Saccharomyces strains were used widely and traditionally for industrial
ethanol production because of its ability to produce high concentrations of
ethanol from hexoses and its high tolerance to ethanol and other inhibitory
compounds. However, S. cerevisiae is naturally unable to metabolize pentoses,
such as xylose, arabinose and also polysaccharides (starch). Fribous (lignocellulosic)
is the second major source of carbohydrates in hard woods and vegetable biomass,
so its fermentation is essential for the economic conversion of starch to ethanol,
which may provide an ideal alternative fuel source in the future (Lin
and Tanaka, 2006). The use of renewable resources, such as lignocellulosic
biomass, to produce ethanol offers several environmental benefits and averts
competition (Kumar et al., 2009; Lin
and Tanaka, 2006).
Many microorganisms, including Saccharomyces cerevisiae, are not able
to degrade starch and they do not produce starch decomposing enzymes as amylase,
pullulanase or isoamylase and glucoamylase (Gupta et
al., 2003). The ethanol-fermenting microorganisms, such as S. cerevisiae
are lack of amylolytic enzymes and unable to directly convert the starch into
ethanol (Ang et al., 2001).
Considering that the main potential feedstock (biomass) for producing bioethanol
is composed of carbohydrates, which include starch, cellulose and hemicelluloses,
among others, the use of enzymes to break down these oligosaccharides into easily
fermentable sugars is a requirement, previous to conducting the fermentation
(Kim and Dale, 2004).
It is necessary either to use starch enzymes producing strains in order to
get free carbohydrates monomers for using as carbon source (Altintas
et al., 2003; Gupta et al., 2003; Nakamura et al., 2002).
Starch fermentations with S. cerevisiae transformed with amylase and
glucoamylase genes showed ethanol productivity similar to that observed when
starch decomposing enzymes were added to the medium (Ulgen
et al., 2002; Eksteen et al., 2003;
Kang et al., 2003; Shigechi
et al., 2004). Starch is converted into ethanol in starch plants
where the raw material, is milled and then treated with a combination of heat
and enzymes without prior separation of its constituents (Bothast
and Schlicher, 2005). Industrial processes such as starch liquefaction demands
the process to be carried out at high temperature so economical application
of amylase to such process, its thermostability is of a prime importance (Joshi,
Several agricultural wastes have been tested for their bioethanol-producing
potential. In the present study, the utilization of some agricultural residues
(like mango residues) containing carbohydrates for the production of bioethanol
was evaluated. Also, In Burkina Faso, enormous quantities of mango are lost
per year and contribute to increase the environmental rate residues. In order
to valorize these residues, it was important to research the possible ways.
The aim of this study was to improve the bioethanol production by silmultaneous
fermentation of yeasts using amylasic properties from Bacillus licheniformis
in the way to valorize mango biomass.
MATERIALS AND METHODS
Collection and processing of samples: Mango residues (70 samples as 25 kg) were collected from waste dumping sites in the principals mango production areas. Three regions (Banfora, Houet, Orodara) and peripheral area of Ouagadougou (Capital of Burkina Faso) were concerned to the sampling. The sampling was done during, April to May, 2010.
Isolation and selection of microorganisms: The selection was carried out on a total of 8 yeasts strains. The strains codified as A1 to A4 were S. cerevisiae strains isolated from wine cultures. Also the strains codified as S1 to S4 the Baker's yeast microorganism commonly used in local drinking beer (dolo).
The yeasts strains were isolated in maintenance medium (used in agar plates) contained 20 g of glucose, 20 g of agar, 5 g of peptone, 5 g of MgSO4•7H2O per liter. In the way to select the strains having best growth; the in the liquid inoculation contained (growth medium) 50 g glucose, 5 g of yeast extract, 1 g of KH2PO4, 0.3 g of NH4Cl and 2 g of MgSO4•7H2O per liter has been utilized. The fermentation medium contained 0.05 M citrate buffer pH 4.8; l,5 g peptone, 5 g of yeast extract, 1 g of KH2PO4, 0.3 g of NH4Cl and 2 g of MgSO4•7H2O per liter. It was used to select the best strains producing alcohol.
Carbohydrate (sugar) fermentation: The ability of the yeasts to ferment
various carbohydrates using glucose, fructose, sucrose, maltose, lactose and
arabinose was determined by growing the isolate in liquid standard medium containing
1% (w/v) of the particular carbohydrate. Durham-tubes were inverted into the
culture tubes for gas collection. The incubation was at 30°C for 24 h and
uninoculated broths were used as control. The standard medium used for fermentation
was it recommended by Konlani et al. (1996).
Bacillus licheniformis isolation: Bacillus licheniformis has been isolated from and characterized in laboratory (CRSBAN).
Also it was performed to improve amylolitic property using different physiologicals conditions (Temperature, pH, percentage of starch). This process is driven to select the Bacillus licheniformis strains having a best enzymatic hydrolysis activity.
Bioethanol production: Methods used for production of bioethanol include enzymatic hydrolysis, fermentation and fractional distillation.
Enzymatic hydrolysis: An optimization of temperature and enzyme activity
was performed. The performance during starch hydrolysis was evaluated based
on the reducers sugars production and the liquefaction yield for the substrate
(peel mango) (Miller, 1959).
One hundred grams of mango peel was weighed into seven 1 L conical flasks and 2 mL of solution containing amylase from B. licheniformis was added to each conical flask.
The flasks were covered with cotton wool, wrapped in aluminum foil, heated for 2 h in a water bath and then autoclaved for 30 min at 121°C. The Flasks were allowed to cool, filtered trough Whatman filter paper and the pH was adjusted to 4.5 with acetate tampon medium.
The amylasic hydrolysis activity of B. licheniformis was setting in evidence by following of kinetic of reducers sugars released through carbohydrates hydrolysis during 5 h.
Fermentation: The fermentation was carried out along with saccharification
(Simultaneous Saccharification and Fermentation (SSF)), as described by Kroumov
et al. (2006) and Oghgren et al. (2006).
The flasks containing the hydrolyzed samples were covered with cotton wool,
wrapped in aluminium foil, autoclaved for 15 min at 121°C and allowed to
cool at room temperature. Yeasts performing strains A1 and S3 were each aseptically
inoculated into each flask and incubated at 30°C. Two flasks of each sample
(containing mango peel) were removed after every 24 h, up to 7 days.
Fractional distillation: The fermented broth was dispensed into round-bottom flasks fixed to a distillation column enclosed in running tap water. A conical flask was fixed to the other end of the distillation column to collect the distillate. A heating mantle with the temperature adjusted to 78°C was used to heat the round-bottomed flask containing the fermented broth.
Determination of quantity of ethanol produced: The distillate collected
over a slow heat at 78°C was measured using a measuring cylinder and expressed
as the quantity of ethanol produced in g L-1 by multiplying the volume
of distillate collected at 78°C by the density of ethanol (0.8033 g mL-1).
The g L-1 is equivalent to the yield of 100 g of dried substrate
(Humphrey and Okafoagu, 2007).
Determination of percentage ethanol: A standard ethanol density curve was prepared by taking series of percentage (v/v) ethanol solutions, which were prepared in volumetric flasks and the weight was measured. The density for each of the prepared ethanol solutions was calculated and a standard curve of density against percentage ethanol was plotted. The percentage ethanol concentration of ethanol produced was obtained by comparing its density with the standard ethanol density curve.
Selection of performed yeasts: Among 8 yeasts coming in different biotope, only two yeasts (S3, A1) were retained for ethanol fermentation process.
Peel saccharification optimization: The amylasic activity of B. licheniformis show the kinetic of reducing sugars released as shown in Fig. 1. The optimal rate of reducing sugars attains 78% (g g-1).
Ethanol production from peel mango sugar performed by simultaneous saccharification and fermentation (SSF): The kinetic of ethanol production by two yeasts strains (S3; A1) is shown in Fig. 2.
||Following of reducers sugars percentage releasing through
carbohydrates hydrolysis on peel mango, incubated with α-amylase of
||Ethanol produced (g L-1) from mango peel using
yeasts strains S3 and A1 separately and each with amylase presence simultaneously
Selection of performed yeasts: The yeasts strains S3, A1 were selected according to their specific growth performance, that varies respectively at 0.31 to 0.24. And yet it can be noticed that specific growth parameters must not be directly link or attach to the ethanol high production.
Peel saccharification optimization: On mango peel containing 22.62 %
(g g-1) of carbohydrates (polysaccharides), the hydrolysis was conducted
up to 4 hours and the results presented in Fig. 1, shows that
releasing of reducing sugars increased with enzymatic activity of B. licheniformis.
This result is demonstrated while varying the volume of the inoculum (contained
the strains). Likewise it is noticed that there has not significant change or
variation of reducers sugars rate without enzyme presence. Kim
and Hamdy (1985) showed that the same trend was observed when the hydrolysis
of potato was studied.
So liquefaction is a preliminary step for saccharification, by which large quantities of D-glucose can be produced from inexpensive sources.
The liquefaction conduct at middle temperature (55°C) result a higher yield
of reducing sugars released (78% g g-1), which might be related to
the type of enzyme utilized. This result was greater than that obtained by Lazic
et al. (2004) in the two-step enzyme hydrolysis (61 %).
Thus, fermentescibles sugars percentage obtained (78% g g-1) by using B. licheniformis enzymatic activity is relatively very higher than at which found in Bacillus absence.
This value is slightly superior to which reported by Somda
et al. (2010) using other species of Bacillus on peel mango
(62% g g-1). It can be explained that the enzyme has the capacity
of decomposing into hexose, all polysaccharides which are built up of glucose
residues united by α-1, 4 glycosidic bonds and also it is thermostable.
And yet the incomplete utilization of polysaccharides (starch) by enzymes may
due to lack of enough oxygen or feedback inhibition of amylase activity by glucose
released, as reported by Abouzied and Reddy (1986).
Others authors have explained mechanism of polysaccharides hydrolysis with some
Lagzouli et al. (2007) showed that the production
of glucoamylase in presence of starch and of glucose suggests that the glucoamylase
produced by Candida guilliermondii is an enzyme greatly led by the starch.
And that its activity is probably under the effect of a glucose repression catabolic.
The same phenomenon of repression catabolic has been observed at Clostridium
thermohydrosulfuricum (Hyun and Zeikus, 1985) and
Bacillus sp. (Kiran et al., 2005).
As B. licheniformis expressing both α-amylase and glucoamylase activities. Therefore this potentiality Bacillus could be used to fermentation process in medium containing polysaccharides substrates like mango peel.
Kinetic of amylasic hydrolysis of peel starch and simultaneous fermentation to ethanol: The hydrolyzate obtained by amylase of B. licheniformis was used as substrates for bioethanol production respectively by yeasts strains A1 and S3. This effect is shown in Fig. 2. Peel mango was used to produce ethanol through enzymatic hydrolysis and SSF with respectively two yeasts strains (S3, A1).
In SSF, the two different microorganisms behaved differently, according to their nutrient requirements, but synergistically in the degradation of organic substrate. An enzyme (carbohydrate hydrolases produced by Bacillus licheniformis) was able to hydrolyze peel mango. The saccharification products were simultaneously utilized by yeasts strains (S3, A1) for ethanol production.
These yeasts are able to produced ethanol due to the presence of Pyruvate Decarboxylase
(PDC) and alcohol dehydrogenase (ADH), which are key enzymes in ethanol formation,
as reported by Gunasekaran and Chandra (2007). Figure
2 shows that the maximum volume of ethanol (16 g L-1) produced
from peel mango by S3 and B1 (14.4 g L-1) in this study at the 120th
h is lower than the results found by Agulejika et al.
(2005) who also reported maximum ethanol yield at 120th hour from fresh
fruit (64.01 g L-1) and waste fruits (21.14 g L-1) using
Z. mobilis. The higher ethanol yield from fresh fruit was due
to higher presence of fructose and glucose in fresh fruits, as stated by Micheal
and Rosaline (2000). The maximum volume of ethanol produced from peel mango
is lower than the 59 g L-1 reported by Gunasekaran
and Chandra (2007) at 120th h from cassava starch hydrolysate. This is due
to cassava containing more carbohydrates, which could be fermented to ethanol.
Thus, Sree et al. (2000) reported about the ethanol
production by SSF of wheat products using Saccharomyces cerevisiae. Those authors
were able to produce up to 44.2 g-ethanol L-1 when, fine wheat flour
was used as substrate and 34.1 g L-1 using damaged wheat flour. Present
results are in agreement with those found previously by these authors quoted.
Also, Tasic et al. (2008) have conducted fermentation
on potato tuber mash and found during 18 to 33 h of incubation, an ethanol concentration
growing at 31.2 to 32.9 g L-1.
Ethanol produced from treated sample (bitter kola, pulp agrowaste) by Humphrey
and Okafoagu (2007) was 11.2 g L-1 at 96 h and 12.9 g L-1
at 216 h. These values were lower than our values.
The decline and stabilization in ethanol noticed at some stages may be due
to the inhibitory effect of ethanol on growth and transport metabolism of the
yeast (D'Amarc and Stewart, 1987; Xu
et al., 1996).
Present results revealed that ethanol could be produced from agricultural residues, such as peel mango using performed yeasts strains (S3, A1) as fermenting organisms.
Considering the cost-effectiveness, in addition to being a means to control environmental pollution, the use of peel mango for ethanol production is concluded as a worthwhile venture.
Present results demonstrate that the simultaneous saccharification and fermentation of mango peel carbohydrates with B. licheniformis and yeasts (S3 and A1) is sufficient to increase bioethanol production. This led us to suggest that it would be interesting to use this carbohydrates fermentation process in repeated fed-batch or continuous fermentation for further improvements of bioethanol production.
Finally, studies using B. licheniformis expressing α-amylase are promising and will lead to valuable applications in ethanol production from renewable feedstocks and the conversion of mango peel into ethanol.
This research was supported by IFS (International Foundation of Science) and ISP/IPICS (International Sciences Programm/International Programm in the Chemicals Sciences).