water Pretreatment of Olive Tree Pruning
Residues. In: Applied Biochemistry and
Biotecnology, ABAB Symposium, pp:136‒140. Mielenz JR, KT Klasson, WS
Adney, JD McMillan (Eds). Humana Press, New York, USA
Chandra RP, R Bura, WE Mabee, A
Berlin, X Pan, JN Saddler (2007). Substrate pretreatment: The key to effective
enzymatic hydrolysis of lignocellulosics? Adv
Biochem Eng Biotechnol 108:67‒93
Coimbra MA, SM Cardoso, JALD
Silva (2010). Olive Pomace, a Source for Valuable Arabinan-Rich Pectic
Polysaccharides. In: Carbohydrates in
Sustainable Development I. Topics in Current Chemistry. pp:129‒141. Rauter A, P Vogel, Y
Queneau (Eds). Springer, Berlin, Heidelberg, Germany
Deswal D, YP Khasa, RC Kuhad (2011).
Optimization of cellulase production by a brown rot fungus Fomitopsis spp. RCK2010 under
solid state fermentation. Bioresour
Technol 102:6065‒6072
Dhillon GS, HS Oberoi, S Kaur, S
Bansal, SK Brar (2011). Value-addition of agricultural wastes for augmented
cellulase and xylanase production through solid-state tray 632 fermentation
employing mixed-culture of fungi. Ind
Crops Prod 34:1160‒1167
Dubois M, KA Gilles, TK
Hamilton, PA Rebers, E Smith (1956). Colorimetric method for determination of
sugar and related substances. Anal Chem 28:350‒356
Ducom G, M Gautier, M Pietraccini, JP Tagutchou, D
Lebouil, N Dumont, R Gourdon (2019). Caractérisation de grignons d’olives en
vue d’une valorisation thermochimique 637 par gazéification. Déch Sci Tech 82:41‒54
El-Ghonemy DHI, TH Ali, ME
Moharam (2014). Optimization of culture conditions 639 for the production of
extracellular cellulases via solid
state fermentation. Braz Microbiol Res
J 4:698‒714
Erdocia X, E Ruiz, I Romero, MJ
Diaz, E Castro, J Labidi (2017). Lignin characterization from two different
pretreatments in bioethanol production processes from olive tree pruning. Chem Eng Trans 61:421‒426
Fernández-Bolaños J, B Felizon,
A Heredia, R Rodriguez, R Guillen, A Jimenez (2001). Steam-explosion of olive
stones: Hemicellulose solubilization and enhancement of enzymatic 64 hydrolysis
of cellulose. Bioresour Technol 79:53‒61
Fonseca LM, LS Parreiras, MT
Murakami (2020). Rational engineering of the Trichoderma reesei RUT-C30 strain into an industrially relevant platform
for cellulase production. Biotechnol Biof
13:93–107
Galbe M, G Zacch (2007). Pretreatment
of lignocellulosic materials for efficient bioethanol production. Adv Biochem Eng Biotechnol 108:41‒65
Gandla ML, C Martin, LJ Jonsson
(2018). Analytical enzymatic saccharification of lignocellulosic biomass for
conversion to biofuel and bio-based chemicals. Energies 11:2936–2955
Gordillo-Fuenzalida F, A Echeverria-Vega, S
Cuadros-Orellana, C Faundez, T Kähne, R Morales-Vera (2019). Cellulases production by a Trichoderma spp. using food manufacturing wastes.
Appl Sci 9:4419–4420
Grethlein HE (1985). The effect
of pore size distribution on the rate of enzymatic hydrolysis of cellulose
substrates. Biol Technol 3:155‒160
Gudynaite-Savitch L, TC White
(2016). Fungal biotechnology for industrial enzyme production: Focus on (hemi)
cellulose production strategies, advances and challenges. In: Gene Expression Systems in Fungi: Advancements and Applications,
Fungal Biology, pp:395‒439. Schmoll M, C, Dattenböck (Eds). Springer, Switzerland
Haddadin MS, J Haddadin, OI
Arabiyat, B Hattar (2009). Biological conversion of olive pomace into compost
by using Trichoderma harzianum and Phanerochaete chrysosporium. Bioresour Technol
100:4773‒4782
Haddadin MS, SM Abdulrahim, GY
Al-Khawaldeh, RK Robinson (1999). Solid-state fermentation of waste pomace from
olive processing. J Chem Technol
Biotechnol Intl Res Process Environ
Clean Technol 74:613‒618
Hendriks ATWM, G Zeeman (2009).
Pretreatments to enhance the digestibility of lignocellulosic biomass. Bioresour Technol 100:10‒18
Hinterdobler W, G Li, K Spiege,
S Basyouni-Khamis, M Gorfer, M Schmoll (2021). Trichoderma reesei isolated from Austrian soil with high potential
for biotechnological 670 application. Front
Microbiol 12:552301
Imran M, Z Anwar, M Irshad, MJ
Asad, H Ashfaq (2016). Cellulase production from species of fungi and bacteria
from agricultural wastes and its utilization industry. Adv Enz Res 4:44‒55
Jun H, T Kieselbach, LJ Jönsson
(2011). Enzyme production by filamentous fungi: Analysis of the secretome of Trichoderma reesei grown on
unconventional carbon source. Microb Cell
Fact 10:1–10
Kaur J, SC Bhupinder, SS
Harvinder (2006). Regulation of cellulase production in two thermophilic fungi Melanocarpus spp. MTCC 3922 and Scytalidium thermophilum MTCC 4520. Enz Microb Technol 38:931‒936
Kim JS, YY Lee, TH Kim (2016). A
review on alkaline pretreatment technology for bioconversion of lignocellulosic
biomass. Bioresour Technol 199:42‒48
Kristensen JB, LG Thygesen, C
Felby, H Jørgensen, T Elder (2008). Cell–wall structural changes in wheat straw
pretreated for bioethanol production. Biotechnol
Biof 1:1–9
Kumar AK, S Sharma (2017).
Recent updates on different methods of pretreatment of lignocellulosic
feedstocks: A review. Bioresour Bioproc
4:7
Kumar P, DM Barrett, MJ
Delwiche, P Stroeve (2009). Methods for pretreatment of lignocellulosic biomass
for efficient hydrolysis and biofuel production. Ind Eng Chem Res 48:3713‒3729
Leite P, JM Salgado, A Venâncio,
JM Domínguez, I Belo (2016). Ultrasounds pretreatment of olive pomace to
improve xylanase and cellulase production by solid-state fermentation. Bioresour Technol 214:737‒746
Linke R, GT Gerhard, T Haarmann,
J Eidner, M Schreiter, P Lorenz (2015). Restoration of female fertility in Trichoderma reesei QM6a provides the
basis for inbreeding in this industrial cellulose producing fungus. Biotechnol Biofuels 8:155
López-Linares JC, I Romero, M
Moya, C Cara, E Ruiz, E Castro (2013). Pretreatment of olive tree biomass with
FeCl3 prior enzymatic hydrolysis. Bioresour
Technol 128:180‒187
Menon V, M Rao (2012). Trends in
bioconversion of lignocellulose: Biofuels, platform chemicals & biorefinery
concept. Progr Ener Combust Sci 38:522‒550
Miller GH (1959). Use of
dinitrosalicylic acid reagent for determination of reducing sugar. Anal Chem 31:426‒429
Mirabella N, V Castellani, S
Sala (2014). Current options for the valorization of food manufacturing waste. J Clean Prod 65:28‒41
Moftah OAS, S Grbavčić, M Žuža, N
Luković, D Bezbradica, Z Knežević-Jugović (2012). Adding value
to the oil cake as a waste from oil processing industry: Production of lipase
and protease by Candida utilis in solid state fermentation. Appl Biochem Biotechnol 166:348‒364
Mosier N, C Wyman, B Dale, R
Elander, YY Lee, M Holtzapple, M Ladisch (2005). Features of promising
technologies for pretreatment of lignocellulosic biomass. Bioresour Technol 96:673‒686
Mtui GYS (2009). Recent advances
in pretreatment of lignocellulosic wastes and production of value added
products. Afr J Biotechnol 8:1398‒1415
National Research Council (1999).
Committee on Biobased Industrial Products. Biobased Industrial
Products-Priorities for Research and Commercialization. National
Academy Press, Washington DC, USA
Nefzaoui A (1991). Valorisation des sous-produits de
l’olivier. In: Fourrages et Sous-produits
Méditerranéens, pp:101‒108.
Tisserant JL, X Alibés (Eds). CIHEAM Options Méditerranéennes, Zaragoza
Neifar M, A Jaouani, A Ayari, O
Abid, HB Salem, A Boudabous (2013). Improving the nutritive value of olive cake
by solid-state cultivation of the medicinal mushroom Fomes fomentarius. Chemosphere
91:110‒114
Oke MA, MM Ishola, MJ
Taherzadeh, MSM Annuard, K Simarani (2016). Effects of pretreatments of single
and mixed lignocellulosic substrates on production of endoglucanase by Bacillus aerius S5-2. Bioresources 11:6708‒6726
Ouyang X, L Chen, S Zhang, Q
Yuan, W Wang, LJ Linhardt (2018). Effect of simultaneous steam explosion and
alkaline depolymerization on Corncob lignin and cellulose structure. Chem Biochem Eng Quart 32:177‒189
Paloheimo M, T Haarmann, S
Mäkinen, J Vehmaanperä (2016). Production of industrial enzymes in Trichoderma reesei. In: Gene Expression
Systems in Fungi: Advancements and Applications, Fungal Biology, pp:23‒58. Schmoll M, C Dattenböck
(Eds). Springer International Publishing, Cham, Switzerland
Papaioannou EH, SI Patsios, AJ
Karabelas, NA Philippopoulos (2013). Characterization of condensates from an
indirect olive oil pomace drying process: The effect of drying temperature. J Environ Chem Eng 1:831‒837
Pellera FM, S Santori, R Pomi, A Polettini, E Gidarakos
(2016). Effect of alkaline pretreatment on anaerobic
digestion of olive mill solid waste. Waste
Manage 58:160‒168
Pirota RDPB, PS Delabona, CS
Farinas (2014). Simplification of the biomass to ethanol conversion process by
using the whole medium of filamentous fungi cultivated under solid-state
fermentation. Bioener Res 7:744‒752
Ravindran R, AK Jaiswal (2016).
A comprehensive review on pre-treatment strategy for lignocellulosic food
industry waste: Challenges and opportunities. Bioresour Technol 199:92‒102
Ray RC, SS Behera (2017).
Biotechnology of Microbial Enzymes. In:
Solid-state Fermentation for Production of Microbial Cellulases, pp:45‒55. Goutam B, AL Demain, JL
Adrio (Eds). Academic Press, New York, USA
Rives O (2021). Consommation et production de l'huile
d'olive en Algérie. In: Programmed’Appui
au Secteur de l’Agriculture, p:39. Pasa Ue (Ed.), Algérie
Rodhe AV, B Venkateswarlu, L
Sateesh, J Sridevi, LV Rao (2011). Enzymatic 798 hydrolysis of sorghum straw
using native cellulose produced by T.
reesei NCIM 992 under solid 799 state fermentation using rice straw. 3 Biotech 1:207‒215
Rodríguez-Zúñiga UF, D Cannella, RDC Giordano, RDLC
Giordano, H Jørgensen, C Felby (2015). Lignocellulose pretreatment
technologies affect the level of 749 enzymatic cellulose oxidation by LPMO. Green Chem 16:2896‒2903
Rodríguez-Zúñiga UF, VB Neto, S
Couri, S Crestana, CS Farinas (2014). Use of spectroscopic and imaging
techniques to evaluate pretreated sugarcane bagasse as a substrate 752 for
cellulase production under solid-state fermentation. Appl Biochem Biotechnol 172:2348‒2362
Roig A, ML Cayuela, MA
Sánchez-Monedero (2006). An overview on olive mill wastes and their valorization
methods. Waste Manage
26:960‒969
Roussos S, I Perraud-Gaime, H Lakhtar, F Aouidi, Y
Labrousse, N Belkacem, H Macarie, J Artaud (2009). Valorisation
biotechnologique des sous-produits de l'olivier par Fermentation en Milieu
Solide. Olivebioteq 26:52–59
Roussos SG, GL Garcia, M Raimbault (1983). Valorisation
de la cossette de betterave par culture de Trichoderma
harzianum en milieu liquide et solide. Industr
Aliment Agric 100:449–452
Sanchez C (2009).
Lignocellulosic residues: Biodegradation and bioconversion by fungi. Biotechnol Adv 27:185‒194
Sansoucy R, X Alibes, F Martilotti, A Nefzaoui, P
Zoïopoulos (1984). Utilisation des sous-produits de l’olivier en alimentation
animale dans le bassin méditerranéen. Etude FAO Prod Anim 121:23–43
Sharma B, R Agrawal, RR
Singhania, A Satlewal, A Mathur, D Tuli, M Adsul (2015). Untreated wheat straw:
Potential source for diverse cellulolytic enzyme secretion by Penicillium janthinellum EMS-UV-8
mutant. Bioresour Technol 196:518‒524
Silveira MHL, RS Aguiar, M
Siika-Aho, LP Ramos (2014). Assessment of the enzymatic hydrolysis profile of
cellulosic substrates based on reducing sugar release. Bioresour Technol 151:392‒396
Soccol CR, ESFD Costa, LAJ
Letti, SG Karp, AL Woiciechowski, LPDS Vandenberghe (2017). Recent developments
and innovations in solid state fermentation. Biotechnol Res Innov 1:52‒71
Soest PJV, JB Robertson (1979). Systems
of analysis for evaluating fibrous feeds. Cornell University Press. New
York, USA
Srivastava N, M Srivastava, PK
Mishra, VK Gupta, S Rodriguez-Couto, A Manikanta, PW Ramteke, G Molina (2018).
Applications of fungal cellulases in biofuel production: Advances and
limitations. Renew Sustain Ener Rev
82:2379‒2386
Stamatelatou K, PS Blika, I
Ntaikou, G Lyberatos (2012). Integrated Management Methods for the Treatment
and/or Valorization of Olive Mill Wastes. In:
Novel Technologies in Food Science. Integrating Food Science and Engineering
Knowledge into the Food Chain, pp:65‒118. McElhatton A, P Sobral
(Eds). Springer, New York USA
Sun S, S Sun, X Cao, R Sun
(2016). The role of pretreatment in improving the enzymatic hydrolysis of
lignocellulosic materials. Bioresour
Technol 199:49‒58
Sun WC, CH Cheng, WC Lee (2008).
Protein expression and enzymatic activity of cellulases produced by Trichoderma reesei Rut C-30 on rice
straw. Proc Biochem 43:1083‒1087
Ugwuanyi JO, B McNeil, LM Harvey
(2009). Production of protein-enriched feed using agro-industrial residues as
substrates. In: Biotechnology for
Agro-Industrial Residues Utilization, pp:77‒103. Nigam PSN, A Pandey (Eds).
Springer, Dordrecht, Netherlands
Vancov T, S McIntosh (2011).
Alkali pretreatment of cereal crop residues for second-generation biofuels. Ener Fuels 25:2754‒2763
Vasco-Correa J, X Ge, Y Li
(2016). Biological Pretreatment of Lignocellulosic Biomass. In: Biomass Fractionation Technologies for a
Lignocellulosic Feedstock Based Biorefinery, pp:561‒585. Mussatto SI (Ed). Elsevier
Amsterdam, Netherlands
Wu L, M Arakane, M Ike, M Wada,
T Takai, K Tokuyasu, M Gau, K Tokuyasu (2011). Low temperature alkali
pretreatment for improving enzymatic digestibility of sweet sorghum 802 bagasse
for ethanol production. Bioresour Technol
102:4793‒4799
Xiao LP, GY Song, RC
Sun (2017). Effect of Hydrothermal Processing on 804 Hemicellulose Structure. In: Hydrothermal Processing in
Biorefineries, pp:45‒94. Ruiz HA, MH Thomsen, HL Trajano (Eds.). Springer International Publishing,
Cham, Switzerland
Yoon LW, TN Ang, GC Ngoh, ASM
Chua (2014). Fungal solid-state fermentation and various methods of enhancement
in cellulase production. Biomass Bioener
67:319‒338
Zhang HY, LR Lynd (2004). Toward
an aggregated understanding of enzymatic hydrolysis of cellulose: Noncomplexed
cellulase systems. Biotechnol Bioeng
88:779‒797
Zheng Y, Z Pan, R Zhang (2009).
Overview of biomass pretreatment for cellulosic ethanol production. Intl J Agric Biol Eng 2:51