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. 2021 Jan 3;11(1):21. doi: 10.1007/s13205-020-02595-6

Co-culture of Saccharomyces cerevisiae (VS3) and Pichia stipitis (NCIM 3498) enhances bioethanol yield from concentrated Prosopis juliflora hydrolysate

Shaik Naseeruddin 1, Suseelendra Desai 2,✉, L Venkateswar Rao 1
PMCID: PMC7779385  PMID: 33442519

Abstract

Biphasic acid hydrolysates and enzymatic hydrolysates from carbohydrate-rich Prosopis juliflora, an invasive perennial deciduous shrub of semi-arid regions, were used for bioethanol production. Saccharomyces cerevisiae and Pichia stipitis were used for fermentation of hexoses and pentoses. P. juliflora acid hydrolysate with an initial sugar concentration of 18.70 ± 0.16 g/L was concentrated to 33.59 ± 0.52 g/L by vacuum distillation. The concentrated hydrolysate was pretreated and fermented by mono- and co-culture methods either singly or in combination with enzyme hydrolysate and ethanol yields were compared. Monoculture with S. cerevisiae (VS3) and S. cerevisiae (NCIM3455) yielded maximum ethanol of 36.6 ± 1.83 g/L and 37.1 ± 1.86 g/L with a fermentation efficiency of 83.94 ± 4.20% and 84.20 ± 4.21%, respectively, after 36 h of fermentation. The ethanol yield obtained was 0.428 ± 0.02 g/g substrate and 0.429 ± 0.02 g/g substrate with a productivity of 1.017 ± 0.051 g/L/hand 1.031 ± 0.052 g/L/h, respectively. P. stipitis (NCIM3498) yielded maximum ethanol of 24 g/L with ethanol yield of 0.455 ± 0.02 g/g substrate and a productivity of 1.004 ± 0.050 g/L/h after 24 h of fermentation. With concentrated acid hydrolysate as substrate, S. cerevisiae (VS3) produced ethanol of 8.52 ± 0.43 g/L, whereas S. cerevisiae (NCIM3455) produced 5.96 ± 0.30 g/L of ethanol. P.stipitis (NCIM3498) produced 4.52 ± 0.23 g/L of ethanol by utilizing 14.66 ± 0.87 g/L of sugars. Co-culture with S. cerevisiae (VS3) addition after 18 h of addition of P. stipitis (NCIM3498) to the mixture of concentrated acid hydrolysate and enzyme hydrolysate produced 13.86 ± 0.47 g/L of ethanol with fermentation efficiency, ethanol yield and productivity of 87.54 ± 0.54%, 0.446 ± 2.36 g/g substrate and 0.385 ± 0.014 g/L/h, respectively. Hence, it is concluded that co-culture with S. cerevisiae and P. stipitis is feasible, further scaling up of fermentation of P. juliflora substrate for bioethanol production.

Keywords: Prosopis juliflora, Lignocellulose, Co-culture, Bioethanol, Fermentation efficiency, Ethanol yield

Introduction

Lignocellulosic substrates are in great abundance, easily available, of relatively low-cost and have potential to produce clean fuel when compared to sugar or starch containing feed stocks (Naseeruddin et al. 2013; Romani et al. 2013). Hence, exploitation of these sources may provide sustainable energy supply at local, regional and national levels and thereby could contribute to the economy (Balat and Balat 2009) without hampering food security. Moreover, bioethanol has got the adaptability to existing engines and if used in transportation can replace 30% of gasoline use (Krishnan et al. 2010; Oleskowicz et al. 2011).

Prosopis juliflora is a perennial deciduous xerophytic invasive weed, commonly found in semi-arid regions of the Indian subcontinent, Saudi Arabia and the United States of America (Gupta et al. 2009). It can grow on even poor and degraded soils and withstand salinity equal to sea water. The species grows rapidly and yields of 75–100 tons per ha in 15-year rotations and it is the only tree that yields 2.5 tons of wood/ha/year where nothing else can grow (Felker 2009). The tree is valued for its shade, forage, drought tolerance, salinity as well as grazing (Pasha et al. 2008). Due to its invasion along the roadsides, irrigation channels, community lands and productive agricultural lands, it has become a social menace. To eradicate this tree, mechanical, chemical and biological control programs were employed. The viable options to manage the menace of these invasive plants for human welfare include its use as fuel wood and application in biofuel industry (Rilov and Crooks 2008) as its carbohydrate content is 69.25% (on dry weight basis). As the tree does not form a part of main food or feed cycle, it qualifies to be a suitable substrate for long-term sustainable production of bioethanol (Pasha et al. 2008).

The various steps involved in bioethanol production from lignocellulosic substrates involve (1) drying of the substrate, (2) size reduction by milling/grinding; (3) pretreatment of the substrates; (4) hydrolysis (acid/enzymatic); (5) detoxification; (6) fermentation; and (7) separation of bioethanol by distillation. The hydrolysate obtained after acid saccharification of lignocellulosic biomass contains toxic compounds, generated mainly due to the degradation of sugars from hemicellulose and cellulose. These inhibitors interfere with the physiology of yeast cells resulting in decreased cell viability, ethanol yield and productivity (Chandel et al. 2007; Hahn-Hagerdal et al. 2007). To minimize the effect of these inhibitory compounds several detoxification methods have been studied, but the effectiveness of each method depends on the type of hydrolysate and microorganism used (Anish and Rao 2009). Moreover, each detoxification method has specificity for certain compounds; therefore, combining one or more methods can yield better results for detoxification of the acid hydrolysate (Carvalho et al. 2006). Overliming followed by neutralization and charcoal treatment method is most widely used and is considered as a promising and economic method of detoxification (Pasha et al. 2008; Jun-jun et al. 2009; Chandel et al. 2011a).

A two-step acid hydrolysis of the biomass with 20–80% (v/v) concentrated sulphuric acid at 40–50 °C for 2–4 h in a reactor followed by dilution with water (Pasha et al. 2008) completely hydrolyzed cellulose to dextrose rapidly, whereas hemicellulose was converted to xylose with a little degradation. In the second step, further hydrolysis of cellulose required dewatering of the solid residue followed by soaking in 30–40% sulfuric acid for 50 min at 100 °C (Pasha et al. 2008). Iranmahboob et al. (2002) reported maximum sugar recovery (78–82% of theoretical yields) from mixed wood chips by using 26% (v/v) sulfuric acid treated for 2 h. The advantage in this process was high sugar recovery due to hydrolysis of both hemicellulose and cellulose fraction into their monomeric fractions with nearly 90% of yields (Chandel et al. 2007).

Enzymatic hydrolysis is also adopted in bioethanol production and the degree of crystallinity of cellulose mainly affects the initial rate of enzymatic hydrolysis. Although enzymatic hydrolysis is slower than chemical method of hydrolysis, it does not require elaborate infrastructure (Dale and Moelhman 2005). The factors that govern enzymatic hydrolysis of lignocellulosic substrates are selection of suitable substrate and pretreatment method prior to enzymatic hydrolysis; cellulase activity; reaction conditions; and optimizing process parameters like temperature, pH enzyme loading, use of surfactant and end product inhibition to improve the enzymatic hydrolysis (Valchev et al. 2009).

For realizing higher ethanol yields, it is imperative that the initial sugars in the hydrolysate should be high. Concentration of initial sugars in the water-soluble fraction of hydrolysate can be achieved by evaporation of water or addition of less water to the hydrolysis process or concentrating the hydrolysate at 70 ºC under vacuum (Srilekha Yadav et al. 2011; Naseeruddin et al. 2016). Carvalho et al. (2006) concentrated the acid hydrolysate of Eucalyptus shavings and reported proportional increase in the sugar content as a function of the concentration factor employed.

Choice of organism is very important in ethanol production from lignocellulosic materials as the hydrolysate contains mixture of both hexoses and pentoses. S. cerevisiae and Zymomonas mobilis are most commonly used in industrial processes for ethanol production because of several advantages such as high ethanol productivity, high tolerance of high ethanol concentrations and inhibitors associated with it (Hickert et al. 2013). However, both of them lack the ability to utilize pentose sugars (Hector et al. 2011). Other yeasts like P. stipitis, Candida shehatae and Pachysolen tannophilus have also attracted interest as choice of organisms as they can convert pentose sugars into ethanol but are less tolerant to ethanol and inhibitors and also require a small and well-controlled supply of oxygen for maximum ethanol production (Hickertet al. 2013).

S. cerevisiae ferments only hexoses and P. stipitis ferments pentoses. As both pentoses and hexoses are present in the substrate, using only one organism could result in more of left-over sugars and thereby reduce ethanol yield. Most of the native microorganisms lack efficiency to ferment both hexose and pentose sugars present in the hydrolysate. For economical production of lignocellulose-based ethanol, simultaneous utilization of these biomass-derived sugars is required (Lin and Tanaka 2006). Though engineered strains of S. cerevisiae with an ability both sugars have been reported (Hou et al. 2017), they might show genomic instability (Wu et al. 2020). Such strains also showed preference for one sugar over another as a result of catabolite repression leading to longer time for the complete utilization and fermentation of all the sugars. On the other hand, standardized co-culture method with native strains could efficiently ferment both sugars resulting in increased ethanol yields and also reduce recovery costs as compared to monoculture. Co-culture for the fermentation of mixture of sugars was shown as a viable option (Srilekha Yadav et al. 2011), which circumvents the problems associated with monoculture of the wild strains or engineered yeast strains (Hickert et al. 2013). Hence, in the present study, an attempt was made to utilize concentrated hydrolysate of P. juliflora pre-treated with dilute acid where both the sugars can simultaneously get fermented to ethanol by co-culturing with Saccharomycs cerevisiae (VS3) and Pitchia stipitis (NCIM3498). This study aims at developing a viable fermentation technology to effectively use the lignocellulosic biomass of P. jliflora by combining the hydrolysates derived from biphasic acid and enzymatic hydrolyses. The novelty in this study lies in its approach to convert P. juliflora, a societal nuisance, into an opportunity by fermenting to produce bioethanol to meet the national energy security needs. Unlike the other substrates, this shrub grows under very hardy conditions without any external inputs including human labour. Also, once the method is standardized, its cultivation could be promoted in degraded lands and used for bioethanol production.

Materials and methods

Bioethanol production from 2nd generation biomass involves various steps from preparation of the raw material and its pre-treatment: hydrolysis of the biomass to release sugars; selection of suitable microorgainsms for fermentation of hexoses and pentoses; standardizing fermentation conditions and ethanol estimation. The processes followed in this study are described below in detail and presented using a flow diagram (Fig. 1).

Fig. 1.

Fig. 1

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Ethanol production and hexose sugars by S. cerevisiae VS3 and S. cerevisiae NCIM3455 in synthetic medium utilization at 10% and 15% level over time

Raw material, delignification and hydrolysis

Dried and chopped small pieces of 4–5 cm of Prosopis juliflora stem were used as raw experimental material for bioethanol production. They were ground in a laboratory mill fitted with a sieve of desired pore size in front of the sample outlet to ensure that the sample is retained till ground to the desired mesh size of 2–3. The ground substrate was again sieved so that starter material of uniform size,2–3 mm, is obtained. Later, the substrate was washed with tap water to make it dust-free and then dried in hot air oven at 65 ± 2 °C overnight to constant weight. From the raw material, both acid- and enzyme-hydrolysates were prepared as described by Naseeruddin et al. (2017) after the pre-treatment of the substrate (Naseeruddin et al. 2013). The methods of pre-treatment and hydrolysis followed are briefly described here.

One hundred grams of P. juliflora milled substrate was taken and pretreated with 2% (w/v) 0.5 M sodium dithionite (Na2S2O4) with a solid to liquid ratio of 1:10 for 18 h at room temperature (30 ± 2 °C). The biomass was neutralized under tap water, dried to constant weight at 65 ± 2 °C and used for further hydrolysis (Naseeruddin et al. (2017).

Alkali-pretreated and dried P. juliflora substrate was subjected to biphasic dilute acid saccharification. In the first phase, the substrate was autoclaved at 110 °C (110 PSI) for 30 min using 1% (v/v) sulphuric acid. In the second phase, the substrate was autoclaved at 121 °C (115 psi) for 45 min with 2% (v/v) sulphuric acid for maximum hydrolysis of holocellulose. The liquid phase containing the soluble materials from both phases were removed between the treatments and used for further studies (Naseeruddin et al. (2013).

Concentration of acid hydrolysate

Concentration of acid hydrolysate 1845 ml of obtained was carried out to increase the sugar concentration by vacuum distillation (assembled in the lab). During the vacuum distillation, the boiling temperature of the liquid was maintained at 70 °C. Concentration and fermentability of the hydrolysates were assessed in accordance with the Dehkhoda and Brandberg (2008). Sugars, phenolics and furans were checked before and after the concentration process.

Pretreatment of concentrated acid hydrolysate

The concentrated hydrolysate was detoxified by over-liming with calcium oxide (CaO), followed by pH adjustment to 6.00 ± 0.5 using 6 N H2SO4 and treated with charcoal. Sugars, phenolics and furans were estimated before and at each step of detoxification process. CaO addition at room temperature under fast stirring till the pH reached 10.5 ± 0.5 detoxified the acid hydrolysate. The mixture was incubated for 1 h with intermittent stirring to precipitate the inhibitors. The slurry was then filtered under vacuum to get clear filtrate by removing precipitates following the protocol of Chandel et al. (2007). The pH of the clear filtrate obtained after treating the hydrolysate with CaO was adjusted to 6.00 ± 0.5 by using 6 N H2SO4 and again filtered under vacuum to remove traces of salt precipitates as described by Chandel et al. (2007). After over-liming and neutralization, the hydrolysate was treated with 3.5% (w/v) of activated charcoal and stirred for 1 h as described by Martinez et al. (2000). The mixture was centrifuged at 3000 rpm for 20 min followed by filtration under vacuum to remove traces of precipitates.

Enzymatic saccharification

Enzymatic saccharification of the neutralized and dried left-over biomass after acid hydrolysis was done following the method of Govumoni et al. (2013) with slight modifications as described here. A sample of 32 g was drawn and subjected to enzymatic hydrolysis using cellulase enzyme (Advanced Enzyme Technologies) in sodium acetate buffer of pH 4.8–5.0. The enzyme loading was 19.46 IU of FPase, 28.66 IU of CMCase and 12.06 IU of b-glucosidase per 1 g of the substrate (equivalent to 4% (w/v) of enzyme dosage (g enzyme/g glucan of the pretreated solids × 100). To the reaction mixture, 1% (v/v) Tween-80 at 6% (w/v) substrate level was added and incubated at 50 °C, 150 rpm and for 36 h. The enzymatic saccharification efficiency was calculated as follows:

Saccharification%=Reducingsugarconcentrationobtained×0.98×100sugarconcentrationinthepretreatedsubstrate.

Microorganisms

Two hexose utilizing strains viz. Saccharomyces cerevisiae (VS3) and S. cerevisiae (NCIM3455) and a pentose utilizing strain Pichia stipitis (NCIM3498) were used in this study (Srilekha Yadav et al. 2011). VS3 originated from the soil samples collected within the hot regions Thermal Power Plant, Kothagudem India (Kiran Sree et al. 2000). S. cerevisiae (NCIM3455) and Pichia stipitis (NCIM3498) were obtained from National Chemical Laboratory, Pune, India. Both yeast cultures were maintained on yeast extract-peptone-dextrose-agar (pH 5.5) consisting of yeast extract, peptone, dextrose and agar at 10, 20, 20 and 25 g/L, respectively. P. stipitis was maintained on medium containing malt extract, glucose, xylose, yeast extract, peptone and agar at 5, 5, 30, 5 and 25 g/L, respectively.

Preparation of inoculum

VS3 and NCIM3455 inocula were first grown aerobically on YEPD broth (pH 5.5) containing yeast extract, peptone and dextrose at 10, 20 and 20 g/L, respectively, for 48 h at 37 ± 0.5 °C and 100 rpm in Erlenmeyer flasks on a rotary shaker (Pasha et al. 2007). NCIM3498 inoculum was grown aerobically in the broth containing 5 g/L each of xylose, malt extract, yeast extract, peptone and dextrose in Erlenmeyer flasks on a rotary shaker for 24 h at 30 °C and 100 rpm. Both inoculation broths were adjusted to pH 5.5. The inoculum of the test strains was adjusted to O.D. of 0.6 at 600 nm and the fermentation medium was inoculated at 10%, v/v by diluting with YEPD broth (Chandel et al. 2007; Gupta et al. 2009).

Fermentation of pentoses and hexoses by monoculture

All the fermentation studies were carried out using the test organisms either by monoculture or co-culture methods. In all the fermentations, the substrates included (1) synthetic medium, (2) concentrated acid hydrolysate medium and (3) a mixture of acid and enzyme hydrolysates. Hexose (dextrose) or pentose sugar s(xylose) was separately used as substrate for monoculture studies and both the sugars were mixed at optimum concentrations (tolerable to microorganisms) for co-culture studies. When concentrated acid hydrolysate and enzyme hydrolysate were used as substrate, no sugar was added but the fermentation medium was supplemented with additional nutrients viz. yeast extract, peptone, NH4Cl, KH2PO4 each at 1 gL−1; and MgSO4.7H2O, MnSO4.5H2O, CaCl2.2H2O, FeCl3.2H2O and ZnSO4.7H2O at 0.5 gL−1 before fermentation.

Synthetic fermentation medium

Twenty mL of medium in 100 mL Erlenmeyer flasks amended with either 10% or 15% sugar was inoculated with either VS3 or NCIM3455 separately and fermentation was carried out at 30 ± 2 °C, 150 rpm for 72 h. Similarly, another set of flasks containing synthetic medium amended with 2%, 4%, 6%, 8% 10% and 12% (w/v) xylose was inoculated NCIM3498 and fermentation was carried out at 30 ± 2 °C, 150 rpm for 48 h. In addition to sugars, the synthetic medium was further supplemented with yeast extract, peptone, NH4Cl, KH2PO4 each at 1 gL−1; and MgSO4.7H2O, MnSO4.5H2O, CaCl2.2H2O, FeCl3.2H2O and ZnSO4.7H2O each at 0.5 gL−1. The pH of the medium was adjusted to 5.5 ± 0.1 and autoclaved at 110 °C for 20 min (Pasha et al. 2007), cooled to 30 ± 2 °C and used for shake flask fermentation. The pH of medium was adjusted to 5.5 ± 0.1 and autoclaved at 110 °C for 20 min (Pasha et al. 2007), cooled to 30 ± 2 °C and used for fermentation. The fermentation medium was inoculated with test strains as prepared and shown above at 10%, v/v by diluting with YEPD broth (Chandel et al. 2007; Gupta et al. 2009). Samples were collected at regular intervals and centrifuged at 6000 rpm for 10 min at 4 °C for the analysis of residual sugar and ethanol produced.

Concentrated acid hydrolysate

Monoculture fermentation with test microorganisms was carried out in 100 mL Erlenmeyer flasks using concentrated acid hydrolysate following the method of Naseeruddin et al. (2013). Six-hundred mL of detoxified and concentrated hydrolysate was supplemented with nutrients as described above before fermentation. Thirty mL of this mixture was drawn from 600 mL hydrolysate in three replications for fermentation. The pH of the mixture was adjusted to 5.5 ± 0.1 and autoclaved and inoculated with each organism separately at 10% (v/v). The shake flask fermentation was carried out at 30 ± 2 °C, 150 rpm for 72 h and samples were collected at regular intervals to estimate residual sugar and ethanol concentration.

Enzyme hydrolysate

The enzyme hydrolysate (533 mL) obtained at 100 g level of substrate was supplemented with nutrients and the pH of the medium was adjusted to 5.5 ± 0.1. For fermentation, 30 mL of hydrolysate was taken, obtained at 100 g level of substrate in two sets of 100 mL Erlenmeyer flasks, supplemented with additional nutrients as described above and autoclaved at 110 °C for 20 min (Pasha et al. 2007). The flasks were inoculated with 10% (v/v) inoculum of either VS3 or NCIM3455 cultures and shake flask fermentation was carried out at 30 ± 2 °C, 150 rpm for 72 h. Samples were collected at regular intervals to estimate residual sugar and ethanol concentration.

Fermentation of pentoses and hexoses by monoculture

Synthetic medium

The different combinations of co-culture used in the study were the following: co-culture a [both P. stipitis (NCIM3498) and S. cerevisiae (VS3) added at the same time], co-culture b [both P. stipitis (NCIM3498) and S. cerevisiae (NCIM3455) added at the same time; co-culture c [S. cerevisiae(VS3) added to the medium after 24 h of addition of P. stipitis (NCIM3498)] and co-culture d [S. cerevisiae (NCIM3455) added after 24 h of addition of P. stipitis (NCIM3498)]. In each of the four 150 mL conical flasks, 75 mL of synthetic medium was taken; the pH was adjusted to 5.5 ± 0.1 and autoclaved (Pasha et al. 2007). Later, the medium was cooled to room temperature (30 ± 2 °C) and inoculated with various combinations of co-cultures as described above. Both the inocula (pentose and hexose yeast) were added in 1:1 ratio at 10% level (5% pentose yeast + 5% hexose yeast) to the fermentation medium and fermentation was carried out at 30 ± 2 °C, 150 rpm for first 18 h and then in static mode till the end of fermentation, i.e. 72 h. Samples were collected at regular intervals, centrifuged at 6000 rpm for 10 min at 4 °C and analyzed for residual sugars and ethanol concentration.

Concentrated acid hydrolysate

About 150 mL of detoxified concentrated acid hydrolysate was taken from 600 mL of detoxified acid hydrolysate concentrate and supplemented with nutrients as described above. The pH of medium was adjusted to 5.5 ± 0.1 and autoclaved (Pasha et al. 2007). From the previous experiment, out of the four combinations of co-cultures tested, co-culture C [P. stipitis (NCIM3455) + S. cerevisiae (VS3)] yielded maximum ethanol yield and hence was was tested for ethanol production from concentrated acid hydrolysate. The inoculated flasks were incubated on a rotary shaker; samples were collected at regular intervals, centrifuged at 6000 rpm for 10 min at 4 °C and analyzed for residual sugars and ethanol.

Mixture of concentrated acid and enzyme hydrolysates

About 300 mL of concentrated acid hydrolysate obtained at 100 g (600 mL) level of substrate was mixed with equal volume of enzyme hydrolysate obtained (533 mL) and supplemented with nutrients as described above. The pH of medium was adjusted to 5.5 ± 0.1 and autoclaved. From the previous experiment, out of the four combinations of co-cultures tested, co-culture C [P. stipitis (NCIM3455) + S. cerevisiae (VS3)] was used here also. The inoculated flasks were incubated on a rotary shaker; samples were collected at regular intervals, centrifuged at 6000 rpm for 10 min at 4 °C and analyzed for residual sugars and ethanol.

Total reducing sugars, phenolics and furans assay

The total reducing sugars released during delignification and after acid hydrolysis were estimated by DNS method (Miller 1959). For estimation of phenolics, Folin–Ciocalteu method as described by (Singleton and Rossi 1965) was followed with gallic acid as standard. For estimation of total furans, the method of Martinez et al. (2000) was adopted with fufuraldehyde as standard. Cell density was measured at 600 nm using a turbidometer. Turbidity of the fermentation broth measured using UV–VIS spectrophotometer 117 (Systronics India).

Ethanol estimation

Ethanol production was estimated by gas chromatograph (Shimadzu GC-2011- Japan) using ZB-Wax column (30 mm × 0.25 mm) with a flame ionization detector. The sample used for ethanol estimation was filtered by 0.22 µm cellulose acetate filter for GC analysis. The analysis was performed according to National Renewable Energy Laboratory procedure LAP #001 (Templeton 1994). The column temperature was maintained at 150 °C (isothermal) and the carrier gas nitrogen was run at pressure of 16 kPa with a run time of 5.5 min. The injector temperature was at 175 °C and the detector temperature was at 250 °C. The other parameters used were flow rate: 40 mL/min, spilt ratio: 1/50, velocity of H2 flow: 60 mL/min, sample quantity: 1 µL.

Data analysis

All the experiments were replicated three times, repeated and mean and standard deviation (SD) values were calculated using MS Excel software. Fermentation efficiency was calculated as follows:

Frementationefficiency =PracticalyeildTheoriticalyeild×100,

where Practical yield is the ethanol produced and the theoretical yield is 0.511 per gram of sugar consumed.

Results and discussion

Concentration of acid hydrolysate

Vacuum distillation of the acid hydrolysate resulted in nearly twofold increase in sugar content to 33.59 ± 0.52 g/L. However, the phenolics content also got nearly doubled and reached 3.94 ± 0.18 g/L and the furans concentration increased to 1.72 ± 0.12 g/L with a decrease in total volume of hydrolysate from 1845 to 922 mL. Pretreatment of the concentrated hydrolysate obtained (922 mL) by detoxification, neutralization and charcoal addition significantly reduced phenolics content to 0.56 ± 0.11 g/L and furans content to 0.21 ± 0.08 g/L. However, there was a marginal reduction in sugar content of 1.52 g/L with a resulting final volume of 600 mL of the concentrate (Table 1). Srilekha Yadav et al. (2011) could concentrate fermentable sugars up to 88% in the rice straw hydrolysate which resulted in increased overall ethanol yield. Among three methods of concentration tried, high-pressure evaporation yielded maximum levels of fermentable sugars in dilute-acid hydrolysate of spruce wood while evaporation and vacuum evaporation were on a par (Dehkhoda and Brandberg 2008).

Table 1.

Sugars, phenolics and furans present in acid hydrolysate before and after concentration and detoxification process

Hydrolysate Volume (ml) Sugars Phenolics Furans
Total (mg) (g/L) Total (mg) g/L Total (mg) g/L
Acid hydrolysate 1845 34,501.5 ± 36.12 18.7 ± 0.16 3985.2 ± 145.00 2.2 ± 0.10 2029.5 ± 1.11 1.1 ± 0.03
After concentration 922 30,970.0 ± 22.84 33.6 ± 0.52 3540.5 ± 135.11 3.9 ± 0.18 2933.0 ± 1.94 1.7 ± 0.12
After detoxification 600 19,200.0 ± 1.43 32.0 ± 0.48 335.0 ± 12.46 0.6 ± 0.11 126.0 ± 2.31 0.2 ± 0.08
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Monoculture studies

Synthetic medium

At 10% (w/v) sugar concentration, ethanol yield increased with the fermentation time and both VS3 and NCIM3455 produced maximum ethanol of 36.6 ± 1.83 g/L and 37.1 ± 1.86 g/L by utilizing 85.50 ± 4.28 g/L and 86.40 ± 4.32 g/L of sugars. The fermentation efficiency was 83.94 ± 4.20% and 84.20 ± 4.21%, respectively, after 36 h of fermentation. The ethanol yield obtained was 0.428 ± 0.02 g/g and 0.429 ± 0.02 g/g with productivity of 1.017 ± 0.051 g/L/h and 1.031 ± 0.052 g/L/h, respectively. After 36 h, the ethanol yield and productivity reduced for both strains (Table 2).

Table 2.

Ethanol production by S. cerevisiae (VS3) and S. cerevisiae (NCIM3455) in synthetic medium amended with 10% and 15% sugars

Time (h) Sugar conc (%) S. cerevisiae (VS3) S. cerevisiae (NCIM3455)
Ethanol yield (g/L) Left over sugar (g/L) Ethanol yield (g/L) Left over sugar (g/L)
12 10 7.4 62.3 7.2 60.2
15 9.1 91.2 8.1 88.3
24 10 17.2 42.3 17.6 39.8
15 24.2 56.4 23.9 51.6
36 10 36.6 14.5 37.1 13.6
15 54.1 17.6 53.8 18.1
48 10 36.7 14.2 37.3 13.1
15 54.3 17.2 53.9 17.6
60 10 36.4 13.8 37.0 12.7
15 54.3 17.0 54.1 17.3
72 10 36.5 13.6 37.2 12.1
15 54.4 16.7 53.8 17.2
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Similar trend was observed with 15% (w/v) sugar concentration too. Both VS3 and NCIM3455 strains produced maximum ethanol of 54.1 ± 2.71 g/Land 53.8 ± 2.69 g/L by consuming 132.40 ± 6.62 g/L and 131.90 ± 6.60 g/L of sugars with fermentation efficiency of 80.12 ± 3.51% and 79.98 ± 2.46%, respectively, after 36 h of fermentation. The ethanol yield obtained was 0.409 ± 0.01 g/g and 0.408 ± 0.02 g/g with productivity of 1.503 ± 0.03 g/L/h and 1.494 ± 0.02 g/L/h, respectively. Here also, after 36 h, the ethanol yield and productivity reduced for both strains (Table 2). Both the strains produced the highest ethanol after 36 h of fermentation. Maximum ethanol production of 24 g/L by utilizing 53 ± 2.65 g/L of D-xylose with ethanol yield of 0.455 ± 0.02 g/g and productivity of 1.004 ± 0.050 g/L/h was obtained with P. stipitis (NCIM3498) after 24 h of fermentation. The ethanol yield and productivity reduced after 24 h. (Fig. 2). After 24 h of fermentation, highest ethanol production was observed. The fermentation efficiency of the P. stipitis (NCIM3498), S. cerevisiae (VS3) and S. cerevisiae (NCIM3455) was first evaluated in synthetic medium individually to select best strains and to optimize the strains and fermentation conditions for fermentation of test substrate. Chandel et al. (2011b) reported that monoculture with P. stipitis NCIM3498 yielded higher ethanol in a simulated synthetic medium as compared to S. cerevisiae (VS3). However, in our studies, significant ethanol yield was realized in synthetic medium when monocultured with S. cerevisiae VS3 either at 10% or 15% sugar levels.

Fig. 2.

Fig. 2

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Ethanol production and pentose sugars’ utilization at different levels of xylose by P. stipitis NCIM3498 in synthetic medium over time

Concentrated acid hydrolysate

Fermentation of concentrated and detoxified acid hydrolysate with initial sugar concentration of 32 ± 0.48 g/L by Saccharomyces cerevisiae (VS3) resulted in production of 8.52 ± 0.43 g/L ethanol by consuming 25.62 ± 1.28 g/L of sugars. The fermentation efficiency, ethanol yield and productivity were 66.21 ± 3.26%, 0.33 ± 0.02 g/g and 0.24 ± 0.01 g/L/h, respectively. However, S. cerevisiae (NCIM3455) could consume only 22.33 ± 1.12 g/L of sugars and produced 5.96 ± 0.30 g/L of ethanol, thereby resulting in fermentation efficiency, ethanol yield and productivity of 52.33 ± 2.62%, 0.27 ± 0.01 g/g and 0.17 ± 0.01 g/L/h, respectively. The pentose sugar fermenting yeast P. stipitis (NCIM3498) produced 4.52 ± 0.23 g/L of ethanol by utilizing 14.66 ± 0.87 g/L of sugars with fermentation efficiency, ethanol yield and productivity of 60.46 ± 3.02%, 0.31 ± 0.02 g/g and 0.13 ± 0.01 g/L/h, respectively (Fig. 3). Though the fermentation was continued for 72 h, all the strains produced the highest ethanol after 36 h. Our findings are in line with that of Gupta et al. (2009) who also could successfully produce ethanol from monoculture of hemicellulosic hydrolysate of P. juliflora with P. stipitis. Canilha et al. (2010) could get a maximum of 0.30 g/g ethanol yield and 0.13 g/L/h ethanol productivity from sugarcane bagasse hemicellulosic hydrolysate after 48 h by monoculture of P. stipitis DSM3651. Batch fermentation using S. cerevisiae (OVB 11) produced maximum ethanol of 7.5 g/L after 36 h of fermentation with ethanol yield 0.3 g/g (Srilekha Yadav et al. 2011). The low ethanol efficiency, i.e. 55% obtained may be due to the reason that most of the xylose was left unfermented by S. cerevisiae (OVB 11) out of xylose and dextrose in hydrolysate.

Fig. 3.

Fig. 3

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Ethanol production and sugar utilization by S. cerevisiae strains VS3 and NCIM3455 and P. stipitis NCIM3498 with acid concentrated hydrolysate over time

Enzyme hydrolysate

Fermentation of enzyme hydrolysate with an initial sugar concentration of 37.37 ± 0.81 g/L by S. cerevisiae (VS3) resulted in production of 11.56 ± 0.78 g/L of ethanol after consuming 30.53 ± 0.34 g/L sugars. The fermentation efficiency, ethanol yield and productivity were 74.25 ± 0.58%, 0.379 ± 3.71 g/g and 0.321 ± 0.02 g/L/h, respectively. On the other hand, slightly less ethanol (9.61 ± 0.74 g/L) was produced when the enzyme hydrolysate was fermented with S. cerevisiae (NCIM3455) by consuming 28.94 ± 0.42 g/L of sugars. The fermentation efficiency, ethanol yield and productivity were 65.12 ± 0.48%, 0.332 ± 3.26 g/g and 0.267 ± 0.02 g/L/h, respectively (Fig. 4). Highest ethanol outturn was recorded after 36 h fermentation by both the strains. Láinez et al. (2019) could successfully ferment enzymatic hydrolysate of Agave salmiana hydrolysate with 50 g/L sugars by employing Kluyveromyces marxianus and the efficiency was about 94%. Similarly, Shokrkar et al. (2017) saccharified both fresh and dried algae with thermostable enzymes with highest recovery of 0.951 g of extracted glucose per gram of total sugar.

Fig. 4.

Fig. 4

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Ethanol production and sugar utilization by S. cerevisiae VS3 and S. cerevisiae NCIM3455 with enzyme hydrolysate over time

Co-culture studies

Synthetic medium

The synthetic medium amended with 6% dextrose and 2% xylose was used for co-culture studies. Among the four co-culture protocols tried, co-culture C produced maximum ethanol followed by co-culture A, whereas the co-culture B was least efficient. In co-culture C, where S. cerevisiae VS3 was added after 18 h of inoculation with P. stipitis NCIM3498, the highest amount of 3.06 ± 0.13 g/L ethanol was produced after utilizing 6.74 ± 0.34 g/L of sugars. The fermentation efficiency, ethanol yield and productivity were 89.02 ± 4.21%, 0.454 ± 0.02 g/g and 0.085 ± 0.003 g/L/h, respectively. Simultaneous inoculation of S. cervisiae VS3 and P stipitis NCIM3498 (co-culture A) resulted in 2.51 ± 0.13 g/L of ethanol yield by consuming 6.14 ± 0.31 g/L sugars. The fermentation efficiency, ethanol yield and productivity were 80.16 ± 4.01%, 0.409 ± 0.02 g/g and 0.070 ± 0.003 g/L/h, respectively. Co-culture D, where S. cerevisiae NCIM3455 was added after 18 h of addition of P. stipitis NCIM3498), produced 2.47 ± 0.14 g/L of ethanol by utilizng 5.88 ± 0.28 g/L of sugars with fermentation efficiency, ethanol yield and productivity of 82.37 ± 3.63%, 0.420 ± 0.01 g/g and 0.069 ± 0.001 g/L/h, respectively. In co-culture B, where S. cerevisiae NCIM3455 and P. stipitis NCIM3498 were used simultaneously as inoculum, 2.21 ± 0.11 g/L of ethanol was produced by consuming 5.52 ± 0.28 g/L of sugars. The fermentation efficiency, ethanol yield and productivity were 78.50 ± 3.93%, 0.400 ± 0.01 g/g and 0.061 ± 0.001 g/L/h, respectively (Fig. 5). In all configurations of co-cultures, maximum ethanol yield was realized after 36 h of fermentation. Co-culture method of fermentation facilitates simultaneous utilization of pentoses and hexose sugars present in the hydrolysate to improve bioethanol yields and thus reduces recovery costs. Monoculturing with Bacillus cereus and B. thuringiensis, Ire et al. (2016) reported ethanol yield of 18.40 and 15.27 g/L from bagasse, respectively, while co-culture with same strains yielded 19.08 g/L ethanol. Similarly, co-culture with two engineered strains of S. cerevisiae could produce 2.1 g/L ethanol from brown microalgae (Sasaki et al. 2018).

Fig. 5.

Fig. 5

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Ethanol yields on synthetic medium fermented with four co-culture configurations over time

Concentrated acid hydrolysate

Co-culture with S. cerevisiae VS3 after 18 h of addition of P. stipitis NCIM3498 (co-culture C) could utilize up to 81% of the sugars present in the concentrated acid hydrolysate with initial sugar levels of 32 ± 0.48 g/L sugars and yield 10.94 ± 0.53 g/L of ethanol. The fermentation efficiency, ethanol yield and ethanol productivity were 83.11 ± 0.42%, 0.420 ± 0.01 g/g and 0.30 ± 0.01 g/L/h, respectively, (Fig. 6). Highest levels of ethanol could be realized after 36 h of fermentation and after that the ethanol yield did not improve any further. Sornvoraweat and Jirasak (2009) also reported that acid hydrolysate from cassava peels yielded maximum ethanol yield when co-cultured with S. cerevisiae and Candida tropicalis as compared to monoculture with S. cerevisiae. Batch fermentation of concentrated and detoxified acid hydrolysate of rice straw using co-culture of S. cerevisiae (OVB 11) and P. stipitis (NCIM3498) resulted in maximum ethanol production after 36 h of incubation with an efficiency of 95% (Srilekha Yadav et al. 2011). In the present study also, 10.94 ± 0.53 g/L of ethanol was obtained and the increased ethanol yield and ethanol efficiency could be due to the total conversion of both hexoses and pentoses present in the hydrolysate. Chandel et al. (2011b) reported that co-culture helped higher yields of bioethanol from sweet sorghum as compared to monoculture. Fu et al. (2009) reported maximum ethanol production of 0.49 g/g from sugarcane bagasse hydrolysate by co-culturing with Z. mobilis (ATCC 10,988) and P. stipitis (CBS 5773) within 48 h of incubation. On the contrary, Ferrari et al. (1992) reported 12.6 g/L ethanol concentration and attributed the same to the complete conversion of dextrose and xylose mixture (50 g/L) by a respiratory deficient mutant of S. diastaticus co-cultivated with P. stipitis in continuous culture up to 75 h.

Fig. 6.

Fig. 6

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Ethanol yields from the medium amended with concentrated acid hydrolysate and fermented by co-culture C over time

Mixture of concentrated acid hydrolysate and enzyme hydrolysate

The mixed hydrolysate (containing 300 mL each of concentrated and detoxified acid hydrolysate and enzyme hydrolysate obtained) had an initial sugar concentration of 34.68 ± 1.73 g/L. Inoculation of this substrate medium with S. cerevisiae VS3 after 18 h of inoculation with P. stipitis NCIM3498 (co-culture c) utilized 31.04 ± 0.11 g/L sugars to produce 13.86 ± 0.47 g/L of ethanol with fermentation efficiency, ethanol yield and productivity of 87.54 ± 0.54%, 0.446 ± 2.36 g/g and 0.385 ± 0.014 g/L/h, respectively, (Fig. 7). When a mixture of acid and enzyme hydrolysate of wheat straw substrate was fermented with S. cerevisiae VS3, an ethanol yield of 0.44 g/g and productivity of 0.6 g/L/h were realized by Govumoni et al. (2013). Similar results were reported by Pasha et al. (2008) using S. cerevisiae VS3 and a mixture of acid and enzyme hydrolysate of P. juliflora. While the present findings are in line with that of Canilha et al. (2010), Srilekha Yadav et al. (2011) and Govumoni et al. (2013), the ethanol yields were marginally less in monoculture with hexose fermenting yeast as compared to the results of Gupta et al. (2009).

Fig. 7.

Fig. 7

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Ethanol yields from the medium amended with mixture of concentrated acid hydrolysate and enzyme hydrolysate and fermented by co-culture C over time

Conclusion

Two species of Saccharomyces and one species of Pitchia could successfully utilize concentrated Prosopis juliflora acid hydrolysate as well as enzyme hydrolysate to ferment both hexose and pentose sugars into ethanol. Monoculture with S. cerevisiae and P. stipitis in synthetic medium, concentrated acid hydrolysate and enzyme hydrolysate converted pentose and hexose sugars into bioethanol. A mixture of concentrated acid hydrolysate and enzyme hydrolysate taken in 1:1 proportion and supplemented with nutrients could be used successfully as substrate for co-culturing with S. cerevisiae VS3 and P. stipitis (NCIM3498) which yielded 10.94 ± 0.53 g/L of ethanol with ethanol yield of 0.420 ± 0.01 g/g and volumetric ethanol productivity of 0.30 ± 0.01 g/L/h. These results could be scaled up for the production of ethanol from lignocellulosic concentrated acid hydrolysate of P. juliflora after pre-treatment for second-generation ethanol production by employing co-culture of S. cerevisiae (VS3) and P. stipitis (NCIM3498) and mitigate environmental hazards due to this invasive species in semi-arid regions.

In this initial study, the feasibility of utilizing a 2nd generation biomass like P. juliflora, which is also an invasive weed, was found to be promising. However, the main limitations in use of sustained viability of use of 2nd generation biomasses for ethanol production are huge capital and operational costs. It is also important to note here that the intangible returns in the form of use of invasive weed should be valued. The left-over solids after fermentation is a value-added product and could be used as soil amendment for improving physico-chemical properties and also supplement crop nutrition.

Acknowledgements

We thank the Department of Biotechnology (DBT), Ministry of Science and Technology, UGC-MANF and DRS-I SAP (Government of India) for the financial assistance.

Author contributions

SN, as research scholar, performed all the experiments, analyzed the data and prepared the manuscript. SD assisted in design of experiments, provided critical inputs, supervised the work as the major thesis supervisor and edited the manuscript. LVR, as Principal Investigator of the projects funded by different agencies, contributed to the technical vetting of the experiments, provided technical and financial facilities and contributed to editing of the manuscript. All authors have read and approved the manuscript.

Data availability

There are no supporting data.

Compliance with ethical standards

Conflict of interests

The authors declare that they have no competing interest.

Consent of publication

All authors have read and approved the manuscript.

Ethics approval and consent to participate

Not applicable.

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