Acetone-butanol-ethanol (ABE) fermentation using the root hydrolysate after extraction of forskolin from Coleus forskohlii

Renewable Energy 86 (2016) 594e601

Contents lists available at ScienceDirect

Renewable Energy journal homepage: www.elsevier.com/locate/renene

Acetone-butanol-ethanol (ABE) fermentation using the root hydrolysate after extraction of forskolin from Coleus forskohlii Shirish M. Harde a, b, Swati B. Jadhav a, b, Sandip B. Bankar b, Heikki Ojamo b, € m b, Rekha S. Singhal a, Shrikant A. Survase b, * Tom Granstro a b

Food Engineering and Technology Department, Institute of Chemical Technology, Matunga, Mumbai 400019, India Department of Biotechnology and Chemical Technology, Aalto University School of Chemical Technology, P.O. Box 16100, 00076 Aalto, Finland

a r t i c l e i n f o

a b s t r a c t

Article history: Received 1 September 2014 Received in revised form 14 August 2015 Accepted 20 August 2015 Available online xxx

The biomass obtained after the extraction of forskolin from the roots of Coleus forskohlii was evaluated as a substrate for the production of acetone-butanol-ethanol (ABE). The spent biomass constituting more than 90% of the raw material showed 50e70% carbohydrates with starch and cellulose being the major constituents. This study was undertaken to optimize enzymatic hydrolysis of C. forskohlii roots for maximum release of fermentable sugars and subsequent fermentation to ABE. The root biomass was hydrolyzed using the Stargen® 002 and Accellerase® 1500. Cocktail of both enzymes (16U Stargen® 002 and 60 FPU Accellerase® 1500) could produce 41.2 g/l of total reducing sugars (glucose equivalent to 32.33 g/l). The production of ABE was optimized in a batch fermentation using Clostridium acetobutylicum NCIM 2877. The maximum ABE production using the root hydrolysates was 0.55 g/l. Pretreatment with lime and Amberlite XAD-4 increased the production of total solvent to 5.33 g/l. © 2015 Elsevier Ltd. All rights reserved.

Keywords: Coleus forskohllii root ABE fermentation Pretreatment Clostridium acetobutylicum Biobutanol

1. Introduction Coleus forskohlii Briq, a herbal plant belonging to the Lamiaceae family, is native to India and is reported in Ayurvedic Materia Medica under the Sanskrit name Makandi and Mayini [1]. Presently, about 40,000 acres of land is under cultivation of C. forskohlii in India, Africa and South East Asia for its tuberous roots [2]. The cultivation may provide an average yield of 800e1000 kg/ha of dry tubers which can be improved to 2000e2200 kg/ha of dry tubers by applying proper practices. The cultivation of C. forskohlii has been on an increase due to its commercial utilization [3]. The roots of the C. forskohlii plant are a unique source of forskolin (FSK), a labdane diterpene compound. FSK has been shown to be useful in the treatment of asthma, glaucoma, cardiovascular diseases and certain types of cancer [4]. The spent biomass obtained after extraction of forskolin from the root has no commercial value. It constitutes more than 90% of carbohydrate-rich raw material which could be used as a substrate for the production of value added chemicals and fuels such as ABE solvents. Currently, tonnes of C. forskohlii root biomass is either

* Corresponding author. Tel.: þ358 400368375; fax: þ358 9 462 373. E-mail address: [email protected] (S.A. Survase). http://dx.doi.org/10.1016/j.renene.2015.08.042 0960-1481/© 2015 Elsevier Ltd. All rights reserved.

dumped or burnt which are environmentally hazardous. There is an increased interest among the researchers to develop various strategies to utilize waste biomass for useful and value-added purpose. Increase in petroleum prices and depletion of fossil fuels are the key reasons for ongoing search of energy alternatives worldwide [5]. Bioconversion of waste biomass to alcoholic fuels such as bioethanol, biobutanol, and biodiesel is rapidly emerging as an area of interest among researchers [6]. Currently, countries like USA and Brazil contribute 20e30% of biobutanol production in fuel market, while in Asia the bioethanol production is at a very early stage of development [7,8]. Biobutanol has attracted the attention of researchers and investors due to its various advantages over other biofuels such as like high heating value, low freezing point, high hydrophobicity, and low heat of vaporization that are closer to gasoline [9e11]. The current production of nbutanol is about 5e6 million tons per year with a worldwide market sale of US$7e8.4 billion [12]. The market demand is anticipated to increase dramatically, if n-butanol can be produced cost-effectively [13]. Butanol can be obtained from renewable biomass by ABE fermentation [14]. Clostridium acetobutylicum utilizes a range of carbon sources to produce butyric acid and acetic acid in the first phase (acidogenesis), and acetone, butanol and ethanol in the second phase (solventogenesis). The process of biobutanol production

S.M. Harde et al. / Renewable Energy 86 (2016) 594e601

consists of several unit operations such as pretreatment of the biomass, clostridial ABE fermentation, and recovery of the desired product. The pretreatment of the biomass varies with its chemical makeup (starch rich, sugar rich or lignocellulosic materials). The cost of the fermentation substrate conclusively decides the economics of biobutanol production [15]. The use of inexpensive, abundantly available, sustainable and renewable feedstock such as lignocellulosic materials (agricultural waste, paper waste, wood chips, etc.) has the potential to reduce the cost of biobutanol production. These agricultural biomass sources cost much less (US$24e75/ton) than the traditional substrates [16]. The use of lignocellulosic biomass such as wheat straw [17], wheat bran [18], barley straw [19], corn stover and switch grass [20], corn fiber [21] and wood [22] for solvent production has been reported earlier. The lignocellulosic biomass is pretreated to fractionate the lignin, hemicelluloses and cellulose prior to hydrolysis and subsequent ABE fermentation. The types of pretreatments and their merits and demerits have been reviewed by Alvira et al. [23] and Sun and Cheng [24]. The hydrolysis of lignocelluloses results in the formation of some inhibitory components such as formic acid, acetic acid, levulinic acid, furfural and hydroxymethyl furfural [25] which limits the ABE fermentation. Several detoxification methods have been previously investigated for the removal of these inhibitors. These include lime treatment, evaporation, and adsorption using ion exchange resin or activated charcoal to biological treatment including laccase and peroxide enzymes [26e29]. Sun and Liu [30] investigated the application of membrane filtered sugar maple wood extract hydrolysates for ABE production and found that overliming treatment can significantly improve the ultimate butanol concentration from 0.8 g/l to 7 g/l. The objective of this study was to investigate ABE production from C. forskohlii root hydrolysate using C. acetobutylicum. The root biomass of C. forskohlii was characterized to determine the carbohydrate content which was further hydrolyzed enzymatically to fermentable sugars. The phenolic compounds present in the root hydrolysates were removed by pretreatment using Ca(OH)2 and resin i.e. Amberlite XAD-4. The pretreated root hydrolysate was subsequently used for ABE fermentation. 2. Materials and methods 2.1. Materials C. forskohlii roots were procured from Salem, Tamilnadu, India. Dried roots were ground in a mill fitted with 18 mesh to a particle size <1 mm and stored in air tight containers for further studies. Peptone, meat extract, yeast extract, glucose, starch, NaCl, sodium acetate, L-cysteine, ammonium acetate, K2HPO4, KH2PO4, p-amino benzoic acid, thiamine HCl, biotin, FeSO4, MnSO4, MgSO4, Ca(OH)2, NaOH and HCl was purchased from Hi Media Laboratories, Mumbai, India. Amberlite XAD-4 resin was purchased from Sigma Aldrich, Mumbai, India. Glucose oxidase-peroxidase (GOD-POD) kit was purchased from Accurex Biomedical Pvt. Ltd. Mumbai, India. Enzymes (Stargen® 002 and Accellerase® 1500) were gifted by Genencor International, Mumbai, India. All the chemicals used were of analytical grade. 2.2. Proximate analysis and carbohydrate profiling of C. forskohlii roots Moisture, ash, fat and nitrogen content were determined by standard AOAC Official methods [31]. Total carbohydrate content of sample was calculated by difference [32], rather than by direct analysis. Dried and powdered C. forskohlii roots samples were used for carbohydrate profiling as per the earlier reports [33,34].

595

Carbohydrate components were fractionated into free sugars, oligosaccharides, starch, pectin, hemicellulose, cellulose and lignin. 2.3. Optimization of parameters for enzymatic saccharification of C. forskohlii roots The principal components of C. forskohlii roots are starch and cellulose. Hence Stargen® 002 and Accellerase® 1500 were used individually and/or in combination in order to determine the optimum conditions for complete saccharification of starch and cellulose. Slurry of C. forskohlii roots (10% w/v) was prepared in sodium citrate buffer by gentle stirring using a magnetic stirrer. Starch breakdown was carried out by using Stargen® 002 which contains Aspergillus kawachi a-amylase expressed in Trichoderma reesei and a glucoamylase from Trichoderma reesei that work synergistically to hydrolyze granular starch substrate to glucose. The pH of slurry (3.5e5.5), hydrolysis temperature (30  Ce70  C) and substrate concentration (5e25% w/v), Stargen® 002 (4e16 U/g dry weight of C. forskohlii roots) and incubation time (6e30 h) were optimized for maximum release of reducing sugars. Cellulose breakdown was carried out by using Accellerase® 1500 which is a mixture of cellulase and glucosidase that works synergistically to hydrolyze cellulosic substrate to glucose. The pH of slurry (3.5e5.5), hydrolysis temperature (30  Ce60  C), incubation time (8e40 h), Accellerase® 1500 (30e120 FPU/g dry weight of C. forskohlii roots) and substrate concentration (5e25% w/v) were optimized for maximum release of reducing sugars. Individual and cocktail effect of enzymes on starch and cellulose hydrolysis was performed under the optimum parameters of saccharification. Reducing sugars released was analyzed by 3,5-dinitrosalicylic acid (DNS) method and expressed as % w/w of C. forskohlii root powder. Residual glucose was detected by glucose oxidase-peroxidase (GOD-POD) kit [35]. 2.4. Detoxification of C. forskohlii root hydrolysates Two pretreatments viz. overliming and passing through hydrophobic polymeric resin, were evaluated for detoxification of C. forskohlii root hydrolysate. In the pretreatment by overliming, the hydrolysate was placed on a magnetic stirrer and heated to 50  C. Ca(OH)2 was added gradually and mixed by using magnetic stirrer until the pH reached 9 to 10. The hydrolysate was then maintained at 50  C for 30 min [36]. The hydrolysate was vacuum filtered to remove the recovered solids (CaSO4). The pH of the overlimed filtrate was adjusted to 6.5 with 0.1 N HCl and used for ABE fermentation. Amberlite XAD-4 resin was obtained from Sigma Aldrich (Mumbai, India). Amberlite XAD resins are nonionic macroreticular cross linked polymers of styrene divinyl benzene with a mean pore diameter of approximately 140 Å. Amberlite XAD-4 (1.5:1) was added to the hydrolysate and allowed to equilibrate at room temperature (28 ± 2  C) for 3 h. The resin-hydrolysate slurry was then filtered through Whatman #1 filter paper. The filtered hydrolysate was then adjusted to pH 6.5 with 0.1 N HCl and used for ABE fermentation [37]. A combined detoxification wherein the root hydrolysate was first treated with lime and then followed by Amberlite XAD-4 resin was also carried out. 2.5. Determination of total phenolic content of C. forskohlii root hydrolysate The amount of total phenolics before and after the pretreatment of C. forskohlii root hydrolysates was determined with the Folin-

596

S.M. Harde et al. / Renewable Energy 86 (2016) 594e601

Ciocalteu reagent. Gallic acid was used as a standard (0.05e0.5 mg/ ml) and the total phenolics were expressed as g/l. The hydrolysates (0.2 ml) were mixed with 10-fold diluted Folin-Ciocalteu reagent (1 ml) and 20% sodium carbonate (0.8 ml). The tubes containing the samples were covered with parafilm and allowed to stand for 30 min in a dark at room temperature (28 ± 2  C). Distilled water (3 ml) was added and the absorbance was measured spectrophotometrically at 760 nm [38]. 2.6. Microorganism and inoculum preparation C. acetobutylicum NCIM 2877 was obtained from NCIM, Pune, India (National Collection of Industrial Microorganisms). Initially, sporulated cells were activated by heat shock at 80  C for 10 min. The activated spore culture (2.5 ml) was inoculated in 100 ml sterile Reinforced Clostridial Medium (RCM) in 125 ml air tight, anaerobic glass bottles and grown for 20 h at 37  C. After 20 h, the prepared inoculum (5% v/v) was used for batch experiments. 2.7. Medium The medium reported by Tripathi et al. [39] was used as a production medium and RCM for the inoculum preparation. The inoculum medium (RCM) contained (in g/l) meat extract 10, peptone 5, yeast extract 3, glucose 5, starch 1, sodium chloride 5, sodium acetate 3, and L-cysteine hydrochloride 0.5 (pH 6.8 ± 0.2). The standard production medium contained (in g/l) glucose 60, magnesium sulfate 0.2, sodium chloride 0.01, manganese sulfate 0.01, iron sulfate 0.01, potassium dihydrogen phosphate 0.5, potassium hydrogen phosphate 0.5, ammonium acetate 2.2, biotin 0.01, thiamin 0.1, and p-aminobenzoic acid 0.1. C. forskohlii root hydrolysates medium was supplemented with all the standard production medium constituents except glucose. The medium was adjusted to pH 6.5 with 0.1 N HCl. After preparation, the medium was purged with nitrogen and autoclaved at 105 Pa (121  C) for 20 min. 2.8. Batch fermentation Batch fermentations were carried out in 125 ml air tight, anaerobic glass bottles with 50 ml volume of production medium. Three different hydrolysates were used for ABE fermentation viz. enzyme hydrolysates (glucose equivalent to 31.46 g/l; made to 50 g/ l with synthetic glucose and; concentrated to 50 g/l). All the hydrolysates were pretreated with overlimimg, resin, and a combination of overliming and resin (Section 2.4) prior to ABE fermentation. It was purged with nitrogen, autoclaved at 105 Pa (121  C) for 20 min. It was inoculated (5% v/v) with 20 h actively growing seed culture and incubated for 96 h at 37  C. All experiments were performed on the basis of glucose content of the root hydrolysates. Samples were drawn at regular intervals and analyzed for solvents and acids using gas chromatography. Residual glucose was detected by a glucose oxidase-peroxidase (GOD-POD) kit [35]. 2.9. Analytical method The solvents and acids were quantified by using gas chromatography as described by Survase et al. [22]. The gas chromatograph (Hewlett Packard series 6890) equipped with a flame ionization detector and DB-WAXetr capillary column (30 m  0.32 mm  1 mm) was used. The injector temperature was 200  C and detector temperature was 250  C. The injector volume was 10 ml. Residual glucose was detected by a glucose oxidase-peroxidase (GOD-POD) kit [35].

3. Results and discussion 3.1. Carbohydrate determination of C. forskohlii roots The samples were analyzed for proximate composition as the raw material influence the bioconversion process which further decides the approaches to be followed for the processing of the material. The carbohydrate profiling of C. forskohlii roots [40] showed that it to be a rich source of starch (18e20%), cellulose (14e15%), hemicelluloses (6e7%) and pectin (3e4%). This suggested the possible use of root biomass for production of biobutanol. 3.2. Optimization of parameters for enzymatic saccharification of C. forskohlii roots Enzymatic hydrolysis offers more specific hydrolysis of samples under mild operating conditions. Hence it is the most useful approach for production of biobutanol. 3.2.1. Optimization of parameters for enzymatic hydrolysis of C. forskohlii roots using Stargen® 002 system The optimum pH for maximum release of reducing sugars was studied with varying pH from 3.5, 4.0, 4.5, 5.0 and 5.5 (Fig. 1a) with constant temperature, substrate loading and enzyme concentration at 50  C, 10% w/v and 16 U/g dry weight of C. forskohlii roots, respectively. The maximum reducing sugars (% w/w of substrate) of 20.18 ± 0.15% w/w was released at pH 4.5 beyond which no further increase in reducing sugars was observed. Therefore pH 4.5 was considered as the optimum pH and used for further experiments. The starch breakdown was carried out on 10% w/v of the substrate for 6 h using pH 4.5 at temperature ranging from 30  C to 70  C with enzyme concentration of 16 U/g dry weight of C. forskohlii roots (Fig. 1b). A maximum release of reducing sugars of 20.22 ± 0.18% w/w was achieved at 50  C. Hence, 50  C was selected as the optimum saccharification temperature for further experiments. Stargen® 002 activity was estimated in our laboratory and was found to be 2000 U/ml. Variable levels of Stargen® 002 (4e16 U/g dry weight of C. forskohlii roots) and incubation time (6e30 h) with 10% w/v substrate loading and pH 4.5 were optimized to get maximum release of reducing sugars. An enzyme concentration of 16U found to be beneficial for reducing sugar yield of 33.75% w/w with an optimum incubation time of 24 h beyond which no further increase in reducing sugars was observed (Fig. 1c). Thus 16U of Stargen® 002 and incubation time of 24 h resulted in optimum starch hydrolysis. The yield of reducing sugar of samples was comparable with 5% w/v with an increase in substrate load to 10% w/v (Fig. 1d). The yield of reducing sugars decreased further with an increase in substrate to 25% w/v. Therefore, 10% w/v substrate load was considered to be optimum for enzyme hydrolysis of starch with reducing sugar yield of 32.8 ± 0.35% w/w (Fig. 1d). At high substrate concentration, the viscosity of the medium increased which could have reduced the interaction of enzyme with the substrate. The initial free sugar content of biomass was used as control value to compare the further increase in reducing sugars after changing different parameters of hydrolysis. 3.2.2. Optimization of parameters for enzymatic hydrolysis of C. forskohlii roots using Accellerase® 1500 system The optimum pH for maximum release of reducing sugars was studied with varying pH from 3.5, 4.0, 4.5, 5.0 and 5.5 (Fig. 2a) with constant temperature, substrate loading and enzyme concentration at 50  C, 10% w/v and 30 FPU/g dry weight of C. forskohlii roots, respectively. The maximum reducing sugars (% w/w of substrate) of

Reducing sugar (% w/w of substrate)

25

(a)

20

15

10

5

Reducing sugar (% w/w of substrate))

S.M. Harde et al. / Renewable Energy 86 (2016) 594e601 40

(c)

35 30

25 20 15 10 5 0

6

0 control

3.5

4

4.5

5

12

18 Time (h)

5.5

4 U/g

pH

(b)

20

15

10

5

8 U/g

12 U/g

24

30 16 U/g

35 Reducing sugar (% w/w of substrate)

25

Reducing sugar (% w/w of substrate)

597

(d)

30

25 20 15 10 5

0

0 control

30

40

50

60

70

Temperature (˚C)

control

5

10 15 20 Substrate loading (% w/v)

25

Fig. 1. Optimization of parameters for an enzymatic hydrolysis of starch in the spent biomass of C. forskohlii roots using Stargen® 002 (a) pH (b) Temperature (c) Stargen® 002 (U/g of substrate) concentration and incubation time (d) Substrate loading.

14.12 ± 0.24% w/w was released at pH 4.5 beyond which no increase in reducing sugars was observed. Therefore pH 4.5 was considered as the optimum pH and used for further experiments. The initial free sugar content of biomass was used as control value to compare the further increase in reducing sugars after changing different parameters of hydrolysis. The cellulose breakdown was carried out on 10% w/v of the substrate for 16 h using pH 4.5 at temperature ranging from 30  C to 60  C (Fig. 2b). A maximum release of reducing sugars of 14.22 ± 0.41% w/w was achieved at 50  C. Hence, 50  C was selected as the optimum saccharification temperature for further experiments. The effect of reaction time was investigated by varying it from 8 to 40 h. The yield of reducing sugar increased progressively with time up to 24 h, and maximum reducing sugar released was found to be 17.89 ± 0.34% w/w (Fig. 2c). Accellerase® 1500 activity was estimated in our laboratory and was found to be 1500 FPU/ml. Variable levels of Accellerase® 1500 (30e120 FPU/g dry weight of C. forskohlii roots) were optimized to get maximum release of reducing sugars. An enzyme concentration of 60 FPU found to be beneficial for reducing sugar yield of 21.22 ± 0.55% w/w beyond which no further increase in reducing sugars was observed (Fig. 2d). Thus 60 FPU of Accellerase® 1500 and incubation time of 24 h were selected as the optimum incubation time for further experiments. The yield of reducing sugar increased with an increase in substrate load from 5% w/v to 10% w/v. Further increase in substrate concentration to 25% w/v decreased the yield of reducing sugars. Hence, 10% w/v substrate load was considered to be the optimum for enzyme hydrolysis of cellulose with reducing sugar yield of 21.76 ± 0.43% w/w (Fig. 2e). Combination of Stargen® 002 and Accellerase® 1500 under optimized conditions could produce 41.2 ± 0.12 g/l of total reducing sugars synergistically (Fig. 3). Glucose content of these hydrolysates was found to be 32.23 ± 1.33 g/l. All the fermentation experiments were performed on the basis of glucose content of the root hydrolysates. The synergistic use of both the enzymes for hydrolysis of root proved to be better than the individual enzymes.

3.3. Batch fermentation ABE fermentation of enzymatic hydrolysates of C. forskohlii roots (glucose equivalent to 31.46 g/l) was performed by using C. acetobutyllicum NCIM 2877. It was found that fermentation could produce only 0.55 g/l of total solvents and 4.29 g/l of total acids as compared to 6.95 g/l of total solvents and 1.59 g/l of total acids in the control production medium containing only glucose. The concentration of probable inhibitors i.e. phenolic compounds in root hydrolysates may be a possible reason for the low production of solvent. C. forskohlii Briq is a herbal plant belonging to the Lamiaceae family. This plant is known to contain higher phenolic contents as compared to other Lamiaceae species. The plant extract with higher phenolic compounds are reported to be antimicrobial [41]. Other investigators have also reported the extracts from these plants to be rich in phenolic compounds [42]. After extraction of forskolin, these phenolic compounds might remain in the roots which could be released in the hydrolysate medium after enzymatic saccharification. Therefore, phenolic content in the enzyme hydrolysates were quantified by using Folin-Ciocalteu method. Total phenolic content of root hydrolysates was found to be 10.95 g/l. This could have a significant effect on the ABE fermentation. The Folin-Ciocalteu method does not analyze any specific phenolic compounds but is based on the reducing power of phenolic hydroxyl groups [43] and detects total phenols with varying sensitivity. Therefore, pretreatment of enzyme hydrolysates to remove phenolic compounds before ABE fermentation was thought to be worthwhile. Phenolic content of the root hydrolysates did decrease from 10.95 g/l to 3.65 g/l and 1.20 g/l after Ca(OH)2 pretreatment and resin (Amberlite XAD-4) pretreatment, respectively. Resin treatment followed by Ca(OH)2 pretreatment could reduce phenolic content from 10.95 g/l to 0.78 g/l. ABE fermentation after detoxification could increase the total ABE concentration from 0.55 g/l to 3.76 g/l (Fig. 4a). The yield obtained was 0.24 g/g as compared to that of 0.34 g/g obtained from the control. Therefore, it was found that overliming and resin pretreatment were essential

S.M. Harde et al. / Renewable Energy 86 (2016) 594e601

Reducing sugar (% w/w of substrate)

16

20

Reducing sugar (% w/w of substrate)

598

(a)

14 12 10

8 6 4 2

16 14 12 10

8 6 4

2

0 control

3.5

4

4.5

5

(c)

18

0

5.5

8

16

16

(b)

14 12 10

8 6

4 2 0

control

30

40

32

40

50

60

25

d)

20 15 10 5 0 control

Temperature (˚C)

Reducing sugar (% w/w of substrate)

24 Time (h)

Reducing sugar (% w/w of substrate))

Reducing sugar (% w/w of substrate)

pH

30

60

90

120

Accellerase concentration (FPU/g of substrate)

25

(e)

20

15 10 5

0 5

10

15

20

25

Substrate loading (% w/v)

Reducing sugar (% w/w of substrate)

Fig. 2. Optimization of parameters for an enzymatic hydrolysis of cellulose in the spent biomass of C. forskohlii roots using Accellerase® 1500 (a) pH (b) Temperature (c) Incubation time (d) Accellerase® 1500 (FPU/g of substrate) concentration (e) Substrate loading.

45

40 35 30

25 20 15

10 5 0

Control

Stargen

Accellerase

Cocktail

Enzyme used Fig. 3. Individual and cocktail effect of enzymes on starch and cellulose hydrolysis in the spent biomass of C. forskohlii roots.

and suitable for ABE fermentation of C. forskohlii root hydrolysates. Roberto et al. [44] reported overliming to be both efficacious and cost-effective method in removing phenolic compounds. The detoxifying effect of overliming is due to both the precipitation of toxic components and to the instability of some inhibitors at high pH [45]. Palmqvist et al. [45] observed the pre-adjustment to pH 10 with NaOH and Ca(OH)2 to decrease the concentration of Hibbert's

ketones in a dilute acid hydrolysate of spruce from by 22% and that of both furfural and HMF by 20%. In the next experiment, pretreated [Amberlite XAD4 þ Ca(OH)2] enzyme hydrolysates (made to glucose equivalent of 50 g/l with synthetic glucose) was used for ABE fermentation. ABE production from this hydrolysate was found to be 5.32 g/l as compared to 8.09 g/l in standard production medium (Fig. 4b). The ABE yield from this hydrolysate was 0.20 g/g as compared to 0.29 g/g obtained from standard production medium. The pretreated [(Amberlite XAD-4 þ Ca(OH)2] concentrated enzyme hydrolysate (glucose equivalent to 50 g/l) was used for ABE fermentation. ABE production from this hydrolysate was found to be 5.14 g/l as compared to 8.09 g/l in standard production medium (Fig. 4c). Fig. 5 shows a representative HPLC chromatogram of ABE analysis. The ABE yield from this hydrolysate was 0.21 g/g as compared to 0.29 g/g obtained in standard production medium. Table 1 represents the ABE production from different substrates [46e50]. The ratio of butanol, acetone and ethanol production was found to be 6:3:1 which is accordance with the stoichiometric conversion of glucose into solvents. Gapes [51] reported the stoichiometric equations of ABE production. Detoxification of lignocellulosic hydrolysates has been reported by using ionexchange resins [52]. Buchert et al. [53] reported the detoxification of birch wood hemicellulosic hydrolysate using a cation exchanger prior to fermentation with Gluconobacter oxydans.

S.M. Harde et al. / Renewable Energy 86 (2016) 594e601

599

Fig. 4. The effect of pretreatments of hydrolysate obtained from the spent biomass of C. forskohlii roots on the overall solvent and acid formation in batch culture of C. acetobutylicum NCIM 2877 (a) Glucose equivalent to 30 g/l (b) Glucose equivalent to 50 g/l (made with synthetic glucose) (c) Glucose concentrated to 50 g/l.

Fig. 5. HPLC chromatogram of ABE analysis.

Overliming in combination with the mixed-bed ion resin was used for improving the fermentability of a hemicellulose hydrolysate of red oak with the yeast Pichia stipitis [54]. A bagasse hydrolysate was treated by a combination of anion and cation exchangers prior

to fermentation with Pachysolen tannophilus and found increase in ethanol production [55]. Treatment with ion-exchange resin was also found to decrease the toxicity of a waste paper hydrolysate [56].

600

S.M. Harde et al. / Renewable Energy 86 (2016) 594e601

Table 1 Production of ABE from various feedstock. Microorganism

Substrate

Total ABE yield (g/g)

Reference

C. acetobutylicum NCIM 2877 C. acetobutylicum NCIM 2337 C. acetobutylicum MTCC 481 C. acetobutylicum ATCC 824 C. beijerinkii ATCC 55025 C. beijerinckii BA101

Coleus forskohlii roots Rice straw Rice straw Marine macroalga þ mannitol Wheat bran Starch

0.21 0.52 0.39 0.16 0.54 0.41

This work [45] [46] [47] [48] [49]

4. Conclusion C. forskohlii root hydrolysate was successfully utilized for batch fermentation of acetone, n-butanol, and ethanol (ABE) using cells of C. acetobutylicum NCIM 2877. Enzyme hydrolysates without pretreatment could produce only 0.55 g/l of total solvents. Pretreatment of C. forskohlii root hydrolysates (CaOH2 and Amberlite XAD4) increased the production of total solvents to 5.33 g/l. This method can be potentially useful in utilizing C. forskohlii root hydrolysate as a substrate for ABE production, and could be undertaken by industrial houses processing the said roots for extraction of forskolin. This work also contributes towards efficient utilization of renewable agro-industrial waste as a bioresource for ABE production. References [1] V. Shah, Cultivation and Utilization of Medicinal Plants (Supplement), RRL and CSIR, Jammu e Tawai, 1996, pp. 385e411. [2] Anonymous. http://www.samilabs.com/services/farming/2012; Accessed on 22 June, 2012. [3] R. Singh, S.P. Gangwar, D. Singh, R. Singh, R. Pandey, A. Kalra, Medicinal plant Coleus forskohlii Briq.: disease and management, Med. Plants 3 (2011) 1e7. [4] M. Suryanayanan, J.S. Pai, Studies in micropropagation of Coleus forskohlii, J. Med. Aromat. Plant Sci. 20 (1998) 379e382. [5] L. Tao, A. Aden, The economics of current and future biofuels, In Vitro Cell Dev. Biol. Plant 45 (2009) 199e217. [6] L. Lin, Z. Cunshan, S. Vittayapadung, S. Xiangqian, D. Mingdong, Opportunities and challenges for biodiesel fuel, Appl. Energy 88 (2011) 1020e1031. [7] J. Yan, T. Lin, Biofuels in Asia, Appl. Energ 86 (2009) 1e10. [8] A. Zhou, E. Thomson, The development of biofuels in Asia, Appl. Energ 86 (2009) 11e20. [9] P.H. Pfromm, V.A. Boadu, R. Nelson, P. Vadlani, R. Madl, Bio-butanol vs. bioethanol: a technical and economic assessment for corn and switchgrass fermented by yeast or Clostridium acetobutylicum, Biomass Bioenerg. 34 (2010) 515e524. [10] G. Jurgens, S. Survase, O. Berezina, E. Sklavounos, J. Linnekoski, A. Kurkij€ arvi, €keva €, A. van Heiningen, T. Granstro €m, Butanol production from lignoM. Va cellulosics, Biotecnol. Lett. 34 (2012) 1415e1434. €m, Biobutanol: the outlook of [11] S.B. Bankar, S.A. Survase, H. Ojamoa, T. Granstro an academic and industrialist, RSC Adv. 3 (2013) 24734e24757. [12] B.Y. Cheng, F.Y. Chuang, C.L. Lin, J.S. Chang, Deciphering butanol inhibition to Clostridial species in acclimatized sludge for improving biobutanol production, Biochem. Eng. J. 69 (2012) 100e105. [13] C.L. Cheng, P.Y. Che, B.Y. Chen, W.J. Lee, C.Y. Lin, J.S. Chang, Biobutanol production from agricultural waste by an acclimated mixed bacterial microflora, Appl. Energ 100 (2012) 3e9. [14] A. Ranjan, V.S. Moholkar, Biobutanol: science, engineering, and economics, Int. J. Energ Res. 36 (2012) 277e323. [15] L.R. Lynd, C.E. Wyman, T.U. Gerngross, Biocommodity engineering, Biotechnol. Prog. 15 (1999) 777e793. [16] N. Qureshi, B.C. Saha, M.A. Cotta, V. Singh, An economic evaluation of biological conversion of wheat straw to butanol: a biofuel, Energy Convers. Manage 65 (2013) 456e462. [17] N. Qureshi, B.C. Saha, M.A. Cotta, Butanol production from wheat straw hydrolysate using Clostridium beijerinckii, Bioprocess Biosyst. Eng. 30 (2007) 419e427. [18] Z. Liu, Y. Ying, F. Li, C. Ma, P. Xu, Butanol production by Clostridium beijerinckii ATCC 55025 from wheat bran, J. Ind. Microbiol. Biotechnol. 37 (2010) 495e501. [19] N. Qureshi, B.C. Saha, B. Dien, R.E. Hector, M.A. Cotta, Production of butanol (a biofuel) from agricultural residues: part IeUse of barley straw hydrolysates, Biomass Bioenerg. 34 (2010) 559e565. [20] N. Qureshi, B.C. Saha, R.E. Hector, B. Dien, S. Hughes, S. Liu, L. Iten, M.J. Bowman, G. Sarath, M.A. Cotta, Production of butanol (a biofuel) from agricultural residues: part IIeuse of corn stover and switchgrass hydrolysates,

Biomass Bioenerg. 34 (2010) 566e571. [21] N. Qureshi, T.C. Ezeji, J. Ebener, B.S. Dien, M.A. Cotta, H.P. Blaschek, Butanol production by Clostridium beijerinckii Part I: use of acid and enzyme hydrolyzed corn fiber, Bioresour. Technol. 99 (2008) 5915e5922. € m, [22] S.A. Survase, E. Sklavounos, G. Jurgens, A. van Heiningen, T. Granstro Continuous acetoneebutanoleethanol fermentation using SO2eethanolewater spent liquor from spruce, Bioresour. Technol. 102 (23) (2011) 10996e11002.  , M. Ballesteros, M.J. Negro, Pretreatment technologies [23] P. Alvira, E. Tom as-Pejo for an efficient bioethanol production process based on enzymatic hydrolysis: a review, Bioresour. Technol. 101 (13) (2010) 4851e4861. [24] Y. Sun, J. Cheng, Hydrolysis of lignocellulosic materials for ethanol production: a review, Bioresour. Technol. 83 (2002) 1e11. [25] E. Palmqvist, B. Hahn-Hagerdal, Fermentation of lignocellulosic hydrolysates II: inhibitors and mechanisms of inhibition, Bioresour. Technol. 74 (2000a) 25e33. [26] W.D. Carvalho, L. Canilha, S.I. Mussatto, G. Dragone, M.L.V. Morales, A.I.N. Solenzal, Detoxification of sugarcane baggase hemicellulosic hydrolysate with ion exchange resins for xylitol production by calcium alginate entrapped cells, J. Chem. Tech. Biotechnol. 79 (2004) 863e868. [27] S. Larsson, A. Reimann, N.O. Nilvebrant, L.J. Josson, Comparison of different methods for the detoxification of lignocelluloses hydrolyzates of spruce, Appl. Biochem. Biotechnol. 91 (1999) 77e79. [28] S.I. Mussatto, I.C. Roberto, Alternatives for detoxification of diluted acid lignocellulosic hydrolyzates for use in fermentative processes, Bioresour. Technol. 93 (2004) 1e10. [29] E. Palmqvist, B. Hahn-Hagerdal, Fermentation of lignocellulosic hydrolysates I: inhibition and detoxification, Bioresour. Technol. 74 (2000b) 17e24. [30] Z. Sun, S. Liu, Production of n-butanol from concentrated sugar maple hemicellulosic hydrolysate by Clostridia acetobutylicum ATCC824, Biomass Bioenergy 39 (2012) 39e47. [31] AOAC, Official Methods of Analysis, twelth ed., Association of Official Analytical Chemists, Washington DC, 1975. [32] A.L. Merrill, B.K. Watt, Energy Value of Foods-basis and Derivation. Agriculture Handbook No. 74, U.S. Government Printing Office, Washington, 1973, p. 105. [33] W.C. Monte, J.A. Maga, Extraction and isolation of soluble and insoluble fiber fractions from the pinto bean (Phaseolus vulgaris), J. Agric. Food Chem. 28 (1980) 1169e1174. [34] S.P. Ng, C.P. Tan, O.M. Lai, K. Long, H. Mirhosseini, Extraction and characterization of dietary fiber from coconut residue, J. Food Agric. Environ. 8 (2010) 172e177. [35] R. Sathish, R. Madhavan, H.R. Vasanthi, A. Amuthan, In-vitro alpha-glucosidase inhibitory activity of abraga chendhooram, a siddha drug, Int. J. Pharmacol. Clin. Sci. 1 (3) (2012) 79e81. [36] A. Mohagheghi, M. Ruth, D.J. Schell, Conditioning hemicellulose hydrolysates for fermentation: effects of overliming pH on sugar and ethanol yields, Process Biochem. 41 (2006) 1806e1811. [37] J.Y. Zhu, W. Zhu, P. OBryan, B.S. Dien, S. Tian, R. Gleisner, X.J. Pan, Ethanol production from SPORL-pretreated lodgepole pine: preliminary evaluation of mass balance and process energy efficiency, Appl. Microbiol. Biotechnol. 86 (2010) 1355e1365. [38] S. Maurya, D. Singh, quantitative analysis of total phenolic content in adhatoda vasica nees extracts, Int. J. Pharm. Tech. Res. 2 (2010) 2403e2406. [39] A. Tripathi, H. Sami, S.R. Jain, M. Viloria-Cols, N. Zhuravleva, G. Nilsson, H. Jungvid, A. Kumar, Improved bio-catalytic conversion by novel immobilization process using cryogel beads to increase solvent production, Enzyme Microb. Technol. 47 (2010) 44e51. €m, R.S. Singhal, S.A. Survase, [40] S.M. Harde, S.B. Bankar, H. Ojamo, T. Granstro Continuous lignocellulosic ethanol production using Coleus forskohlii root hydrolysate, Fuel 126 (2014) 77e84. [41] N.G. Baydar, G. Ozkan, O. Sadic, Toal phenolic contents and antimicrobial activities of grape (Vitis vinifera L.) extracts, Food Control. 15 (2004) 335e339. [42] K. Sunitha, K. Bhaskara chary, C.C. Nimgulkar, B. Dinesh kumar, D. Manohar rao, Identification, quantification and antioxidant activity of secondary metabolites in leaf and callus extracts of coleus forskohlii, Int. J. Pharm. Bio Sci. 4 (3) (2013) 1139e1149. [43] D.H. Hahn, L.W. Rooney, C.F. Earp, Tannins and phenols of sorghum, Cereal Food World 29 (1984) 776e779. [44] I.C. Roberto, M.G.A. Felipe, L.S. Lacis, S.S. Silva, I.M. de Mancilha, Utilization of sugar cane bagasse hemicellulosic hydrolyzate by Candida guilliermondii for xylitol production, Bioresour. Technol. 36 (1991) 271e275.

S.M. Harde et al. / Renewable Energy 86 (2016) 594e601 [45] E. Palmqvist, B.H. Hagerdal, Fermentatiom of lignocellulosic hydrolysates. I: inhibition and detoxificaton, Bioresour. Technol. 74 (2000) 17e24. [46] A. Ranjan, S. Khanna, V.S. Moholkar, A Corrigendum to “Feasibility of rice straw as alternative substrate for biobutanol production by Ranjan et al.” [Appl. Energy, 103 (2013) 32e38], Appl. Energ 116 (2014) 436e438. [47] A. Ranjan, R. Mayank, V.S. Moholkar, Process optimization for butanol production from developed rice straw hydrolysate using Clostridium acetobutylicum MTCC 481 strain, Biomass Conv. Bioref 3 (2013) 143e155. [48] M.H. Huesemann, L.J. Kuo, L. Urquhart, G.A. Gill, G. Roesijadi, Acetone-butanol fermentation of marine macroalgae, Bioresour. Technol. 108 (2012) 305e309. [49] Z. Liu, Y. Ying, F. Li, C. Ma, P. Xu, Butanol production by Clostridium beijerinckii ATCC 55025 from wheat bran, J. Ind. Microbiol. Biotechnol. 37 (2010) 495e501. [50] T.W. Jesse, T.C. Ezeji, N. Quereshi, H.P. Blaschek, Production of butanol from starch-based waste packing peanuts and agricultural waste, J. Ind. Microbiol. Biotechnol. 29 (2002) 117e123. [51] J.R. Gapes, The economics of acetone-butanol fermentation: theoretical and

601

market considerations, J. Mol. Microbiol. Biotechnol. 2 (2000) 27e32. [52] L. Olsson, B. Hahn-H€ agerdal, Fermentation of lignocellulosic hydrolysates for ethanol production, Enzyme Microb. Technol. 18 (1996) 312e331. €, J. Puls, K. Poutanen, Improvement of the fermentability [53] J. Buchert, K. Niemela of steamed hemicellulose hydrolysate by ion exclusion, Proc. Biochem. 25 (1990) 176e180. [54] A.V. Tran, R.P. Chambers, Ethanol fermentation of red oak prehydrolysate by the yeast Pichia stipitis, Enzyme Microb. Technol. 8 (1986) 439e444. [55] N.E. Watson, B.A. Prior, P.M. Lategan, M. Lussi, Factors in acid treated bagasse inhibiting ethanol production from D-xylose by Pachysolen tannophilus, Enzyme Microb. Technol. 6 (1984) 451e456. [56] C.J. Rivard, R.E. Engel, T.K. Hayward, N.J. Nagle, C. Hatzis, G.P. Philippidis, Measurement of the inhibitory potential and detoxification of biomass pretreatment hydrolysate for ethanol production, Appl. Biochem. Biotechnol. 57e58 (1996) 183e191.