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Kinetic study of batch and fed-batch enzymatic saccharification of pretreatedsubstrate and subsequent fermentation to ethanol

Biotechnology for Biofuels 2012, 5:16 doi:10.1186/1754-6834-5-16

Rishi Gupta ([email protected])Sanjay Kumar ([email protected])

James Gomes ([email protected])Ramesh Chander Kuhad ([email protected])

ISSN 1754-6834

Article type Research

Submission date 10 January 2012

Acceptance date 20 March 2012

Publication date 20 March 2012

Article URL http://www.biotechnologyforbiofuels.com/content/5/1/16

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Kinetic study of batch and fed-batch enzymatic saccharification of pretreated substrate and subsequent fermentation to ethanol

Rishi Gupta,Aff1 Email: [email protected]

Sanjay Kumar,Aff2 Email: [email protected]

James Gomes,Aff2 Email: [email protected]

Ramesh Chander Kuhad,Aff1 Corresponding Affiliation: Aff1 Email: [email protected]

Aff1 Lignocellulose Biotechnology Laboratory, Department of Microbiology, University of Delhi South Campus, New Delhi 110021, India

Aff2 Kusuma School of Biological Sciences, Indian Institute of Technology Delhi, New Delhi 110016, India

Abstract

Background

Enzymatic hydrolysis, the rate limiting step in the process development for biofuel, is always hampered by its low sugar concentration. High solid enzymatic saccharification could solve this problem but has several other drawbacks such as low rate of reaction. In the present study we have attempted to enhance the concentration of sugars in enzymatic hydrolysate of delignified Prosopis juliflora, using a fed-batch enzymatic hydrolysis approach.

Results

The enzymatic hydrolysis was carried out at elevated solid loading up to 20% (w/v) and a comparison kinetics of batch and fed-batch enzymatic hydrolysis was carried out using kinetic regimes. Under batch mode, the actual sugar concentration values at 20% initial substrate consistency were found deviated from the predicted values and the maximum sugar concentration obtained was 80.78 g/L, which on using fed-batch strategy was implemented to enhance the final sugar concentration to 127 g/L. The batch and fed-batch enzymatic hydrolysates were fermented with Saccharomyces cerevisiae and ethanol production of 34.78 g/L and 52.83 g/L, respectively, were achieved. Furthermore, model simulations showed that higher insoluble solids in the feed resulted in both smaller reactor volume and shorter residence time.

Conclusion

Fed-batch enzymatic hydrolysis is an efficient procedure for enhancing the sugar concentration in the hydrolysate. Restricting the process to suitable kinetic regimes could result in higher conversion rates.

KeywordsEnzymatic hydrolysis, Fed-batch, Kinetic model, Fermentation, Delignified substrate, Bioethanol

BackgroundProduction of cellulosic ethanol from lignocellulosic biomass represents a potential alternative to the petroleum fuel due to its renewable nature and sustainable availability. Currently, the major strategy used for cellulosic ethanol production includes three main steps i.e., biomass pretreatment, enzymatic hydrolysis and ethanol fermentation [1,2]. The enzymatic hydrolysis contributes significantly to the cost of cellulosic ethanol and from the process economics perspective, the improvement in the enzymatic hydrolysis step is a prerequisite [3,4]. The main obstacles for enzymatic hydrolysis are low rate of reaction, high cost of enzyme, low product concentration and lack of understanding of cellulase kinetics on lignocellulosic substrates [5,6]. One way to overcome this problem is to operate the enzymatic hydrolysis using high insoluble solid consistency [7-9]. However, the saccharification reaction at high insoluble solid consistency will have to encounter the problems of increased viscosity, higher energy requirement for mixing, shear inactivation of cellulases, and poor heat transfer due to rheological properties of dense fibrous suspension [9,10].

Interestingly in fed-batch enzymatic hydrolysis such problems could be avoided by adding the substrate and/or enzymes gradually to maintain the low level of viscosity [11]. The fed-batch enzymatic saccharification process has several other economic advantages over conventional batch process such as lower capital cost due to reduced volume, lower operating costs and lower down-stream processing cost due to higher product concentration [6,7]. There are several reports on fed-batch enzymatic saccharification which mainly deal with the development of appropriate kinetic models for mechanistic description of the phenomena [9,12,13]. However, the reports on process operation, optimization and control for fed-batch enzymatic saccharification are scarce [14]. Till date, the strategies used for fed-batch enzymatic saccharification are categorized into three main groups i.e., (i) to recycle enzyme; (ii) fed-batch SSF to mitigate inhibitory effect and (iii) fed-batch saccharification to increase the cumulative substrate in a reactor [7]. Here, the present study falls within the third category and our main emphasis was to enhance the total solid content and sugar concentration, which eventually resulted in higher ethanol production.

The experimental data on cellulose hydrolysis by cellulases point to various bottlenecks that decrease the rate of conversion. Mathematical modeling of the enzymatic hydrolysis process is an important tool for analyzing these bottlenecks [5]. Use of mathematical modeling can lead to several advantages viz. the effect of feeding profiles on sugar conversion can be evaluated apriori, kinetics of the hydrolysis process can be studied and process simulations can be made to understand the kinetic regimes. Recently, Hodge and colleagues [7] have used

model based fed-batch approach to develop a feeding profile for the fed-batch enzymatic saccharification, while, Morales-Rodriguez and coworkers [14] used a modeling approach to reduce the amount of enzyme during the fed-batch enzymatic saccharification.

The present study deals with the development of the feeding profile and a mathematical model for the understanding of the enzymatic saccharification kinetics in a stirred tank reactor (STR). Moreover, the hydrolysates obtained after batch and fed-batch enzymatic hydrolysis has subsequently been fermented to ethanol, and an overall comparison between batch and fed-batch process has been presented.

Results

Kinetics of batch and fed-batch enzymatic hydrolysis

A series of batch experiments were performed using the initial substrate concentrations 5%, 10%, 15% and 20% (w/v). Then applying the kinetic model the rate constants of cellulose hydrolysis, ki (i=1–4) was calculated in each case. It was observed that the rate constant for enzymatic hydrolysis decreases with an increase in the initial insoluble solid concentration (Figure 1). However, the ki values have shown good correlation with the initial substrate consistencies used for enzymatic hydrolysis with a regression coefficient R2 of~0.9.

Figure 1 Plot between substrate concentrations versus time for the enzymatic saccharification of delignified lignocellulosic biomass at different initial substrate consistencies

The maximum rate constant (k1=0.0421 h-1) was obtained when the hydrolysis was carried out with 5% initial substrate concentration. The rate constants ki were then validated using the glucose concentration measurements obtained during the hydrolysis experiments. The root mean square error (RMSE) values between the predicted and the experimental values for enzymatic saccharification carried out at 5, 10, 15 and 20% initial substrate consistency were 0.997, 0.779, 1.843 and 1.995, respectively (Figure 2 a-d). The results also indicated that the maximum deviation of the experimental data from the model prediction was observed when the enzymatic saccharification was carried out at 20% initial substrate consistency. Moreover, the experimental values also depicted that the sugar concentration increased significantly only upto 15% substrate consistency and declined thereafter at 20% substrate level (Figure 2 a-d). The maximum sugar concentration obtained at each substrate concentration were 41.10 g/L (S1,0 =5%), 72.47 g/L (S2,0 =10%), 90.07 g/L (S3,0 =15%) and 80.05 g/L (S4,0 =20%) (see Figure 2 a-d).

Figure 2 Plots between the actual and the predicted values of glucose concentration released during the enzymatic hydrolysis of delignified lignocellulosic biomass at 5% (A), 10% (B), 15% (C) and 20% (D) initial substrate consistencies

Kinetics of fed-batch enzymatic hydrolysis

The kinetic parameters determined from the batch experiments was used to simulate the hydrolysis profile during the fed-batch enzymatic hydrolysis. Fed-batch hydrolysis was performed employing discrete feeding policy. Insoluble solid substrate concentration (50 g) was added at 24, 56 and 80 h. The insoluble solid concentration was measured at 4 h

intervals. A similar pattern of pulse responses were observed in both the experimental and predicted values for every instance of addition of 50 g feed to insoluble substrate (Figure 3). However, the final insoluble substrate concentrations for both the predicted and experimental values were 58.52 and 65.59 g/L, respectively (Figure 3).

Figure 3 Plot between the experimental versus predicted insoluble solids during the fed-batch enzymatic hydrolysis

Comparison between batch and fed-batch enzymatic hydrolysis

A comparison between batch and fed-batch enzymatic hydrolysis was made to determine which had a higher cellulose conversion. The results showed that in fed-batch operation at Sc=20%, the remaining insoluble solids in the reaction slurry was 65.59 g/L. While in contrast, during the batch enzymatic hydrolysis at S4,0=20% (w/v), the concentration of remaining insoluble solid was 107.29 g/L. The time profiles of glucose concentration and cellulose conversion levels for both batch (S4,0=20%) and fed-batch (Sc=20%) were plotted in Figure 4 a - b. The final sugar concentration for batch and fed-batch were 80.78 g/L and 127. 0 g/L (Figure 4 a), while the cellulose conversion was 40.39% and 63.56%, respectively (Figure 4 b). These results showed that intermittent addition of solids in a repeated fed-batch mode, resulted in better conversion compared to the addition of an equal amount once at the beginning of a batch.

Figure 4. Comparative profiling of batch and fed-batch enzymatic saccharification (A) glucose production, (B) cellulose conversion

Simulation of kinetic model

The kinetic parameters determined from the batch experiments were used to simulate different feeding policies. These simulations provide an insight about the operational protocol that may be implemented to obtain the best hydrolysis results. The simulation of discussed kinetic model under the fed-batch optimization approach has been shown in Figure 5 and 6. The Figure 5 depicted four different feeding policies developed from simulations with the target cumulative insoluble solids in the reactor as 20%. The simulation results showed that the cumulative insoluble solid concentration increases with time and saturates at different final values depending on the initial feed concentration at dilution rates of 0.1-0.4 h-1 (Figure 5). The results indicated that long residence times are required to reach these higher solids levels, when solids were controlled at 5% or lower (Figure 5). While, the simulation results in Figure 6 indicated that higher insoluble solids levels in the feed resulted in both smaller reactor volumes and shorter residence times to achieve a given feeding objective. However, it has been predicted from the simulation results that the feed controlled at 10% initial solid levels resulted in maximum saccharification.

Figure 5 Kinetic simulation profile for cumulative insoluble solids feeding at different initial substrate concentration (5-15%) at varied dilution rate of (A) 0.1 h-1, (B) 0.2 h-1,(C) 0.3 h-1 and (D) 0.4 h-1

Figure 6 Kinetic simulation profile for the volume of bioreactor at different initial substrate consistency (5-15%) using dilution rate of (A) 0.1 h-1, (B) 0.2 h-1, (C) 0.3 h-1 and (D) 0.4 h-1

Fermentation of enzymatic hydrolysate

The fermentation profiles of batch (S4,0=20%) and fed-batch (Sc=20%) enzymatic hydrolysates containing 76.52±2.82 and 117.35±1.14 g/L initial sugars have been shown in Figure 7 and 8. The fermentation of batch enzymatic hydrolysate brought about the production of 34.78±1.10 g/L ethanol with yield and productivity of 0.45 g/g and 3.16 g/L/h, respectively, after 11 h of incubation (Figure 7). Moreover, the biomass production during the fermentation of batch enzymatic hydrolysate increased till 8 h (1.86±0.04 g/L) and then remained almost constant (Figure 7). While, the fed-batch enzymatic hydrolysate when fermented with S. cerevisiae, produced 52.83±1.70 g/L ethanol and 4.50±0.004 g/L biomass with an ethanol yield of 0.45 g/g and ethanol productivity of 4.40 g/L/h after 12 h of incubation (Figure 8).

Figure 7 Fermentation profile of batch enzymatic hydrolysate

Figure 8 Fermentation profile of fed-batch enzymatic hydrolysate

DiscussionThe main aim of the present investigation was to achieve high ethanol concentration as the final ethanol concentration in the fermentation broth is critical to make a cost-effective ethanol production process. Since the ethanol concentration is directly proportional to the sugar concentration, hence high concentration sugar syrup is a prerequisite. In the present study, the process modeling consisting of mass balance and kinetic models were used to provide insights into the process performance and to optimize the process for enhanced enzymatic hydrolysis. During the batch saccharification at different consistencies, a regular decrease in the rate constant with increase in the substrate concentration was observed (Figure 1) and the reaction was assumed to be a first order reaction. This decrease in rate may be attributed to the product inhibition, improper heat and mass transfer and the thermal deactivation of enzymes [7, 12]. The difference between the experimental values and those predicted through simulation for our batch experiments at 20% insoluble solid consistency may be attributed to the same reasons (Figure 2d).

To overcome this problem in batch operation, fed-batch enzymatic hydrolysis was implemented. This approach exploits the property of cellulose solubilization during the enzymatic hydrolysis to increase the solid loading to the reactor, which otherwise would be difficult to handle if the entire insoluble solid was added initially. Interestingly, considering the fact that there are two phases present in the slurry, in the present study, the cellulose conversion has been mentioned in terms of g/L of actual liquid present in the slurry, which was a major pit fall in the earlier report [7], who reported the conversion in terms of g/Kg of total slurry. The later was further amended by correcting the measurement of glucose in the liquid phase (which may represent only 80-90% of the total mass of the slurry) for the content of insoluble solids in order to accurately estimate conversion [15].

The present study demonstrated that fed-batch hydrolysis resulted in higher solid saccharification with high saccharification yield. The results in the (Figures 4a and 4b) depicted a final sugar concentration of 127 g/L with~64% cellulose conversion, which was significantly higher than the cellulose conversion at batch operation (S4,0=20%). It is estimated that an increase in solid substrate consistency from 5 to 8% in simultaneous

saccharification and fermentation process (SSF) reduced the process cost by 19% [16]. While according to report by National Renewable Energy Laboratory (NREL), Department of Energy (DOE), US, an increase in solid consistency from 20 to 30% can reduce the minimum ethanol selling price by $0.10/gallon ethanol [17]. Therefore, the high final sugar concentrations obtained in this work may lead to an economically competitive process.

Comparison of the accuracy of the model prediction validated that a well-designed fed-batch approach could be used to allow an STR reactor capable of handling pretreated P. juliflora at below than 10% insoluble solids to operate at cumulative initial insoluble solids as high as the set goal of 20% (Figure 5). Moreover such validation are also in accordance with the earlier reports of Hodge and coworkers [7], according to whom, using fed-batch strategy the STR, was able to achieve very high cumulative solid loading (S4,0=20%), thus improving its working capability. In addition, the model may also be used to determine a fed-batch feeding policy required to maintain proper mixing and temperature control necessary for high cumulative insoluble solids.

The fermentation of the enzymatic hydrolysate obtained from batch and fed-batch operation also indicates the significance of the study. The fermentation of enzymatic hydrolysate from fed-batch operation brought about approximately 50% and 40% increment in the ethanol concentration and the ethanol productivity, respectively. As there have been estimations that by doubling the ethanol concentration from 2.5 to 5%, the energy required to distill a fermentation broth to 93.5% ethanol using conventional distillation techniques can be reduced by 33% [9]. The enhanced ethanol concentration and productivity from fed-batch operation also made the process more industrially realistic.

ConclusionTo produce higher concentration sugar syrup and subsequently the high ethanol concentration, fed-batch enzymatic saccharification was conducted with the pretreated P.juliflora. Through the fed-batch process, the cumulative solid loading (Sc) up to 20% in a stirred tank reactor increased the sugar released by 56% compared to the batch process with an initial insoluble solid loading of 20%. This model used here provided additional insight into the effect of the operational conditions on productivity. This may be refined by including the degree of polymerization of substrate, accessible cellulose fraction, crystallinity of substrate and enzyme adsorption to distinguish the various causes of the decreasing rate of reaction.

Methods

Raw material and chemicals

Prosopis juliflora wood, collected from University of Delhi South Campus, New Delhi, India, was comminuted by a combination of chipping and milling to attain a particle size of 1–2 mm using a laboratory knife mill (Metrex Scientific Instrumentation, Delhi, India). The processed wood of P. juliflora was delignified with 4% sodium chlorite at 120 C for 30 minutes as described earlier [18].

Commercial cellulases and 3,5-di nitro salicylic acid (DNS) were purchased from Sigma (St. Louis, Missouri, U.S.A.). Ethanol was purchased from Merck (Darmstadt, Germany). Rest of the chemicals and media components of highest purity grade were purchased locally.

Microorganism and culture conditions

The yeast Saccharomyces cerevisiae HAU procured from the culture collection of C.C.S. Haryana Agricultural University, Hisar, Haryana, India was maintained on agar slants containing (g/L): glucose, 30.0; yeast extract, 3.0; peptone, 5.0; agar, 20.0 at pH 6.0�±�0.2 and temperature 30oC, as described earlier [1,8]. While the S. cerevisiae inoculum was grown for 24 h at 30oC in a culture medium containing (g/L): glucose, 30.0; yeast extract, 3.0; peptone, 5.0; (NH4)2HPO4, 0.25 at pH 6.0�±�0.2 [1,19]. Cells were cultured to an absorbance of 0.6–0.8 at 600 nm.

Enzymatic hydrolysis

Batch enzymatic hydrolysis

Enzymatic hydrolysis of pretreated substrate was carried out at different substrate consistency (5–20% w/v) in 0.05 M citrate phosphate buffer (pH 5.0) in a 3.0 L stirred tank bioreactor (Scigenics Pvt. Ltd, Chennai, India) fitted with Rushton impellors, heating jacket and heat exchangers for proper agitation and temperature control. Before enzyme loading, slurry was acclimatized by incubating at 50oC at 150 rpm for 2 h. Thereafter, an enzyme (lyophilized) dosage of 22 Filter paper cellulase activity (FPU) /g dry substrate (gds), 68 U �-glucosidase/gds was added to preincubated cellulose slurry, and reaction was continued for 48 h. One percent Tween 80 and 1 mM CuCl2 were also added to facilitate the enzymatic reaction. The samples were withdrawn at regular intervals, centrifuged at 10,000 rpm for 10 min and the supernatants were used for further analysis.

Fed-batch enzymatic hydrolysis

Fed-batch enzymatic saccharification of pretreated substrate was carried out in the same bioreactor with an initial substrate consistency of 5% (w/v) in the suspension. Before enzyme loading, the slurry was acclimatized by incubating at 50oC at 150 rpm for 2 h. Thereafter, an enzyme dosage of 22 FPU/gds and 68 U �-glucosidase/gds, 1% Tween 80 and 1 mM CuCl2 was added to preincubated cellulose slurry. The equal amount of initial substrate and half of the initial enzyme (lyophilized) was added to the enzymatic suspension thrice after 24, 56 and 80 h to get a final substrate concentration of 200 g/L. The samples were withdrawn at regular intervals, centrifuged at 10,000 rpm for 10 min and the supernatant was subjected to sugar estimation. After incubation, the hydrolysate was harvested, centrifuged to remove the un-hydrolyzed residues and the filtrate was used for fermentation studies.

Fermentation of enzymatic hydrolysate

The fermentation studies of both the enzymatic hydrolysates from batch operation (S4,0=20%) and fed-batch operation (Sc=20%) were carried out. The batch and fed-batch enzymatic hydrolysates containing 37 g/L and 120 g/L sugars, respectively, supplemented with 3 g/L yeast extract and 0.25 g/L (NH4)2HPO4, were inoculated with 6% (v/v) S. cerevisiae. The fermentation was carried out at 30oC, 200 rpm and initial pH 6.0±0.2. Aeration of 0.4 vvm

was maintained throughout the study. The pH was adjusted with 2 N HCl and 2 N NaOH. The samples withdrawn were centrifuged at 10,000 rpm for 10 min at 4oC and the cell free supernatant was used for the determination of ethanol produced and sugar consumed.

Kinetics and theoretical aspects of batch and Fed-batch enzymatic hydrolysis

Cellulose conversion is commonly used as a measure of the effectiveness of enzymatic hydrolysis of cellulose. The conversion efficiency (�) is described in terms of cellulose conversion to glucose (G) and the initial cellulose concentration (Si,0), given by

��

��

��

0,

111.1 iS

G (1)

The insoluble solids level can be measured from the change in sugar concentrations relative to the initial cellulose concentration. This change in sugar levels can be related stoichiometrically to the amount of cellulose removed from the solid phase to estimate an insoluble solids level.

11.10,GSS ii � (2)

The equations describing the dynamic changes in Si and G are

iii Sk

dtdS

� (3)

where ki, i = 1 - 4, are the rate constants for different loadings, and

GkSkdtdG

iii � 0,11.1 (4)

These equations can be solved analytically to obtain following relation

� �� tkStG ii � exp111.1)( 0, (5)

Mass balance equation for prediction of fed-batch capabilities

Mass balances were performed on the reaction system to evaluate the fed-batch procedure. A key assumption was that the insoluble solids (Si) are fed at a fixed flow rate (F). The final insoluble solid consistency obtained from the mass balance on insoluble solids at any time point is given by

iiiF SkSSS � (6)

Where S is the final insoluble solid concentration and SF is the concentration of solids fed. The cumulative insoluble solid (Sc) is the sum of the total amount of insoluble solids present initially and the amount of substrate fed to the reactor. It would represent the level of solid

Competing interests The authors declare that they have no competing interests.

Authors’ Contributions RG carried out the experimental work, analyzed the results and drafted the manuscript. SK helped in preparation of kinetic data and in technical checking of manuscript paper. JG designed the kinetic model and discussed the analysis of their results RCK coordinated the overall study and helped to analyze the results and finalize the paper. All authors read and approved the final manuscript.

AcknowledgementsThe authors are thankful to Department of Biotechnology, Government of India, New Delhi, Council of Scientific and Industrial Research, New Delhi, India and University of Delhi, Delhi, for the financial support.

References1. Gupta R, Sharma KK, Kuhad RC: Separate hydrolysis and fermentation (SHF) of Prosopis juliflor, a woody substrate, for the production of cellulosic ethanol by Saccharomyces cerevisia and Pichia stipiti-NCIM 3498. Bioresour Technol 2009, 100:1214–1220.

2. Kuhad RC, Gupta R, Khasa YP: Bioethanol production from lignocellulosics: an overview. In Wealth from waste. Edited by Lal B and Sharma PM. New Delhi, India:Teri press; pp 53–106.

3. Galbe M, Zacchi G: A review of the production of ethanol from softwood. Appl Microbiol Biotechnol 2002, 59:618–628.

4. Lynd LR, Laser MS, Bransby D, Dale BE, Davison B, Hamilton R, et al: How biotech can transform biofuels. Nat Biotechnol 2008, 26:169–172.

5. Bansal P, Hall M, Realf MJ, Lee JH, Bommarius AS: Modeling cellulase kinetics on lignocellulosic substrates. Biotechnol Adv 2009, 27:833–848.

6. Zheng Y, Pan Z, Zhang R, Jenkins BM: Kinetic modeling for enzymatic hydrolysis of pretreated creeping wild ryegrass. Biotechnol Bioeng 2009, 102:1558–1569.

7. Hodge DB, Karim MN, Schell DJ, McMillan JD: Model-based fed-batch for high-solids enzymatic cellulose hydrolysis. Appl Biochem Biotechnol 2009, 152:88–107.

8. Kuhad RC, Gupta R, Khasa YP, Singh A: Bioethanol production from Lantana camar (red sage): Pretreatment, saccharification and fermentation. Bioresour Technol 2010, 10:8348–8354.

9. Chandra RP, Au-Yeung K, Chanis C, Roos AA, Mabee W, Chung PA, Ghatora S, Saddler JN: The influence of pretreatment and enzyme loading on the effectiveness of batch and fed-batch hydrolysis of corn stover. Biotechnol Prog 2011, 27:77–85.

10. Kristensen JB, Felby C, Jorgensen H: Yield-determining factors in high-solids enzymatic hydrolysis of lignocellulose. Biotechnol Biofuel 2009, 2:11. doi:10.1186/1754-6384-2-11.

11. Kuhad RC, Mehta G, Gupta R, Sharma KK: Fed batch enzymatic saccharification of newspaper cellulosics improves the sugar content in the hydrolysates and eventually the ethanol fermentation by Saccharomyces cerevisia. Biomass Bioenergy 2010, 34:1189–1194.

12. Schell DJ, Farmer J, Newman M, McMillan JD: Dilute sulfuric acid pretreatment of corn stover in pilot-scale reactor. Appl Biochem Biotechnol 2003, 105–108:69–85.

13. Kadam KL, Rydholm EC, McMillan JD: Development and validation of a kinetic model for enzymatic saccharification of lignocellulosic biomass. Biotechnol Prog 2004, 20:698–705.

14. Morales-Rodriguez, Capron M, Huusom JK, Sin G: Controlled fed-batch operation for improving cellulose hydrolysis in 2 G bioethanol production. 2010, 20th European Symposium on Computer Aided Process Ebgineering-ESCAPE20.

15. Zhu Y, Malten M, Torry-Smith M, McMillan JD, Stickel JJ: Calculating sugar yields in high solids hydrolysis of biomass. Bioresour Technol 2011, 2011(102):2897–2903.

16. Wingren A, Galbe M, Zacchi G: Techno-economic evaluation of producing ethanol from softwood: comparison of SSF and SHF and identification of bottlenecks. Biotechnol Prog 2003, 19:1109–1117.

17. Aden A, Ruth M, Ibsen K, Jechura J, Neeves K, Sheehan J, Wallace B, Montague L, Slayton A, Lukas J. Lignocellulosic biomass to ethanol process design and economics utilizing co-current dilute acid prehydrolysis and enzymatic hydrolysis for corn stover. 2002, National Renewable Energy Laboratory Technical Report. NREL/TP-510-32438.

18. Gupta R, Khasa YP, Kuhad RC: Evaluation of pretreatment methods in improving the enzymatic saccharification of cellulosic materials. Carbohydr Polym 2011, 84:1103–1109.

19. Chen M, Xia L, Xue P: Enzymatic hydrolysis of corncob and ethanol production from cellulosic hydrolysate. Int Biodeter Biodeg 2007, 59:85–89.

20. Ghose TK: Measurement of cellulase activity. Pure Appl Chem 1987, 59:257–268.

Figure 1

y = 0.0421x

R2 = 0.9284

y = 0.0297x

R2 = 0.9812

y = 0.0216x

R2 = 0.9622

y = 0.012x

R2 = 0.9554

0

0.5

1

1.5

2

2.5

0 10 20 30 40 50 60

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ln (S

i,0/

S i)

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Figure 1

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cose

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Figure 2

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0 24 48 72 96 120Time (h)

Inso

lubl

e so

lids

(g/L

)

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Experimental

Figure 3

Figure 4

�

0

20

40

60

80

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0 24 48 72 96 120

Time (h)

Suga

r co

ncen

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(g/L

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�A

0

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Cel

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e co

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(g/

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Figure 4

that would be present if the entire solid were added initially and the reactor was operated in batch mode to enable comparison of fed-batch performance with the batch reactor performance on an equivalent basis.

For fed-batch operation, the model provides the rate expression for concentration of cellulose in the insoluble solid, glucose and cellulase enzyme. In addition to these variables, the dilution rate (D) was introduced to account for the changing mass and concentration due to a feed stream and is defined as the feed flow rate (F) per volume (V). The equations for fed-batch operations are as follows:

FdtdV

EEDdtdE

GGDSkdtdG

SSDSkdtdS

F

Fii

iFiii

�

�

�

�

)(

)(11.1

)(

(7)

Fed-batch saccharification model simulation

Using the above kinetic model, a feeding policy was developed based upon controlling the insoluble solids below a defined critical value during the saccharification reaction. This is possible by feeding a stream of pretreated substrate at a rate that approximately matches the rate of saccharification. Using the kinetic model equations, the rate of change of insoluble solids can be determined with the set of initial operating conditions and the insoluble solids at any given time point [Si(t)] will be,

� �� tkDk

DStS ii

Fi

� exp1

)()( (8)

Using this algorithm, fed-batch feeding policies were developed by generating a set of feeding curves over various reactor solids concentration and initial conditions generated to determine within the theoretical physical limitation of the system and the potential for using a fed-batch approach.

Analytical methods

The cellulase activities were determined following International Union of Pure and Applied Chemistry (IUPAC) methods [20]. The hydrolysates were analysed using high performance liquid chromatography (HPLC) (Waters, USA) for the presence of carbohydrates. Carbohydrate-ZX (Agilent Technologies, USA) column (300.0 x 7.8 mm) was used with Milli-Q water as an eluent with flow rate of 1.0 mL/min keeping oven temperature at 30 C with RID detector. Ethanol was estimated by gas chromatography (GC) (Perkin Elmer, Clarus 500) with an elite-wax (cross bond-polyethylene glycol) column (30.0 m�×�0.25 mm), at oven temperature 90oC and flame ionization detector (FID) at 200oC. Nitrogen with a flow rate of 0.5 mL min�1 was used as carrier gas.

��

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��

A B

��

��

��

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��

C D

0

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Sc (%

)

5%10%15% 0

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)5%10%15%

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Carbohydrate Polymers 84 (2011) 1103–1109

Contents lists available at ScienceDirect

Carbohydrate Polymers

journa l homepage: www.e lsev ier .com/ locate /carbpol

Evaluation of pretreatment methods in improving the enzymatic saccharificationof cellulosic materials

Rishi Gupta, Yogender Pal Khasa, Ramesh Chander Kuhad ∗

Lignocellulose Biotechnology Laboratory, Department of Microbiology, University of Delhi South Campus, Benito Juarez Road, New Delhi 110021, India

a r t i c l e i n f o

Article history:Received 14 October 2010Received in revised form20 December 2010Accepted 23 December 2010Available online 8 January 2011

Keywords:Cellulosic feedstocksPretreatmentAcidAlkaliChloriteEnzymatic hydrolysis

a b s t r a c t

The effectiveness of alkali, acid and chlorite pretreatment of lignocellulosic feedstocks for improvingthe enzymatic saccharification of cellulose has been evaluated. The feedstocks such as Corncob, Prosopisjuliflora and Lantana camara were pretreated with varied concentration of sulfuric acid, sodium hydrox-ide and sodium chlorite at 121 ◦C for 15–60 min. Among different methods used, chlorite pretreatmentremoved maximum lignin with ∼90% (w/w) residual holocellulose content in all the substrates tested.Moreover, irrespective of the substrates used, the chlorite treated substrates were enzymatically sac-charified from 86.4% to 92.5% (w/w). While, the alkali treated substrates containing 66.0–76.0% (w/w)holocellulose could be enzymatically saccharified up to 55% (w/w). The acid pretreated substrates werefound to contain almost 54–62% (w/w) holocellulose, which on enzymatic hydrolysis could result in39.5–48% (w/w) saccharification.

© 2011 Elsevier Ltd. All rights reserved.

1. Introduction

Lignocellulosics, the most abundant biomass available on earth,have attracted considerable attention as an alternative feedstockfor the production of various value added products due to theirrenewable nature and low cost availability (Kuhad & Singh, 2007).Various technological developments have improved the biocon-version of these substrates into bioethanol (Kapoor, Chandel,Kuhar, Gupta, & Kuhad, 2007). Enzymatic saccharification is oneof the promising strategy to convert cellulosic biomass into sugarsbecause of low energy requirement and less pollution. However,the primary challenge in enzymatic hydrolysis of cellulose is its lowaccessibility due to association with lignin. Therefore, efficient pre-treatment of lignocellulosic substrates has become a pre-requisiteto improve enzymatic saccharification (Zhao, Zhang, & Liu, 2008).The main focus of different pretreatment methods is to remove thelignin content and to decrease the cellulose crystallinity (Mosieret al., 2005). Although various physical (comminution, hydrother-molysis), chemical (acid, alkali, solvents, ozone), and biologicalpretreatment methods have been investigated over the years(Gupta, Mehta, Khasa, & Kuhad, 2010; Kuhar, Nair, & Kuhad, 2008;Kumar, Barrett, Delwiche, & Stroeve, 2009), thermo-chemical pre-treatment of biomass has been a pretreatment of choice to enhance

∗ Corresponding author. Tel.: +91 11 24112062; fax: +91 11 24115270.E-mail address: [email protected] (R.C. Kuhad).

substrate accessibility for efficient enzymatic hydrolysis (Himmelet al., 2007).

The thermo-chemical pretreatment strategies such as acid,alkali and oxidation are commonly used for lignocellulosic biomass.The dilute mineral acids have been reported to remove thehemicellulosic fraction from substrates to improve enzymatic sac-charification of cellulose (Gupta, Sharma, & Kuhad, 2009; Kuhad,Gupta, Khasa, & Singh, 2010; Schell, Farmer, Newman, & McMillan,2003). It has dual advantage of solubilizing hemicellulose and sub-sequently converting it into fermentable sugars. Whereas, the alkalipretreatment remove lignin and various uronic acid substitutionsresponsible for inhibiting the cellulose accessibility for enzymaticsaccharification (Chang & Holtzapple, 2000). Moreover, alkali treat-ment is also reported to increase the biodegradability of the cellwalls due to cleavage of the lignin bonds with hemicellulose andcellulose (Spencer & Akin, 1980). In contrast to acid and alkali treat-ments, sodium chlorite, a powerful oxidizing agent has been usedfrequently to delignify wood for cellulose isolation (Sun, Sun, Zhao,& Sin, 2004). The chlorine dioxide produced in this pretreatmentmethod oxidizes lignin to the phenolic compounds and in turnmakes cellulose accessible.

Since there is no universal and economically viable pretreat-ment method available, which could be used to pretreat variedcellulosic biomass, in the present study, it has been attempted toevaluate the suitably used three pretreatment methods (acid, alkaliand chlorite treatment) for lignocellulosic feedstocks viz., Prosopisjuliflora (PJ, a woody biomass), Lantana camara (LC, a shrub and

0144-8617/$ – see front matter © 2011 Elsevier Ltd. All rights reserved.doi:10.1016/j.carbpol.2010.12.074

1104 R. Gupta et al. / Carbohydrate Polymers 84 (2011) 1103–1109

weed) and Corncob (CC, agricultural residue). The pretreatmentsof the substrates were attempted at varied chemical dosage andpretreatment time. The pretreated plant materials were enzymat-ically hydrolysed and the cellulosic saccharification efficiency wasdetermined to evaluate the efficacy of these methods. Irrespectiveof the substrates used, the chlorite treatment was found to be anefficient method for delignification and producing cellulose richplant material, which was almost 90% hydrolysable by cellulasesinto glucose.

2. Materials and methods

2.1. Raw materials

The lignocellulosic substrates: Corncob (CC), P. juliflora (PJ) andL. camara (LC) were collected locally, dried in sunlight and thencut into small pieces. The dried material was ground and passedthrough a 40–60 mesh size screen using a laboratory knife mill(Metrex Scientific Instrumentation, Delhi, India). The processedsubstrate was thoroughly washed, dried at 60 ◦C and stored insealed plastic bags at room temperature.

2.2. Pretreatments

2.2.1. Acid pretreatmentThe dilute sulfuric acid pretreatment of lignocellulosic substrate

(100.0 g) was carried out using varied acid concentration (1–5%(w/v)) and incubation time (15–60 min) at 121 ◦C. The hydrolysatesafter treatment were separated by filtering the contents throughdouble layered muslin cloth. The residual biomass was washed withtap water till neutral pH and dried in a hot air oven at 60 ◦C.

2.2.2. Alkali pretreatmentThe substrate (100.0 g) was presoaked in different concentra-

tions of alkali (NaOH) ranging from 1% to 5% (w/v) for 2 h andthereafter thermally pretreated at 121 ◦C for 15, 30, 45 and 60 min.The pretreated material was filtered through double layered muslincloth, washed extensively with tap water until neutral pH and driedat 60 ◦C.

2.2.3. Chlorite pretreatmentThe lignocellulosic substrate (100.0 g) was treated with different

concentrations of sodium chlorite (1–5% (w/v)) at 121 ◦C for 15,30, 45 and 60 min. The pretreated material was filtered throughdouble-layered muslin cloth, washed thoroughly till neutral pH anddried in a hot air oven at 60 ◦C.

3. Enzymatic saccharification of pretreated substrates

Cellulase from Trichoderma reesei (ATCC 26921) with an activ-ity of 6.5 FPU/g, supplemented with �-glucosidase from Aspergillusniger (Novozyme 188) having 250 U/g was used for saccharifyingthe cellulosic material obtained after each pretreatment type.

Enzymatic hydrolysis of each type of pretreated plant materials(10.0 g each) was carried out at 5% (w/v) substrate consistency in50 mM citrate phosphate buffer (pH 5.0). The substrate with bufferwas pre-incubated at 50 ◦C on a rotatory shaker (Innova-40, NewBrunswick Scientific, Germany) at 150 rpm for 2 h and thereafterthe slurry was added with cellulase (3 FPU/ml) and �-glucosidase(9 U/ml). Tween 80 (1% (v/v)) was also added to the reaction mix-ture and the reaction continued up to 36 h. Samples of enzymatichydrolysate were withdrawn at regular intervals and analysed foramount of glucose released.

Table 1Proximate chemical composition analysis of Corncob, P. juliflora and L. camara usingTAPPI (1992) protocols.

Components (% (w/w)) Corncob P. juliflora L. camara

Cellulose 37.4 ± 4.18 47.5 ± 3.27 44.1 ± 1.72Pentosans 34.2 ± 1.02 18.7 ± 1.09 17.0 ± 0.81Holocellulose 71.6 ± 3.21 66.2 ± 4.54 61.1 ± 2.53Klason lignin 19.2 ± 0.83 29.1 ± 2.05 32.3 ± 1.57Moisture 7.4 ± 0.42 2.7 ± 0.28 4.4 ± 0.13Ash 1.8 ± 0.17 2.0 ± 0.12 2.3 ± 0.13

4. Analytical methods

The chemical composition (�-cellulose, klason lignin, pen-tosans, moisture and ash) of all the three substrates and theirresidual solid fraction post pretreatment were determined follow-ing standard TAPPI (1992), protocols. The reducing sugars releasedwere estimated using the DNS method (Miller, 1959) and the yieldof reducing sugars in enzymatic hydrolysate (YRSEH) was calcu-lated as follows:

YRSEH (%)

= Sugars released in enzymatic hydrolysateTotal carbohydrate content in pretreated substrates

× 100

The percent loss and gain of different components in pretreatedsubstrates were calculated using following equations:

Loss (%) = Mi −Mf

Mi× 100 (1)

Gain (%) = Mf −Mi

Mi× 100 (2)

where Mi is the amount of component in the untreated substrateand Mf is the amount of the component in the substrate after pre-treatment.

5. Statistical analysis

All the experiments were performed in triplicate and the resultsare presented as mean± standard deviation.

6. Results and discussion

6.1. Compositional analysis of different lignocellulosic substrates

The chemical composition analysis of different lignocellulosicbiomass revealed that the holocellulose content was in the range of61.1–71.6% (w/w), where P. juliflora (PJ) contained maximum cellu-lose content (47.5±3.27%) followed by L. camara (LC; 44.1±1.72%)and Corncob (CC; 37.4±4.18%). The lignin contents observed inCC, PJ and LC were 19.2±0.83%, 29.1±2.05% and 32.3±1.57%(w/w), respectively. However, the hemicellulose content was max-imum in CC (34.2±1.02%) compared to PJ (18.7±1.09%) andLC (17.0±0.81%). Moreover, all the substrates had almost equalamount of ash content ranging from 1.8% to 2.3%, and the mois-ture content in CC, PJ and LC were 7.4±0.42%, 2.7±0.28% and4.4±0.13%, respectively (Table 1). The considerably high carbohy-drate content in all the three lignocellulosic substrates qualifiedthem as potential feedstocks for bioethanol production.

6.2. Effect of chemical pretreatment

6.2.1. Acid treatmentThe dilute acid treatment is the most commonly used method

to pretreat the lignocellulosic biomass to hydrolyse hemicellulosic

R. Gupta et al. / Carbohydrate Polymers 84 (2011) 1103–1109 1105

Table 2Effect of acid treatment on the chemical composition of lignocellulosic substrates.

Time(min)

Acid (%(v/v))

Corncob P. juliflora L. camara

RL (% (w/w)) RH (% (w/w)) Hemicelluloseremoval (%(w/w))



15 1 22.4 ± 0.5 70.1 ± 1.2 31.9 ± 1.5 32.4 ± 0.9 64.1 ± 2.7 41.8 ± 2.3 33.5 ± 3.1 60.2 ± 3.3 23.4 ± 3.12 24.6 ± 0.6 66.7 ± 2.8 57.3 ± 2.7 33.7 ± 0.5 62.5 ± 3.4 61.1 ± 2.8 34.0 ± 3.1 59.5 ± 1.1 32.2 ± 2.83 25.7 ± 1.1 65.1 ± 3.5 67.6 ± 2.9 34.5 ± 1.8 61.5 ± 4.0 72.3 ± 3.4 34.5 ± 2.3 58.8 ± 4.2 40.9 ± 5.14 26.6 ± 0.4 63.7 ± 4.8 75.7 ± 3.1 34.7 ± 2.2 61.2 ± 4.3 64.9 ± 6.2 34.7 ± 3.8 58.6 ± 5.1 43.3 ± 6.35 27.3 ± 1.8 62.5 ± 4.5 82.4 ± 2.7 34.8 ± 2.2 61.1 ± 1.8 62.6 ± 1.5 34.8 ± 2.7 58.4 ± 2.5 45.6 ± 2.9

30 1 26.6 ± 1.2 63.5 ± 1.7 76.5 ± 2.8 32.9 ± 1.7 63.5 ± 1.9 49.2 ± 4.2 34.1 ± 1.0 59.4 ± 3.0 33.7 ± 4.22 27.3 ± 1.5 62.4 ± 2.6 82.2 ± 3.5 34.4 ± 2.2 61.6 ± 5.7 71.2 ± 2.9 34.8 ± 2.0 58.5 ± 2.8 44.6 ± 0.83 27.4 ± 0.9 62.4 ± 1.2 82.8 ± 3.1 35.0 ± 1.0 60.8 ± 3.1 79.9 ± 3.4 35.5 ± 1.2 57.6 ± 3.2 55.5 ± .14 27.4 ± 1.5 62.3 ± 3.8 83.1 ± 2.8 35.2 ± 0.6 60.5 ± 4.2 72.0 ± 1.8 35.6 ± 3.8 57.3 ± 2.4 58.5 ± 1.95 27.4 ± 1.6 62.3 ± 4.2 83.0 ± 4.9 35.4 ± 0.9 60.1 ± 3.6 68.2 ± 1.0 35.8 ± 4.2 57.1 ± 2.9 61.5 ± 0.4

45 1 26.8 ± 1.4 63.2 ± 5.4 74.6 ± 2.3 33.8 ± 2.2 62.3 ± 4.6 63.4 ± 0.8 36.5 ± 1.3 56.2 ± 2.7 70.7 ± 5.32 27.5 ± 0.9 62.3 ± 6.1 84.0 ± 4.1 35.3 ± 1.7 60.5 ± 3.1 83.3 ± 2.4 37.2 ± 0.8 55.2 ± 3.3 81.7 ± 3.13 27.5 ± 0.8 62.2 ± 3.0 84.0 ± 6.0 35.9 ± 0.6 59.7 ± 4.1 91.2 ± 1.1 38.0 ± 2.6 54.1 ± 3.4 92.6 ± 2.94 27.6 ± 1.1 62.1 ± 3.8 84.1 ± 3.8 35.9 ± 2.7 59.5 ± 3.0 86.0 ± 3.2 38.1 ± 3.1 54.1 ± 2.5 92.8 ± 5.15 27.6 ± 0.6 62.1 ± 4.9 84.2 ± 2.8 36.1 ± 1.4 59.4 ± 4.7 85.7 ± 3.8 38.1 ± 3.4 54.0 ± 1.6 93.5 ± 2.7

60 1 27.8 ± 1.5 61.7 ± 1.1 87.6 ± 5.2 34.1 ± 1.3 62.0 ± 3.0 66.7 ± 2.9 36.8 ± 2.6 55.8 ± 2.6 75.2 ± 1.02 28.0 ± 1.2 61.5 ± 6.5 86.4 ± 4.9 35.4 ± 0.9 60.3 ± 2.4 85.1 ± 3.5 37.6 ± 3.1 54.7 ± 2.3 86.6 ± 1.93 28.1 ± 0.4 61.5 ± 3.6 87.5 ± 6.5 36.1 ± 1.2 59.5 ± 4.7 93.5 ± 3.8 38.0 ± 1.9 54.2 ± 4.2 92.6 ± 1.04 28.3 ± 1.9 61.4 ± 5.9 86.1 ± 2.4 36.2 ± 2.3 59.2 ± 3.4 90.2 ± 4.9 38.0 ± 3.1 54.2 ± 1.9 92.1 ± 2.05 28.3 ± 0.9 61.4 ± 4.8 87.9 ± 3.0 36.2 ± 0.6 59.1 ± 1.8 84.7 ± 2.1 37.9 ± 3.4 54.1 ± 3.5 90.9 ± 2.5

RL, residual lignin; RH, residual holocellulose.

Fig. 1. Lignin removal from Corncob (A), P. juliflora (B) and L. camara (C) at different concentrations of alkali ( 1% NaOH, 2% NaOH, 3% NaOH, 4% NaOH and 5%NaOH).


Fig. 2. Holocellulose enrichment in Corncob (A), P. juliflora (B) and L. camara (C), after treatment with different concentrations of alkali ( 1% NaOH, 2% NaOH, 3% NaOH,4% NaOH and 5% NaOH).

fraction of lignocellulosic substrates, which is a critical parame-ter for process efficacy (Ishizawa, Davis, Schell, & Johnson, 2007).The hemicellulose fraction in CC was optimally hydrolysed to82.2±3.5% (w/w) with 2.0% (v/v) H2SO4 for 30 min. While, theoptimum acid saccharification in PJ (91.2±1.1% (w/w)) and LC(92.6±2.9% (w/w)) was achieved with 3.0% (v/v) sulphuric acidfor 45 min (Table 2). The higher acid concentration and pretreat-ment time requirement for optimal hydrolysis of PJ and LC maybe attributed to the woody nature of these substrates. Irrespec-tive of the substrates, increase in the acid dosage or pretreatmenttime beyond optimal conditions resulted in decrease of sugar yield,which may be because of the formation of sugar degradation prod-ucts such as furfurals and hydroxyl–methyl furfurals (Chandel,Kapoor, Sigh, & Kuhad, 2007; Gupta et al., 2009). However, theacid hydrolysed CC contained 62.4±2.6% (w/w) holocellulose and27.3±1.5% (w/w) lignin, while the acid treated PJ and LC werefound to have 59.7±4.1% (w/w) and 54.1±3.4% (w/w) holo-cellulose with 35.9±0.6% (w/w) and 38.0±2.6% (w/w) lignin,respectively (Table 2). As compared to untreated substrates (con-trol), the higher residual lignin and lower residual holocellulose inthe pretreated substrates may be due to the removal of acid sol-uble carbohydrate fraction (hemicellulose). Our results are well inaccordance with the previous reports. Chen, Zhao, and Xia (2009)reported an increase in lignin content from 19.3% to 28.4% (w/w)in acid hydrolysed corn stover. While, an increase in lignin contentfrom 21.8% to 28.5% (w/w) was observed when switch grass was

hydrolysed with 1.2% sulphuric acid at 160 ◦C for 20 min (Li et al.,2010).

6.2.2. Alkali treatmentDuring the alkali pretreatment, an increase in alkali concentra-

tion up to 5.0% (w/v) caused a regular increase in removal of ligninand in turn holocellulose gain in PJ and LC, while a 4.0% (w/v) alkaliconcentration was found to be optimum for CC (Figs. 1 and 2). Inall three substrates, the lignin removal increased with increase inpretreatment time till 30 min and remained almost constant there-after. Irrespective of the substrates, lignin content was observed tobe reduced in a range of 28–36% (w/w) (Fig. 1) with a concomitantenrichment in holocellulose content (5.2–7.1% (w/w)) (Fig. 2). Theresidual lignin in alkali treated CC, PJ and LC were 12.4%, 20.6% and19.4% (w/w), while the holocellulose content were 75.8%, 71.3% and65.8% (w/w), respectively. The increased lignin removal and enrich-ment of holocellulose in alkali treated substrates may be due tothe cleavage of the ester bonds between hydroxycinnamic acids,and the �-benzyl ether linkages of the cell wall of plant materialsby alkali (Mosier et al., 2005; Silverstein, Chen, Sharma-Shivappa,Boyette, & Osborne, 2007; Zhao et al., 2008). Similarly, other studieson alkali pretreatment have also reported approximately 40–60%delignification in crofton weed stem (Zhao et al., 2008), rice straw(Jeya, Zhang, Kim, & Lee, 2009) and switch grass (Nlewem & Thrash,2010).


Table 3Enzymatic hydrolysis of pretreated (under optimized conditions) lignocellulosic substrates.

Substrate Pretreatment type Reagent concentration Pretreatment time (min) RL (% (w/w)) RH (% (w/w)) YRSEH (% (w/w))

Corncob Untreated – – – – 38.5 ± 2.5Alkali treatment 4% (w/v) 30 12.4 ± 1.0 75.8 ± 4.8 55.4 ± 3.7Chlorite treatment 4% (w/v) 30 5.4 ± 0.5 90.3 ± 6.7 91.5 ± 3.1Acid treatment 2% (v/v) 30 27.3 ± 1.5 62.4 ± 2.6 39.5 ± 4.2

P. juliflora Untreated – – – – 33.2 ± 2.1Alkali treatment 5% (w/v) 30 20.6 ± 2.7 71.3 ± 3.5 52.2 ± 1.8Chlorite treatment 4% (w/v) 30 3.1 ± 0.2 90.7 ± 5.8 92.5 ± 4.3Acid treatment 3% (v/v) 45 35.9 ± 0.6 59.7 ± 4.1 47.7 ± 3.7

L. camara Untreated – – – – 31.2 ± 0.9Alkali treatment 5% (w/v) 30 19.4 ± 1.1 65.8 ± 2.6 50.8 ± 2.4Chlorite treatment 4% (w/v) 30 6.5 ± 0.2 90.0 ± 7.2 86.4 ± 1.6Acid treatment 3% (v/v) 45 38.0 ± 2.6 54.1 ± 3.4 48.0 ± 3.4

RL, residual lignin; RH, residual holocellulose; YRSEH, yield of reducing sugars in enzymatic hydrolysate.

6.2.3. Chlorite treatmentAll the three substrates when pretreated with 4% (w/v) sodium

chlorite for 30 min, maximum lignin removal as well as gainin holocellulose content were observed. Thereafter, any furtherincrease, either in the sodium chlorite concentration or pre-treatment time did not cause any significant improvement indelignification (Fig. 3). Moreover, irrespective of the substratestested, approximately 80–90% (w/w) delignification was obtainedwhen pretreated under optimal conditions (Fig. 3). The pretreated

substrates were found to have an increase in their holocellulosecontent by 26.2±0.8%, 37.2±5.2% and 47.3±2.6% (w/w) in CC,PJ and LC, respectively, as compared to control, with almost 90%(w/w) holocellulose recovery (Fig. 4). Earlier reports with sodiumchlorite treatment for the preparation of cellulose rich residue alsoshowed similar trends (Sevenson, Cheng, Jameel, & Kadla, 2005;Sun et al., 2004; Wi, Kim, Mahadevan, Yang, & Bae, 2009). Theextensive delignification of lignocellulosic substrates with chloritetreatment was due to the generation of chlorine dioxide (ClO2),

Fig. 3. Lignin removal from Corncob (A), P. juliflora (B) and L. camara (C) at different concentrations of Na-chlorite ( 1% Na-chlorite, 2% Na-chlorite, 3% Na-chlorite,4% Na-chlorite and 5% Na-chlorite).


Fig. 4. Holocellulose enrichment in Corncob (A), P. juliflora (B) and L. camara (C), after treatment with different concentrations of Na-chlorite ( 1% Na-chlorite, 2%Na-chlorite, 3% Na-chlorite, 4% Na-chlorite and 5% Na-chlorite).

an oxidation product of chlorous acid (HClO2) and hypochlorousacids (HOCl) produced during the thermal degradation of sodiumchlorite, which degraded the lignin either by side-chain displace-ment or hydroxylation/de-alkylation reaction (Hamzeh, Mortha, &Lachenal, 2006).

6.3. Comparison of different pretreatments for improvingenzymatic saccharification of cellulosic substrates

The enzymatic saccharification of all the three pretreated sub-strates showed an improved conversion of cellulose to glucosebecause of lignin and/or hemicellulose removal during pretreat-ments. The acid pretreated samples with minimum lignin removalshowed lowest enzymatic hydrolysis (39.5–48.0% (w/w)), while,the alkali and sodium chlorite pretreated substrates caused higherenzymatic saccharification, which could be because of compara-tively lower lignin content in the pretreated substrates (Table 3).Similar observations on pretreatment of lignocellulosic substratesfollowed by enzymatic hydrolysis have also been reported by Chenet al. (2009), Gupta et al. (2009) and Kuhad et al. (2010). The lim-ited enzymatic saccharification in presence of higher lignin may bedue to the high affinity of cellulases towards lignin, which resultedin unavailability of cellulase to cellulose moieties and led to poorenzymatic saccharification yields (Yang & Wyman, 2004).

Among different pretreated substrates evaluated here for theenzymatic hydrolysis, the chlorite pretreated substrates were

observed to be more vulnerable to the enzymatic hydrolysis andresulted in maximum saccharification efficiency i.e., from 86.4%to 92.5% (w/w) (Table 3). The higher enzymatic saccharifica-tion in chlorite pretreated substrates may be attributed to thepresence of higher holocellulose content (∼90.0% (w/w)) withminimum amount of residual lignin (3.1–6.5% (w/w)). It hasbeen shown by several workers that the delignification treat-ments not only remove the lignin but also act as a swellingagent, which in turn enhances the surface area of the substrateand make the substrate more amenable for enzymatic action(Gupta et al., 2009; Kuhad, Manchanda, & Singh, 1999; Kuhadet al., 2010). In contrast, the alkali treated substrates whensubjected to enzymatic saccharification brought about approxi-mately 51–55% (w/w) substrate hydrolysis (Table 3). As comparedto chlorite pretreated substrates, the lower enzymatic saccha-rification efficiency in alkali treated substrates might be dueto the lower delignification of the substrates by alkali treat-ment, which could be explained as the resistance of ligninremoval due to strong lignin–cellulose attraction in the cell wall(Zhao et al., 2008). Our results of higher enzymatic saccharifi-cation with chlorite pretreated substrates strongly agree withthe previous reports of achieving more than 65–67% sacchari-fication in chlorite pretreated sugarcane bagasse (Adsul et al.,2005) and approximately 70% (w/w) hydrolysis from chloritepretreated seaweed Ceylon moss (Gelidium amansii) (Wi et al.,2009).


7. Conclusion

Among different chemical pretreatments studied, the sodiumchlorite pretreatment was found to be the most effective in ligninremoval and led to the enrichment of the holocellulose content intreated substrates. This method offers the possibility of producingcellulosic material largely free from lignin, which eventually wouldbe a good substrate for bioethanol production. However, there isa need to develop efficient biological delignification methods tomake the process environmentally safe.

Acknowledgement

The authors are grateful to Department of Biotechnology, Gov-ernment of India and Council of Scientific and Industrial Research,India, for the financial support.

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ORIGINAL PAPER

Fungal delignification of lignocellulosic biomass improvesthe saccharification of cellulosics

Rishi Gupta • Girija Mehta • Yogender Pal Khasa •

Ramesh Chander Kuhad

Received: 20 April 2010 / Accepted: 5 August 2010 / Published online: 14 August 2010

� Springer Science+Business Media B.V. 2010

Abstract The biological delignification of lignocel-

lulosic feedstocks, Prosopis juliflora and Lantana

camara was carried out with Pycnoporus cinnabari-

nus, a white rot fungus, at different scales under solid-

state fermentation (SSF) and the fungal treated

substrates were evaluated for their acid and enzymatic

saccharification. The fungal fermentation at 10.0 g

substrate level optimally delignified the P. juliflora by

11.89% and L. camara by 8.36%, and enriched their

holocellulose content by 3.32 and 4.87%, respec-

tively, after 15 days. The fungal delignification when

scaled up from 10.0 g to 75.0, 200.0 and 500.0 g sub-

strate level, the fungus degraded about 7.69–10.08%

lignin in P. juliflora and 6.89–7.31% in L. camara,

and eventually enhanced the holocellulose content by

2.90–3.97 and 4.25–4.61%, respectively. Further-

more, when the fungal fermented L. camara and

P. juliflora was hydrolysed with dilute sulphuric acid,

the sugar release was increased by 21.4-42.4% and the

phenolics content in hydrolysate was decreased by

18.46 and 19.88%, as compared to the unfermented

substrate acid hydrolysis, respectively. The reduction

of phenolics in acid hydrolysates of fungal treated

substrates decreased the amount of detoxifying mate-

rial (activated charcoal) by 25.0–33.0% as compared

to the amount required to reduce almost the same level

of phenolics from unfermented substrate hydrolysates.

Moreover, an increment of 21.1–25.1% sugar release

was obtained when fungal treated substrates were

enzymatically hydrolysed as compared to the hydro-

lysis of unfermented substrates. This study clearly

shows that fungal delignification holds potential in

utilizing plant residues for the production of sugars

and biofuels.

Keywords Bioethanol � Lignocellulose � Fungaldelignification � Solid-state fermentation �Pycnoporus cinnabarinus � Saccharification

Introduction

Lignocellulosic biomass is a potential source of

carbohydrate polymers for fermentation but the

conversion of its structural polysaccharides into

simple sugars is highly problematic due to the

recalcitrancy of lignin. Lignin acts as a cementing

material, which together with hemicelluloses forms

an amorphous matrix in which the cellulosics fibrils

are embedded and protected against chemical or

enzymatic degradation (Kuhad et al. 1997; Himmel

et al. 2007). Therefore, removal of lignin has become

a prerequisite for the efficient utilization of carbohy-

drate from lignocellulosics.

R. Gupta � G. Mehta � Y. P. Khasa � R. C. Kuhad (&)

Lignocellulose Biotechnology Laboratory, Department

of Microbiology, University of Delhi South Campus,

Benito Juarez Road, New Delhi 110021, India

e-mail: [email protected]

123

Biodegradation (2011) 22:797–804

DOI 10.1007/s10532-010-9404-6

Pretreatment of lignocellulosics is the most impor-

tant step required to remove lignin and disrupt the

crystallinity of carbohydrate fraction without losing

too much of the structural sugars (Mosier et al. 2005;

Wyman 2007; Kumar et al. 2009). Physicochemical

pretreatments such as acid treatment (Gupta et al.

2009), alkali treatment (Carrillo et al. 2005), steam

explosion (Ohgren et al. 2006) and ammonia fiber

explosion (Teymouri et al. 2005) are well recognized

for enhancing the conversion of cellulosic biomass

into monomeric sugars. However, these pretreatment

methods require high energy and often generate toxic

compounds, which makes the process economically

unviable and environment unfriendly (Teymouri et al.

2005; Silverstein et al. 2007). Therefore, environ-

mentally benign pretreatment methods are required to

reduce release of pollutants, costs and improve

cellulose saccharification.

The biological pretreatments of plant residues to

improve the accessibility of cellulosic fraction have

been attracting the extensive interest of researchers

(Taniguchi et al. 2005; Zhang et al. 2007a; Yu et al.

2009b). The potential of biological pretreatments has

been explained by the ability of certain microbes to

disrupt the plant cell wall by partial breakdown of the

lignin/carbohydrate complex (Keller et al. 2003). The

most promising microorganisms for biological pre-

treatment are basidomycetes, and among them the

selective lignin degrading white rot fungi holds

immense importance (Akhtar et al. 1997; Kuhad

et al. 1997; Kuhad and Singh 2007; Yu et al. 2009a).

Pycnoporus cinnabarinus, a selective lignin degrad-

ing white rot fungus, has been reported to produce

laccase, degrade lignin and to transform lignin derived

compounds (Eggert et al. 1996; Falconiner et al. 1994;

Galhaup et al. 2002). Recently, P. cinnabarinus,

Phanerochaete chrysporium and Crinepellis sp. RCK-

1 have been tested for their lignin degradation and

improvement in acid hydrolysis of the delignified

substrates (Kuhar et al. 2008). However, the applica-

tion of P. cinnabarinus for delignifying the lignocel-

lulosic materials and subsequently its effect on

enzymatic hydrolysis of the fermented substrate

(mycosubstrate) has scarcely been studied.

In the present study, biological delignification of

two different weeds (lignocellulosic substrate) viz.

Lantana camara and Prosopis juliflora using

P. cinnabarinus was attempted under solid-state

fermentation (SSF) conditions with pretreatment

scalability up to 500.0 g substrate level. The pre-

treated substrates were further evaluated for acid as

well as enzymatic hydrolysis. The acid and enzymatic

hydrolysis of fungal fermented material resulted in

significant increase in the release of sugars as

compared to unfermented ones. Interestingly, the

acid hydrolysates of fungal fermented substrates were

found to have lesser amount of toxic compounds as

compared to hydrolysate from unfermented material.

Materials and methods

Raw material and chemicals

The woods of L. camara and P. juliflora were

collected locally and processed through a combination

of chipping and milling to attain a particle size of

1–2 mm using a laboratory knife mill (Metrex Scien-

tific Instrumentation, Delhi, India). The grounded

materials washed thoroughly with water and dried

overnight at 60�C were used throughout the study.

Cellulase from Trichoderma reesei (ATCC

26921), b-glucosidase (Novozyme 188) from Asper-

gillus niger, and 3,5-dinitrosalicylic acid (DNS) were

brought from Sigma, St. Louis, Missouri, USA,

while, all other chemicals were purchased locally.

Microorganism and inoculum preparation

Pycnoporus cinnabarinus ATCC 2004378, a kind gift

from Late Dr. K. E. L. Eriksson, Professor Emeritus,

Department of Biochemistry and Molecular Biology,

University of Georgia, Athens, USA, was grown at

30�C on malt extract agar (MEA, Dhawan and Kuhad

2003). The medium contained (g l-1): malt extract

20.0, KH2PO4 0.5, MgSO4 0.5, Ca(NO3)2 0.5 and agar

20.0 (pH 5.5), while the stock cultures were main-

tained on MEA slants at 4�C with periodic transfer.

The fungal cultures were grown statically at 30�C in

250 ml Erlenmeyer flasks with 50 ml of malt extract

broth (MEB, Dhawan and Kuhad 2003), which were

inoculated with 2 fungal discs (8 mm diameter each)

from a 6 day old colony grown on MEA. The cultures

were harvested and the aseptically collected fungalmat

was homogenized and transferred into a fresh 50 ml

MEB in 250 mlErlenmeyer flasks. Theywere grown at

30�C and 150 rpm in a rotatory incubator shaker

(Innova-40, New Brunswick Scientific, USA) for

798 Biodegradation (2011) 22:797–804

123

another 6 days to develop the inoculum for solid state

fermentation of lignocellulosic materials.

Fungal treatment of L. camara and P. juliflora

under SSF

The pretreatment was carried out in 500 ml Erlen-

meyer flasks with 10.0 g of the oven dried lignocellu-

losic substrate (L. camara or P. juliflora), moistened

with mineral salt solution (MSS) containing (g l-1):

KH2PO4 0.5, MgSO4 0.5, Ca(NO3)2 0.5 and pH 5.5 to

obtain a substrate to moisture ratio of 1:2.5 and

autoclaved (121�C for 30 min). Each flask was then

inoculated with 7.5 mg fungal mass (dry weight) per

gram dry substrate (gds) and incubated at 30�C for

25 days. The solid state fungal fermentation of both the

substrates was also studied at 75.0, 200.0 and 500.0 g

substrate levels in enamel trays of different sizes:

21.5 cm 9 16.5 cm 9 5.0 cm, 31.5 cm 9 26.5 cm

6.0 cm and 41.5 cm 9 34.5 cm 9 8.0 cm, respec-

tively. The samples (mycosubstrate) were harvested

periodically, washed thoroughly and dried overnight at

60�C. The dried mycosubstrates were then analysed for

biochemical changes, acid and enzymatic saccharifica-

tion. The flasks containing sterilized uninoculated

substrates served as control.

Evaluation of delignified substrates for acid

hydrolysis

The fungal pretreated and control (non-fungal

treated) substrates were hydrolyzed at 10% (w/v)

substrate consistency with dilute sulfuric acid (3.0%,

v/v) at 120�C for 45 min (Gupta et al. 2009). The

hydrolysate was vacuum filtered and analyzed for

reducing sugars and total phenolics. The residual

substrate after hydrolysis was washed thoroughly to a

neutral pH, dried at 60�C in a hot air oven till

constant weight was achieved.

Detoxification of acid hydrolysates

Acid hydrolysates of unfermented and fungal treated

substrates were detoxified by adding varied dosages

of activated charcoal (0–2.5%, w/v) under constant

stirring at room temperature for 30 min. After

incubation, the hydrolysate was vacuum filtered and

analysed for total sugars and phenolics.

Enzymatic hydrolysis of fermented

and non-fermented plant residues

Enzymatic hydrolysis of fermented and control (non-

fermented) substrate samples was carried out at 5.0%

(w/v) substrate consistency in 50 mM citrate phos-

phate buffer (pH 5.0). Prior to the enzyme loading,

the slurry was incubated at 50�C for 2 h at 150 rpm.

Thereafter, cellulase (24 FPU gds-1), b-glucosidase(120 U gds-1) and 1.0% (v/v) Tween 80 were added

to the preincubated substrate suspension and the

saccharification was carried out at 50�C and 150 rpm

for 48 h. Samples were withdrawn after every 6 h,

centrifuged at 10,000 rpm for 15 min and the super-

natants were analysed for total reducing sugars.

Analytical methods

The total phenolics present in hydrolysates were

determined by the method of Singleton et al. (1999)

using Vanillin as standard, while, the reducing sugars

were estimated by DNS method (Miller 1959).

The plant material was extracted with alcohol-

benzene (1:2 v/v) to remove wax and resins etc. The

extractive free, oven dried plant material was pro-

cessed for biochemical analysis following the TAPPI

(1992) protocols. The holocellulose to lignin ratio

(H/L) was calculated as follows:

H/L¼Amount of holocellulose Hð Þ present in the substrate

Amount of lignin Lð Þ present in the substrate�100

The percent loss and gain of different components

such as total organic matter (TOM) loss, lignin and

holocellulose in fungal treated substrates were cal-

culated using Eqs. 1 and 2, respectively.

Loss %ð Þ ¼ Mi �Mf

Mi� 100 ð1Þ

Gain %ð Þ ¼ Mf �Mi

Mi� 100 ð2Þ

where Mi is the amount of component in the control

(non-fermented) substrate and Mf is the amount of the

component left in the mycosubstrate.

All the experiments and the analysis were carried

out in triplicates and the data presented is the mean

Biodegradation (2011) 22:797–804 799

123

value of the triplicates. The standard deviation was

calculated using the mean values and remained

within the range of ±10.0%.

Results and discussion

Solid state fermentation (SSF) of lignocellulosic

substrates

Irrespective of the substrates used, the increase in loss

of total organic matter (TOM) correlated positively

with the increase in lignin degradation, and holocel-

lulose to lignin (H/L) ratio. While, the holocellulose

content increased till 15 days of fungal fermentation

and marginally declined thereafter (Table 2). The

SSF of L. camara and P. juliflora by P. cinnabarinus

after 25 days caused 18.87 and 15.40% loss of TOM,

respectively (Table 1). The loss in TOM may be

attributed to the CO2 evolution due to the metabolic

activities of fungus on the substrates, however, the

weight loss in substrates includes the loss of lignin

and holocellulose. The fungus degraded more amount

of lignin in P. juliflora (13.13%) than in L. camara

(8.87%). The fungal delignification in both the

substrates was higher during the first 15 days, and

thereafter no significant improvement in lignin deg-

radation was observed (Table 1). Our results are in

accordance with the earlier reports on biological

delignification of lignocellulosic materials with white

rot fungi (Bustamente et al. 1999; Taniguchi et al.

2005; Meza et al. 2006). P. cinnabarinus was

reported to degrade 8.5% lignin in sugarcane bagasse

(Meza et al. 2006), whereas, Ceriporiopsis subver-

mispora and Pleurotus ostreatus degraded 9.0 and

17.0% lignin in depithed sugarcane bagasse, respec-

tively (Bustamente et al. 1999). The fungal cultures

have been observed to degrade more lignin in wheat

straw than in woody material and this low ability of

lignin degradation in woody substrate by the fungi

could be attributed to the difference in the lignin

properties of wood and straw (Taniguchi et al. 2005).

Thus lignin structure could be seen as a detrimental

factor for economic exploitation of plant residues.

Besides delignification, the availability of carbo-

hydrates is also an important criterion for evaluating

the biological pretreatment performance because the

higher cellulose content in fermented substrates

eventually provides higher accessibility of Table

1Analysisofcompositional

changes

duringthefungal

treatm

entofL.camara

andP.juliflora

usingP.cinnabarinusat

differenttimeintervals(5–25days)

Components

Substrates

L.camara

P.juliflora

Tim

e(days):

Control

510

15

20

25

Control

510

15

20

25

Weight(g)

10.0

8.91(10.87)a

8.67(13.27)a

8.47(15.33)a

8.37(16.27)a

8.11(18.87)a

10.0

9.23(7.73)a

9.05(9.47)a

8.81(11.87)a

8.63(13.67)a

8.46(15.40)a

Klason-Lignin

(%)

33.18

31.64(4.63)a

30.91(6.85)a

30.41(8.36)a

30.28(8.74)a

30.24(8.87)a

30.71

28.71(6.51)a

27.72(9.72)a

27.06(11.89)a

26.89(12.44)a

26.68(13.13)a

Holocellulose

(%)

59.82

60.94(1.87)b

61.70(3.14)b

62.73(4.87)b

62.45(4.40)b

62.25(4.06)b

63.58

64.44(1.35)b

64.92(2.11)b

65.69(3.32)b

65.42(2.90)b

65.07(2.34)b

H/L

1.80

1.93(6.99)b

2.00(10.90)b

2.06(14.62)b

2.06(14.58)b

2.06(14.37)b

2.07

2.24(8.42)b

2.34(13.12)b

2.43(17.28)b

2.43(17.54)b

2.44(17.83)b

aPercentageloss

infungal

treatedsubstrateswithrespectto

control(untreated)substrates

bPercentagegainin

fungal

treatedsubstrateswithrespectto

control(untreated)substrates

800 Biodegradation (2011) 22:797–804

123

carbohydrates for acid or enzymatic saccharification

(Shi et al. 2008). There are several reports where

P. chrysosporium, the most widely studied white rot

fungus for lignin degradation, resulted in higher

amounts of lignin degradation but the fungus was also

found to cause a significant loss in carbohydrate

fraction (Taniguchi et al. 2005; Kuhar et al. 2008).

However our results showed an increase in holocel-

lulose content in both the fungal fermented substrates

by 3.32 and 4.87% compared to control (unfer-

mented) substrates after 15 days and thereafter it

declined gradually (Table 1). The fungal delignifica-

tion also increased the H/L ratio by 14.62 and 17.28%

in L. camara and P. juliflora, respectively after

15 days (Table 1). The increase in H/L ratio may be

due to the reason that the fungus produces a fairly

good amount of laccase with very low activity of

carbohydrate hydrolyzing enzymes (Alves et al.

2004), which might have resulted in higher degrada-

tion of lignin as compared to holocellulose in the

P. cinnabarinus treated substrates. Similar results

were also obtained during the biological pretreatment

of wheat straw using Streptomyces cyaneus, which

caused 16 and 8% degradation of lignin and holo-

cellulose, respectively, after 15 days of SSF and

eventually enhanced the H/L ratio of the fermented

wheat straw (Berrocal et al. 2000).

Scale up of biological delignification

The fungal delignification of lignocellulosic substrates

was scaled up under similar conditions such as particle

size, substrate to moisture ratio, pH, temperature, tray

volume to substrate ratio. The study revealed that

irrespective of the scale of treatment, the lignin

degradation was found to be comparatively more or

less same. The P. cinnabarinus when grown on

L. camara and P. juliflora at larger scales (75.0, 200.0

and 500.0 g) caused 15.19–17.41 and 12.05–15.56%

weight loss, respectively. Moreover, the fungal pre-

treatment of L. camara and P. juliflora removed lignin

by 6.89–7.31 and 7.69–10.08% and also enhanced the

holocellulose content by 4.25–4.61 and 2.9–3.97%,

respectively, after 15 days of incubation. Furthermore,

the H/L ratio of fermented L. camara and P. juliflora

remained in the range of 2.02–2.03 and 2.31–2.39%,

respectively (Table 2). The insignificant difference in

H/L ratios and compositional changes in the fermented

substrates at different scale experiments demonstrated

the possibility to further scale up of the process.

Evaluation of biologically delignified substrates

for acid saccharification

Acid hydrolysis of fungal fermented and the control

(unfermented) substrates was evaluated. The acid

hydrolysis of fungal fermented L. camara and

P. juliflora resulted in 131.4 and 189.7 mg gds-1

sugar yield as compared to sugar release of 108.2 and

133.2 mg sugar gds-1 from control (unfermented)

substrates, respectively (Fig. 1). The acid saccharifi-

cation of fungal fermented materials has significantly

improved in sugar release (21.4–42.4%) as compared

to the unfermented substrates. The increased sugar

release in fungal treated substrates might be because

of the preferential lignin degradation ability in the

Table 2 Compositional changes during the scale up of biological delignification of L. camara and P. juliflora using P. cinnabarinusat different levels (75.0, 200.0 and 500.0 g)

Compound Substrates (g)

L. camara P. juliflora

75 200 500 SD 75 200 500 SD

Weight (%) 61.94 (17.41)a 167.04 (16.48)a 424.05 (15.19)a 1.11 63.33 (15.56)a 174.68 (12.66)a 439.75 (12.05)a 1.88

Klason-Lignin

(%)

30.75 (7.31)a 30.89 (6.89)a 30.85 (7.01)a 0.22 28.35 (7.69)a 27.61 (10.08)a 27.69 (9.84)a 1.32

Holocellulose

(%)

62.58 (4.61)b 62.36 (4.25)b 62.45 (4.39)b 0.18 65.42 (2.90)b 66.10 (3.97)b 65.86 (3.58)b 0.54

H/L 2.03 (13.04)b 2.02 (12.14)b 2.03 (12.44)b 0.006 2.31 (11.49)b 2.39 (15.64)b 2.38 (14.90)b 0.004

a Percentage loss in fungal treated substrates with respect to control (untreated) substratesb Percentage gain in fungal treated substrates with respect to control (untreated) substrates

Biodegradation (2011) 22:797–804 801

123

substrates by the fungus P. cinnabarinus, which in

turn would have resulted in enrichment of carbohy-

drate content in the fermented substrates. Similar

results have been reported in fungal treated wheat

straw by Kuhar et al. (2008).

Acid hydrolysis not only release the reducing

sugars but also release some lignin degradation

compounds (phenolics), which are toxic to the fer-

menting microorganisms (Palmqvist et al. 1999;

Mussatto and Roberto 2004; Chandel et al. 2007).

The acid hydrolysates obtained from fungal treated

L. camara and P. juliflora were found to have

18.5–19.9% less phenolics as compared to the hydrol-

ysates of unfermented substrates (Fig. 1). This reduction

in phenolics can be attributed to the fungal degradation

of phenolic compounds (Kuhar et al. 2008).

An attempt has been made to evaluate the effect of

fungal pretreatment on reduction in the usage of

chemicals in detoxification of acid hydrolysates.

Interestingly, the activated charcoal even when used

at a lower concentration (1 and 1.5% w/v, 25–33%

lower than control) removed more than 90% phen-

olics from acid hydrolysates of fungal treated

substrates as compared to the acid hydrolysates of

unfermented ones (Table 3). Our observations clearly

indicated that fungal fermentation (delignification) of

lignocellulosic material resulted in holocellulosic

materials more vulnerable to acid hydrolysis, which

in turn gives hydrolysates rich in pentose sugars and

lesser phenolics.

Evaluation of biologically delignified substrates

for enzymatic saccharification

In comparison to the unfermented substrates, the

fungal fermented L. camara and P. juliflora when

hydrolysed with cellulases for 48 h, an increase of

25.1 and 21.1% (w/w) saccharification with release of

reducing sugars (389.1 and 402.1 mg gds-1), respec-

tively, was achieved (Fig. 2). The enhanced enzy-

matic saccharification of fungal fermented substrates

may be attributed to the partial degradation of lignin

seal responsible for preventing the penetration of

cellulase molecules (Taniguchi et al. 2005). More-

over, the improvement in enzymatic hydrolysis could

be because the biological treatment increases the

initial adsorption of cellulases to the cellulose, which

in turn enhances the sugars released during the

enzymatic hydrolysis (Yu et al. 2009a).

It has been observed that fungal fermentation with

P. cinnabarinus has considerable potential in improv-

ing enzymatic saccharification of cellulosics into

sugar rich syrup. Taniguchi et al. (2005) have

reported 330 mg gds-1 sugar released from rice

straw fermented with P. ostreatus after 60 days.

Zhang et al. (2007b) when enzymatically hydrolysed

A B C D

0

20

40

60

80

100

120

140

160

180

200

suga

rs a

nd p

heno

lics

rele

ased

(m

g/g)

Phenolics

Sugars

Fig. 1 Release of sugars and phenolics during the acid

hydrolysis (3% H2SO4, 120�C and 45 min) of P. cinnabarinustreated and control (untreated) substrates (where A, untreated

L. camara; B, SSF-treated L. camara; C, untreated P. juliflora;D, SSF-treated P. juliflora)

Table 3 Detoxification of

phenolics from the acid

hydrolysates obtained from

different substrates

(untreated and fungal

treated L. camara and

P. juliflora) using varied

dosage of activated charcoal

a The values in parenthesis

is the percentage amount of

phenolics removed

Amount of

charcoal

used (%, w/v)

Amount of phenolics (mg g-1)

L. camara P. juliflora

Untreated SSF-treated Untreated SSF-treated

0.00 24.00 (0.00) 19.23 (0.00) 18.26 (0.00)a 14.84 (0.00)

0.50 18.69 (22.13) 14.20 (26.15) 13.25 (27.44) 7.13 (51.97)

1.00 12.51 (47.88) 6.04 (68.60) 5.74 (68.57) 0.73 (95.11)

1.50 4.67 (80.54) 0.95 (95.04) 0.85 (95.37) 0.06 (99.59)

2.00 0.09 (99.62) 0.08 (99.56) 0.05 (99.72) 0.05 (99.64)

2.50 0.03 (99.87) 0.04 (99.78) 0.05 (99.74) 0.05 (99.64)

802 Biodegradation (2011) 22:797–804

123

Trametes versicolor and Echinodontium taxodii fer-

mented bamboo culms for 120 days, the sugar

released was approximately 360 and 200 mg gds-1,

respectively. Recently Yu et al. (2009a) have also

reported 255.75 and 70.4 mg gds-1 sugar yield from

E. taxodii fermented chinese willow and china fir

after 120 days, as calculated from their data. Thus the

sugars released in our study from both the fungal

fermented substrates after 15 days was higher (389.1

and 402.1 mg gds-1) than the above cited studies.

Although it’s difficult to make exact comparison in

studies where lignocellulosic substrates are different

and moreover fermentation period is variable. But

comparatively it seems that the fungus used in this

study holds considerably more potentials.

Conclusion

The conventional non-biological pretreatment meth-

ods require high energy, often generate toxic sub-

stances inhibitory to fermentation, which makes the

processes uneconomical and environmentally inimi-

cal. The results of this fungal fermentation based

study indicate that in order to increase the acid or

enzymatic saccharification of hemicellulose and cel-

lulose, it is prerequisite to degrade the lignin in the

plant material to be used for producing value added

compounds. The fungal fermentation have led to

significant improvement in acid hydrolysis of

hemicellulosic content and enzymatic hydrolysis of

cellulose content in L. camara and P. juliflora.

Moreover, fungal fermentation of plant material has

resulted into material, which on acid hydrolysis

produces hydrolysates with lesser amount of toxic

compounds. Thus, the fungal delignification if opti-

mized can be used as a potential alternative pretreat-

ment method for improving the enzymatic

saccharification of lignocellulosic residues and even-

tually for economizing the ethanol production as

biofuel.

Acknowledgments The authors are thankful to Ms. Urvashi

Kuhad, Department of English, University of Delhi for her help

during the preparation of manuscript. The financial support

from the Department of Biotechnology, Ministry of Science

and Technology, New Delhi, India is highly acknowledged.

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Renewable and Sustainable Energy Reviews 15 (2011) 4950– 4962

Contents lists available at SciVerse ScienceDirect

Renewable and Sustainable Energy Reviews

jo ur n al hom ep a ge: www.elsev ier .com/ locate / rser

Bioethanol production from pentose sugars: Current status and future prospects

Ramesh Chander Kuhada,∗, Rishi Guptaa, Yogender Pal Khasaa, Ajay Singhb, Y.-H. Percival Zhangc

a Lignocellulose Biotechnology Laboratory, Department of Microbiology, University of Delhi South Campus, Benito Juarez Road, New Delhi 110021, Indiab Lystek International Incorporation, 107-279 Weber Street North, Waterloo, Ontario, Canada N2J3H8c Biological Systems Engineering Department, Virginia Polytechnic Institute and State University, Blacksburg, VA 24061, USA


Article history:Received 19 January 2011Received in revised form 11 May 2011Accepted 7 July 2011Available online 15 September 2011

Keywords:HemicellulosePentose sugarsBioethanol, Fermentation

a b s t r a c t

The utilization of hemicellulose, the second most abundant polysaccharide, is must for the cost-efficientproduction of ethanol from second generation feedstocks. Xylan, the major hemicellulose in plant biomassyields mainly xylose as pentose sugars on hydrolysis. The progress in fermentation of pentose sugars hasgone on slow pace as there are few microorganisms known, which are capable of pentose metabolism.The future perhaps lies in finding organisms that would ferment high density hydrolysates withoutpurification. This obviously has to use the genetic and metabolic engineering routes. Either a direct or asequential fermentation system needs to be worked out. This review provides an overview of the currentpentose bioconversion processes and future prospects for bioethanol production.

© 2011 Elsevier Ltd. All rights reserved.

Contents

1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 49512. Hemicellulose . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4951

2.1. Softwood hemicellulose . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 49512.2. Hardwood hemicellulose . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4951

3. Pentose-fermenting microorganisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 49513.1. Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 49513.2. Filamentous fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 49523.3. Yeast . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4952

4. Xylose metabolizing pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 49525. Fermentation of pentose sugars . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4953

5.1. Pentose fermentation using whole cell immobilization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 49545.2. Pentose fermentation using recycling of cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 49555.3. Pentose fermentation using simultaneous saccharification and fermentation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4955

6. Strain improvement for pentose fermentation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 49556.1. Strain improvement through mutagenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 49556.2. Strain improvement through protoplast fusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 49566.3. Strain improvement through adaptation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 49566.4. Strain improvement through genetic manipulation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4956

6.4.1. Genetic engineering of E. coli . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 49566.4.2. Genetic engineering of Z. mobilis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 49586.4.3. Genetic engineering of S. cerevisiae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4958

7. Future prospects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4959Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4959References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4959

∗ Corresponding author. Tel.: +91 11 24112062; fax: +91 11 24115270; mobile: +91 9871509870.E-mail address: [email protected] (R.C. Kuhad).

1364-0321/$ – see front matter © 2011 Elsevier Ltd. All rights reserved.doi:10.1016/j.rser.2011.07.058

R.C. Kuhad et al. / Renewable and Sustainable Energy Reviews 15 (2011) 4950– 4962 4951

1. Introduction

Utilization of lignocellulosic biomass for ethanol production isone of the most promising alternatives for liquid fuel, which hasnear zero green house gas emission with great socio-economic ben-efits [1–4]. Lignocellulose is the most abundant, renewable organicmaterial on the biosphere, but by virtue of its structural proper-ties they are resistant to bioconversion [5–7]. For cost-effectiveproduction of bioethanol from lignocellulosic biomass, the high-efficiency utilization of both carbohydrate fractions, i.e., celluloseand hemicellulose is required.

Bioethanol production from lignocellulosic biomass involvesseveral steps such as pretreatment, hydrolysis of complex carbo-hydrates, fermentation, and distillation for product recovery [8,9].Various pretreatment methods have been explored to enhancethe accessibility of lignocellulosic substrates. Among them, diluteacid pretreatment is the method of choice which has been stud-ied widely [9–13]. However, it has drawback of non-selectivityand formation of fermentation inhibitory compounds such asformic acid, levulinic acid, furfural, 5-hydroxymethylfurfural andphenolics [9,14,15]. In order to remove these toxic compoundsfrom acid hydrolysates, strategies such as overliming, charcoaltreatment, steam stripping, ion-exchange resin treatment, and bio-logical methods such as enzymatic and microbial detoxificationhave been employed [1,16–18].

A number of cellulolytic microorganisms are known to pro-duce cellulases and hemicellulases, which can convert cellulose andhemicellulose respectively into soluble monomeric or oligomericsugars. These sugars further can be fermented into ethanol by anumber of bacterial, yeast and filamentous fungi. The fermentationprocess would be economically viable only if both hexose and pen-tose sugars present in the hydrolysates are converted to ethanol.The ability to ferment pentoses is not widespread among microor-ganisms. The most promising yeast species identified so far, areCandida shehatae, Pichia stipitis and Pachysolen tannophilus [19–22].But, the use of these yeasts for ethanol production from xyloseat commercial level is limited mainly due to their slow fermen-tation rates, carefully regulated oxygen requirement, sensitivityto inhibitors and low ethanol tolerance [22]. Although significantimprovements have been made in the microorganisms for theefficient fermentation of pentose sugars into ethanol, however,the bioconversion of pentoses to ethanol is still one of the majorbottlenecks for ethanol commercialization effort. In this paper,an overview of bioconversion of pentose sugars into bioethanolusing fermentation and molecular strategies for improvement ofpentose-fermenting strains utilizing hemicellulosic sugars are dis-cussed.

2. Hemicellulose

Among the different structural units of lignocellulosics, hemi-cellulose comprises almost 15–30% of the total dry weight andhence represents an important energy and material resourcefor bioconversion [6,23–25]. The hemicelluloses comprised bothlinear and branched hetero-polymers of d-xylose, l-arabinose, d-mannose, d-glucose, d-galactose and d-glucuronic acid (Fig. 1A).The xylose-rich hemicelluloses in both soft and hard wood areusually termed as xylans. The hemicellulose from hardwoods andagricultural residues are typically rich in xylan, while, softwoodcontains more mannan and less xylan [6,7,24,26].

2.1. Softwood hemicellulose

Galacto-mannans are the principal hemicelluloses in soft-woods. Their backbone is a linear chain built up by 1,4-linked

�-d-gluco-pyranose and �-d-manno-pyranose units (Fig. 1B).The mannose and glucose units in the backbone are partiallysubstituted at C-2 and C-3 positions by acetyl groups, approx-imately 1 per 3–4 hexose units [6]. Arabino-glucuronoxylan isanother major hemicellulosic sugar and is composed of 1,4-linked-�-d-xylopyranose units. This chain is substituted at C-2by 4-o-methyl-�-d-glucuronic acid group with approximately twosuch units per ten xylose units. The xylose backbone is also sub-stituted by �-l-arabino-pyranose units, on the average 1.3 residueper ten xylose units [6,25]. Arabino-galactan is a minor componentin both softwoods and hardwoods. The backbone of this galactanis built up by 1,3-linked �-d-galacto-pyranose units, and almostevery galactose unit is substituted at C-6 position.

2.2. Hardwood hemicellulose

The o-acetyl-4-o-methyl-glucurono-�-xylan (commonlyknown as glucuronoxylan) is the major component of hard woodhemicelluloses [6,27] (Fig. 1B). The xylan content varies between15 and 30% in different hardwood species. The backbone of xylanconsists of �-d-xylo-pyranose units linked by 1,4-bonds, whileseven of ten xylose units are substituted by acetyl group at C-2or C-3 position and in one of ten xylose units, the 4-o-methyl-�-d-glucuronic acid residue unit is linked at C-1, 2 positions tothe hemicellulose backbone [6,25,28]. Gluco-mannan is anotherhemicellulose in hard woods (Fig. 1B), comprises 2–5% of thewood and is composed of �-d-gluco-pyranose and �-d-manno-pyranose units by 1,4-bonds. Depending on the wood species, theglucose:mannose ratio varies between 1:1 and 1:2 [6,24,25,29].

3. Pentose-fermenting microorganisms

During the last more than three decades, several laboratoriesround the world have examined the utilization of pentose sugars bydifferent bacteria, fungi and yeasts for production of acids, alcoholsand other fermentation products under cultivation conditions [30].

3.1. Bacteria

Most of the fungi cannot ferment pentoses anaerobically, whilemany bacteria can readily convert xylose to various products underanaerobic fermentation. The common pentose fermenting bacteriainclude Bacillus macerans, Bacillus polymyxa, Klebsiella pneumo-niae, Clostridium acetobutylicum, Aeromonas hydrophila, Aerobactersp., Erwinia sp., Escherichia sp., Leuconostoc sp., Lactobacil-lus sp., Thermoanaerobacterium saccharolyticum, and Zymomonasmobilis [22,31]. Among bacteria, thermophiles may be the bestsuited for the production of alcohol, polyols and ketones dueto decreased cooling energy and low risk of contamination.Promising thermophilic pentose fermenting bacteria includeT. saccharolyticum, Clostridium thermohydrosulfurium, Clostridiumthermosaccharolyticum, Clostridium thermosulfurogenes, Clostridiumtetani and Thermoanaerobacter ethanolicus [32–34]. These ther-mophiles have many industrially important properties such as widesubstrate range, less biomass, no specific oxygen requirement, lessrisk of contamination and continuous recovery of volatile products.But, the low product tolerance and byproduct formation during thefermentation make these bacteria commercially unviable; how-ever, high temperature fermentation may save cooling energy ascompared to low temperature fermentation. Moreover, the geneticmodification of thermophilic organisms would solve the problemof product tolerance.

4952 R.C. Kuhad et al. / Renewable and Sustainable Energy Reviews 15 (2011) 4950– 4962

Fig. 1. (A) Structure of monosaccharides commonly present in xylan backbone: (i) �-d-glucopyranose; (ii) �-l-rhamnopyranose; (iii) �-l-fucopyranose; (iv) �-d-xylopyranose; (v) �-d-mannopyranose; (vi) �-d-galactopyranose and (vii) �-l-arabinofuranose; (B) structures of polymeric units of softwood and hardwood xylans.

3.2. Filamentous fungi

The filamentous fungi have been known to ferment sugarsfor more than 80 years. Several fungal species belonging to gen-era Chalara [35], Fusarium [36], Rhizopus [37], Neurospora [38],Paecilomyces [39] and Trichoderma [40] have potential for fer-menting xylose. Some other useful fungal strains have also beenstudied that can ferment more complex natural cellulosics sub-strates as well. Monilia sp., Neocallimastix sp., Trichoderma reeseiand Fusarium oxysporum have shown the ability for direct conver-sion of cellulose/hemicellulose to ethanol/acetic acid in single stepfermentation [40,41]. Despite the pentose fermentation character-istics, these fungi have several physiological drawbacks such as,long fermentation period, low ethanol productivity, high viscosityfermentation broth, requirement of low critical oxygen levels andformation of byproducts in large amounts. However, a filamentousfungal system might be interesting because of their ability to growon natural plant biomass, which yeast systems usually lack [22].

3.3. Yeast

The use of yeasts in conversion of carbohydrates to ethanolis known for generations. However, only a few strains are

capable of converting pentoses [21]. The extensively studied yeastspecies for xylose fermentation are P. tannophilus, C. shehatae, P.stipitis and Kluveromyces marxianus [19,20,22]. Many other yeastspecies are also reported for their xylose-fermenting capabilities,which include Brettanomyces, Clavispora, Schizosaccharomyces, sev-eral other species of Candida viz., C. tenius, C. tropicalis, C. utilis,C. blankii, C. friedrichii, C. solani and C. parapsilosis, and speciesfrom Debaromyces viz., D. nepalensis and D. polymorpha [42,43].Suh and coworkers have isolated a novel xylose-fermenting yeast‘Enteroramus dimorbhus’ from the microflora in the hindgut of bee-tles ‘Odontotaenius disjunctus’ [44]. Our group has also screened 20yeast strains for pentose fermentation recently, where only fewstrains of Pichia and Pachysolen showed pentose fermenting capa-bilities [45]. The pentose fermenting yeasts were observed to be lesstolerant to pH, ethanol and hydrolysate inhibitors when comparedto Saccharomyces cerevisiae. Moreover their inability to produceethanol as major end-product from xylose is a major drawback forethanol production [21,22,25].

4. Xylose metabolizing pathways

The initial metabolic pathway of d-xylose in microorganismsinvolves its conversion to d-xylulose followed by xylulose kinase


Fig. 2. Schematic diagram of d-xylose metabolism. (A) Xylose reductase; (B) xylitol dehydrogenase; (C) xylulokinase; (D) phosphoketolase; (E) transaldolase; (F) transketolase;(G) pyruvate decarboxylase; (H) alcohol dehydrogenase and (I) xylose isomerase.

reaction to d-xylulose-5-phosphate and then directed to pentosephosphate pathway (PPP). For d-xylose to d-xylulose conversion,bacteria generally use xylose isomerase (XI) enzyme, whereasyeasts and mycelial fungi employ a two step oxidation–reductionpathway [46]. In yeasts and fungi, the d-xylose is first reduced toxylitol by d-xylose reductase (XR) and subsequently oxidized to d-xylulose by xylitol dehydrogenase (XDH), which is further oxidizedto form d-xylulose-5-phosphate. d-Xylulose is further metabolizedvia pentose phosphate pathway (PPP), in which the non-oxidativerearrangements of �-xylose-5-phosphate by ribulosephosphate-3-epimerase, transaldolase (TAL) and transketolase (TK) resultsin the formation of glyceraldehyde-3-phosphate and fructose-6-phosphate, which can be converted to ethanol by fermentativereactions of the Embden–Meyerhoff–Parnas (EMP) pathway [47](Fig. 2).

Fermentative yeasts generally possess both aerobic and anaer-obic pathways along with adaptive regulatory mechanisms. Evenif they can metabolize d-xylose anoxically, d-xylose-fermentingyeasts require functional mitochondria and oxygen for growth,regardless of the carbon source. During typical aerobic xylose fer-mentation, ethanol concentration peaks sharply and then declines

as consumption exceeds production. However, ethanol uptake isnot observed in anaerobic xylose fermentation, where the TCA cycleis not operational [22,48].

5. Fermentation of pentose sugars

The general requirements of an organism for ethanol produc-tion from pentose sugar hydrolysate should be high ethanol yield,high productivity, good tolerance against inhibitors as well as highethanol concentrations and ability to ferment at relatively low pH.S. cerevisiae is one of the most commonly used yeasts for ethanolfermentation using glucose. However, it does not have the abilityof fermenting pentose sugars. The most promising yeast speciesidentified so far for the pentose fermentation are, C. shehatae, P.stipitis and P. tannophilus [19,20]. Various studies have been carriedout for the fermentation of xylose rich hydrolysates from differ-ent lignocellulosic materials (Table 1). Moniruzzaman achieved78% theoretical ethanol yield during the fermentation of enzy-matic hydrolysate of steam exploded rice straw, however, a 2–3 hlag due to diauxic phenomenal metabolic shift from glucose to


Table 1Summary of research work on pentose fermentation using lignocellulosic biomass.

Substrates Organism Sugar (g/l) Ethanol (g/l) Ethanol yield (g/g) Reference

Corn stover P. stipitis 40 15.92 0.4 [97]Prosopis juliflora P. stipitis 18 7.1 0.39 [9]Rice straw P. stipitis 33 14.9 0.45 [155]Sun flower seed hull P. stipitis 34 11 0.32 [52]Sugar cane bagasse Pachysolen tannophilus 63.5 19 0.34 [156]Sugar maple P. stipitis 35 12.4 0.35 [51]Corn stover P. stipitis 40 15 0.37 [50]Sugar cane bagasse Candida shehatae 30 8.67 0.29 [1]Corn stover P. stipitis 60 25 0.42 [157]Red oak wood chips P. stipitis 36 14.5 0.4 [158]Red oak spent sulfite liquor P. stipitis 49 20.2 0.41 [159]Wheat straw P. stipitis 52 22.3 0.43 [160]Corn cob Pichia stipitis 30 10.4 0.34 [161]Poplar P. stipitis 39 12 0.31 [162]Switch grass P. stipitis 39 14 0.36 [162]Rice straw Candida shehatae 20 9 0.45 [19]Rice straw P. stipitis 15 6 0.4 [49]L. camara P. stipitis 16.8 5.16 0.33 [12]

xylose was also observed [49]. In another study using the acid andthe auto-hydrolysate of rice straw, C. shehatae NCIM 3501 showedenhanced ethanol production in auto-hydrolysate (23.1 g/L) thanin acid hydrolysate (20.0 g/L) because of lower inhibitor concen-tration [19]. The pentose fermentation process does not requireintensive aerobic fermentation because of high cell mass synthesis,low ethanol yields and higher aeration energy consumption. How-ever, aeration is required for the biomass production, which couldbe a major problem during the fermentation of non-detoxifiedhydrolysate. Interestingly, the fermentation of non-detoxified cornstover hydrolysate at higher aeration improved the ethanol produc-tion which was due to the higher xylose consumption translatinghigher biomass concentration [50]. In a similar study the degree ofaeration showed a prominent effect on xylose utilization, ethanolproduction and xylitol minimization during the fermentation ofmembrane treated sugar maple hydrolysate using P. stipitis NRRLY-7124 [51]. Further, different detoxification strategies were usedby various researchers to enhance the ethanol production [1,52].The removal of toxic inhibitors from fermentation broth signif-icantly improved the ethanol yield (2.4-fold) and productivity(5.7-fold), compared to neutralized hydrolysate. Similarly, the fer-mentation of sugarcane bagasse acid hydrolysate with C. shehataeNCIM 3501 showed maximum ethanol yield (0.48 g/g) from ionexchange treated hydrolysate, followed by treatment with acti-vated charcoal (0.42 g/g), laccase (0.37 g/g), overliming (0.30 g/g)and neutralization (0.22 g/g) [1]. While in another study, thesequential application of overliming with sodium sulfite additionwas observed to be the best detoxification method for the sunflower seed hull acid hydrolysate, for maximum ethanol yield(0.32 g/g) and ethanol productivity (0.065 g/L/h) [52]. Recently,P. stipitis NCIM 3498 was used to ferment the detoxified acidhydrolysates of two lignocellulosic feedstocks (Prosopis juliflora andLantana camara), resulting in an ethanol yield ranging from 60 to70% of the theoretical yields [9,12].

Although significant work has been carried out on pentosefermentation, but no economically feasible process has been devel-oped so far. Therefore, in order to achieve improved ethanolproduction, the major focus of research on pentose fermentation isshifting toward the exploration of improved fermentation strate-gies and strain improvement.

5.1. Pentose fermentation using whole cell immobilization

The immobilization technique offers many advantages suchas cell recycling, economic extraction of the product, easy

maintenance of specific growth and dilution rate, maintenance ofhigh cell density, high productivity, good mixing and mass transferwith low risk of contamination [53–56]. Though the immobilizedmicroorganisms showed enhanced volumetric productivities ascompared to free microbes, however, in most of the cases, thexylose and glucose are not utilized at the same time due to catabo-lite repression [47]. In an attempt to ferment glucose and xylosesimultaneously, Grootjen and coworkers used a co-culture cul-tivation of alginate immobilized S. cerevisiae and P. stipitis in aconventional bioreactor, where P. stipitis cells inoculum were takenin comparatively higher amount, which allowed more xylose uti-lization under anaerobic conditions and the fermentation appearsto be simultaneous [53]. Furthermore, the modified stirred tankreactor (STR) system equipped with two teflon-made HPLC filtersair diffusers with improved mixing and less shearing during co-culture strategy improved the ethanol yield up to 80% with calciumalginate immobilized P. stipitis and S. cerevisiae [54]. Interestingly,an agar sheet sandwiched between two chambered bioreactor hasalso been used for the co-immobilization of S. cerevisiae and C. she-hatae during the mixed sugar (glucose and xylose) fermentation,however, the cell proliferation in the gel clogged the microporousmembrane, which in turn limited the mass transfer [57]. In anotherstudy to overcome the problem of glucose catabolite repression,sieve plates adjusted STR with a movable device was used for thecoculture of immobilized Z. mobilis and free P. stipitis to improve thefermentation efficiency [58]. Recently, a calcium alginate immobi-lized recombinant S. cerevisiae strain ZU-10 has been used for thefermentation of detoxified corn stover hemicellulosic hydrolysate,which showed the consumption of more than 92% xylose with anenhanced ethanol yield and productivities with higher tolerance tofermentation inhibitors [56].

Despite various improvements of these fermentations, therepeat culture with the same batch of immobilized microorgan-ism under the same conditions resulted in decreased performance[58]. Alternatively yeast cells immobilized by self flocculation haveshown many advantages such as no requirement of support matrix,maintained biomass and enhanced ethanol tolerance. Moreoverflocculated yeast cells can be recovered by sedimentation from fer-mentation broth. However, CO2 bubbles produced during ethanolfermentation can alter the settling zone and disturb the sedimenta-tion of yeast floc, whereas a specially designed baffle can overcomethis problem [59]. There are also some reports of yeast cells immo-bilization for biocatalysts development for simultaneous sacchari-fication and fermentation (SSF). Fujita and coworkers constructeda yeast-based whole-cell biocatalyst displaying T. reesei xylanase II


on the cell-surface and showed xylan degradation by recombinantcells [60]. Significant attempts have also been made for the develop-ment of yeast cells displaying cellulase activities on the cell surfaceso as to decrease the usage of exogeneously added cellulase [61,62].

5.2. Pentose fermentation using recycling of cells

Increasing the cell density by cell recycling and cell retentionhas been a suitable way of increasing the volumetric productivityfor slow growing microorganisms [63,64]. The cell recycling oper-ation requires lesser amount of nutrients, achieves high dilutionrate in continuous operations, decreases cell mass synthesis, andincreases ethanol yields [57,65]. A high productivity system thatinvolved a membrane bioreactor with cell recycling of Z. mobilisZM4 capable of converting both glucose and xylose to ethanolhad been developed [66]. Similar strategy was also applied dur-ing the continuous cultivation of a recombinant xylose fermentingS. cerevisiae TMB 3001 on a xylose-glucose mixture [64]. Interest-ingly, the recycled cells get adapted to the fermentation inhibitorspresent in the hydrolysate and showed better results in ligno-cellulose hydrolysate containing mixed sugars [63]. Pruwadi andcoworkers found similar results, where continuous cultivation ofhigh cell density flocculating yeast in toxic dilute acid hydrolysateof spruce residues in a single and serial bioreactor adapted to themedium and reduced the requirement of any detoxification [67].Recently, a fuzzy optimization of continuous fermentation withcell recycling for ethanol production was carried out [68]. From thecomputational results, the overall productivity of the continuousfermentation process with cell recycling allowed a higher dilutionrate with 7.3-fold higher productivity [68].

Though in most of the earlier reports, the cell enrichment wascarried out by cell resettling, centrifugation, microfiltration andultrafiltration, but recently hydroclone has been used to recoverthe cells of S. cerevisiae IROST 5209 from the fermentation broth[69]. The hydroclone system offers an advantage of negligible celldisruption [69]. Furthermore, implementation of cell recycling inindustrial process is possible only if the effluent passes the mem-brane without fouling. Industrial effluents are not suitable formembrane filtration due to suspended particles, solid residues andhigh viscosity. However, decreasing the viscosity in industrial efflu-ents by enzymatic treatment is one of the possible solutions of thisproblem [69,70].

5.3. Pentose fermentation using simultaneous saccharificationand fermentation

Simultaneous saccharification and fermentation (SSF) approachhas also been employed to decrease sugar inhibition to cellulase,achieve improved ethanol production, and to reduce the overallprocess cost since both the processes took place in single reactor.The SSF technique reduces the processing time, which in turn leadsto increase the apparent volumetric productivity of ethanol [71].The other advantages of this approach are shorter fermentationtime and reduced risk of contamination with external microfloradue to the high temperature of the process, the presence of ethanolin the reaction medium and the anaerobic conditions [30,71,72].Interestingly, in a comparison between the separate hydrolysis andfermentation (SHF) and SSF of steam exploded corn stover, it wasobserved that SSF gave a 13% higher overall ethanol yield than SHF(72.4% versus 59.1% of the theoretical) [72]. However, the perfor-mance of the SSF process is limited by the competence betweenoptimum temperatures for enzymatic saccharification and micro-bial fermentation [73]. Hence, the coexistence of two processconditions for low-temperature microbe and high-temperaturefungal enzyme may hinder the efficacy of the SSF process.

To improve process efficiency, many microorganisms have beentested with cellulases produced by T. reesei mutants [30]. Besides,there are some microorganisms such as Clostridium, Cellulomonas,Trichoderma, with high cellulolytic and hemicellulolytic activityand are highly capable of fermenting mono- or oligo-saccharidesinto ethanol. Application of thermotolerant yeast strains maybe beneficial in high temperature SSF processes, where fungalcellulase can also exhibit higher activity [73]. Ballesteros andcolleagues tested different treatments to improve the thermotol-erance of some species belonging to the genera Saccharomyces, andKluyveromyces and the best results were obtained with K. marxi-anus LG [74]. The strain was further mutagenised and the mutatedK. marxianus strain CECT 10875 achieved a SSF yield of 50–72% in72–82 h using various lignocellulosic feedstocks such as Populussp. and Eucalyptus sp., Sorghum sp. bagasse, wheat straw and Bras-sica carinata residue [75]. The efficacy of Kluyveromces was furtherconfirmed by other researchers as well, where Kluveromyces strainNCIM 3358 in SSF experiments of softwood resulted in more than70% ethanol conversion [76]. Similarly, during the SSF of lignocel-lulosic wastes with this thermotolerant yeast at 10% (w/v) initialsubstrate consistency, a final ethanol concentration of 2–2.5% wasobtained after 72 h [77].

In another strain improvement report, a thermophilic anaerobicbacterium T. saccharolyticum, capable of utilizing xylan and biomassderived sugars, was engineered by removing the genes encodingacetate kinase, phosphate acetyl transferase and d-lactate dehy-drogenase. The engineered thermophilic strain ALK2 thus obtainedwas able to reduce the requirement of externally added fungal cel-lulase by 2.5-fold [34]. Recently, a respiratory-deficient mutant ofthe thermotolerant yeast Candida glabrata (Cgrd1), was subjectedto ethanol production by high-temperature SSF under aerobic con-ditions to achieve maximum ethanol (17.0 g/L) within 48 h at 66.6%of its theoretical yield and with 0.35 g/L h productivity [78].

However, more efforts are required in the development ofmicroorganisms for economically feasible industrial ethanol pro-duction from pentose sugars. Since enzymatic hydrolysis is therate-limiting step in SSF, increasing the rate of hydrolysis will lowerthe cost of ethanol production via SSF [30].

6. Strain improvement for pentose fermentation

6.1. Strain improvement through mutagenesis

There are several reports where mutagenised recombinantstrains showed enhanced ethanol production over their parentstrains [4,79–81]. In an early report, a recombinant S. cerevisiaestrain TJ1 mutagenised with ethyl methane sulfonate (EMS) wasfound to have lower XR activity but high XD and xylulokinase(XKS) activities than the parent strain, which in turn resulted in1.6-fold increase in ethanol production [82]. Further, a mutant ofS. cerevisiae TMB 3001 capable of utilizing xylose under anaero-bic condition was developed by sequential EMS mutagenesis andadaptation of the mutant strain under microaerobic and anaero-bic conditions [83]. Later on similar strategy was used to developtwo EMS mutagenised S. cerevisiae strains 3399 and 3400 withimproved growth on xylose [80]. Since the most efficient xyloseutilizing microbes are not able to metabolize the pentose sug-ars anaerobically, the strategies of natural selection and randommutations were also tested [84]. Besides EMS, several other muta-gens had also been used to obtain mutants derepressed for pentosemetabolism. Sreenath and Jeffries used 2-deoxyglucose (2-DOG)mutated strain, showing considerable improvement in xylose uti-lization [63]. While in another report, a UV mutagenised P. stipitisNRRL Y-7124 strain was found to produce higher ethanol than thewild strain [85].


Site-directed mutagenesis is another strategy used to obtain themutants for better xylose fermentation. Watanabe and coworkersused multiple site-directed mutagenesis of the NAD+-dependentXDH from P. stipitis and introduce a structural zinc atom for thecomplete reversal of the coenzyme specificity [81]. The selectedmutants were found to exhibit significant thermostability andenhanced catalytic activity with NADP+. Similarly, several PsXDHmutants were generated with complete reversal of coenzymespecificity toward NADP+ by multiple site-directed mutagenesiswithin the coenzyme-binding domain and with increased ther-mostability by refining the structural zinc-binding loop withoutaffecting their activities [86]. In addition one of the S. cerevisiaemutant (MA-R5) under the control of a strong constitutive pro-moter showed particularly high ethanol production from xyloseand low xylitol yield by fermentation of not only xylose as the solecarbon source, but also a mixture of glucose and xylose [87–89].Additionally using this approach, an ethanologenic Escherichia colimutant, devoid of foreign genes, has also been developed bycombining the activities of pyruvate dehydrogenase and the fer-mentative alcohol dehydrogenase and the mutant was found able toferment glucose or xylose to ethanol with 82% ethanol yield underanaerobic conditions [79].

6.2. Strain improvement through protoplast fusion

Protoplast fusion provides characteristic advantage such aspromotion of high frequencies of genetic information betweenorganisms for which poor or no genetic exchange has been demon-strated or which are genetically uncharacterized [90–92]. In thepresence of a fusogenic agent such as polyethylene glycol (PEG),protoplasts are induced to fuse and form transient hybrids ordiploids. Several reports on protoplast fusion between pentose andhexose utilizing yeasts showed efficient utilization of both sug-ars with higher biomass yield. Heluane and coworkers successfullytransferred the genes of xylose utilization from P. tannophilus toS. cerevisiae [90]. The hybrids reassembled the S. cerevisiae parentmorphologically but displayed the ability to use the pentose sug-ars (xylose) similar to P. tannophilus. The same has been supportedby other workers, where a fusant of Schizosaccharomyces pombeand Lentinula edodes were found to utilize xylan as carbon source[91]. In an another study, the protoplasts of thermotolerant S. cere-visiae and mesophilic xylose-utilizing C. shehatae were fused byelectrofusion and the fusant yeast gave an ethanol yield of approx-imately 0.459 g/g with productivity of 0.67 g/L/h and fermentationefficiency of 90% and showed higher temperature tolerance up to40 ◦C as well [92]. Moreover, using a combinatorial approach, axylose fermenting fusant (F6) of C. shehatae and S. cerevisiae wasdeveloped, showing improved ethanol production (28%) than itsparental strain [93]. In this strategy the C. shehatae was first adaptedfor ethanol tolerance and then mutagenised by UV irradiation andthus a respiration deficient mutant RD-5 was selected. Further, theprotoplasts of RD-5 and S. cerevisiae were fused and the resultantfusant strain F6 showed 28% higher ethanol production than theparent C. shehatae stain, with the production level of 18.75 g/L from50 g/L xylose. Recently a strategy of genome shuffling was also used,in which the genomes of 6 UV mutagenised P. stipitis strain (WT,PS302, GS301, GS302, GS401 and GS402) were shuffled and afterthe 3rd and 4th rounds of genome shuffling, putative improvedmutant colonies were pooled, re-grown and spread on hardwoodspent sulphite liquor (HWSSL) gradient plate again [94]. P. stipitisWT and PS 302 could not grow in any of the HWSSL concentrations,while 2 mutants (GS401 and GS402) from the 4th round could growin 80% (v/v) HWSSL while another 2 mutants (GS301 and GS302)from the 3rd round could grow in 85% (v/v) HWSSL. Thus the studyconcluded that the mutated strains showed improved inhibitorstolerance against HWSSL [94].

6.3. Strain improvement through adaptation

Fermentation of wood-derived hydrolysates is sometimesproblematic because of the toxic inhibitors released during thermo-chemical hydrolysis. However, the adaptation approach can be analternative means to improve the microbial strains [95–97]. Thereare several reports on enhancement of ethanol yield and produc-tivity using adapted strains of P. stipitis and C. shehatae for thefermentation of undetoxified or partially detoxified hydrolysates[97–99]. For instance, an ethanologenic yeast when adapted againstinhibitors by repeated sub-culturing in a medium with furfural andHMF up to a concentration of 10–20 mM was found to grow moreefficiently than its parent strain in the presence of inhibitors [98].Another strategy of natural selection and breeding was used todevelop non-recombinant strains of S. cerevisiae that could growefficiently on xylose [99]. By breeding and natural selection over 23mating cycles and 1463 selection days, a non-genetically modifiedS. cerevisiae (MBG-2303) was obtained, which grew aerobically onxylose and demonstrated 57-fold higher biomass production thanthe control strain [99]. Later on, it was demonstrated that adap-tation of P. stipitis CBS 6054 in solid agar produce more ethanol(19.4 g/L) than liquid adapted (18.4 g/L) and unadapted strains(16.3 g/L) [95]. Recently, studies were carried out on adaptation of P.stipitis CBS 5776 strain which on fermentation of steam explodedprehydrolysate of corn stover showed improved ethanol yield of15.92 g/L with 80.34% theoretical yield [97].

Moreover, the evolutionary adaptation approaches have alsobeen applied to recombinant strains to improve their fermenta-tion capability. Lawford and group improved the xylose-fermentingrecombinant strain Z. mobilis 39767 to tolerate higher concentra-tion of acetic acid by subculturing in a medium containing 10–50%of hydrolysate and the adapted isolates demonstrated a signifi-cant improvement in ethanol productivity compared to un-adaptedstrains [100]. Similarly, an engineered E. coli KO11 was developedto tolerate high ethanol concentration using a long term adapta-tion strategy of alternative serial selections for liquid and solidmedium. The mutants thus developed, i.e., LY01, LY02 and LY03demonstrated more than 50% survival rate in 10% ethanol (0.5 minexposure) and also reduced the fermentation time [96]. In almost allprevious efforts of evolutionary adaptation the organism was firstsubjected to genetic engineering, which was followed by adaptiveselection [83,101,102]. However, recently a new strategy consistinggenetic engineering, mutation with EMS followed by two-step evo-lutionary adaptation (under sequential aerobic and oxygen limitedconditions) has also been attempted [4]. The strain thus developedshowed fourfold increase in its specific growth rate compared tothe parental strain. Interestingly the activity of critical enzymes ofxylose metabolism (XR, XDH and XK) remain unchanged suggest-ing that chemical mutagenesis and evolutionary adaptation mighthave created a new genetic traits making the mutants capable ofxylose metabolism [4].

6.4. Strain improvement through genetic manipulation

Substantial progress in the genetic engineering of differentmicrobes for the conversion of xylose or pentose sugars to ethanolhas been achieved [4,88,103]. Although the genetically engineeredhost strains of bacteria and yeast showed tremendous improve-ment in final ethanol yields and efficient utilization of pentosesugars (Table 2), the information about the usage of geneticallymodified organisms for large scale pentose fermentation is scarcelyavailable [25].

6.4.1. Genetic engineering of E. coliIngram’s group have done extensive work on the develop-

ment of efficient recombinant E. coli strains for ethanol production.


Table 2List of pentose utilizing recombinant yeasts and bacterial strains.

Strain Sugar/sugar mix*

used (g/l)Ethanolproduction (g/l)

Ethanol yield (g/g) Ethanolproductivity (g/l/h)

Reference

E. coli KO11 80 X 41.6 102 0.87 [107]E. coli KO11 90 X 41 89 0.85 [96]E. coli KO11 140 X 59.5 [96]E. coli LY01 140 X 63.2 88 0.66 [96]E. coli FBR5 95 X 41.5 90 0.59 [110]E. coli FBR5 A:X:G

15:30:3034.0 90 0.92 [110]

E. coli SE2378 60 X 27.8 82 NA [79]T. saccharolyticumALK-2

70X 33.1 92 2.2 [41]

Z. mobilis CP4 25 X 11.0 86 0.57 [115]Z. mobilis CP4 G:X

65:6524.2 95 0.81 [115]

Z. mobilis ATCC 39767 G:X:A30:30:20

33.5 82-84 0.82-0.65 [117]

Z. mobilis CP4 60 X 23 94 0.32 [163]Z. mobilis ZM4 G:X

65:6562 90 1.29 [66]

Z. mobilis AX101 A:G:X20:40:40

42 84 0.61 [119]

S. cerevisiae PRD1 21.7 X 1.6 14 0.07 [164]S. cerevisiae TJ1 50 X 2.7 10.6 0.02 [165]S. cerevisiae 1400 G:X:A:Gal

31:15:10:222 90 0.92 [166]

S. cerevisiae 1400 80 X 27 66 1.12 [166]S. cerevisiae 1400 50 X 1.5 6 [121]S. cerevisiae H158 80 X 21.6 54 NA [123]S. cerevisiae TMB 3008 50 X 19 76 0.27 [48]S. cerevisiae TMB 3251 50 X 17 68 0.24 [48]S. cerevisiae TMB 3255 50 X 20.5 82 0.29 [48]S. cerevisiae H 2684 50 X 20.5 82 NA [131]S. cerevisiae TMB 3400 20 X 5 50 NA [80]S. cerevisiae TMB 3001 10 X 2.4 48 NA [83]S. cerevisiae BH42 G:X

50:5028 56 NA [167]

S. cerevisiae TMB 3120 10 X 4.6 92 0.064 [168]S. cerevisiae TMB 3050 50 X 14.5 58 NA [103]S. cerevisiae RWB 217 20 X 8.6 86 NA [101]S. cerevisiae TMB 3270 50 X 18 72 0.32 [135]S. cerevisiae TMB 3400 50 X 17 68 0.12 [128]S. cerevisiae L2162 40 X 5.2 0.13 0.032 [130]S. cerevisiae DR PHO13 40 X 10 0.25 0.093 [130]S. cerevisiae TMB 3066 50 X 21.5 86 0.073 [128]S. cerevisiae CMB JHV 20 X 6.4 0.32 NA [169]S. cerevisiae BP10001 20 X 6.8 0.34 NA [139]S. cerevisiae MA-N5 45 X 15.8 0.36 0.24 [88]S. cerevisiae RBW202-AFX

50 X 19.5 78 NA [170]

S. cerevisiae INVSC1 50 X 19.5 0.39 0.043 [171]S. cerevisiae ADAP8 20 X 8.6 0.43 0.07 [172]S. cerevisiae MA-R4 45 X 15.75 0.35 0.36 [84]S. cerevisiae MA-R5 45 X 16.65 0.37 0.50 [84]S. cerevisiae ZU-10 80 X 30.2 75.6 0.50 [56]S. cerevisiae LEK 513 50 X 8.13 32.5 0.113 [4]

* A, arabinose; G, glucose; Gal, galactose; M, maltose; and X, xylose.

They eliminated the dependence of host on alcohol dehydrogenase(ADH) activity by combining adh B and pdc (coding for pyru-vate decarboxylase, PDC) genes of Z. mobilis to form pet operon[104,105]. Considering a number of factors influencing ethanol pro-duction such as substrate range and growth conditions, E. coli strainATCC 11303 was chosen as the host for the pet plasmid [106]. Fur-ther, to improve the genetic stability, the pet operon was integratedinto the chromosome of ATCC11303 [99]. Since the strain contain-ing the integrated genes produced only low levels of ethanol, thespontaneous mutation strategy generated a hyper ethanol produc-ing KO4 strain. This strain was further modified by deletion of thesuccinate production gene (frd) to prevent the formation of succi-nate, a major byproduct of E. coli metabolism. The finally developed

strain KO11 was able to convert glucose and xylose to ethanol attheoretical yields of 100% in rich media containing ample yeastextract [107]. To further improve pentose fermentation by KO11,a number of spontaneous mutants defective in glucose transportwere selected and two such strains SL28 and SL40 when fermentedusing individual or mixture of xylose and glucose produced ethanolmore efficiently (by 20%) than the parent strain KO11 [108]. Therecombinant E. coli strain was further improved to achieve bet-ter xylose fermentation, the glycolytic flux and the growth rateof recombinant strain [109]. However, from an industrial point ofview, these recombinant strains still had the drawback of requiringnutrient rich medium for ethanol production. In order to overcome,a lactate producing recombinant of KO11 was reengineered for


ethanol production by deleting genes encoding for fermentativeroutes for NADH and randomly inserting a promoter-less cassettecontaining the complete Z. mobilis ethanol pathway into KO11 chro-mosome [96].

Using different approach, Dien and coworkers developed newethanologenic strains of E. coli such as FBR3, FBR4 and FBR5 [110].These strains were generated by transforming a xylose-utilizingisolate of strain E. coli FMJ39 with plasmid pLOI297 having Z. mobilispyruvate to ethanol converting enzymes. Alternatively a homo-ethanologenic strain of E. coli SE2738 from wild type E. coli K-12W3110 was also developed, where the mutant strain exhibited 82%theoretical ethanol yield when grown on xylose under anaerobicconditions [79]. In a recent study, E. coli cells for efficient ethanolproduction from hexoses and pentoses were developed using ele-mentary mode analysis to dissect the metabolic network into itsbasic building blocks [111]. During this strategy the functionalspace of the central metabolic network was reduced, with eightgene knockout mutations, from over 15,000 pathway possibilitiesto 6 pathway options that support cell function. Furthermore theelimination of three additional genes resulted in a strain that selec-tively grows only on pentoses, even in the presence of glucose, witha high ethanol yield. Later on using the similar approach, a glycerolto ethanol converting E. coli strain was designed by reducing thefunctional space of central carbon metabolism to a total of 28 glyc-erol utilizing pathways [112]. More recently an attempt has beenmade to engineer E. coli for the production of ethanol from fatty acidfeedstocks, resulting in ethanol yield higher than the theoreticalmaximum obtained from sugars [113].

6.4.2. Genetic engineering of Z. mobilisThe xylose utilization in Z. mobilis was developed by inte-

grating XI and XKS from E. coli, Xanthomonas campestris and K.pneumoniae in its genome [114,115]. Using the similar approach,Zhang and coworkers developed recombinant Z. mobilis CP4 show-ing activity for all four enzymes (XI, XKS, tal and tkt), whichwhen grew on xylose as the sole carbon source resulted in 86%theoretical ethanol yield [115]. In another work, the constructedoperons encoding xylose assimilation and pentose phosphate path-way enzymes were transformed into Z. mobilis to generate pZB5strain for the effective fermentation of xylose to ethanol [36,115].Further to construct improved strains with higher ethanol produc-tivities and yields, pZB5 was transformed into Z. mobilis ethanolproducing strain ZM4; ATCC 31821, which showed the capabilityof converting a mixture of 65 g/L of glucose and 65 g/L of xylose to62 g/L ethanol in 48 h with an overall yield of 0.46 g/g [66]. Fol-lowing similar approach, another group incorporated five genesof arabinose utilization from E. coli ara A (coding for l-arabinoseisomerase), ara B (coding for l-ribulokinase), ara D (coding for l-ribulose-5 phosphate-4-epimerase), tal and tkt in Z. mobilis ATCC39767 [116]. The resultant strain showed more than 90% ethanolyield from l-arabinose. However, 40% of the cells lost their abilityto ferment arabinose when grown on complex medium. A numberof other improvements have also been made in Z. mobilis strainsand the new strain Z. mobilis AX101 fermented both arabinose,xylose and glucose and carried seven necessary recombinant genesas part of chromosomal DNA [117–119]. The co-fermentation pro-cess yield from Z. mobilis AX101 was about 84%, with preferentialorder in sugar utilization as glucose followed by xylose and arabi-nose [119]. However, these strains showed acetic acid sensitivity[119]. To address the problem of sensitivity to toxic fermenta-tion inhibitors, a new strain of Z. mobilis ZM4/AcR (pZB5) wasdeveloped with increased acetate resistance that has enhancedperformance in the presence of 12 g sodium acetate per litre atpH 5 [61]. Z. mobilis ZM4 produced near theoretical yields ofethanol with high specific productivity and was able to fermentboth C-5 and C-6 sugars. The transformed Z. mobilis ZM4 performed

best under anaerobic conditions, but also exhibited tolerance toaerobic conditions. However, the genetic and physiological basisof ZM4’s response to various stresses has only been understoodpoorly. Recently, transcriptomic and metabolomic profiles for ZM4under aerobic and anaerobic fermentations have been elucidatedusing microarray, high-performance liquid chromatography andgas chromatography–mass spectrometry (GC–MS) analysis [120].

6.4.3. Genetic engineering of S. cerevisiaeS. cerevisiae produces ethanol from hexose sugars but cannot fer-

ment xylose or arabinose. However, the yeast is able to metabolizea xylose isomer, xylulose and recombinant DNA technologists havetaken advantage of this in creating xylose-fermenting strains. Hoand coworkers were the first to successfully create a recombinant S.cerevisiae strain capable of effective xylose fermentation and xyloseand glucose co-fermentation [121]. The recombinant plasmids withXR and XDH genes from P. stipitis and XKS gene from S. cere-visiae were transformed into S. cerevisiae for the co-fermentationof glucose and xylose. In contrasting report the overexpression ofendogenous XKS from S. cerevisiae was found to inhibit its growthon xylulose [122]. Similarly some other workers have also reportedabout lower consumption of xylose in such strains [123–125].However, Toivari and coworkers reported successful xylose fer-mentation to ethanol through over-expression of the endogenousXKS 1 and PsXR and XDH genes [126]. Recently following similarstrategies, the improved xylose utilization and high ethanol pro-duction have been reported by various groups [88,127–129]. Basedon these reports it can be concluded that the controlled overexpres-sion of XKS gene in S. cerevisiae improved the xylose consumption aswell as ethanol production. Jeffries and coworkers identified someinteresting spontaneous or chemically induced mutants of recom-binant S. cerevisiae that can overcome the growth inhibition causedby overexpression of ScXKS and PsXKS gene [130].

In order to achieve reduction in xylitol formation during xylosefermentation, recombinant S. cerevisiae strains expressing PsXR andPsXDH and overexpression of ScXKS were constructed that loweredthe oxidative PPP activity through the GND1 (6-phosphogluconatedehydrogenase) and ZWF1 genes (glucose-6-phosphate dehydro-genase) [48]. These mutants showed increase in ethanol yieldand xylose consumption rate compared to the parent strain. Fur-thermore an attempt to overexpress Kluyveromyces lactis GDP1gene (NADP-dependent glyceraldehyde 3-phosphate dehydroge-nase, GAPDH) in a xylose-fermenting S. cerevisiae strain was alsofound to enhance ethanol production [131].

In another approach to enhance ethanol yields, the metabolicflux toward ethanol formation appeared to be a significant strategyto improve the intracellular cofactor concentrations in S. cerevisiae[132]. The impact of over-expression of NADH kinase (encodedby the POS5 gene) on glucose and xylose metabolism in recom-binant xylose-utilizing S. cerevisiae has also been studied [133].The expression of NADH kinase in cytosol instead of mitochon-dria redirected the carbon flow from CO2 to ethanol during aerobicgrowth on glucose, whereas under anaerobic growth the fluxdirected toward ethanol and acetate fermentation. The cytosolicNADH kinase appeared to revert these effects during anaerobicmetabolism of xylose by channeling carbon flow from ethanol toxylitol [133]. The heterologous expression of a xylose isomerase(XI) may also be a good approach to enable yeast cells to metabo-lize xylose. In this aspect, Brat and coworkers screened nucleic aciddatabases for sequences encoding putative XIs and cloned them toexpress a highly active XI from the anaerobic bacterium Clostrid-ium phytofermentans in S. cerevisiae, which resulted in an efficientmetabolism of d-xylose as the sole carbon and energy source byrecombinant yeast cells [134].

Alternative protein engineering approach has also been inves-tigated to reduce xylitol formation and enhancing ethanol yield


using recombinant S. cerevisiae. Using this approach an improvedethanol production accompanied by decreased xylitol formationwas achieved in recombinant S. cerevisiae expressing mutated PsXR(having reduced affinity for NADPH), PsXDH and ScXKS [135]. In thisaspect, several NADH-preferring XR mutants from Candida tenuishave been developed [136–139]. Recently one such S. cerevisiaestrain harboring the K274R-N276D CtXR double mutant showedenhanced ethanol production with decreased xylitol formation[140]. The heterologous expression of xylose specific transportersin recombinant S. cerevisiae for improved ethanol production hasalso been tested. The SUT1 gene [141] coding a sugar trans-porter in P. stipitis, has been successfully expressed in S. cerevisiae[142]. Moreover, the glucose/xylose-facilitated diffusion trans-porter and glucose/xylose symporter from Candida intermedia,encoded by Gxf1 and Gxf2 genes [143] have been expressed inS. cerevisiae [144], where the recombinant xylose-fermenting S.cerevisiae strain harboring Gxf1 showed faster xylose uptake andethanol production [145]. Recently, a combinatorial approach ofgenetic engineering, chemical mutagenesis and evolutionary adap-tation has been used to improve the xylose utilization. The S.cerevisiae strain W303-La was introduced with XI and XKS genefrom P. stipitis NRRL7124 to make S. cerevisiae LEK 122. Thereafter,the selected strain was chemically mutagenised with EMS followedby their evolutionary adaptation for xylose utilization and growthunder oxygen-limited conditions [4].

7. Future prospects

Ethanol has always been considered a better alternate to gaso-line, as it reduces the dependence on fossil fuel reserves andpromises cleaner combustion leading to a healthier environment.Interestingly, the world’s focus is switching over from corn and sug-arcane to second generation feedstocks as renewable raw materialfor production of bioethanol [146–149]. In recent years, signifi-cant developments in hemicellulose to ethanol conversion havebeen achieved. However the industrial activities for bioethanolproduction are limited mainly because of the cost of raw mate-rial (lignocellulosic biomass) processing to obtain high yield offermentable sugars and unavailability of efficient fermentationstrategies. The process of hemicellulose conversion to bioethanolrequires adequate breakdown of lignocellulosic biomass with max-imum pentose sugar yield and efficient utilization of pentose sugarsduring fermentation. Although various pretreatment methods andtheir effects on biomass composition and sugar yields have widelystudied and described in literature but efficient fermentation ofhigh yield hemicellulosic hydrolysates is essential to maximizethe ethanol productivity and subsequent cost effective ethanolrecovery [25]. The generation of microbial inhibitors during thepretreatment process is a major concern, which affects the eco-nomics of bioethanol production from lignocellulosic materials.Different detoxification strategies have been applied to removethese inhibitors from hemicellulosic hydrolysates to improve theirfermentability [9,12,21,150], this additional step increases theoverall ethanol production cost. There is an imperative need forbioprospecting of new robust microbial strains capable of convert-ing the pentose sugars efficiently from un-or partially detoxifiedhydrolysates [151].

Since the efficient utilization of pentose sugars is a prerequi-site for cost effective and sustainable ethanol production, the majorresearch has now focused on further improving ethanol productiv-ities using genetic modification and chemical additives [152,153].Although several research groups have taken lead in developinga number of improved xylose utilizing recombinant organisms,it appears that xylose utilization and fermentation capabilitiesare commercially unattractive. Therefore, the genetic engineering

approaches should be more focused on developing new improvedstrains with higher substrate tolerance and improved productionkinetics. Additionally, the better elucidation of pentose sugarstransport at the molecular level and characterization of kineticand regulatory properties, including quorum-sensing mechanisms,should be given high priority because it may provide the basisfor developing improved strains that can simultaneously utilizepentose and hexose sugars released during biomass hydrolysis.High-throughput screening techniques and better expression sys-tems for efficient production of membrane proteins, and enzymecomplexes such as cellulosomes are in progress.

Recently, the chromosomal integration of genes encoding thexylose utilization pathway enzymes into industrially applied yeaststrains are reported necessary for large scale fermentation of xylosefrom lignocellulosic biomass to ethanol [89]. Therefore, furtherstudies at micro levels are also required to combine the func-tional genomics with metabolic engineering strategies, which mayprovide further clues for developing robust strains. Moreover, theapproach of flux analysis to divert or increase the activity of cer-tain crucial enzymes in ethanol production with efficient xyloseutilization needs more attention.

To make the bioethanol production process successful at indus-trial scale with reduction in capital and operation cost, someintegrated unit operations using robust microorganisms for betterproduct yields should be adopted [154]. An ideal up-scaling strat-egy needs to be fully integrated to evaluate the complete system(e.g., enzymes, nutrients, product yields and titers, and fermentingyeasts) with sufficient flexibility to investigate alternative processconfigurations. From a process scale-up perspective, the challengeslie not only with finding the most efficient organism for hemicel-lulose conversion but also to make an intelligent use of the entireprocess integration, a biorefinery concept.

Acknowledgements

The authors are thankful to Prof. P. Tauro, Manglore Biotech,Manglore, India and Prof. P. Gunasekaran, Department of Micro-bial Genetics, Madurai Kamaraj University, India for their help inediting the manuscript. The authors also acknowledge the finan-cial assistance from Department of Biotechnology (DBT), Councilof Scientific and Industrial Research (CSIR) and University of Delhi,Delhi India.

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Bioethanol production from Lantana camara (red sage): Pretreatment,saccharification and fermentation

Ramesh Chander Kuhad a,*, Rishi Gupta a, Yogender Pal Khasa a, Ajay Singh b

aDepartment of Microbiology, University of Delhi South Campus, Benito Juarez Road, New Delhi 110021, Indiab Lystek International Incorporation, 107-279 Weber Street North, Waterloo, Ontario, Canada N2J3H8


Article history:Received 18 March 2010Received in revised form 26 May 2010Accepted 7 June 2010Available online 2 July 2010

Keywords:Lantana camaraCellulosic ethanolDilute acid hydrolysisEnzymatic hydrolysisFermentation

a b s t r a c t

Lantana camara contains 61.1% (w/w) holocellulose and can serve as a low-cost feedstock for bioethanolproduction. Acid hydrolysis (3.0%, v/v H2SO4, 120 �C for 45 min) of L. camara produced 187.14 mg/g totalsugars along with fermentation inhibitors such as phenolics (8.2 mg/g), furfurals (5.1 mg/g) and hydroxymethyl furfurals (6.7 mg/g). Sequential application of overliming (pH 10.0) and activated charcoal (1.5%,w/v) adsorption was used to remove these toxic compounds from the acid hydrolysate. The acid-pre-treated biomass of L. camara was further delignified through combined pretreatment of sodium sulphite(5.0% w/v) and sodium chlorite (3.0% w/v), which resulted in about 87.2% lignin removal. The enzymatichydrolysis of delignified cellulosic substrate showed 80.0% saccharification after 28 h incubation at 50 �Cand pH 5.0. Fermentation of acid and enzymatic hydrolysates with Pichia stipitis and Saccharomyces cere-visiae gave rise to 5.16 and 17.7 g/L of ethanol with corresponding yields of 0.32 and 0.48 g/g after 24 and16 h, respectively.

� 2010 Elsevier Ltd. All rights reserved.

1. Introduction

Lantana camara, commonly known as red sage, is one of theworld’s top 100 worst invasive species. It has invaded millions ofhectares of grazing land globally (Day et al., 2003). In Australiaalone, since its introduction as an ornamental plant in the 1840’sit has spread to infest four million hectares (www.weeds.org.au).While in India, the weed has invaded most of the tropical and sub-tropical parts and is found in areas from the seacoast to 5000 ft inaltitude (Sankaran, 2007). The approximate total biomass pro-duced by L. camara per year ranges from 15 to 17 tonnes/ha (Bhattet al., 1994), which projects the availability of Lantana biomass inhuge quantity. Therefore, the abundance of L. camara biomass (lig-nocellulosic material) is likely to offer a potential feed stock forethanol production.

Currently, the biomass to ethanol conversion technology reliesmainly on chemical and enzymatic treatments. Chemical hydroly-sis of biomass with dilute sulphuric acid has long been recognizedas a critical step for removing the hemicellulosic fraction from thelignocellulosic substrate to economize the biological conversion ofcellulosic biomass to ethanol. However, the pentose sugar-rich acidhydrolysate also contains toxic byproducts such as furfural,

hydroxy methyl furfural (HMF) and phenolics, which significantlyaffect yeast cell metabolism during fermentation (Palmqvist andHahn-Hagerdal, 2000; Chandel et al., 2007). Although variousdetoxification methods have been investigated for the removal offermentation inhibitory compounds (Palmqvist and Hahn-Hager-dal, 2000; Chandel et al., 2007), of which overliming and activatedcharcoal adsorption methods are widely used (Miyafuji et al.,2003; Gupta et al., 2009).

An appropriate delignification strategy is essential for the effi-cient enzyme hydrolysis of cellulosic biomass as lignin hindersthe saccharification process. The cellulase components such asb-glucosidase and endoglucanase showed higher binding affinitytowards lignin compared to carbohydrates, which in turn loweredthe saccharification efficiency (Kaya et al., 2000). Various delignifi-cation approaches have been exploited in the past such as alkalipretreatment (Carillo et al., 2005), hydrogen peroxide pretreat-ment (Saha and Cotta, 2007), sulphite pretreatment (Kuhad et al.,1999), ammonia fiber expansion pretreatment (Teymouri et al.,2005) and sodium chlorite pretreatment (Gupta et al., 2009).

The efficient conversion of biomass into ethanol requires opti-mum utilization of both pentose and hexose sugars. The widelystudied yeast for hexose fermentation, Saccharomyces cerevisiae,is unable to utilize pentose sugars while, among the pentose fer-menting yeasts only Pichia stipitis, Pachysolen tannophilus and Can-dida shehatae are proven to be highly efficient in the conversion ofxylose to ethanol (Abbi et al., 1996a,b). The fermentation of bothtypes of sugars can be carried out either by co-cultivation or

0960-8524/$ - see front matter � 2010 Elsevier Ltd. All rights reserved.doi:10.1016/j.biortech.2010.06.043

* Corresponding author. Address: Lignocellulose Biotechnology Laboratory,Department of Microbiology, University of Delhi South Campus, Benito JuarezRoad, New Delhi 110021, India. Tel.: +91 124 24112062; fax: +91 11 24115270.

E-mail address: [email protected] (R.C. Kuhad).

Bioresource Technology 101 (2010) 8348–8354


Bioresource Technology

journal homepage: www.elsevier .com/locate /bior tech

through separate fermentation. Co-cultivation has several draw-backs such as the diauxic phenomenon (inefficient utilization ofxylose in presence of glucose) (Grootjen et al., 1991), and competi-tion between organisms for nutrients requirements. Recently somestudies on separate fermentation of individual hydrolysates usingsugar specific yeast strains have been reported (Gupta et al.,2009; Laser et al., 2009).

The aim of the present work was to extract maximum sugarswith minimum inhibitory compounds from lignocellulosic sub-strate, L. camara, using less stringent acid hydrolysis conditions.We attempted to remove the toxic substances produced duringacid hydrolysis employing sequential overliming and activatedcharcoal adsorption. The delignification of pretreated biomasswas carried out by a combinatorial approach using sodium chloriteand sodium sulphite. The pentose and hexose sugar hydrolysateswere subsequently fermented separately using P. stipitis and S.cerevisiae.

2. Methods

2.1. Biomass collection and preparation

L. camara was collected from Aravali Hills, University of DelhiSouth Campus, New Delhi, India. Dried substrate was comminutedby the combination of chipping and milling to attain a particle sizeof 1–2 mm using laboratory Knife mill (Metrex Scientific Instru-mentation Pvt. Ltd., New Delhi, India). The chopped plant materialwas washed thoroughly with tap water and dried overnight at60 �C.

2.2. Biomass composition analysis

The chemical composition of L. camara was analysed for holo-cellulose, Klason lignin, pentosans, ash and moisture content. Theraw material was consecutively extracted with alcohol–benzene(1:2, v/v) mixture. The extractive-freeL. camara dust was processedfor chemical analyses following the TAPPI (1992) protocols.

2.3. Optimization of dilute acid pretreatment

The optimization of dilute acid hydrolysis was carried out at dif-ferent temperatures (100–140 �C), time periods (30 and 45 min)and H2SO4 concentration (1–5%, v/v) at 10.0% (w/v) solid content.The hydrolysates recovered were filtered through double-layeredmuslin cloth, and the remaining biomass was washed properlywith tap water to a neutral pH. The hydrolysate was analysed forthe amounts of sugar, furans and phenolics released; and the left-over biomass was dried overnight at 60 �C prior to delignification.

2.4. Detoxification of acid hydrolysate

The acid hydrolysate of L. camara was mixed with calciumhydroxide to raise the pH of the hydrolysate to 10.0. The wholeslurry was stirred for 30 min by an overhead stirrer (Remi MotorsLtd., Mumbai, India). After bringing to room temperature, thehydrolysate was neutralized with concentrated H2SO4 and thencentrifuged (10,000g, 15 min). The overlimed hydrolysate was fur-ther detoxified by mixing 1.5% (w/v) activated charcoal with con-tinuous stirring for 30 min at room temperature and theresulting sugar was recovered using vacuum filtration.

2.5. Delignification

The acid-pretreated residue of L. camara was treated with dif-ferent dosages of sodium sulphite (5.0–20.0% w/v) alone and in

combination with sodium chlorite (3.0% w/v) at different temper-atures (100–140 �C) and time intervals (30 and 45 min). The del-ignified biomass was then filtered through double-layered muslincloth and the cellulosic residue was washed thoroughly with tapwater until a neutral pH was achieved and dried overnight at60 �C.

2.6. Enzyme hydrolysis

Commercial cellulase (6.0 FPU/mg) from Trichoderma reesei(ATCC 26921) and b-glucosidase (250 U/mL) from Aspergillus ni-ger (Novozyme 188) were purchased from Sigma (St. Louis,MO, USA).

The delignified cellulosic residue of L. camara was suspended in0.05 M citrate phosphate buffer (pH 5.0) at 50 �C with a solid con-tent of 5% (w/v) and soaked in a rotary incubator shaker (Innova-40, New Brunswick Scientific, NJ, USA) for 2 h. The soaked suspen-sion was further supplemented with cellulase (3 FPU/mL) and b-glucosidase (Novozyme 188) (9 U/mL). Enzymatic hydrolysis wasperformed at 50 �C and 150 rpm for 36 h. A dose of 0.005% sodiumazide was introduced to avoid any microbial contamination; and1.0% (v/v) of Tween 80 was added to facilitate the enzymatic ac-tion. Samples were withdrawn every 4 h and subsequently ana-lysed for glucose released in the reaction mixture.

2.7. Ethanol fermentation

2.7.1. MicroorganismsP. stipitis NCIM 3498 was obtained from National Collection of

Industrial Microorganism (NCIM), National Chemical Laboratory(NCL), Pune, India and was maintained in a medium having (g/L):xylose, 20; yeast extract, 3; peptone, 5; malt extract, 3; agar, 20at pH 5.0 ± 0.2 and temperature 30 �C (Nigam, 2001). The S. cerevi-siae strain from our laboratory was maintained in medium contain-ing (g/L): glucose, 30; yeast extract, 3; peptone, 5; agar, 20 at pH6.0 ± 0.2 and temperature 30 �C (Chen et al., 2007).

The P. stipitis inoculum was prepared as described by Nigam(2001) using (g/L): xylose, 50.0; yeast extract, 3.0; malt extract,3.0; peptone, 5.0; at pH 5.0 ± 0.2 and temperature 30 �C. S. cerevi-siae inoculum was developed by growing the cells at 30 �C for 24 hin a culture medium containing (g/L): glucose, 30.0; yeast extract,3.0; peptone, 5.0; (NH4)2HPO4, 0.25; pH 6.0 ± 0.2 (Chen et al.,2007). Cells were grown to an optical density (OD600) of 0.6.

2.7.2. FermentationThe fermentation of acid and enzymatic hydrolysates was car-

ried out separately in an in situ sterilizable 12 L fermentor (B-LiteSartorius India Ltd., Bengaluru, India) with a working volume of10 L. The acid hydrolysate (16.83 g/L sugars) supplemented with(g/L): NH4Cl, 0.5; KH2PO4, 2.0; MgSO4�7H2O, 0.5; yeast extract,1.5; CaCl2�2H2O, 0.1; FeCl3�2H2O, 0.1; ZnSO4�7H2O, 0.001 (pH5.5 ± 0.2), was inoculated with 10% (v/v) inoculum of P. stipitis(OD600 0.6) and incubated at 30 �C for 36 h under shaking condi-tions (150 rpm). The cellulosic hydrolysate (34.75 g/L sugars, pH6.0 ± 0.2) aided with 3 g/L yeast extract and 0.25 g/L (NH4)2HPO4

was inoculated with S. cerevisiae (10.0% v/v). The pH of the mediumwas adjusted with 2 N HCl and 2 N NaOH. The silicone-based anti-foaming agent (10.0% v/v) was used to control the foaming, when-ever required. The dissolved oxygen concentration was monitoredcontinuously throughout the process using a dissolved oxygenprobe and a air flow at 0.4 L/min (lpm) was maintained throughout the study. Samples were withdrawn at regular intervals of4 h and centrifuged at 10,000g for 15 min at 4 �C. The cell freesupernatant was used to determine the ethanol and residual sugarconcentration.

R.C. Kuhad et al. / Bioresource Technology 101 (2010) 8348–8354 8349

2.8. Fourier Transform Infrared Spectroscopy (FTIR) characterization ofbiomass samples

The untreated, acid-pretreated and delignified L. camaramateri-als were characterized according to Singh et al. (2005). A spectrumFTIR (FTIR-1, Perkin–Elmer, MA, USA) was used to characterize theKBr-discs of different samples, which were prepared by grinding1.0 mg sample/100 mg pre-dried KBr. The spectra were recordedin the range of 450–4000 cm�1.

2.9. Analytical methods

The hydrolysates were analysed using high performance liquidchromatography (HPLC) (Shimadzu Corporation, Kyoto, Japan) forestimating the presence of carbohydrates. An Aminex (Bio-Rad,Hercules, CA, USA) column (300.0 � 7.8 mm) was used with0.04 M H2SO4 as an eluent with the flow rate of 0.5 mL/min keep-ing the oven temperature at 60 �C with a RID detector. Furans, furf-urals and lignin derivatives were estimated with Luna 5U C18column (250.0 � 4.6 mm), using acetonitrile:phosphoric acid(1.0%) (22:78) as a mobile phase with a flow rate at 0.5 mL/min,keeping the oven temperature at 35 �C with a UV-detector. Theethanol content was determined by Gas Chromatography (Per-kin–Elmer, Clarus 500) with an elite-wax (cross bond-polyethyleneglycol) column (30.0 m � 0.25 mm) at an oven temperature of85 �C and flame ionization detector (FID) at 200 �C. The ethanolstandards were prepared using commercial grade ethanol (Merck,Darmstadt, Germany). Nitrogen with a flow rate of 0.5 mL/min wasused as the carrier gas.

Total reducing sugars were estimated by the DNS method ofMiller and the enzymatic saccharification efficiency was calculatedas:

Saccharification ð%Þ

¼ Amount of glucose releasedTotal sugar concentration in the pretreated substrate

� 100

Moreover, the total phenolics released were determined by theFolin–Ciocalteu reagent method (Singleton et al., 1999) using van-illin as a standard. The OD600 of culture filtrate was measured witha double beam spectrophotometer (Specord 200, Analytical Jena,Germany) and the dry biomass of yeast cells was measured afterdrying the pellets at 70 �C until constant weight.

2.10. Statistical analysis

All the experiments were performed in triplicate and the resultsare presented as mean ± standard deviation.


3.1. Compositional analysis of L. camara

The comminuted and oven dried wood pieces (40–60 meshsize) of L. camara were found to contain holocellulose(61.1 ± 2.53%) comprising cellulose (44.1 ± 1.72%) and pentosans(17.0 ± 0.81%); lignin (32.25 ± 1.57%), ash (2.30 ± 0.11%) and mois-ture (4.35 ± 0.13%). The presence of 61.1% of total carbohydrates(holocellulose) makes it a potential and renewable material forbioethanol production.

3.2. Dilute acid hydrolysis

Among the different temperatures, the maximum sugar yield(191.43 ± 1.91 mg/g) was obtained when L. camara was hydrolysedwith 5% (v/v) H2SO4 at 140 �C for 45 min. No significant differencein sugars were observed when the substrate was treated with 3.0%(v/v) H2SO4 at 120 �C for 45 min (Table 1). Increasing the pretreat-ment period beyond 45 min, resulted in a decrease in the final su-gar concentration (data not shown). This may be due to thedegradation of sugars at severe conditions. The acid hydrolysateobtained at optimum condition of hydrolysis when analysed wasfound to be majorly consisted of (g/L): xylose (14.29 ± 0.82), glu-cose (2.21 ± 0.09), arabinose (1.49 ± 0.08), HMF (0.67 ± 0.004), fur-fural (0.51 ± 0.03) and phenolics (0.82 ± 0.05). The furfural andHMF are degradation products of pentose and hexose sugars,respectively, while, phenolics are lignin degradation byproducts.Furfural and HMF are known to decrease fermentability by reduc-ing the activities of several yeast enzymes e.g., alcohol dehydroge-nase, aldehyde dehydrogenase and pyruvate dehydrogenase(Modig et al., 2002). The furfurals have more severe effect thanHMF as they were believed to have competitive inhibition to acet-aldehyde, the natural substrate of aldehyde dehydrogenase. It hasalso been reported that the accumulation of acetaldehyde alsoinhibits the microbial growth (Taherzadeh et al., 2000).

Among different detoxification treatments, overliming (Chandelet al., 2007) and activated charcoal (Miyafuji et al., 2003; Chandel

Table 1Effect of different variables (time, temperature and acid concentration) on the release of sugars and phenolics during the acid pretreatment of L. camara.

Temperature (�C) Acidconcentration(% v/v)

Time (min)

30 45

Sugarconcentration(g/L)

Sugar yield(mg/g drysubstrate)

Phenolicsconcentration(g/L)

Phenolics yield(mg/g drysubstrate)

Sugarconcentration(g/L)

Sugar yield(mg/g drysubstrate)

Phenolicsconcentration(g/L)

Phenolics yield(mg/g drysubstrate)

100 1 7.00 ± 0.040 70.00 0.66 ± 0.016 6.58 9.57 ± 0.048 95.71 0.73 ± 0.001 7.282 9.38 ± 0.271 93.77 0.70 ± 0.007 6.94 11.79 ± 0.035 117.86 0.74 ± 0.043 7.413 11.75 ± 0.068 117.54 0.73 ± 0.011 7.29 14.00 ± 0.008 140.00 0.75 ± 0.014 7.534 12.23 ± 0.212 122.36 0.80 ± 0.004 7.97 14.43 ± 0.029 144.29 0.82 ± 0.011 8.225 12.71 ± 0.254 127.17 0.86 ± 0.013 8.64 14.86 ± 0.015 148.57 0.89 ± 0.002 8.90

120 1 8.71 ± 0.050 87.14 0.72 ± 0.028 7.24 15.00 ± 0.345 150.00 0.79 ± 0.018 7.892 10.56 ± 0.161 105.67 0.76 ± 0.012 7.63 16.86 ± 0.422 168.57 0.81 ± 0.012 8.073 12.41 ± 0.143 124.19 0.80 ± 0.008 8.01 18.71 ± 0.870 187.14 0.82 ± 0.046 8.244 12.92 ± 0.325 129.24 0.83 ± 0.008 8.27 18.75 ± 0.066 187.52 0.86 ± 0.017 8.655 13.42 ± 0.134 134.29 0.85 ± 0.034 8.53 18.87 ± 0.057 188.69 0.90 ± 0.008 9.05

140 1 8.86 ± 0.051 88.57 0.83 ± 0.007 8.27 16.29 ± 0.163 162.86 0.87 ± 0.007 8.722 10.66 ± 0.062 106.51 0.84 ± 0.030 8.39 17.36 ± 0.174 173.58 0.90 ± 0.018 8.973 12.45 ± 0.213 124.45 0.85 ± 0.008 8.51 18.73 ± 0.108 187.29 0.92 ± 0.016 9.214 13.44 ± 0.616 134.37 0.84 ± 0.012 8.39 18.79 ± 0.564 187.86 0.92 ± 0.007 9.145 14.42 ± 0.382 144.29 0.83 ± 0.026 8.26 19.14 ± 0.191 191.43 0.91 ± 0.010 9.06

8350 R.C. Kuhad et al. / Bioresource Technology 101 (2010) 8348–8354

et al., 2007) are commonly used. However, no single method is effi-cient at removing all the inhibitors present in acid hydrolysates.Thus, we attempted a sequential treatment of overliming and acti-vated charcoal adsorption for the detoxification of acid hydroly-sate. Overliming reduced furfurals (34.28%), HMF (70.59%) andphenolics (14.04%), while subsequent detoxification with activatedcharcoal resulted in further reduction of furfural (65.0%), HMF(29.41%) and phenolics (85.61%). Our findings are in agreementwith our earlier work on the detoxification of acid hydrolysatesfrom Prosopis juliflora by overliming followed by activated charcoaladsorption (Gupta et al., 2009).

3.3. Delignification of L. camara

Since a significant negative correlation exists between the per-centage of lignin in plant material and its enzymatic digestibility(Kaya et al., 2000), in order to improve the enzymatic hydrolysisof L. camara, the lignin removal is essential. The chemical deligni-fication of acid treated L. camara displayed a continuous increase inphenolics with the increase in sodium sulphite from 5.0% to 20.0%(w/v) (Table 2). The acid treated Lantana when treated with 20.0%sodium sulphite at 140 �C for 45 min resulted in 77.0% delignifica-tion with phenolics yield (248.33 ± 7.59 mg/g). However, the use of3.0% sodium chlorite along with sodium sulphite (5.0% w/v), re-duced the sodium sulphite requirement from 20.0% to 5.0% anddelignified the L. camara by 87.16% with phenolics yield(281.1 ± 9.37 mg/g). Sodium chlorite is an elemental chlorine freecompound, which on heating produces chlorine dioxide that actsas a delignifying agent and depolymerizes the lignin (Hamzehet al., 2006; Gupta et al., 2009). Earlier we had used sodium sulph-ite for the pretreatment of sugarcane bagasse (Kuhad et al., 1999)and sodium chlorite for P. juliflora (Gupta et al., 2009) in our labo-ratory. The chemical delignification pretreatments not only re-move the lignin but also act as a swelling agent, which in turnenhances the surface area of the substrate made accessible forenzymatic action (Kuhad et al., 1999).

3.4. FTIR characterization

The FTIR spectroscopy is extensively used for lignocellulosecharacterization since it presents a relatively easy method ofobtaining direct information on chemical changes that occurs dur-ing various chemical pretreatments (Ristolainen et al., 2002). The

FTIR spectra of untreated, acid treated and delignified L. camara de-picted that as compared to hemicellulose and cellulose, a large dif-ference was found in the fingerprint region (1830–730 cm�1) forlignin’s IR spectrum (Electronic Annexure I). All spectras had asharp peak at 898 cm�1, which is characteristic of b-glycosidic link-ages between the sugar units, as it has been earlier reported byGupta et al. (1987). A prominent band of hemicellulose was ob-served at 1370 cm�1, which was subsequently removed in acidtreated substrate. The acid treated substrate showed a reductionin the band at 1238 and 1738 cm�1, which are indicative of hemi-cellulose–lignin linkage and C@O stretching due to carbohydratelinked with lignin, respectively (Lin and Dence, 1992). Results alsoshowed a band at 655 cm�1 that is a characteristic feature of ligno-sulphonates (Nada et al., 1998) occurring after the action of sul-phuric acid over lignin. The absence of this band in delignified sub-strate shows removal of lignin (Singh et al., 2005). The aromaticC@C stretch from aromatic ring of lignin gave two peaks at 1508and 1458 cm�1 in untreated and at 1509 and 1457 cm�1 in acidtreated substrates. The FTIR spectra of delignified substrate afterchlorite treatment of acid-pretreated substrate revealed noticeablechanges at the bands relating to aromatic ring vibration at 1508–

0

100

200

300

400

500

600

700

800

900

0 4 8 12 16 20 24 28 32 36

Time (h)

Sacc

hari

fica

tion

Yie

ld (

mg

g-1)

0

10

20

30

40

50

60

70

suga

r co

ncen

trat

ion

(g L

-1)

and

Sacc

hari

fica

tion

rat

e (m

g g-1

h-1)

Saccharification yield

Sugar concentration

Saccharification rate

Fig. 1. Enzymatic saccharification profile of delignified L. camara. Where, darksquare is saccharification rate (mg/g/h), white square is saccharification yield (mg/g)and the triangle is sugar concentration (g/L).

Table 2Release of phenolics during the delignification of acid treated biomass of L. camara with different concentration of Na2SO3 (A) and NaClO2 (B).

Temperature (�C) Delignifying agents (% w/v) Time (min)

30 45

Phenolics yield(mg/g dry substrate)

Delignification (% w/w) Phenolics yield(mg/g dry substrate)

Delignification (% w/w)

100 5 A 98.40 ± 2.04 30.51 119.83 ± 3.67 37.1610 A 137.76 ± 6.79 42.72 167.76 ± 1.94 52.0215 A 158.87 ± 13.69 49.26 174.43 ± 1.74 54.0920 A 190.64 ± 7.70 59.11 188.38 ± 4.74 58.415 A + 3 B 243.81 ± 8.57 75.60 252.74 ± 9.53 78.37

120 5 A 117.93 ± 5.82 36.57 136.50 ± 6.26 42.3310 A 165.10 ± 9.09 51.19 191.10 ± 9.43 59.2615 A 193.32 ± 12.42 59.94 206.65 ± 13.77 64.0820 A 231.98 ± 9.66 71.93 223.18 ± 11.45 69.205 A + 3 B 281.10 ± 9.37 87.16 280.47 ± 14.29 86.97

140 5 A 142.05 ± 8.64 44.05 165.07 ± 10.31 51.1810 A 198.87 ± 19.18 61.67 231.10 ± 7.43 71.6615 A 238.87 ± 11.94 74.07 241.10 ± 13.28 74.7620 A 246.64 ± 13.58 76.47 248.33 ± 7.59 77.005 A + 3 B 281.14 ± 14.29 87.18 280.37 ± 10.28 86.94


1509 and 1457–1458 cm�1. The disappearance of these bandsshowed that lignin was largely removed in comparison to polysac-charides during the chlorite pretreatment (Sun et al., 2005). Theabsorbances at 1428, 1331, 1281, 1111, 1056 and 899 cm�1 wereassociated with the typical value of cellulose (Sun et al., 2005).

3.5. Enzymatic saccharification of delignified L. camara

During the course of enzymatic saccharification, a perpetualincrease in sugar concentration was observed till 28 h, which on

prolonged incubation remained almost constant (Fig. 1). However,the optimal saccharification yield (777.13 mg/g) was achievedafter 28 h with a saccharification rate of 24.29 g/L/h (Fig. 1). Themaximum rate of saccharification was achieved after 4 h of incuba-tion, which thereafter it started declining as reported earlier (Gup-ta et al., 2009). The decline in hydrolysis rate could be due to theincreasing resistance of the substrate during the course of hydroly-sis or other factors. End product inhibition of the enzymes by glu-cose and cellobiose may also have a major impact on cellulosehydrolysis (Kuhad et al., 1999). Moreover, the enzymatic sacchari-

Table 3Fermentation profile of detoxified acid hydrolysate of L. camara by P. stipitis.

Time (h) Ethanol (g/L) Sugar (g/L) Ethanol yield (g/g) Ethanolproductivity (g/L/h)

Biomass (g/L) Biomassyield (g/g)

Biomassproductivity (g/L/h)

0 0.05 ± 0.003 16.83 ± 0.044 0.00 0.00 0.15 ± 0.004 0.01 0.004 0.27 ± 0.009 15.83 ± 0.047 0.02 0.07 1.42 ± 0.058 0.08 0.368 1.49 ± 0.060 14.81 ± 0.203 0.09 0.19 2.78 ± 0.083 0.17 0.3512 2.87 ± 0.057 13.13 ± 0.162 0.17 0.24 3.93 ± 0.228 0.23 0.3316 3.31 ± 0.076 9.68 ± 0.011 0.20 0.21 5.36 ± 0.386 0.32 0.3420 4.11 ± 0.234 7.29 ± 0.030 0.24 0.21 5.91 ± 0.242 0.35 0.3024 5.16 ± 0.371 5.23 ± 0.198 0.33 0.23 6.20 ± 0.546 0.37 0.2628 4.49 ± 0.174 4.37 ± 0.095 0.27 0.16 6.44 ± 0.509 0.38 0.2332 4.01 ± 0.084 4.27 ± 0.136 0.24 0.13 6.65 ± 0.499 0.40 0.21

Table 4Fermentation profile of enzymatic hydrolysate of L. camara by S. cerevisiae.

Time (h) Ethanol (g/L) Sugar (g/L) Ethanol yield (g/g) Ethanolproductivity (g/L/h)

Biomass (g/L) Biomassyield (g/g)

Biomassproductivity (g/L/h)

0 1.24 ± 0.029 34.75 ± 1.538 0.03 0.00 0.32 ± 0.015 0.01 0.006 14.4 ± 0.245 8.68 ± 0.102 0.39 2.40 2.25 ± 0.063 0.06 0.388 15.7 ± 0.769 6.92 ± 0.023 0.43 1.96 4.82 ± 0.096 0.14 0.6012 16.5 ± 0.462 3.25 ± 0.007 0.45 1.38 6.64 ± 0.325 0.19 0.5516 17.7 ± 0.955 1.53 ± 0.018 0.48 1.11 7.48 ± 0.434 0.22 0.4720 15.9 ± 0.845 1.51 ± 0.003 0.43 0.80 7.52 ± 0.233 0.22 0.3824 15.0 ± 0.540 1.49 ± 0.000 0.41 0.63 7.51 ± 0.188 0.22 0.31

Acid hydrolysis

Delignification Enzymatic hydrolysis

Hexose Fermentation

Detoxification Pentose Fermentation

Lantana Camara(1000.0 g)

461.1±17.98 g Cellulose

177.7±8.46 g Hemicellulose

337.1±16.41 g Lignin

24.01±1.15 g Ash

6.7±0.038 g Hydroxy-methyl Furfurals

5.1±0.29 g Furfurals

8.2±0.46 g Phenolics

3.16±0.23 g Others



6.7±0.029 g Hydroxy-methyl Furfurals



3.16±0.004 g Others

51.6±3.72 g Ethanol



37.77±0.025 g Xylose

429.42±12.83g Cellulose

328.90±14.29 g Lignin

23.89±1.08 g Ash

7.02±0.87 g Hemicellulose

414.62±12.72 g Cellulose

47.8±1.24 g Lignin

23.70±0.11 g Ash

14.80±0.45 g Cellulose7.0±0.28 g Hemicellulose281.1±9.37 g Lignin

325.12±10.44 g Glucose

89.53±1.42 g Cellulose47.8±2.54 g Lignin23.42±.19 g Ash

148.14±7.99 g Ethanol

13.5±0.98 g Glucose

142.80±8.21 g Xylose

22.11±0.87 g Glucose

14.99±0.76 g Arabinose

7.50±0.54 g Other sugars

10.35±0.024 g Xylose

5.37±0.247 g Glucose1.78±0.074 g Arabinose

1.60±0.059 g Other sugars

132.45±8.47 g Xylose16.75±1.08 g Glucose

13.21±0.95 g Arabinose

esonibarAg10.0±41.01sragusrehtOg520.0±98.5

4.32±0.02 g other sugars

Solid Solid Filtrate

Filt

rate

Filt

rate

Solid

Solid

Filtrate

Fig. 2. Mass balancing of the complete bioconversion process for ethanol production from L. camara.


fication efficiency (77.7%, w/w) obtained was in agreement withother earlier reports about saccharification of pretreated substrateswith a saccharification efficiency of 67.6%, 70.0% and 80.0% (w/w)from chlorite pretreated sugarcane bagasse (Adsul et al., 2005), al-kali treated rice straw (Jeya et al., 2009) and chlorite pretreated P.juliflora (Gupta et al., 2009), respectively.

3.6. Fermentation

Both acid and enzymatic hydrolysates were fermented with P.stipitis and S. cerevisiae, respectively. The detoxified xylose richhydrolysate obtained after dilute acid hydrolysis (16.83 ± 0.04 g/Lsugars), when fermented with P. stipitis, resulted in ethanol pro-duction of 5.16 ± 0.37 g/L with yield 0.33 g/g and productivity of0.23 g/L/h after 24 h. The biomass yield and productivity werefound to be 0.37 g/g and 0.26 g/L/h, respectively (Table 3). How-ever, fermentation of cellulosic hydrolysate (34.75 ± 1.54 g/L sug-ars) using S. cerevisiae resulted in a maximum ethanol content of17.7 ± 0.96 g/L with yield of 0.48 g/g and productivity of 1.11 g/L/h with a biomass yield (0.22 g/g) and productivity (0.47 g/L/h) after16 h incubation (Table 4). Our results were in agreement with pre-vious reports on fermentation with P. stipitis (Kapoor et al., 2008;Gupta et al., 2009). The ethanol yield obtained from cellulosichydrolysate using S. cerevisiae in our case (0.48 g/g) was higheror comparable to some of the previous reports of ethanol yieldfrom corncob (0.48 g/g) (Chen et al., 2007) and from P. juliflora(0.49 g/g) (Gupta et al., 2009). The S. cerevisiae produced maximumethanol after 16 h and it declined thereafter (Table 4). The declinein ethanol production after 16 h incubation can be due to the con-sumption of accumulated ethanol by the organism. According toRamon-Portugal et al., 2004, when the ethanol accumulated inthe medium, the microbial population was adapted to consumesimultaneously sugar and ethanol. The increased yeast biomasscould be because of the utilization of the yeast extract present infermentation medium. The increase biomass in pentose and hexosefermentation is in accordance with the observations of Wang et al.(2004) and Lau and Dale (2009).

Moreover, a mass balance study of the ethanol production fromL. camara has also been elucidated in Fig. 2.

4. Conclusion

Bioethanol from lignocellulosics is a globally accepted alterna-tive fuel. The production of ethanol from L. camara would havethe dual advantage of producing energy and serving as an effectivemethod of weed management. The present study demonstratesthat the pretreatment (acid hydrolysis and delignification) of L. ca-mara can provide cellulosic material, which eventually is hydroly-sed to hexose sugars. Thus pentose and hexose sugars can beseparately fermented to ethanol. Developing a fermentation sys-tem with yeast strains capable of fermenting both hexose and pen-tose sugars simultaneously would economize ethanol productionfrom lignocellulosic biomass.

Acknowledgements

The authors are thankful to Dr. A.J. Varma, Polymer ScienceDivision, National Chemical Laboratory, Pune for FTIR analysis ofour samples. The authors also thank Ms. Urvashi Kuhad, Depart-ment of English, University of Delhi, Delhi, for editing the manu-script. The financial support from Department of Biotechnology(DBT), Ministry of Science and Technology, Government of Indiais highly acknowledged.

Appendix A. Supplementary data

Supplementary data associated with this article can be found, inthe online version, at doi:10.1016/j.biortech.2010.06.043.

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Fed batch enzymatic saccharification of newspaper cellulosicsimproves the sugar content in the hydrolysates andeventually the ethanol fermentation bySaccharomyces cerevisiae

Ramesh Chander Kuhad*, Girija Mehta, Rishi Gupta, Krishna Kant Sharma

Lignocellulose Biotechnology Laboratory, Department of Microbiology, University of Delhi South Campus, Benito Juarez Road, New Delhi

110021, India


Article history:

Received 7 May 2009

Received in revised form

9 March 2010

Accepted 15 March 2010

Available online 10 April 2010

Keywords:

Newspaper

Cellulose

Cellulases

Fermentation

Ethanol

Saccharomyces cerevisiae

a b s t r a c t

The newspaper is comprised of (w w�1) holocellulose (70.0%) with substantial amount of

lignin (16.0%). Bioconversion of the carbohydrate component of newspaper to sugars by

enzymatic saccharification, and its fermentation to ethanol was investigated. Of various

enzymatic treatments using cellulase, xylanase and laccase, cellulase enzyme system was

found to deink the newspaper most efficiently. The saccharification of deinked paper pulp

using enzyme cocktail containing exoglucanase (20 U g�1), b-glucosidase (60 U g�1) and

xylanase (80 U g�1) resulted in 59.8% saccharification. Among additives, 1% (v v�1) Tween 80

and 10 mol m�3 CoCl2 improved the enzymatic hydrolysis of newspaper maximally,

releasing 14.64 g L�1 sugars. The fed batch enzymatic saccharification of the newspaper

increased the sugar concentration in hydrolysate from 14.64 g L�1 to 38.21 g L�1. Moreover,

the batch and fed batch enzymatic hydrolysates when fermented with Saccharomyces

cerevisiae produced 5.64 g L�1 and 14.77 g L�1 ethanol, respectively.

ª 2010 Elsevier Ltd. All rights reserved.

1. Introduction

Presentlymost of the bioethanol is produced using corn kernel

or sugarcane molasses but lignocellulosics, the second

generation biofuel substrate and the least explored renewable

resource is yet to be tapped. Lignocellulosics are the most

abundant biomass available on earth, comprising mainly of

cellulose and hemicellulose [1,2]. Among various lignocellu-

losics, the recycled books, magazines and newspaper could

have value addition via deinking and reuse of fiber either in

manufacturing of new paper or in ethanol production.

An efficient conversion of newspaper to ethanol depends

mainly on the extent of carbohydrate saccharification,

however, the enzymatic conversion of newspaper is majorly

hindered by toner’s ink, which forms a physical barrier,

restricting the hydrolysable sites as well as release fermen-

tation inhibitory heavy metal ions [3,4]. Among various

enzymes tested to deink the newspaper pulp [5], only cellu-

lases were found to be the most effective [6]. The deinked

paper pulp can be saccharified using either acid [7,8] or

enzymes [9e11], however, enzymes are preferred over acid

because enzymatic hydrolysates are free from any fermenta-

tion inhibitory products. The enzymes such as cellulases have

been found to be the most effective for this purpose [12]. It is

well known that combined action of cellulase and hemi-

cellulase results in a higher sugar production as compared to

* Corresponding author. Tel.: þ91 11 24112062; fax: þ91 11 24115270.E-mail address: [email protected] (R. Chander Kuhad).

Avai lab le a t www.sc iencedi rec t .com

ht tp : / /www.e lsev ier . com/ loca te /b iombioe

b i om a s s a n d b i o e n e r g y 3 4 ( 2 0 1 0 ) 1 1 8 9e1 1 9 4

0961-9534/$ e see front matter ª 2010 Elsevier Ltd. All rights reserved.doi:10.1016/j.biombioe.2010.03.009

cellulase alone [13]. Despite the high extent of saccharification

efficiency, the lower substrate consistency in the enzymatic

suspension resulted into lower sugar concentration during the

enzymatic hydrolysis. Raising the substrate concentration in

batch hydrolysis helps to obtain higher sugar concentration,

but also causes mixing and heat transfer problem due to

rheological properties of dense fibrous suspension [14]. While

in fed batch hydrolysis, such problem could be avoided by

adding the substrate gradually to maintain the low level of

viscosity [15]. In the present study, the optimization of enzy-

matic saccharification using both cellulase and xylanase was

carried out to improve the hydrolysis of the newspaper. In

order to increase the sugar concentration in enzymatic

hydrolysate, the fed batch enzymatic hydrolysis of deinked

newspaper was carried out. Thereafter, the enzymatic

hydrolysates comprising both the hexose and pentose sugars

were fermented to ethanol using Saccharomyces cerevisiae.

2. Materials and methods

2.1. Raw material and chemicals

The newspaper procured locally was shredded mechanically

into small pieces and the processed substrate was dried in an

oven at 80 �C until constant weight was obtained.Commercial cellulase from Trichoderma reesei (ATCC 26921),

b-glucosidase from Aspergillus niger (Novozyme 188), xylanase

fromThermomyces languinosus, 1-hydroxylbenzotriazole (HOBT)

and 3,5-di nitro salicylic acid (DNS) were purchased from

Sigma (St. Louis, Missouri, U.S.A.). Ethanolwas purchased from

Merck (Darmstadt, Germany). Laccase was produced in our

laboratory as described earlier [16]. Rest of the chemicals and

media components of highest purity grade were purchased

locally.

2.2. Microorganism and culture conditions

S. cerevisiae RCK-1 was procured from culture collection of

Department of Microbiology, University of Delhi South

Campus, New Delhi, India and was maintained on agar slants

containing (g L�1): glucose, 30.0; yeast extract, 3.0; peptone,5.0; agar, 20.0 at pH 6.0 � 0.2 and temperature 30 �C. Inoculumof S. cerevisiae was prepared in medium containing (g L�1)glucose, 5.0; yeast extract, 3.0; peptone, 5.0; (NH4)2HPO4, 0.25

media constituents at pH 5.5 � 0.2 [15].

2.3. Compositional analysis of newspaper

The chemical composition of newspaper was analysed for

holocellulose, a-cellulose, pentosans, moisture and ash

content following the Technical Association of Pulp and Paper

Institutes (TAPPI) methods [17].

2.4. Deinking of newspaper

The deinking of newspaper was carried out using floatation

method at 2.0% (w v�1) consistency having various enzymesystems such as cellulase, xylanase, laccase with HOBT (LH)

and laccase without HOBT (LB). The waste paper slurry was

incubated at 50 �C for cellulase [18], 60 �C for xylanase [19] and30 �C for laccase [16].

2.5. Enzymatic hydrolysis

2.5.1. Batch enzymatic hydrolysisEnzymatic hydrolysis of deinked newspaper was carried out

at 2% (w v�1) consistency in a 50 mol m�3 citrate phosphatebuffer (pH 5.0) containing 0.005% (w v�1) sodium azide. Before

enzyme loading, slurrywas preincubated at 50 �C on a rotatoryshaker (Innova-40, New Brunswick Scientific, Germany) at

2.5 Hz for 2 h. Thereafter, to obtain the optimized enzyme

doses for the saccharification of deinked newspaper pulp,

varied doses of exoglucanase (5e30 U g�1), b-glucosidase

(15e90 U g�1), and xylanase (20e100 U g�1) were added to thepreincubated slurry. Different types of additives such as

nonionic surfactants (Tween, Polyethylene glycol and Triton

X-100) andmetal ions were also examined to study their effect

on the extent of saccharification efficiency. Samples were

withdrawn at regular intervals of 4 h, centrifuged at 2.5 Hz for

15 min in a centrifuge (Sigma, Germany) and the supernatant

was analysed for total reducing sugars released.

2.5.2. Fed batch enzymatic hydrolysisFed batch enzymatic saccharification of deinked newspaper

was carried out at optimized conditions of saccharification

with an initial substrate consistency of 2% (w v�1) in the

suspension. The substrate was added twice at 20th and 40th

hour to get a final substrate concentration of 60 g L�1. Thesamples were withdrawn at regular intervals of 4 h and ana-

lysed for the release of total reducing sugars.

2.6. Fermentation of enzymatic hydrolysate

The fermentation studies of enzymatic hydrolysate were

carried out in 2.0 L Erlenmeyer flasks. The batch and fed batch

enzymatic hydrolysates containing 14.64 g L�1 and 38.21 g L�1

sugars, respectively, supplemented with 3.0 g L�1yeast extractand 0.25 g L�1 (NH4)2HPO4 were inoculated with 10% (v v�1) S.cerevisiae. The flasks were incubated at 30 �C, 3.33 Hz and pH5.5 � 0.2. The pH was adjusted with 2 N HCl and 2 N NaOH.

Samples were withdrawn at regular intervals of 4 h and

centrifuged at 2.5 Hz for 15 min at 4 �C. The cell free super-natant was used for the determination of ethanol produced

and sugar consumed.

2.7. Enzyme assays

All the cellulase assays were carried out in 50 mol m�3 citratephosphate buffer (pH 5.0) unless otherwise stated. Endoglu-

canase and exoglucanase activities were determined

following International Union of Pure and Applied Chemistry

(IUPAC) methods [20]. For the estimation of endo and exo-

glucanase, 2.0% carboxymethyl cellulose (CMC) and 50.0 mg

Whatman No. 1 filter paper, respectively, were used as the

substrates. The release of sugars was determined by the DNS

method [21]. One unit of endo and exoglucanase activity is

defined as the amount of enzyme required to produce

1.0 mmol m�3 of glucose equivalents per min under assay

conditions. b-glucosidase activity was determined by taking


0.1% p-nitrophenol b-D-glucopyranoside (pNPG) as substrate

and the activity was calculated as the enzyme required to

liberate 1.0 mmol m�3 of p-nitrophenol (pNP) per minuteunder the assay conditions. Xylanase activity was quantified

by measuring the release of reducing sugar groups by DNS at

60 �C using 1% (w v�1) birch wood xylan in citrate phosphatebuffer (200 mol m�3), pH 6.0 [19]. One unit of xylanase was

defined as amount of enzyme required to release

1.0 mmol m�3 of xylose in 1 min under the assay conditions.The activity of laccase was determined as described previ-

ously [22]. The reaction was set by adding 0.2 mL of culture

supernatant in 1.8 mL of 10 mol m�3 guaiacol in 100 mol m�3

citrate phosphate buffer (pH 5.4). The reaction mixture was

incubated at 26 �C for 10 min and color development was

assayed at 470 nm on a UVeVIS spectrophotometer (Analyt-

ical Jena, Specord 205, Germany). Heat denatured enzyme

served as control. One unit of laccase was defined as the

change in absorbance of 0.01 mL�1 min�1 at 470 nm.


Ethanol was estimated by gas chromatography (GC) (Perkin

Elmer, Clarus 500) with an elite-wax (cross bond-polyethylene

glycol) column (30.0 m � 0.25 mm), at oven temperature 85 �Cand flame ionization detector (FID) at 200 �C. The ethanolstandards were prepared using commercial grade ethanol

(Merck, Germany). Nitrogen with a flow rate of 0.5 mL min�1

was used as carrier gas. The extent of deinking (Chromophore

released) of newspaper was measured with a UVevisible

spectrophotometer at 528 nm, the wavelength at which

treated filtrate gave maximum absorbance peak. Total

reducing sugars were estimated by the DNS method [21] and

the saccharification efficiency was calculated as follows:


3.1. Compositional analysis of newspaper

Newspaper was found to contain (w w�1), a-cellulose (51.0%),pentosans (19.0%), lignin (16.0%), moisture (8.0%) and ash

(2.0%). The high carbohydrate content (holocellulose, 70.0%

w w�1) in the newspaper made it a substrate of choice for

bioethanol production. The results obtained here were similar

to earlier studies, where lignin, holocellulose and moisture

content in newspaper were found to be 16e22%, 60e75% and

7e10%, respectively [7,8].

3.2. Deinking of newspaper cellulosics

The dyes fixed on the repeating units of cellulose backbone

block the sequential hydrolysis of paper by the enzymes

and inhibited the hydrolysis kinetics significantly [4,23].

Therefore, many enzyme systems were used to study their

effect on the deinking of newspaper. The comparative effi-

ciency of different enzyme systems to deink the newspaper

has been shown in Fig. 1. The maximum deinking was

obtained with cellulase system, while the minimum deink-

ing occurred when treated with laccase with out HOBT (LB),

however the deinking efficiency of newspaper using

different enzyme systems were in an order of

cellulase > xylanase > laccase with HOBT (LH) > laccase

with out HOBT (LB) (Fig. 1). The possible mechanism may be

that the enzymatic action of cellulase had released the

cellulose micro-fibrilar matrix in which the toners are

embedded. In earlier reports, a combined action of cellulase

and xylanase enzymes brought about 62.0% deinking [5].

The results showed that when compared, LB enzyme

system caused a significant increase in deinking efficiency

than LH, this may be due to the fact that laccase itself

cannot enter into pulp fibers due to size limitations,

however the presence of mediator could help in trans-

mitting electron between laccase and lignin. In this case,

firstly the mediator was oxidized by laccase in presence of

O2, then the oxidized mediator diffused into pulp fiber and

cause a series of oxidative degradation reactions [24]. With

the removal of lignin, the bonding between fiber and ink

particles is loosened, which aid to the deinking process [24].

Although there are several reports of using alkaline condi-

tions for deinking [25e27], however, in the present work, we

have demonstrated the deinking process at low pH, because,

as the toners are not only associated with cellulose fibrils

but also with the white pigment fillers and coating such as

calcium carbonate, therefore, the slight acidic conditions

could lead to the improvement of the deinking process by

the removal of CaCO3 and fillers [5,28].

3.3. Enzymatic hydrolysis of deinked newspaper

3.3.1. Batch enzymatic hydrolysis

3.3.1.1. Effect of enzyme dosage on the hydrolysis. Differentdosages of exoglucanase (5e30 U g�1), were used to enzy-

matically hydrolyze the deinked newspaper (Fig. 2). The

sugars release increasedwith increase in enzyme dosage upto

20 U g�1, which resulted in 23.94% saccharification after 24 h ofincubation. Where as, an increase in the enzyme dosage

beyond 20 U g�1 did not cause any further improvement in thesaccharification (Fig. 2). In earlier reports, enzymatic hydro-

lysis of paper material at loading of 45 FPU g�1 [29] and120 FPU g�1 [30] have resulted in 45.6 and 90% saccharification,

respectively, after 6 days. The lower saccharification effi-

ciency may be possibly due to feed back inhibition by the

cellobiose moieties, which are released by the action of exo-

glucanase [31]. To overcome the problem of feedback inhibi-

tion, varied doses of b-glucosidase (15e90 U g�1) were tested inthe enzyme system containing exoglucanase (20 U g�1) that

Saccharification efficiencyð%Þ ¼ Reducing sugar concentration obtained� 100Total carbohydrate content in the substrate

b i om a s s a n d b i o e n e r g y 3 4 ( 2 0 1 0 ) 1 1 8 9e1 1 9 4 1191

could hydrolyze the cellobiose moieties and reduce the

cellobiose accumulation, which might have resulted in

enhanced saccharification. Among different dosages of

b-glucosidase used, addition of 60 U g�1 of b-glucosidase

caused 13.8% increase in sugar yield (264.13 mg g�1), whilefurther increase in dosage of b-glucosidase did not enhance

the saccharification (Fig. 3). This is also in accordance with

earlier reports where the addition of b-glucosidase increased

the saccharification of recycled paper sludge from 45 to 100%

was observed [30].

In order to enhance the enzymatic saccharification of dei-

nked newspaper, different doses of xylanase (20e100 U g�1)were added to the optimized enzymatic suspension (20 U g�1

exoglucanase and 60 U g�1 b-glucosidase). Among different

dosages of xylanase used, the addition of 80 U g�1 xylanase to

the enzymatic cocktail resulted in optimum sugar yield

(419.54 mg g�1) (Fig. 4). The effect of combined action of

cellulases and hemicellulases demonstrated that compared to

cellulase alone, the addition of xylanase with cellulase

enhanced the saccharification efficiency from 37.7 to 59.8%.

This is because the enzymatic saccharification with xylanase

cleaves the xylan backbone and releases the lignin from car-

bohydrateelignin complex, which helps in improvement of

saccharification of lignocellulose [19].

3.3.1.2. Effect of additives on the hydrolysis. Out of varioussurfactants used, Tween 80 and Triton X-100 improved the

sugar release by 10e30%, while Tween 20, 40, 60 and PEG 400,

0

20

40

60

80

100

120

140

160

180

200

0 4 8 12 16 20 24Time (h)

Sug

ar r

elea

sed

(mg

g-1)

5 U g-110 U g-115 U g-120 U g-125 U g-130 U g-1

Fig. 2 e Effect of varied dosage of exoglucanase on the

enzymatic saccharification of deinked newspaper at 50 �Cand 2.5 Hz.

0

50

100

150

200

250

300

0 4 8 12 16 20 24

Time (h)

Suga

r re

leas

ed (

mg

g-1)

15 U g-130 U g-145 U g-160 U g-175 Ug-190 U g-1

Fig. 3 e Effect of varied dosage of b-glucosidase along with

20 U of exoglucanase on the enzymatic saccharification of

deinked newspaper at 50 �C and 2.5 Hz.

gg

m(desaeler

raguS1-)

gg

m(desaeler

raguS1-)

0

50

100

150

200

250

300

350

400

450

0 4 8 12 16 20 24

Time (h)

20 U g-140 U g-160 U g-180 U g-1100 U g-1

0

50

100

150

200

250

300

350

400

450

0 4 8 12 16 20 24

Time (h)

20 U g-140 U g-160 U g-180 U g-1100 U g-1

Fig. 4 e Effect of varied dosage of xylanase along with the

optimized cellulase doses (20 U exoglucanase and 60 U

b-glucosidase) on the enzymatic saccharification of

deinked newspaper at 50 �C and 2.5 Hz.

0

10

20

30

40

50

60

70

80

90

100

110

Cel Xyl LH LB Control

Enzymes

)%(

ycneiciffegniknied

evitarapmo

C

Fig. 1 e Comparison of different enzyme systems for the

deinking of newspaper. (Where, Cel, cellulase; Xyl,

xylanase; LH, laccase with HOBT and LB, laccase without

HOBT).


4000 and 6000 at any loading were not statistically distin-

guishable and reduced the sugar release by 30e40% from the

control. The increase in newspaper saccharification with

Tween 80 could be because of (i) the presentation of unpro-

ductive enzyme adsorption to lignin [32] and (ii) the protection

of the enzymes from thermal deactivation [33]. Moreover, on

increasing the concentration of Tween 80 more than 1%

declines the rate of hydrolysis, which might be because of the

detergent property of the surfactant, which denatures the

enzymes [34].

Differentmetal ionswere also tested to study their effect in

enzymatic saccharification and it was observed that the

presence of cobalt (10 mol m�3) increased the saccharification

efficiency by 17.1%, which is in agreement with the earlier

reports [35,36]. While, other metal ions (Cu, Hg, Fe) at

10molm�3 concentrationwere found to inhibit the enzymatichydrolysis of deinked newspaper.

3.3.2. Fed batch enzymatic saccharificationDuring the time course of fed batch enzymatic hydrolysis, the

saccharification continued till 84 h and remained constant on

prolonged incubation (Fig. 5). The maximum amount of sugar

(38.29 g L�1) was released after 84 h of incubationwith a rate of

0.48 g L�1 h�1 and 85.71% efficiency. Our results are also in

accordancewith the earlierworkof Chenet al.where in the fed

batch enzymatic saccharification of corncob 79.5% efficiency

was achieved after 60 h of incubation [15].

3.4. Fermentation

The fermentation of batch enzymatic hydrolysate (14.64 g L�1

sugars) produced 5.64 g L�1ethanol with yield and productivityof 0.39 g g�1 and 0.71 g L�1 h�1, respectively, after 8 h of

incubation (Table 1). While, the fed batch enzymatic hydro-

lysate containing 38.21 g L�1 sugars, when fermented with S.

cerevisiae, produced 14.77 g L�1 ethanol with yield (0.39 g g�1)and productivity (0.74 g L�1 h�1), after 20 h of incubation

(Table 1). The ethanol yield observed in this study is higher

than the value of 0.17 g g�1 [37] and 0.25 g g�1 [30] producedfromyeast extract-amended spent sulphite pulping liquor and

enzymatic hydrolysate of newspaper, respectively. However,

the lower ethanol yield and productivity may be due to the

presence of pentose sugars in the enzyme hydrolysate, which

was not utilized by S. cerevisiae. According to earlier reports, S.

cerevisiae can ferment the glucose sugar readily but could not

metabolize xylose sugars due to lack of xylose reductase and

xylitol dehydrogenase [15]. Irrespective of saccharification

mode, the prolonged incubation in fermentation resulted in

decline in ethanol concentration after attaining themaximum

ethanol yield. This may be due to consumption of accumu-

lated ethanol by the organism for its maintenance [38,39].

4. Conclusion

The reuseofnewspaper for theproductionofbioethanol seems

feasible approach for energy conservation and waste minimi-

zation. The fed batch enzymatic hydrolysis of newspaper

cellulosics improved thereleaseof sugars (pentoseandhexose)

and subsequently the ethanol yield.However, the yeast did not

utilized pentose sugars from the hydrolysates and non-

utilization of pentose sugars limits the economization of

ethanol production from cellulosics. Therefore, in order to

improve the ethanol fermentation from lignocellulosic waste,

0.0

5.0

10.0

15.0

20.0

25.0

30.0

35.0

40.0

45.0

4 8 1216 2024 28 3236 40 4448 5256 60 6468 72 7680 8488 92 96Time (h)

Lg(

noitartnecnocraguS

1-)

0.0

0.2

0.4

0.6

0.8

1.0

1.2

1.4

1.6

/L/g(

noitacifirahccaSfo

etaR

h

Sugar concentration

Rate of saccharification

Fig. 5 e Time course of fed batch enzymatic hydrolysis of

deinked newspaper using 20 U exoglucanase, 60 U b-

glucosidase, 60 U xylanase, 1% Tween 80 and 10 mol mL3

CoCl2 at 50 �C and 2.5 Hz.

Table 1 e Ethanol production profile from batch and fed batch enzymatic hydrolysate of deinked newspaper cellulosics bySaccharomyces cerevisiae RCK-1 3441 at 30 �C and 3.33 Hz.

Time (h) Batch enzymatic hydrolysate Fed batch enzymatic hydrolysate

Sugar(g L�1)

Ethanol(g L�1)

Ethanolyield (g g�1)

Ethanolproductivity(g L�1 h�1)

Sugar(g L�1)

Ethanol(g L�1)

Ethanolyield (g g�1)

Ethanolproductivity(g L�1 h�1)

0 14.64 0.00 0.00 0.00 38.21 0.00 0.00 0.00

4 7.44 3.16 0.22 0.79 30.12 3.24 0.08 0.81

8 2.70 5.64 0.39 0.71 22.45 6.30 0.16 0.79

12 1.92 4.93 0.34 0.41 10.26 11.18 0.29 0.93

16 0.97 3.31 0.23 0.21 4.56 13.46 0.35 0.84

20 0.53 2.58 0.18 0.13 1.29 14.77 0.39 0.74

24 0.38 2.04 0.14 0.08 0.85 13.25 0.35 0.55

28 0.10 1.64 0.11 0.06 0.51 12.08 0.32 0.43

32 nil 1.29 0.09 0.04 0.25 10.18 0.27 0.32

b i om a s s a n d b i o e n e r g y 3 4 ( 2 0 1 0 ) 1 1 8 9e1 1 9 4 1193

it is necessary to develop recombinant yeast strain capable of

fermenting both sugars simultaneously.

Acknowledgement

The authors are thankful to Dr. Y. P. Khasa for his help during

the preparation of thismanuscript. The financial support from

University of Delhi, Delhi, India is highly acknowledged.

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Separate hydrolysis and fermentation (SHF) of Prosopis juliflora,a woody substrate, for the production of cellulosic ethanolby Saccharomyces cerevisiae and Pichia stipitis-NCIM 3498

Rishi Gupta, Krishna Kant Sharma, Ramesh Chander Kuhad *

Lignocellulose Biotechnology Laboratory, Department of Microbiology, University of Delhi South Campus, Benito Juarez Road, New Delhi-110 021, India


Article history:Received 19 April 2008Received in revised form 20 August 2008Accepted 20 August 2008Available online 2 October 2008

Keywords:Lignocellulosic biomassProsopis julifloraAcid hydrolysisEnzymatic hydrolysisFermentation

a b s t r a c t

Prosopis juliflora (Mesquite) is a raw material for long-term sustainable production of cellulosics ethanol.In this study, we used acid pretreatment, delignification and enzymatic hydrolysis to evaluate the pre-treatment to produce more sugar, to be fermented to ethanol. Dilute H2SO4 (3.0%, v/v) treatment resultedin hydrolysis of hemicelluloses from lignocellulosic complex to pentose sugars along with other byprod-ucts such as furfural, hydroxymethyl furfural (HMF), phenolics and acetic acid. The acid pretreated sub-strate was delignified to the extent of 93.2% by the combined action of sodium sulphite (5.0%, w/v) andsodium chlorite (3.0%, w/v). The remaining cellulosic residue was enzymatically hydrolyzed in 0.05 M cit-rate phosphate buffer (pH 5.0) using 3.0 U of filter paper cellulase (FPase) and 9.0 U of b-glucosidase permL of citrate phosphate buffer. The maximum enzymatic saccharification of cellulosic material (82.8%)was achieved after 28 h incubation at 50 �C. The fermentation of both acid and enzymatic hydrolysates,containing 18.24 g/L and 37.47 g/L sugars, with Pichia stipitis and Saccharomyces cerevisiae produced7.13 g/L and 18.52 g/L of ethanol with corresponding yield of 0.39 g/g and 0.49 g/g, respectively.


1. Introduction

Worldwide high demand for energy, uncertainty of petroleumresources and concern about global climatic changes has led tothe resurgence in the development of alternative liquid fuels. Eth-anol has always been considered a better choice as it reduces thedependence on reserves of crude oil and promises cleaner combus-tion leading to a healthier environment. Developing ethanol as fuelbeyond its current role of fuel oxygenate, would require lignocell-ulosics as a feedstock because of its renewable nature, abundanceand low cost (Saha et al., 2005).

Lignocelluloses are mainly comprised of cellulose, a polymer ofsix-carbon sugar, glucose; hemicellulose, a branched polymer com-prised of xylose and other five-carbon sugars and lignin consistingof phenyl propane units. The presence of lignin limits the fullestusage of cellulose and hemicellulose. To convert these energy richmolecules into simpler forms, it is necessary to remove the ligninfrom lignocellulosic materials. A number of pretreatments suchas concentrated acid hydrolysis (Liao et al., 2006), dilute acidhydrolysis (Cara et al., 2008), alkali treatment (Carrillo et al.,2005), sodium sulphite treatment (Kuhad et al., 1999; Kapooret al., 2008), sodium chlorite treatment (Sun et al., 2004), steam

explosion (Ohgren et al., 2005), ammonia fiber explosion (Teymo-uri et al., 2005) lime treatment (Kim and Holtzapple, 2005), and or-ganic solvent treatment (Xu et al., 2006) have been used frequentlyto remove lignin and improve the saccharification of the cell wallcarbohydrates.

Of these methods, dilute acid treatment and enzymatic hydroly-sis have been the most popular ones. Dilute acid hydrolysis is a fastand convenient method to perform but it leads to the accumulationof fermentation inhibitory compounds such as furfurals, hydroxymethyl furfurals (HMF) and phenolics. These compounds, depend-ing on their concentration in the fermentation media, can inhibitmicrobial cell and affect the specific growth rate and cell-massyield. Several treatments e.g., ion exchange (Canilha et al., 2004;Chandel et al., 2007), overliming (Martinez et al., 2001; Chandelet al., 2007), activated charcoal adsorption (Mussatto et al., 2004;Canilha et al., 2004; Chandel et al., 2007), and laccase oxidationtreatment (Chandel et al., 2007) have been reported for thedetoxification of hydrolysate to improve the fermentability of acidhydrolysates into ethanol. However, the combination of pH adjust-ment by overliming followed by activated charcoal adsorption hasbeen shown to improve the detoxification of hemicellulosic hydro-lysate (Converti et al., 1999).

The acid hydrolysis pretreatment removes the hemicellulosicportion and some fraction of lignin but rest of the lignin remainsintact to the cellulosic substrate. Kaya et al. (2000) had reportedthat during enzymatic hydrolysis of lignocellulosic biomass,


* Corresponding author. Tel.: +91 124 24112972; fax: +91 11 24110876.E-mail address: [email protected] (R.C. Kuhad).

Bioresource Technology 100 (2009) 1214–1220



journal homepage: www.elsevier .com/ locate/bior tech

cellulase components, b-glucosidase and endoglucanase have morebinding affinity towards lignin than to the carbohydrates, resultingin lower efficiency of saccharification. Hence, to achieve maximumhydrolysis of cellulosics, which is a prerequisite for ethanol fer-mentation, an appropriate delignification treatment of biomass isrequired. In the present work, the combination of sodium sulphiteand sodium chlorite for the delignification of cellulosic biomasshas been attempted.

The cellulosic and hemicellulosic sugars obtained through acidand enzymatic hydrolysis can efficiently be used for ethanol fer-mentation either by separate fermentation of individual hydroly-sate or fermentation of mixed hydrolysate using co-culture.However, in co-culture cultivation, optimum growth conditionsof the yeasts would be different and might result in lowerefficiency and lower product yield. Hence, for better efficiency ofethanol production, the approach of separate hydrolysis andfermentation (SHF) was preferred (Olsson and Hahn-Hagerdal,1993).

In the present study, Prosopis juliflora (Mesquite), a perennialdeciduous thorny shrub, the common vegetation of semi-aridregion of Indian subcontinent, was used as a raw material for theproduction of cellulosic ethanol. The mesquite has recently beensuggested to be used as raw material for long-term sustainableproduction of cellulosic ethanol (Hopkins, 2007). Its nature totolerate drought, grazing, heavy soil, sand as well as saline dry flatsand no competence with animal feed demand made it a potentiallow value substrate for ethanol production. Here, an attempt wasmade to saccharify P. juliflora into reducing sugars and eventuallyto ethanol fermentation.

2. Methods


Prosopis juliflora wood, collected from University of Delhi SouthCampus, New Delhi, India, was comminuted by a combination ofchipping and milling to attain a particle size of 1–2 mm using alaboratory knife mill (Metrex Scientific Instrumentation, Delhi, In-dia). The processed substrate was washed thoroughly and driedovernight at 60 �C.

Commercial cellulase from Trichoderma reesei (ATCC 26921)(6.5 FPU/mg), b-glucosidase (Novozyme 188) (250 U/g) from Asper-gillus niger and 3,5-dinitrosalicylic acid (DNS) were purchased fromSigma, St. Louis, Missouri, USA. Ethanol was purchased from Merck(Darmstadt, Germany). Rest of the chemicals and media compo-nents of highest purity grade were purchased locally.

2.2. Micro-organisms and culture conditions

Pichia stipitis NCIM 3498 was procured from National ChemicalLaboratory (NCL), Pune, India and was maintained on agar slantscontaining (g/L): xylose, 20.0; yeast extract, 4.0; peptone, 5.0;KH2PO4, 1.5; MgSO4 � 7H2O, 0.5; agar, 20.0 at pH 5.0 ± 0.2 and tem-perature 30 �C. Saccharomyces cerevisiae, was procured from theculture collection of University of Delhi South Campus, New Delhi,India, and maintained on agar slants containing (g/L): glucose,30.0; yeast extract, 3.0; peptone, 5.0; agar, 20.0 at pH 6.0 ± 0.2and temperature 30 �C.

Inoculum of P. stipitis was prepared as described by Nigam(2001) using (g/L): xylose, 50.0; yeast extract, 3.0; malt extract,3.0; peptone, 5.0 at pH 5.0 ± 0.2 and temperature 30 �C. Saccharo-myces. cerevisiae inoculum was grown for 24 h at 30 �C in a culturemedium containing (g/L): glucose, 30.0; yeast extract, 3.0; peptone,5.0; (NH4)2HPO4, 0.25 at pH 6.0 ± 0.2 (Chen et al., 2007). Cells werecultured to an optical density of 0.6–0.8 at 620 nm.

2.3. Proximate chemical composition analysis of the substrate

The chemical composition of P. juliflora wood was analysed forholocellulose, Klason lignin, pentosans, ash and moisture content.The plant material was extracted with alcohol–benzene (1:2 v/v)to remove wax, resin etc. The extractive-free wood dust was pro-cessed for chemical analysis following the TAPPI (1992) protocols,(a-Cellulose–TAPPI Method T203 om–83; Klason lignin–TAPPIMethod T222 om–83; Pentosans–TAPPI Method T223 hm–84;Moisture–TAPPI Method T208 om–84 and Ash–TAPPI method211om–93).

2.4. Dilute acid pretreatment

The dilute sulphuric acid pretreatment of wood dust wasoptimized at varied temperatures (100–140 �C), treatment time(15–60 min) and acid concentrations (1.0–5.0%, v/v) at 10.0%(w/v) consistency, in a 20.0 L plastic vessel (Carboy, Tarson Pvt.Ltd., Kolkata, India), using an autoclave (Russian make). The acidhydrolysate after treatment was recovered by filtering the contentsthrough double-layered muslin cloth. The remaining wood dustwas washed with tap water till neutral pH. The hydrolysate wasanalysed for sugars, phenolics, acetic acid and furans and theleftover plant biomass was dried overnight till constant weightand used for further experiments.


The acid hydrolysate was overlimed at room temperature byadding dried lime [Ca(OH)2] till the pH reached 10.0, with constantstirring for 30 min by an overhead stirrer (Remi Motors Ltd,Mumbai, India). After overliming, the hydrolysate was neutralizedwith concentrated H2SO4 and centrifuged at 10,000g for 15 min toremove the precipitate formed during neutralization. Theoverlimed hydrolysate was further detoxified by treating withactivated charcoal (1.5%, w/v) with constant stirring at room tem-perature for 30 min and the sugar syrup was recovered throughvacuum filtration.

2.6. Chemical delignification of acid pretreated woody biomass

Acid pretreated residue of P. juliflora was delignified by treatingwith sodium sulphite (5.0–20.0%, w/v) alone and combination ofsodium sulphite (5.0%, w/v) and sodium chlorite (3.0%, w/v) andautoclaved at different temperatures (100–140 �C) for differenttime intervals (15–60 min). The delignified material was thenfiltered through double-layered muslin cloth and the leftovercellulosic residue was washed repeatedly with water till neutralpH. The cellulosic residue was dried overnight at 60 �C till constantweight and hydrolyzed enzymatically.

2.7. Enzymatic hydrolysis of delignified cellulosic substrate

Enzymatic hydrolysis of cellulose (delignified acid treated plantmaterial) was carried out at a 5.0% (w/v) consistency in 0.05 Mcitrate phosphate buffer (pH 5.0) containing 0.005% sodium azide.Before enzyme loading, slurry was acclimatized by incubating at50 �C on a rotatory shaker (Innova-40, New Brunswick Scientific,Germany) at 150 rpm for 2 h. Thereafter, a mixture of 3.0 U of filterpaper cellulase (FPase) and 9.0 U of b-glucosidase per mL of citratephosphate buffer was added to preincubated cellulose slurry andreaction continued for 36 h. Samples were withdrawn at regularinterval of 4 h, centrifuged at 10,000g for 15 min and the superna-tant was analysed for total reducing sugars released. The extent ofhydrolysis was calculated as follows:

R. Gupta et al. / Bioresource Technology 100 (2009) 1214–1220 1215

Saccharificationð%Þ

¼ Reducing sugar concentration obtainedPotential sugar concentration in the pretreated substrate

� 100

Different surfactants (1.0%, v/v or w/v) including nonionicsurfactants (Tween 20, Tween 40, Tween 60, Tween 80 andTriton X-100), polyethylene glycols (PEG 4,000, PEG 6,000 andPEG 10,000) and ionic surfactants (sodium dodecyl sulphite(SDS) and cetyl trimethyl ammonium bromide (CTAB)) wereused for studying their effect in improving enzymatic sacchari-fication.

2.8. Ethanol fermentation

The acid and enzymatic hydrolysates were fermented in anin situ sterilizable fermenter (B-Lite Sartorius India Ltd., Banga-lore, India) having geometric volume of 13.5 L and working vol-ume of 10.0 L. The acid hydrolysate (9.0 L) containing 18.24 g/Lsugars was supplemented with nutrients (g/L) NH4Cl, 0.5;KH2PO4, 2.0; MgSO4 � 7H2O, 0.5; Yeast extract, 1.5; CaCl2 � 2H2O,0.1; FeCl3 � 2H2O, 0.1 and ZnSO4 � 7H2O, 0.001 (pH 5.5 ± 0.2) andinoculated with 10.0% (v/v) culture of P. stipitis (OD 0.6). Whilecellulosic hydrolysate having 37.47 g/L sugars supplementedwith yeast extract, 3.0 g/L and (NH4)2HPO4, 0.25 g/L was fer-mented with S. cerevisiae (10.0%, v/v; OD 0.6). The agitation of150 revolutions per min (rpm), aeration 0.4 liter per minute(lpm), temperature 30 �C and pH 5.5 ± 0.2 were maintainedthroughout the process. The pH was adjusted with 2 N HCl and2 N NaOH, as required. A 10.0% (w/v) solution of siliconeantifoaming agent was used for controlling foam as and whenrequired. The dissolved oxygen was monitored continuously.Samples were withdrawn at regular intervals of 4 h and centri-

fuged at 10,000g for 15 min at 4 �C. The cell free supernatantswere used for the determination of ethanol produced and sugarconsumed. An attempt was also made to test the efficacy of theyeast to increase ethanol production by growing it in mediumhaving initial sugar load of 100 g/L.


The hydrolysates were analysed using high performance liquidchromatography (HPLC) (Shimadzu Kyoto, Japan) for the presenceof carbohydrates. Aminex (Bio-Rad, Hercules CA USA) column(300.0 � 7.8 mm) was used with 0.04 M H2SO4 as an eluent withflow rate of 0.5 mL/min keeping oven temperature at 60 �C withRID detector. Furans, furfurals and lignin derivatives were esti-mated with Luna 5 U C18 column (250.0 � 4.6 mm). The analysiswas done by using acetonitrile: 1.0% phosphoric acid (22:78) asmobile phase with flow rate 0.5 mL/min and oven temperature35 �C with UV–detector. Ethanol was estimated by Gas chroma-tography (GC) (Perkin Elmer, Clarus 500) with an elite-wax (crossbond-polyethylene glycol) column (30.0 m � 0.25 mm) at oventemperature of 85 �C and flame ionization detector (FID) at200 �C. The ethanol standards were prepared using commercialgrade ethanol (Merck, Darmstadt, Germany). Nitrogen with a flowrate of 0.5 mL/min was used as carrier gas. Total reducing sugarswere estimated by the DNS method and the total phenolicsreleased were determined by the Folin–Ciocalteu reagent method(Singleton et al., 1999) using vanillin as standard. The optical den-sity (A600 nm) of culture filtrate was measured using a doublebeam spectrophotometer (Specord 200). Dry biomass of yeastcells was measured after drying the yeast pellets at 70 �C till con-stant weight.

Table 1Release of sugars and phenolics during the dilute sulphuric acid (H2SO4) hydrolysis of P. juliflora wood at different temperatures

Time (min) 15 30 45 60

Temperature Acid concentration Sugar(mg/g)

Phenolics(mg/g)

Sugar(mg/g)

Phenolics(mg/g)

Sugar(mg/g)

Phenolics(mg/g)

Sugar(mg/g)

Phenolics(mg/g)

100 �C 1% 94.36 6.38 114.61 7.59 138.71 9.04 142.45 9.60(14.25)a (2.19) (17.31) (2.61) (20.95) (3.11) (21.52) (3.30)

2% 125.75 6.80 150.00 8.13 161.00 9.24 161.83 10.51(18.99) (2.34) (22.66) (2.79) (24.32) (3.18) (24.44) (3.61)

3% 144.29 7.56 159.05 8.30 176.67 4.55 177.35 11.565(21.8) (2.6) (24.03) (2.85) (26.69) (1.56) (26.79) (3.97)

4% 142.42 8.21 149.13 9.48 164.45 10.61 166.27 11.71(21.51) (2.82) (22.53) (3.26) (24.84) (3.64) (25.12) (4.02)

5% 139.17 8.88 142.70 9.71 156.75 10.62 159.29 12.24(21.02) (3.05) (21.56) (3.34) (23.68) (3.65) (24.06) (4.20)

120 �C 1% 108.10 6.71 122.07 7.91 148.56 9.37 154.69 9.37(16.33) (2.30) (18.44) (2.72) (22.44) (3.22) (23.37) (3.22)

2% 144.19 7.07 163.10 8.40 185.80 9.51 189.05 9.51(21.78) (2.43) (24.64) (2.88) (28.07) (3.27) (28.56) (3.27)

3% 165.24 7.83 179.37 8.80 200.50 10.05 204.84 10.05(24.96) (2.69) (27.10) (3.02) (30.29) (3.45) (30.94) (3.45)

4% 151.27 9.00 164.61 9.81 190.88 10.93 198.62 10.93(22.85) (3.09) (24.86) (3.37) (29.34) (3.76) (29.94) (3.76)

5% 146.99 9.21 157.46 10.02 190.34 10.56 188.34 11.06(22.20) (3.16) (23.79) (3.44) (28.75) (3.63) (28.45) (3.80)

140 �C 1% 117.57 7.68 130.75 8.36 140.23 9.67 156.86 9.835(17.76) (2.64) (19.75) (2.87) (21.18) (3.32) (23.69) (3.38)

2% 150.94 8.00 167.67 8.74 182.56 10.61 181.95 10.95(22.80) (2.75) (25.33) (3.00) (27.58) (3.65) (27.48) (3.76)

3% 172.65 8.47 176.77 8.36 169.14 11.47 160.79 11.86(26.08) (2.91) (26.7) (2.87) (25.55) (3.94) (24.29) (4.07)

4% 163.87 9.61 154.05 10.14 149.07 11.75 135.41 12.03(24.75) (3.30) (23.27) (3.48) (22.52) (4.04) (20.45) (4.13)

5% 155.24 10.435 148.87 8.36 126.86 12.23 114.69 12.87(23.45) (3.59) (22.49) (2.87) (19.16) (4.20) (17.32) (4.42)

a Values in parenthesis are sugar and phenolic yield with respect to total carbohydrate and total lignin content of the substrate.

1216 R. Gupta et al. / Bioresource Technology 100 (2009) 1214–1220

3. Results

3.1. Proximate chemical composition of P. juliflora

Prosopis juliflora wood contained 66.20% holocellulose (47.50%a-cellulose and 18.70% pentosans), 29.10% Klason lignin, 2.68%moisture and 2.02% ash content.

3.2. Dilute acid hydrolysis of P. juliflora

The woody material when hydrolyzed with different concentra-tion of dilute H2SO4, at 100 �C and 120 �C, the release in sugarincreased with increase in acid concentration upto 3.0% (v/v)H2SO4 and it declined thereafter. While at 140 �C, the acid concen-tration beyond 2.0% (v/v) resulted in continuous decrease inrelease of sugar. The maximum sugars (204.84 mg/g) werereleased, when the woody material was treated with 3.0% H2SO4

at 120 �C for 60 min. However, no significant difference in sugarreleased was observed when hydrolysis was carried either for 45or 60 min. (Table 1). The acid hydrolysis also resulted in releaseof phenolics ranging from 2.0–4.5% (w/w of phenolics present insubstrate). Under optimized hydrolysis conditions (3.0% H2SO4,120 �C, 60 min), the hydrolysate was found to contain furfural(0.34 mg/mL), HMF (0.58 mg/mL) and caffeic acid (0.13 mg/mL).


The sequential detoxification of acid hydrolysate using overlim-ing and activated charcoal resulted in a drastic decrease in the con-centrations of inhibitors. Overliming resulted in reduction of HMF

(51.18%), furfural (59.0%), acetic acid (12.4%) and caffeic acid(20.08%) followed by further detoxification with activated charcoaladsorption with additional removal of HMF (38.24%), furfural(29.31%), acetic acid (45.26%) and caffeic acid (74.92%).

3.4. Delignification of pretreated wood

A regular increase in delignification of woody material was ob-served with the increase in concentration of sodium sulphite from5.0% to 20.0% (w/v) (Table 2). The treatment of woody biomasswith 20.0% (w/v) sodium sulphite at 140 �C and 60 min resultedin release of 246.65 mg/g of phenolics. However, when sodiumchlorite (3.0%, w/v) was used along with sodium sulphite (5.0%,w/v), it reduced the concentration of sodium sulphite from 20.0%(w/v) to 5.0% (w/v), and released maximum phenolics at 120 �Cfor 30 min. Moreover, at optimized conditions, the release ofphenolics (271.10 mg/g) was found to be 8.4% (w/w) higher thanthe phenolics released by the sodium sulphite alone (20.0%, w/v,140 �C and 60 min).

3.5. Enzymatic saccharification of delignified cellulosic material

During the time course of enzymatic saccharification of deligni-fied cellulosic substrate, a regular increase in release of sugars wasobserved till 28 h of incubation, which remained almost constantthereafter, however, after attaining the maximum rate of sacchar-ification (49.58 mg/g/h, 4 h), the saccharification rate decreasedregularly showing the reciprocal relationship with saccharificationyield (Fig. 1). Moreover, the maximum yield of saccharification,586.16 mg/g, was achieved after 28 h incubation, with saccharifi-cation rate of 20.60 mg/g/h (Fig. 1).

Different ionic and nonionic surfactants were evaluated fortheir ability to improve the enzymatic hydrolysis of delignified P.juliflora. The amount of sugar released in the presence of Tween80 increased maximally by 47.51% compared to control and fol-lowed by PEG 4,000 (43.81%) and Tween 40 (39.17%). However,the quantity of sugar released in presence of ionic surfactantse.g., CTAB and SDS, decreased by 60.12% and 48.24%, respectively(Fig. 2).

3.6. Fermentations of hemicellulosic and cellulosic hydrolysates

The hemicellulosic hydrolysate containing 18.24 g/L sugarswhen fermented with P. stipitis produced 7.13 g/L ethanol with ayield of 0.39 g/g and productivity of 0.30 g/L/h after 24 h. After24 h of fermentation, P. stipitis produced biomass (5.96 g/L) withyield and productivity, 0.33 g/g and 0.25 g/L/h, respectively (Table

Table 2Release of phenolics during the delignification of acid treated biomass of P. juliflora

Time (min) 15 30 45 60

Temperature Chemicalsb

(%, w/v)Phenolics(mg/g)

Phenolics(mg/g)

Phenolics(mg/g)

Phenolics(mg/g)

100 �C S 5 92.21 105.54 149.98 149.98(31.69)a (36.27) (51.54) (51.54)

S 10 125.54 137.76 167.76 170.87(43.14) (47.34) (57.65) (58.72)

S 15 152.21 158.87 174.43 181.1(52.3) (54.6) (59.94) (62.23)

S 20 163.32 169.98 187.76 191.1(56.12) (58.41) (64.52) (65.67)

S + C 5 + 3 187.42 194.11 212.76 216.89(64.41) (66.7) (73.11) (74.53)

120 �C S 5 111.54 126.87 156.65 163.32(38.33) (43.6) (53.83) (56.12)

S 10 154.43 165.1 191.1 201.54(53.07) (56.73) (65.67) (69.26)

S 15 180.21 193.32 206.65 214.65(61.93) (66.43) (71.01) (73.76)

S 20 187.1 200.43 211.1 214.43(64.29) (68.88) (72.54) (73.69)

S + C 5 + 3 255.26 271.1 270.47 270.28(87.72) (93.16) (92.95) (92.8)

140 �C S 5 127.76 143.32 189.98 187.76(43.9) (49.25) (65.29) (64.52)

S 10 189.98 198.87 231.1 234.43(65.29) (68.34) (79.41) (80.56)

S 15 207.76 238.87 241.1 241.1(71.4) (82.09) (82.85) (82.85)

S 20 215.54 244.43 244.43 246.65(74.07) (84) (84) (84.76)

S + C 5 + 3 260.15 270.44 270.37 270.25(89.04) (92.81) (92.6) (92.51)

a Values in parenthesis are yield of phenolics with respect to total lignin contentpresent in substrate.

b Delignifying chemicals used; S, sodium sulphite and C, sodium chlorite.

0

100

200

300

400

500

600

700

12 16 20 24 28 32 360

10

20

30

40

50

60Sugar yield

Sacc

hari

fica

tion

rat

e (m

g/g/

h)

Saccharification rate

Time(h)4 8

Suga

r y

ield

(m

g/g)

Fig. 1. Enzymatic saccharification profile of delignified P. juliflora.


3). Fermentation of cellulosic hydrolysate (37.47 g/L) using S.cerevisiae, gave maximum ethanol (18.52 g/L) with yield (0.49 g/g) and productivity (1.16 g/L/h) after 16 h, whereas, the biomasswas found to be 8.02 g/L having yield (0.21 g/g) and productivity(0.50 g/L/h) (Table 4). However, S. cerevisiae, when grown inmedium containing 100 g/L glucose produced 42.5 g/L ethanolwith a yield of 0.43 g/g and productivity 2.66 g/L/h (Fig. 3).

4. Discussion

Dilute acid pretreatment method is commonly used to sacchar-ification of any lignocellulosic biomass. It has the dual advantage ofsolubilizing hemicellulose and further converting it to fermentablesugars. In the present work, dilute sulphuric acid pretreatment ofmilled wood of P. juliflora was optimized to achieve maximumsugar yield at minimum severity conditions. The optimum pre-treatment conditions i.e., 3.0% (v/v) acid at 120 �C for 45 min, gave

0

100

200

300

400

500

600

700

800

900

Suga

r yi

eld

Surfactants

T-20 T-40 T-60 T-80 T X-100 CTAB SDS PEG4000

PEG10000

PEG20000

Control

Fig. 2. Effect of different surfactants on the enzymatic hydrolysis of delignified P. juliflora after 36 h. (T, Tween (1%, v/v); TX, Triton (1%, v/v); CTAB, cetyl trimethyl ammoniumbromide (1%, w/v); SDS, sodium dodecyl sulphite (1%, w/v); PEG, polyethylene glycol (1%, w/v)).

Table 3Fermentation profile of detoxified acid hydrolysate using P. stipitis

Time (h) Ethanol (g/L) Sugar (g/L) Ethanol yield (g/g) Ethanol productivity (g/L/h) Biomass (g/L) Biomass yield (g/g) Biomass productivity (g/L/h)

0 0.17 18.24 0.01 0.00 0.15 0.01 0.004 0.19 17.43 0.01 0.05 1.47 0.08 0.378 1.08 15.09 0.06 0.14 2.91 0.16 0.3612 3.33 11.17 0.18 0.28 3.59 0.20 0.3016 4.41 9.02 0.27 0.3 4.55 0.25 0.2820 6.24 5.39 0.34 0.31 5.29 0.29 0.2624 7.13 3.51 0.39 0.3 5.96 0.33 0.2528 6.95 2.87 0.38 0.25 6.12 0.34 0.2232 6.71 2.05 0.37 0.21 6.23 0.34 0.19

Table 4Fermentation profile of enzymatic hydrolysate using S. cerevisiae

Time (h) Ethanol (g/L) Sugar (g/L) Ethanol yield (g/g) Ethanol productivity (g/L/h) Biomass (g/L) Biomass yield (g/g) Biomass productivity (g/L/h)

0 0.49 37.47 0.03 0.00 0.17 0.00 0.004 11.61 15.29 0.28 2.44 1.91 0.05 0.488 16.13 5.51 0.43 2.02 3.90 0.10 0.4912 17.47 3.84 0.47 1.46 6.05 0.16 0.5016 18.52 1.86 0.49 1.16 8.02 0.21 0.5020 16.81 1.67 0.45 0.84 7.99 0.21 0.4024 14.90 1.52 0.40 0.62 7.95 0.21 0.33

0

20

40

60

80

100

120

0 12 16 20 24 280.0

1.0

2.0

3.0

4.0

5.0

6.0

Suga

r an

d E

than

ol c

once

ntra

tion

(g/

L)

Time (h)

Eth

anol

Yie

ld (

g/g)

and

Pro

duct

ivit

y (g

/L/h

)

Ethanol production (g/L)Sugar consumption (g/L)Ethanol Yield (g/g)Ethanol Productivity (g/L/h)

4 8

Fig. 3. Time course for the production of ethanol from glucose (100 g/L).


a saccharification yield of 200.50 mg/g. While any further increasein pretreatment stringency caused the increase in release of toxiccompounds without much effect on sugar yield. Increase in toxiccompounds with corresponding increase in acid concentration,suggested to use the minimal acid concentration with reducedinhibitory compound and achieving maximum sugar hydrolysis.

The mechanistic rationale for the inhibitory action of furfuraland HMF on yeast cultures could be the result of decrease in theactivities of aldehyde dehydrogenase (AlDH), pyruvate dehydroge-nase (PDH) and alcohol dehydrogenase (ADH) (Modig et al., 2002).Detoxification of acid hydrolysate using Ca(OH)2 effectivelyreduced the level of inhibitors. This may be the result of eitherpolymerization or chemical transformations of these inhibitors athigher pH (Martinez et al., 2001). Previous overliming reportsshowed the similar trends of decrease in inhibitors present in acidhydrolysates (Martinez et al., 2001; Chandel et al., 2007). Whileactivated charcoal being hydrophobic in nature removes thehydrophobic inhibitory compound i.e., furan and phenolics moreeffectively (Saha, 2004; Chandel et al., 2007). Our results showedthat detoxification of acid hydrolysate using Ca(OH)2 followed byactivated charcoal adsorption had resulted in almost complete re-moval of majority of fermentation inhibitory compounds exceptacetic acid, which was removed partially (58%). However, the leftover concentration of acetic acid was lower than the minimuminhibitory concentration i.e., 5.0 g/L as reported by Taherzadehet al. (1997). Moreover, the yeast can grow on a media containingacetic acid up to 20 g/L at pH (5.5) (Taherzadeh et al., 2000). Con-verti et al. (1999) had also shown the detoxification of hydrolysatethrough sequential steps of overliming and activated charcoaladsorption. However, removal of phenolics using laccase had alsobeen reported (Chandel et al., 2007).

The chemical delignification of lignocellulosic material has pre-viously been reported to achieve better enzymatic saccharificationas compared to untreated sample (Saha, 2004; Kapoor et al., 2008).Earlier, Kuhad and coworkers had reported sodium sulphite (13.7%,w/v) for efficient delignification of biomass (Kuhad et al., 1999;Kapoor et al., 2008). In the present study, the maximum delignifi-cation was observed in P. juliflora when pretreated with combina-tion of sodium sulphite (5.0%, w/v) and sodium chlorite (3.0%, w/v).It is interesting to note that the overall concentration of sodiumsulphite (20.0%, w/v) was drastically reduced to 5.0% (w/v) withaddition of merely 3.0% (w/v) sodium chlorite as delignificationagent. Sodium chlorite is an elemental chlorine free compound,which at high temperature produces chlorous (HClO2) and hypo-chlorous acids (HOCl). The subsequent oxidation of chlorous byhypochlorous acid regenerates chlorine dioxide (ClO2) that furtherreact with lignin (Hamzeh et al., 2006). Chlorine dioxide has hadtremendous success as a replacement for chlorine because it is eas-ily substituted in a conventional chlorination stage without anyspecial modification, produces fewer potentially toxic byproductsand causes less damage to the wood fibers (Svenson et al., 2005).

During the enzymatic saccharification of delignified substrate, aregular decrease in the rate of hydrolysis was observed after 4 h ofsaccharification, which may be due to the end product inhibition ofthe enzymes (Kuhad et al., 1999). However, to weaken the feed-back inhibition caused by the cellobiose accumulation, b-glucosi-dase at a 3-fold concentration to FPase was used. In the presentexperiment, 5.0% (w/v) substrate consistency was found to obtainmaximum hydrolysis, however, further increase in substrate con-sistency led to the reduced extent of hydrolysis (data not shown).This decrease in hydrolysis efficiency may be because of mixingand heat transfer problem due to the rheological properties of adense fibrous suspension, which ultimately cause insufficientadsorption of the cellulase to the cellulose (Chen et al., 2007). Asthe enzymatic saccharification of cellulose involve transport of en-zyme molecules and soluble sugars between the solid substrate

and bulk reaction solution, a modification of the surface and inter-facial properties of the reaction system may improve saccharifica-tion (Hemmatinejad et al., 2002). For further improvement in thesaccharification efficiency of delignified P. juliflora, various ionicas well as nonionic surfactants were investigated. Among differentsurfactants tested, Tween 80, a nonionic surfactant has supportedthe enzymatic saccharification of delignified P. juliflora maximally(Fig. 2). Kaar and Holtzapple (1998) had also proposed that Tweenprotects the enzymes from thermal deactivation during the enzy-matic hydrolysis. This may be the result of reduced contact ofenzyme with the air-liquid interface due to surface activity of thesurfactant. The reduction in surface tension of the solution inhibitsthe non-productive attachment of the exoglucanase to the ligninsurface and allows the saccharifying exoglucanase greater accessto cellulose, which results in increase of sugar release (Hemma-tinejad et al., 2002).

The acid and enzymatic hydrolysates of P. juliflora werefermented by P. stipitis and S. cerevisiae, respectively. Several evi-dences suggested that separate fermentation by substrate specificorganisms work better instead of using mixed hydrolysate witheither single culture or co-culture method (Delgenes et al., 1996).Co-culture fermentation associates both hexose and pentose fer-menting yeasts that trade-off for oxygen requirement betweenthe two microorganisms. Whereas, ethanol production from mixedhydrolysates using single culture, circumvent the fact that utiliza-tion of xylose becomes a subject of glucose catabolite repression(Olsson and Hahn-Hagerdal, 1993). The acid hydrolysate of ligno-cellulosics comprises mainly pentose sugars and very few microor-ganisms e.g., Candida shehatae, P. stipitis and Pachysolentannophilus, which can utilize pentose sugars efficiently have beenidentified so far (Abbi et al., 1996). Among different pentose utiliz-ing yeasts, P. stipitis has shown great potential by having broadsubstrate specificity and no absolute vitamin requirement forpentose utilization (du Preez et al., 1986). In our experiment, theethanol yield (0.39 g/g) from hemicellulosic hydrolysate using P.stipitis was very much in agreement with previous reports havingyield, 0.37 g/g from aspen wood (Delgenes et al., 1996), 0.39 g/gfrom D-xylose (du Preez et al., 1986) and 0.36 g/g from P. juliflora(Kapoor et al., 2008). However, the ethanol yield obtained from cel-lulosic hydrolysate using S. cerevisiae (0.49 g/g) was even higherthan the previous results of ethanol yield 0.40 g/g from sweet sor-ghum (Mamma et al., 1995) and 0.48 g/g from corncob (Chen et al.,2007). A low aeration rate of 0.4 lpm was maintained throughoutthe process, as low aeration conditions are attributed to obligatehypoxic induction of essential fermentative enzymes (Mc Millanand Boynton, 1994) and also in slightly aerobic conditions the pro-duction of ethanol enhanced by lowering down the production ofglycerol as byproduct (Alfenore et al., 2004). However, S. cerevisiaewhen grown in medium having 100 g/L sugar produced 42.5 g/Lethanol, suggesting thereby to increase the initial sugar loadingeither by concentration of the hydrolysates or by supplementationwith sugar to improve cellulosic ethanol production.

5. Conclusion

Mesquite (P. juliflora) wood could serve as novel material for theproduction of ethanol. The separate hydrolysis and fermentationcould prove better in improving the ethanol production fromcellulosic hydrolysates by minimizing the problem of cataboliterepression. The ethanol production may be enhanced by increasingthe initial sugar load in cellulosic hydrolysates. There is still a needto develop; (i) a more efficient and economic pretreatment process(ii) a hyper-cellulase producing strain for improved saccharifica-tion and, (iii) an improved recombinant yeast strain capable ofutilizing both pentose and hexose sugars, which in turn wouldincrease ethanol production.


Acknowledgements

Authors are grateful to Department of Biotechnology (DBT),Ministry of Science and Technology (Government of India) for thefinancial support. Authors are thankful to Dr. Ajay Singh, AdjunctProfessor, Department of Biology, University of Waterloo, Ontario,Canada, for his suggestions during the preparation of the manu-script. The assistance from Mr. Adesh Kumar and Mr. ManwarSingh Shah during the course of this work is highly acknowledged.

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Journal of Molecular Catalysis B: Enzymatic 74 (2012) 170– 177


Journal of Molecular Catalysis B: Enzymatic

j ourna l ho me pag e: www.elsev ier .com/ locate /molcatb

Xylanase production from an alkalophilic actinomycete isolate Streptomyces sp.RCK-2010, its characterization and application in saccharification of secondgeneration biomass

Adesh Kumar, Rishi Gupta, Bhuvnesh Shrivastava, Yogender Pal Khasa, Ramesh Chander Kuhad ∗

Lignocellulose Biotechnology Laboratory, Department of Microbiology, University of Delhi South Campus, New Delhi 110021, India


Article history:Received 30 May 2011Received in revised form 9 September 2011Accepted 3 October 2011Available online 8 October 2011

Keywords:XylanaseStreptomycesOptimizationResponse surface methodologyEnzymatic saccharification

a b s t r a c t

Xylanase production by a newly isolated Streptomyces sp. RCK-2010 was optimized for varying cultureconditions following one factor at a time (OFAT) and response surface methodology (RSM) approaches.An initial medium pH 8.0, agitation 200 rpm, incubation temperature 40 ◦C and inoculum size 1.0% (v/v)were found to be optimal for xylanase production (264.77 IU/ml), after 48 h of incubation. Among variouscarbon sources tested, the actinomycete secreted higher level of xylanase on wheat bran. The productionmedium when supplemented separately with various nitrogen sources, the enhanced xylanase produc-tion was observed with beef extract followed by peptone. RSM employing central composite design(CCD) was used to optimize the xylanase production using wheat bran, beef extract and peptone asmodel factors. The RSM showed that the optimum level of wheat bran (2.5% w/v), peptone (0.2% N2

equivalent) and beef extract (1.2% N2 equivalent) resulted in almost 3.0 fold improvement in xylanaseproduction (2310.18 IU/ml). To the best of our knowledge this is the best xylanase volumetric producti-vity (1155 IU/ml/day) by any Streptomyces spp. reported in the literature. The enzyme was most activeat 60 ◦C and pH 6.0 and almost 40% stable after 4 h at optimum temperature. Saccharification of steamexploded rice straw with xylanase (60 IU/g dry substrate) supplemented with cellulase (24 FPU/g dry sub-strate) and �-glucosidase (60 IU/g dry substrate) resulted in 88% (w/w) saccharification of the cellulosicsubstrate.

© 2011 Elsevier B.V. All rights reserved.

1. Introduction

Economical feasibility of biofuel production from lignocellulosicmaterials, the second-generation biomass, is a major technologicalchallenge. Currently the major attraction for bioethanol produc-tion is cellulose fraction, while, hemicellulose, the second mostabundant natural polymer is yet to be tapped. Xylan, the majorhemicellulose, consists of 1, 4-�-linked d-xylose units substitu-ted with different side groups such as l-arabinose, d-galactose,acetyl, feruloyl, p-coumaroyl and glucuronic acid residue [1,2].Depending on its state, xylan can be used for various pur-poses such as, polymeric xylan as adhesives and emulsifiers [3],arabino-xylooligocaccharides as prebiotics [4,5], whereas mono-meric xylose can be fermented to ethanol [6,7].

Among various methods used to breakdown the xylan backbone,its hydrolysis using xylanases is one of the most environmentalbenign alternative, as xylanases are highly specific in their natureand application [2,8,9]. Xylanases (EC. 3.2.1.8) are also attracting

∗ Corresponding author. Tel.: +91 11 24112062; fax: +91 11 24115270.E-mail address: [email protected] (R.C. Kuhad).

extensive interest due to their wide applications in pulp bleaching,bioethanol production and oligosaccharides production [7,10,11].Furthermore, tolerance to high pH and temperature are desirableproperties of xylanases for their effective use in various industries.The alkalitolerant xylanases reduce the chlorine requirements forpulp bleaching, while the higher temperature will enhance theirrate of reaction. Though a variety of microorganisms have beenreported to produce xylanolytic enzymes [2,8,10], there are fewreports on the production of alkali and thermostable xylanase fromactinomycetes. The present study deals with the production ofthermostable and alkalitolerant xylanase from an alkalistable acti-nomycete isolate Streptomyces sp. RCK-2010.

Since, the production of xylanases is strongly influenced bytheir culture conditions and medium constituents, to maintain abalance among process conditions and to minimize the amountof un-utilized components, optimization of medium compositionis essential [8]. The optimization process is generally carried outusing one factor at a time (OFAT) approach, but it does not consi-der interaction among variables [12]. While, the optimization usingstatistical approaches such as response surface methodology (RSM)offers quick screening of large experimental domain and mode-ling of interactive effects of process variables, which enables each

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A. Kumar et al. / Journal of Molecular Catalysis B: Enzymatic 74 (2012) 170– 177 171

reaction parameters to be optimized incoherence with others forachieving maximum enzyme production [12,13]. Moreover, multi-variate experiments in RSM also reduce the number of necessaryoptimization and give more precise results than those obtained byunivariate strategies [13] and results in significant improvement ofenzyme production.

In the present study, the optimization of xylanase productionfrom an actinomycete isolate Streptomyces sp. RCK-2010 followingOFAT and RSM approach was carried out. Moreover, an attempt hasbeen made to evaluate the efficiency of xylanases in improving thehydrolysis of steam exploded rice straw, the abundantly availablesecond generation feedstock, into sugars.

2. Experimental


The steam exploded rice straw having 77% (w/w) holocellu-lose (18% hemicellulose, 59% cellulose) was procured from Dr. A.J. Varma, Polymer Chemistry Division, National Chemical Labora-tory, Pune, India. The substrate was washed thoroughly and driedat 60 ◦C in a hot air oven [14]. Xylan, cellobiose, glucose, car-boxymethyl cellulose, d-xylose and 3, 5 di-nitrosalicylic acid werepurchased from Sigma (St. louis, U.S.A.). All other media compo-nents and chemicals were purchased locally.

2.2. Isolation, screening of xylanase producing actinomycetes andidentification of the potent isolate

For isolation of alkalo-tolerant actinomycetes, the soil samplescollected from University of Delhi South Campus, New Delhi, Indiawere initially treated with alkali by suspending 1.0 g of each samplein sterile distilled water (pH 9.0) [10]. The samples were thenserially diluted and spreaded on actinomycete isolation agar contai-ning (g/L) sodium caseinate 2.0, l-aspargine 0.1, sodium propionate4.0, K2HPO4 0.5, MgSO4 0.5, FeSO4 0.001 and agar 15.0 (pH 8.0)and incubated at 37 ◦C for 96 h. The actinomycetes colonies deve-loped were purified by repeated transfer of cultures and the isolateswere screened for xylanase production using congo-red plate assaymethod and selected on the basis of hydrolysis zone [15].

For identification of the potent xylanase producing actino-mycete isolate (R-10), a 500 bp region of 16S rRNA gene wasamplified in a thermocycler (G-storm, USA) using the universalprimers (CCAGCAGCCGCGGTAATACG) and (ATCGGCTACCTTGTTACGACTTC). The PCR products were purified and sequenced as des-cribed earlier [13] and the nucleotide sequence has been depositedin the GenBank database (accession no. HQ658475). The sequencedata was analyzed for the homology with the similar existingsequences available in the data bank of National Center for Bio-technology Information (NCBI) using BLAST search.

2.3. Microorganism and maintenance

Streptomyces sp. RCK-2010 was maintained on xylan agar platescontaining (g/L) xylan 5.0, peptone 5.0, yeast extract 5.0, KH2PO41.0, MgSO4 0.1 and agar 20.0 at 37 ◦C, subcultured every fortnightand stored at 4 ◦C. The modified Horikoshi medium having (g/L)KH2PO4 1.0, MgSO4 0.1, yeast extract 5.0, peptone 5.0, and glucose5.0, was used for the xylanase production.

2.4. Optimization of xylanase production

In order to screen the effective parameters for the optimizationof xylanase production from Streptomyces sp. RCK-2010, variousprocess variables such as cultivation time (up to 60 h), tempe-rature (30–42 ◦C), initial pH (5.0–9.0) of the medium, agitation

(100–300 rpm), inoculum size (0.4–1.2% v/v of 18 h old culture),carbon sources and nitrogen sources were studied under submer-ged fermentation conditions using OFAT approach. The time courseof xylanase production was carried out at 37 ◦C, pH 8.0 and 200 rpmwith an inoculum size of 0.4% (v/v). Each factor examined for opti-mization was incorporated further in the subsequent experiments.All other experiment conditions were kept constant unless other-wise stated.

Further to study the interaction among the three effective para-meters selected from OFAT method, i.e., wheat bran (A), beef extract(B) and peptone (C) on xylanase production from Streptomyces sp.RCK-2010, experiments were conducted using RSM approach. Themaxima (+1) and minima (−1) values for wheat bran were 0.5and 2.5% (w/v), while for both beef extract and peptone were 0.2and 1.2% (N2 equivalent). The statistical software Package Design-Expert 6.0 Stat-Ease, USA was used to analyze the experimentaldesign. The design was used for the determination of the opti-mum culture conditions for xylanase production. The analysis wasused to identify the influence of variables on each other on thexylanase production and to determine the optimum fermentationconditions.

2.5. Characterization of xylanase from Streptomyces sp.RCK-2010

The xylanase produced under optimized conditions was puri-fied partially with ammonium sulphate precipitation. The optimalpH for partially purified xylanase activity was determined usingdifferent buffers (0.2 M) (citrate–phosphate, pH 3.0–6.5; phos-phate buffer, pH 6.0–7.5; Tris–HCl, pH 7.5–9.2; glycine–NaOH, pH8.5–10.5 and carbonate–bicarbonate, pH 9.0–10.0) and the opti-mum temperature for xylanase activity was determined between30 and 85 ◦C. The thermostability of the partially purified enzymewas studied by incubating the enzyme at 55 and 60 ◦C up to 4 h atpH 6.0, and the residual activities were estimated periodically.

2.6. Application of xylanase in saccharification of steam explodedrice straw

Enzymatic hydrolysis of steam exploded rice straw (10.0 g)was carried out at 5% (w/v) substrate concentration in 50 mMcitrate phosphate buffer (pH 5.0). The substrate suspension waspre-incubated at 50 ◦C on a rotatory shaker (Innova-40, New Bruns-wick Scientific, Germany) at 150 rpm for 2 h. Thereafter, the slurrywas supplemented with enzyme cocktail; cellulase (24 FPU/g), �-glucosidase (60 IU/g) and crude enzyme extract containing varieddosages of xylanase (20–80 IU/ml). The enzymatic reaction withoutxylanase served as control. Tween 80 (1% v/v) was also added inthe reaction mixture to facilitate the enzyme action. The enzyma-tic hydrolysis was performed at 50 ◦C and 150 rpm for 36 h. Samplesof enzymatic hydrolysate were withdrawn regularly and analyzedfor amount of glucose released.


All the analyses have been carried in triplicates and the data pre-sented here is the mean of the triplicates along with the standarddeviation.

The xylanase activities were determined by measuring theamount of reducing sugars released from xylan (1% w/v in citratephosphate buffer; pH 6.0) at 60 ◦C for 10 min, as described earlier[4]. One international unit (IU) of xylanase activity was defined asamount of enzyme required to release 1 �M of reducing sugars asxylose from xylan per min under reaction conditions. The proteinconcentrations were measured by the Lowry’s method with BSA(Bovine serum albumin) as standard [16]. The reducing sugars

172 A. Kumar et al. / Journal of Molecular Catalysis B: Enzymatic 74 (2012) 170– 177

were estimated following DNS method of Miller [17] and thesaccharification efficiency was calculated as follows:

Saccharification (%)

= Amount of sugar released in enzymatic saccharificationAmount of carbohydrate present in the substrate

×100


3.1. Selection and molecular identification of potentxylanase-producing actinomycete

Out of the 57 alkalo-tolerant actinomycetes isolated from soilsamples, 13 isolates showed the ability to produce xylanase. Amongthese isolates, R-10 was found to exhibit the largest zone of hydro-lysis on xylan agar plate flooded with congo-red [15]. Moreover,the identification of the actinomycete isolate R-10 was done by16S rRNA gene sequences. The nucleotide BLAST similarity searchanalysis based on 16S rRNA gene sequence revealed that the isolateR-10 was closely related to the genus Streptomyces (Fig. 1) and theorganism was termed as Streptomyces sp. RCK-2010.

3.2. Effect of pH, temperature, agitation and inoculum size onxylanase production

Time course of xylanase production by Streptomyces sp. RCK-2010 showed the maximum xylanase production (158.2 IU/ml)after 48 h of incubation and thereafter it gradually declined (Fig. 2).The reduction in xylanase yield could be due to nutrients depletionor due to proteolysis [18]. Among various incubation temperaturesstudied, the maximum xylanase production (170.72 IU/ml) withspecific xylanase activity of 73.59 IU/mg protein was achieved at40 ◦C. The enzyme production was observed to be decreased by

022.6

5101520

Streptomyces sp. MS362Streptomyces variabilis xsd08042Uncultured Streptomyces sp.Enterococcus sp. GS-5Streptomyces griseoflavusStreptomyces pseudogriseolus

Streptomyces sp. J22-G6Streptomyces acrimycini

Streptomyces althioticusStreptomyces sp. ERI 04Uncultured actinobacteriumStreptomyces sp. R10

Nucleotide Substitutions (x100)

Streptomyces flavoviridis HBUM174594

Streptomyces thermocarboxydus RAG-19

Fig. 1. Dendrogram of Streptomyces sp. RCK-2010 (Streptomyces sp. R-10) showingthe similarity with other Streptomyces cultures.

0

20

40

60

80

100

120

140

160

180

6 12 18 24 30 36 42 48 54 600

0.5

1

1.5

2

2.5

3

3.5

4

4.5

5XylanaseProte in

Xyl

anas

e ac

tivity

(IU

/mL

)

Prot

ein

(IU

/mL

)

Time (h)

Fig. 2. Time course of xylanase production from Streptomyces sp. RCK-2010 undersubmerged cultivation conditions.

12.3% on increasing the incubation to 42 ◦C (Table 1). The optimumtemperature of 40 ◦C for xylanase production is also in correlationwith our previous report on Streptomyces cyaneus [19].

The actinomycete isolate when grown under variable shakingconditions, the optimum xylanase of 170.79 IU/ml with specificactivity 73.51 IU/mg protein was observed at 200 rpm after 48 hof incubation (Table 1). Increase or decrease in the agitation ratesbeyond 200 rpm resulted in low xylanase yields. The lower enzymelevel under low agitation conditions may be attributed to the dissol-ved oxygen (DO) limitation, improper mixing of media componentsand cell clumping [20]. However, the decrease in enzyme activityon increasing the agitation might be as a result of the cells shearing[13].

Among wide range of initial pH of the production mediumtested, pH 8.0 was found to be more effective in production ofmaximum xylanase (178.67 IU/ml; specific activity 82.17 IU/mgprotein), which drastically decreased beyond pH 8.0 (Table 1). Ear-lier reports on xylanase production by several fungi and bacteriahave also been shown to be markedly dependent on the initialpH of the medium [21,22]. Interestingly, the preference of higherpH (8.0) by Streptomyces sp. RCK-2010 qualified it as a alkalophilicactinomycete.

An inoculum concentration ranging from 0.4 to 1.20% (v/v)revealed 1.0% (v/v) as optimal inoculum level for maximumxylanase yield (264.77 IU/ml) and specific xylanase activity(115.11 IU/mg protein) (Table 1). However, lower enzyme yield(226.68 IU/ml) at higher inoculum level (1.2% v/v) could be theresult of faster nutrient consumption. Hence, an optimal inoculumlevel is necessary for maintaining balance between the prolifera-ting biomass and available nutrients to produce maximum enzymelevel.

3.3. Effect of carbon and nitrogen sources on xylanase production

Among various carbon sources used Streptomyces sp. RCK-2010exhibited clear preference for lignocellulosic agro-residues as com-pared to pure sugars (Table 2) and wheat bran was observedto be the optimum substrate for maximum xylanase production(761.37 IU/ml). This may be because that wheat bran acts as a com-plete nutritious feed containing various soluble sugars, which arehelpful for the initiation of growth and replication of microorga-nisms and remains loose even under moist conditions providing alarge surface area [8]. In addition, higher xylanase production onwheat bran may be due to low lignin and silica content [11]. Wheatbran has been reported as an ideally suitable substrate for xylanaseproduction for other microorganisms as well [23,24]. Interestin-gly, when monosaccharides and disaccharides were used as carbonsources, xylanase production ranged from 82 to 260 IU/ml. Thisimplies that the enzyme is not specific to xylan-rich substrates andcan be produced constitutively up to a certain level. But the useof pure sugars as substrate is uneconomical for large-scale pro-duction of xylanases, while agricultural residues are cost-effectivesubstrates for xylanase production [25]. Further different concen-trations of wheat bran ranging from 1.0 to 3.0% (w/v) were testedfor xylanase production and the maximum xylanase production(805.12 IU/ml) was obtained when the actinomycete was grownon 2.5% (w/v) wheat bran (Fig. 3)

To optimize the effect of nitrogen sources for xylanase pro-duction, different organic and inorganic nitrogen sources wereadded by replacing the yeast extract and peptone from the pro-duction medium. The maximum activity of xylanase (617.58 IU/ml)with specific activity of 263.92 IU/mg protein was obtained withbeef extract and followed by peptone (340.69 IU/ml) (Table 3).The enhanced production of xylanase in presence of beef extractas well as peptone may be attributed to organic nitrogen sourcemediated regulation of microbial growth and metabolism, as has


Table 1Physiological parameters optimization for xylanase production from Streptomyces sp. RCK-2010 under submerged cultivation conditions following OFAT method.

Variables Temperature (◦C)

30 35 37 40 42

Activity (IU/ml) 90.11 ± 1.80 98.14 ± 3.93 156.34 ± 11.36 170.72 ± 0.68 149.71 ± 9.66Protein (mg/ml) 1.92 ± 0.12 2.07 ± 0.08 2.28 ± 0.13 2.31 ± 0.16 2.36 ± 0.14Specific activity (IU/mg protein) 46.86 ± 2.41 47.36 ± 0.48 68.52 ± 1.24 73.59 ± 1.11 63.42 ± 0.44

Variables Agitation (RPM)

100 150 200 250 300


Variables pH

5 6 7 8 9


Variables Inoculum size (v/v)

0.4 0.6 0.8 1.0 1.2


Table 2Effect of carbon sources on xylanase production from Streptomyces sp. RCK-2010 at 40 ◦C with shaking (200 rpm) after 48 h of incubation following OFAT method.

Carbon source (0.5% w/v) Xylanase yield (IU/ml) Protein mg/ml (mg/ml) Specific activity (IU/mg protein)

Amylose 181.01 ± 12.36 2.59 ± 0.14 69.78 ± 2.44Glucose 260.03 ± 17.22 2.02 ± 0.07 128.71 ± 8.41Pectin 115.31 ± 8.96 2.28 ± 0.06 91.84 ± 5.62Mannose 203.61 ± 15.63 2.22 ± 0.02 97.81 ± 6.57Lactose 151.63 ± 9.87 1.55 ± 0.07 103.0 ± 8.35Fructose 158.42 ± 11.84 1.54 ± 0.05 74.42 ± 6.21Sucrose 119.27 ± 9.14 1.60 ± 0.08 90.51 ± 4.58Maltose 152.34 ± 9.21 1.68 ± 0.10 47.45 ± 2.10Arabinose 82.81 ± 3.52 1.94 ± 0.08 42.57 ± 1.57Xylose 225.33 ± 13.25 1.80 ± 0.11 125.46 ± 10.25Wheat bran 761.37 ± 2.24 2.51 ± 0.14 303.33 ± 8.54Wheat straw 505.11 ± 36.33 2.31 ± 0.12 216.93 ± 12.73Prosopis juliflora 381.05 ± 19.22 1.91 ± 0.14 199.47 ± 14.37Lantana camara 373.35 ± 27.13 1.76 ± 0.08 212.13 ± 15.49Corn cob 325.19 ± 24.66 1.55 ± 0.13 184.85 ± 15.62Corn stover 289.86 ± 22.63 1.89 ± 0.14 153.36 ± 7.58Sugarcane bagasse 423.15 ± 36.51 1.63 ± 0.07 259.60 ± 5.36Birchwood xylan 215.27 ± 13.11 1.53 ± 0.08 140.69 ± 10.24

Table 3Effect of nitrogen sources on xylanase production from Streptomyces sp. RCK-2010 at 40 ◦C with shaking (200 rpm) after 48 h of incubation following OFAT method.

Nitrogen source (0.55% N2 equivalent) Xylanase yield (IU/ml) Protein (mg/ml) Specific activity (IU/mg protein)

OrganicPeptone 340.69 ± 19.74 2.21 ± 0.12 154.15 ± 5.47Beef extract 617.58 ± 36.98 2.34 ± 0.15 263.92 ± 3.65Yeast extract 281.69 ± 22.49 2.22 ± 0.08 126.89 ± 7.46Tryptone 258.34 ± 23.30 2.15 ± 0.10 120.16 ± 7.25InorganicNH4Cl 88.35 ± 5.26 (2.43 36.35) 2.43 ± 0.21 36.35 ± 2.54NaNO2 122.38 ± 6.93(2.13) 2.13 ± 0.15 57.45 ± 4.29NaNO3 205.08 ± 18.73 2.95 ± 0.09 69.51 ± 1.58(NH4)3HPO4 210.13 ± 18.73 2.37 ± 0.11 88.66 ± 3.65

Controla 78.26 ± 5.42 1.87 ± 0.07 41.85 ± 2.17

a The control medium used was devoid of nitrogen source.


Fig. 3. Effect of different dosages of carbon and nitrogen sources on xylanase pro-duction from Streptomyces isolate RCK-2010. Where, A, B, C, D and E are 0.5, 0.6, 0.7,0.8, 0.9% N2 equivalent of beef extract and peptone; and 1.0, 1.5, 2.0, 2.5 and 3.0%(w/v) wheat bran concentration, respectively.

been reported that the complex nitrogen is an essential require-ment for growth and enzyme production [26]. Moreover, it has alsobeen observed that nitrogen can significantly affect the pH of themedium during the course of fermentation [27], thereby influencesthe microbial metabolism. Our results are very much in agree-ment with the earlier reports, where actinomycetes were foundto produce higher xylanase on organic nitrogen sources [24,28,29].In order to estimate the optimum dosage of nitrogen sources, thexylanase production was carried out at different dosage (0.5–0.9%N2 equivalent) of both peptone and beef extract were carried out.Irrespective of the nitrogen sources tested, the maximum xylanaseproduction was observed at 0.7% N2 equivalent. The xylanase pro-duction with optimal level of beef extract and peptone as nitrogensources was observed to be 889.21 and 503.24 IU/ml, respectively(Fig. 3).

3.4. Statistical optimization of xylanase production using RSM

The result of RSM experiment for studying the effect of threeindependent variables; wheat bran (A), beef extract (B) and peptone(C), are presented along with the mean predicted and observed res-ponses in Table 4. The table showed the maximum and minimum

levels of variables resulted in the RSM and also depicted that totalfour center points were set up at runs of 7, 12, 13 and 19 and almostsimilar xylanase activities (∼1100 IU/ml) were observed. However,the maximum xylanase production (2310.18 IU/ml) was achievedat run no. 18 containing (% w/v); wheat bran 2.5, beef extract 1.2and peptone 0.20. While, the minimum xylanase (439.9 IU/ml)was observed in run no. 16 containing minimum amount ofwheat bran (0.50% w/v), beef extract (0.20% w/v) and peptone(0.20% w/v). From multiple regression analysis, it was observedthat the second-order polynomial equation can explain xylanaseproduction regardless of the significance of the coefficients:

Y = 65.69 + 235.79 A + 801.24 B + 406.87 C

+240.83 AB − 794.90 BC

where Y is the response value. In current experiment, Y value isthe level of xylanase production (IU/ml), A, B and C represent thecoded levels of wheat bran, beef extract and peptone, respectively.

The statistical significance of the regression model was checkedby F-test. The model was highly significant, as manifested by theF-value and the probability value [(Ptotal model > F) = >0.0001] wasachieved. The goodness of fit was manifested by the determina-tion coefficient (R2). In this case the R2 value of 0.9775 indicatedthat the response model can explain 97.75% of the total variations.The value of the adjusted determination coefficient (AdjR2) wasalso high enough (0.9695) to indicate the significance of the model.The parameter coefficient and the corresponding P-value sugges-ted that among the independent variables, A (wheat bran) and B(beef extract) have a significant effect on xylanase production.

The 3D response surfaces plots were employed to determinethe interaction of the medium components and the optimumlevels that have the most significant effect on xylanase produc-tion. Fig. 4a describes the effects of wheat bran and beef extracton xylanase production, when peptone was fixed at its middlelevel (0.70 N2 equivalents). The xylanase production yield increa-sed with increase in both the components and the increase inxylanase activity as a function of increasing levels of wheat branseems very similar to the increase as a function of increasing beefextract concentration (Fig. 4a). High levels of xylanase productionwith increase in wheat bran concentration could be due to the factthat wheat bran is nutrient reservoir for xylanolytic microorga-nism and acts as carbon and nitrogen source. While, the interactionbetween beef extract and peptone (wheat bran at middle level)

Table 4Experimental design and results of the RSM for the production of xylanase from Streptomyces sp. RCK-2010.

Run Factor-A wheat bran Factor-B beef extract Factor-C peptone Response xylanase activity (IU/ml)

Predicted value Actual value

1 0.5 1.2 0.2 1134.85 1106.92 2.5 1.2 0.7 1956.23 2015.213 1.5 0.7 0.2 1203.21 1192.654 0.5 0.7 0.7 724.06 677.785 2.5 0.2 0.2 1030.74 969.526 2.5 0.2 1.2 1187.97 1262.037 1.5 0.7 0.7 1128.42 1190.558 0.5 0.2 1.2 710.71 692.229 1.5 0.2 0.7 825.40 801.87

10 1.5 0.7 1.2 1053.64 944.9811 0.5 1.2 1.2 678.49 753.1112 1.5 0.7 0.7 1128.42 1190.2513 1.5 0.7 0.7 1128.42 1101.5214 2.5 1.2 1.2 1637.40 1553.5915 2.5 0.7 0.7 1532.79 1559.3116 0.5 0.2 0.2 372.17 439.9017 1.5 1.2 1.2 1157.94 1223.3518 2.5 1.2 0.2 2275.06 2310.1819 1.5 0.7 0.7 1128.42 1093.2520 1.5 1.2 0.7 1431.45 1347.64


Fig. 4. Response curves of the RSM experiments for the production of xylanasesfrom Streptomyces sp. RCK-2010.

demonstrated that the xylanase production increased with increasein beef extract concentration, but the enhancement in peptoneconcentration did not significantly increased the xylanase produc-tion (Fig. 4b).

The xylanase production capability of Streptomyces sp. RCK-2010 has been compared with the other Streptomyces strainsgrown under optimized conditions (Table 5). To the best of ourknowledge Streptomyces sp. RCK-2010 has been found to produce

0

20

40

60

80

100

120

3 4 5 6 7 8 9 10pH

Res

idua

l act

ivity

Fig. 5. Effect of pH on xylanase activity assayed at 60 ◦C for 10 min The xylanase acti-vity was assayed in pH range 3.0–10.0 using different buffers (citrate–phosphatebuffer, Tris–HCl and glycine–NaOH). 100% xylanase activity was equivalent to2310 IU/ml.

fairly good amount of xylanase (2310 IU/ml) compared to themajority of the Streptomyces strains reported in the literature(8–2360 IU/ml) and comparatively exhibited maximum volume-tric productivity (1155 IU/ml/day) under submerged fermentationconditions (Table 5).

3.5. Effect of pH and temperature on the activity and stability ofxylanase

Among various pH values (3–10) tested, the partially purifiedxylanase was active over a wide range of pH 5–9 with more than80% of residual enzyme activities. The optimal pH for Streptomycessp. RCK-2010 xylanase was 6.0 (Fig. 5). Although, most xylanasesknown today are active at either acidic or neutral pH [30–32],recently several alkalotolerant xylanases have also been reportedin efficient bleaching of paper pulp [4,33]. Moreover the alkalinexylanases, which are operationally stable at higher temperature,are more beneficial because of savings in cooling cost and time [4].The xylanase from Streptomyces sp. RCK-2010 exhibited its tem-perature optima of 60 ◦C at pH 6.0 (Fig. 6). Similar temperatureoptima of 60–65 ◦C have been reported earlier for other xylanases[4,31,33]. While the profiles obtained for thermostability (at 55 and60 ◦C) of partially purified xylanase from Streptomyces sp. RCK-2010revealed that the enzyme was thermostable with approximately 50and 40% residual activity at 55 and 60 ◦C, respectively, after 4 h ofincubation (Fig. 7).

Table 5Comparison of xylanase production by different species of Streptomyces.

Culture Production (U/ml) Time (days) Volumetric productivity (IU/mL/day) Substrate Reference

Streptomyces sp. QG-11-3 84.26 2 42.1 Xylan [36]Streptomyces sp. QG-11-3 2360 5 472.0 Wheat bran [36]Streptomyces galbus ATCC3005 12 6 2.0 Oat splet xylan [37]Streptomyces sp. Ab106 15 5 3.0 Cane bagasse [38,39]Streptomyces sp. AMT-3 28 10 2.8 Wheat bran [40]Streptomyces sp. AMT-3 70 10 7.0 Larchwood xylan [40]Streptomyces sp. Ab106 15 6 2.5 Corn hulls [41]Streptomyces actuosus A-151 10.3 4 2.6 Rice bran [42]Streptomyces olivaceoviridis E-86 1653 5 330.6 Corncob [43]Streptomyces galbus NR 8.75 5 1.8 Wheat bran [44]Streptomyces cyaneus SN32 730 2 365.0 Wheat bran [25]Streptomyces violaceoruber 1500 1.5 1000.0 Wheat bran [13]

Streptomyces sp. RCK-2010 2310 2 1155.0 Wheat bran This study


0

20

40

60

80

100

120

30 40 50 55 60 65 70 75 80 85

Temperature (°C)

Res

idua

l act

ivity

(%)

Fig. 6. Effect of temperature on xylanase activity assayed at different temperaturerange for 10 min. 100% xylanase activity was equivalent to 2310 IU/ml.

3.6. Application of Streptomyces sp. RCK-2010 in saccharificationof steam exploded rice straw

The time course of enzymatic saccharification of steam explodedrice straw with a combination of cellulase and xylanase revealedthat the sugar yield increased continuously till 48 h (Fig. 8). Amongvarious dosage of xylanase used along with cellulases, the enzymedosage of 60 IU/g dry substrate resulted in maximum increase insaccharification (208 mg/g dry substrate) and thereafter it did notresult in any significant improvement (Fig. 8). Though the xylanfraction in steam exploded rice straw was only 18%, the higherrelease of sugar might be due to the loosening of xylan backbonein the steam exploded substrate. The higher sugar yield may alsobe attributed to the release of glucan monomers bound tightly tothe xylan backbone. The synergistic action of cellulases and xyla-nases led to a total saccharification efficiency of 88%. Our resultsare consistent with the earlier works [7,34,35] and indicate highersaccharification of lignocellulosic substrates with combined actionof cellulases and xylanases.

0

20

40

60

80

100

120

0 0.5 1 1.5 2 2.5 3 3.5 4Time (h)

Res

idua

l act

ivity

(%)

Fig. 7. Thermo stability profile of xylanase activity assayed at 55 ◦C (�) and 60 ◦C(N) temperature for 240 min. 100% xylanase activities was equivalent to 2310 IU/ml.

0

50

100

150

200

250

6 12 18 24 30 36 42 48Time (h)

Suga

r re

leas

ed (m

g/g)

20 IU/g40 IU/g60 IU/g80 IU/g

Fig. 8. Effect of different dosage of xylanase (20–80 IU/g substrate) along with 24UFPase, 60U �-glucosidase and 1% (v/v) Tween 80 on the enzymatic saccharificationof steam exploded rice straw.

4. Conclusion

The xylanase from Streptomyces sp. RCK-2010 shows potentialin saccharification of second generation feedstocks into sugars fortheir eventual fermentation to ethanol.

Acknowledgements

The authors are thankful to Dr. A. J. Varma, Polymer ChemistryDivision, National Chemical Laboratory, Pune, India, for providingthe steam exploded rice straw. The financial support from Ministryof Environment and Forest (Government of India), Council of Scien-tific and Industrial Research, New Delhi and University of Delhi,Delhi, India is highly acknowledged.

References

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Application of lignocellulolytic enzymes produced under solid statecultivation conditions

Deepa Deswal, Abha Sharma, Rishi Gupta, Ramesh Chander Kuhad ⇑Lignocellulose Biotechnology Laboratory, Department of Microbiology, University of Delhi South Campus, Benito Juarez Road, New Delhi 110021, India


Article history:Received 2 September 2011Received in revised form 3 October 2011Accepted 8 October 2011Available online xxxx

Keywords:Lignocellulolytic enzymesSaccharificationANFChick feedDecolorization

a b s t r a c t

In this paper, cellulose from brown-rot fungus Fomitopsis sp. RCK2010, thermostable and alkalostablexylanase from Bacillus pumilus MK001 and laccase from Ganoderma sp. rckk-02 were evaluated for (i)saccharification of alkali pretreated rice straw and wheat straw, (ii) upgradation of chick feed and (iii)decolorization of dyes, respectively. The cellulose from brown-rot fungus resulted in a sugar release of151.48 and 214.11 mg/g, respectively, from rice straw and wheat straw, which was comparatively higherthan the earlier reports. While xylan, one of the main anti-nutritional factors (ANFs) present in the chickfeed was removed to an extent of 11.6 mg/g xylose sugars at 50 �C using the thermostable xylanase.Besides, the treatment with thermostable xylanase also brought about a release of 0.85 (mg/g) of solublephosphorous. Moreover, the laccase when used for the decolorization of Remazol Brilliant Blue R (RBBR)and xylidine ponceau cause almost complete decolorization in 2 and 4 h, respectively, depicting high rateof decolorization.


1. Introduction

Enzymes have always played a substantial role in the produc-tion of variety of commercial products from many industries viz.food processing, beverage production, animal feed, leather, textileand detergent (Gavrilescu and Chisti, 2005; Kuhad and Singh,2007). These enzymes not only make the process environmentallybenign but also play an important role in improving the productiv-ity and eventually the cost of product formation. Since last twodecades, among various industrial enzymes, celluloses, xylanasesand laccases (lignocellulolytic enzymes) are gaining enormousattention for their potential applications in bioconversion of cellu-losic materials into value added commodities, biostoning andbiopolishing of jeans, improving efficacy of detergents, macerationand color extraction from juices, enzymatic deinking, pulping,wastewater treatment, to improve the nutritional properties ofanimal feed, retting of flax, production of oligosaccharides, clarifi-cation of juices, biotransformation, treatment of dyes and otherorganic pollutants and development of biosensors, etc. (Xu, 2005;Kuhad and Singh, 2007; Kuhad et al., 2011).

Specially for more than a decade, hydrolysis of crop-byproductssuch as rice and wheat straw, etc. by celluloses has picked upmomentum for their conversion into sugars for ethanol production(Kuhad et al., 2010a; Deswal et al., 2011). The information onhydrolysis of crop byproducts (lignocellulosics) by celluloses from

soft-rot and white-rot fungi is available in plenty, while the reportson hydrolysis of cellulosics with celluloses from brown-rot fungalcelluloses are scanty (Baldrian and Valaskova, 2008). The brownrot-fungi differ substantially from soft-rot and white-rot fungi withrespect to the cellulolytic enzymes produced and the pattern ofcellulose degradation. These fungi are generally reported to lackthe exoglucanases that can hydrolyze crystalline cellulose, yet theycause the most destructive type of wood decay and are importantcontributors to biomass recycling (Kuhad et al., 1997). However,recently we have observed that Fomitopsis sp. RCK2010, a brown-rot fungus, produces exoglucanases (Deswal et al., 2011).

In addition to prominent usage of xylanase in pulp and paperindustry (Beg et al., 2001), they have been reported to improve feedvalue and performance of monogastric animals (Sinha et al., 2011).Xylanases have potential in improving chick feed by hydrolyzingthe anti-nutritional factors present in the feed grains i.e. non-starch polysaccharides such as arabinoxylans. Other advantagesof using xylanases as feed supplements are dehulling of cerealgrains, better emulsification and flexibility of feed materials(Galante et al., 1998).

Textile effluents containing synthetic dyes constitute one of themost problematic wastewaters to be treated not only for their highchemical and biological oxygen demands, suspended solids andtoxic compounds but also for their color (Diwaniyan et al., 2010).Biological decolorization by means of ligninolytic enzymes isconsidered to be a suitable and eco-friendly method to dye degra-dation and color removal. Among ligninolytic enzymes, laccaseshave received much attention due to their ability to oxidize both


⇑ Corresponding author. Tel.: +91 11 24112062; fax: +91 11 24115270.E-mail address: [email protected] (R.C. Kuhad).

Bioresource Technology xxx (2011) xxx–xxx



journal homepage: www.elsevier .com/locate /bior tech

Please cite this article in press as: Deswal, D., et al. Application of lignocellulolytic enzymes produced under solid state cultivation conditions. Bioresour.Technol. (2011), doi:10.1016/j.biortech.2011.10.023

phenolic and non-phenolic lignin related compounds as well ashighly recalcitrant environmental pollutants including syntheticdyes (Kuhad et al., 2004; Couto and Herrera, 2006). This abilitymakes them efficient for waste-water treatment of effluentscontaining chemically diverse and toxic dyes from textile indus-tries. Moreover, the decolorization of dyes with laccases alsoreduces the operational cost and energy required in conventionalchemical methods (Couto and Herrera, 2006).

In the present study, we have specifically studied the applica-tions of lignocellulolytic enzymes produced under solid state culti-vation conditions in the hydrolysis of crop-byproducts,improvement of chick feed and decolorization of industrial dyes.The cellulose from a brown rot fungal isolate Fomitopsis sp.RCK2010 (Accession number GU991381) has been evaluated forthe hydrolysis of pretreated crop-byproducts (wheat straw andrice straw). The highly thermostable and alkalostable xylanasefrom Bacillus pumilus MK001 (Accession number AY389345) hasbeen used for the improvement of chick-feed, while the laccasefrom Ganoderma sp. rckk-02 (Accession number AJ749970) hasbeen tested for its ability to decolorize azo (xylidine ponceau)and anthraquinone dyes (Remazol Brilliant Blue R; RBBR).

2. Methods

2.1. Raw materials and biomass preparation

Lignocellulosic substrates (wheat bran, wheat straw and ricestraw) obtained locally were dried and cut mechanically into smallpieces by a chopper, as described earlier (Kuhad et al., 2010b; Gup-ta et al., 2011). Xylidine ponceau and RBBR dyes were purchasedfrom Sigma Chemical Co. (St. Louis, MO, USA), while 1-hydroxy-benzotriazole (HBT) and other media components were procuredfrom Hi Media Laboratories Pvt. Ltd. (Mumbai, India).

The crop-byproducts were washed properly, dried at 60 �C tillconstant weight and stored in sealed plastic bags till further use.While, the chick feed (pH 5.9) obtained from a local poultry farmwas washed thoroughly with double distilled water and driedovernight at 60 �C.

2.2. Microorganisms and culture conditions

B. pumilus MK001, Fomitopsis sp. RCK2010 and Ganoderma sp.rckk-02 were procured from the culture collection of Lignocellu-lose Biotechnology Laboratory, Department of Microbiology,University of Delhi South Campus, New Delhi, India. The fungalcultures (Fomitopsis sp. RCK2010 and Ganoderma sp. rckk-02) weregrown on malt extract agar (MEA) composed of (g/l): malt extract,20.0; Ca(NO3)2�4H2O, 0.5; MgSO4�7H2O, 0.5; KH2PO4, 0.5; and agar,20.0 (pH 5.5) at 30 �C (Vasdev et al., 2005). While, B. pumilus strainMK001 was maintained on modified Horikoshi xylan–agar med-ium containing (g/l): xylan 5.0, peptone 5.0, yeast extract 5.0,KH2PO4 1.0, MgSO4�7H2O 0.1 and agar, 20.0 (pH 9.0) at 37 �C(Kapoor et al., 2007). The cultures were maintained by periodicalsub culturing and stored at 4 �C.

The fungal inoculum was developed by inoculating four myce-lial discs (0.8 cm dia each) from each fungal culture grown onMEA plate separately in 250 ml Erlenmeyer flasks containing50 ml of malt extract broth (MEB) and incubated at 30 �C understatic cultivation conditions for 7 days. The mycelial mat thusobtained was homogenized with pestle and mortar under sterileconditions and used as inoculum for the production of enzymes.In contrast the inoculum of B. pumilus MK001 was produced byinoculating a loopful of culture in 100 ml Erlenmeyer flasks eachcontaining 20.0 ml modified Horikoshi medium (Kapoor et al.,2007) at 37 �C and 200 rpm.

2.3. Lignocellulolytic enzyme production under solid statefermentation

Among lignocellulolytic enzymes, cellulose and laccase wereproduced from Fomitopsis sp. RCK2010 and Ganoderma sp. rckk-02, respectively, while the xylanase was produced from B. pumilusMK001. All the enzymes were produced under solid-state cultiva-tion conditions. For the production of fungal enzymes (celluloseand laccase), 0.5% (w w�1) of both the fungal cultures having0.25 ± 0.013 g of fungal dry mass were inoculated separately in250 ml Erlenmeyer flasks, each having 5.0 g of dry wheat branmoistened with mineral salt solution (g/l: (NH4)2SO4, 0.5; KH2PO4,0.5; MgSO4, 0.5 and pH 5.5) to attain the final substrate-to-mois-ture ratio of 1:3.5 and incubated at 30 �C for 11 and 6 days, respec-tively. After incubation, the fungal fermented wheat bran wasremoved aseptically and the enzymes were extracted in 50 mlcitrate buffer (100 mM, pH 5.5) (Sharma et al., 2005; Deswalet al., 2011).

For the production of xylanase, each 250 ml Erlenmeyer flaskscontaining 5 g of dry wheat bran moistened with mineral salt solu-tion, containing (g/l): KH2PO4, 1.0; NaCl, 2.5; MgSO4�7H2O, 0.1;(NH4)2SO4, 1.0; CaCl2, 0.1 and pH 9.0, to attain final substrate-to-moisture ratio of 1:4 were inoculated with 10% (v w�1) of seedculture of B. pumilus strain MK001 containing 106 CFU m l�1 andincubated at 37 �C for 5 days under static conditions. The enzymewas extracted in 50 ml citrate phosphate buffer (pH 6.0;100 mM) (Kapoor et al., 2007).

2.4. Applications

2.4.1. Enzymatic hydrolysis of lignocellulosic biomass2.4.1.1. Pretreatment of the substrates. Wheat straw and rice straw,the lignocellulosic, were pretreated with different dosage of so-dium hydroxide (0.5–2.5% w v�1) and sulphuric acid (0.5–2.5%v v�1) to enhance their enzymatic susceptibility. The pretreatedsubstrates were washed until neutral pH and dried in an oven at60 �C till constant weight was achieved.

2.4.1.2. Enzymatic hydrolysis. Prior to the enzymatic hydrolysis, thepretreated substrate at 2.0% substrate consistency in 50 mM citratephosphate buffer (pH 5.0) containing 0.005% (w v�1) sodium azidewas acclimatized at 50 �C and 150 rpm for 2 h. Further the enzymedosage of 25 U FPase/g dry substrate was added to the preincu-bated substrate suspension and the reaction mixture was incu-bated at 50 �C, 150 rpm for 24 h. Samples were withdrawn atregular intervals of 4 h, centrifuged at 10,000 rpm for 10 min andthe filtrate was used for the analysis of reducing sugars followingDNS method of Miller (1959).

2.4.2. Improvement of chick feedFor the pretreatment of chick feed with xylanase, the crude

xylanase dosage was optimized by incubating 10.0 g of chick feed(moistened with 20 ml water) with variable amounts of crudexylanase (10–50 U/g chick feed) for 2 h at 50 �C. After incubationthe suspension was centrifuged at 10,000 rpm for 10 min and thesupernatant was analyzed for the release of sugars and solublephosphorous.

2.4.3. Dye decolorizationTo evaluate the potential of laccase for decolorization efficiency,

50.0 ml of azo (xylidine ponceau) and anthraquinone (RBBR) dyes(100 mg/l; pH 5.5) were mixed separately with the enzyme(10 U/ml of dye solution; unless otherwise specified) (with andwithout HBT), was incubated at 30 �C and 150 rpm in a rotary incu-bator shaker. Dye content was monitored at the maximum visibleabsorbance of each dye (respectively, 542 and 595 nm). A control

2 D. Deswal et al. / Bioresource Technology xxx (2011) xxx–xxx


in which laccase was replaced by citrate phosphate buffer(100 mM, pH 5.5) was conducted in parallel. The decolorizationpercentage was defined as follows:

Decolorization percentageð%Þ ¼ ðA0 � AÞ=A0 � 100

where, A0 is the dye absorbance of the control, A is the dye absor-bance of the reaction sample.

In this study, decolorization of both the dyes was studied withrespect to three operational parameters: effect of HBT and its con-centration, reaction time and enzyme dose. To study the effect ofHBT concentration on dye decolorization, varied doses of the medi-ator (0.05–2.0% w v�1) were added to the reaction mixture. Theoptimum dose of the mediator was subsequently used for dyedecolorization with varied laccase doses (5–30 U/ml) to obtainthe optimal enzyme dosage for improved decolorization in mini-mum time. Decolorization was monitored at an interval of 2 h tillcomplete decolorization was achieved for each enzyme dose.


The compositional analysis of oven dried crop byproducts forlignin and holocellulose was carried out using the protocols ofTechnical Association of Pulp and Paper Industries (TAPPI, 1992).The cellulose activity was determined in accordance with theInternational Union of Pure and Applied Chemistry procedures(Ghose, 1987), while the laccase and xylanase were determined

as described earlier (Sharma et al., 2005). Reducing sugars were as-sayed by dinitrosalicyclic acid (DNSA) method of Miller (1959),while the soluble phosphorous content was analyzed followingthe method of Fiske and Subbarow (1925).

All the experiments were done in triplicates and the valuespresented here are the mean values ± standard deviations withinthe range of 10%.


3.1. Enzymatic saccharification of pretreated substrates

Lignocellulosic biomass cannot be hydrolyzed efficiently by en-zymes without its suitable pretreatment, as the lignin in the cellwall hinders the enzymatic catalysis. Therefore in order to improvethe enzymatic hydrolysis of rice straw and wheat straw the ligninremoval is essential (Gupta et al., 2011). In the present study weevaluated the effect of both acid and alkali pretreatments on theenzymatic saccharification. During the enzymatic saccharificationof alkali pretreated substrates, the sugar release increased withincreasing in NaOH concentration up to 2.0% and thereafterremained almost constant. The enzymatic hydrolysis of rice strawand wheat straw pretreated with 2% (w v�1) alkali resulted in a su-gar release of 151.48 and 214.11 mg/g dry substrate, respectively(Fig. 1a and b). The maximum acidic saccharification from wheatstraw (68.071 mg/g dry substrate) and rice straw (76.05 mg/g dry

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Fig. 1. Enzymatic saccharification profile of alkali treated rice straw (a), alkali treated wheat straw (b), acid treated rice straw (c) and acid treated wheat straw (d) usingcellulose from brown rot fungus Fomitopsis sp. RCK2010.

D. Deswal et al. / Bioresource Technology xxx (2011) xxx–xxx 3


substrate) was observed, when both the substrates were pre-treated with 0.5% (v v�1) sulphuric acid (Fig. 1c and d). As com-pared to untreated substrates (control), the higher residual ligninand lower residual holocellulose in pretreated substrates may bedue to the removal of acid soluble carbohydrate fraction, resultingin a decreased saccharification (Gupta et al., 2011). Also, duringacid pretreatment solubilized lignin can quickly condensate andprecipitate in acidic environments (Gupta et al., 2011). Our resultsare well in accordance with the previous reports where Chen et al.(2009) reported an increase in lignin content in acid hydrolyzedcorn stover. The results obtained in the present study revealed thatthe enzyme system from Fomitopsis sp. RCK2010 has shown com-paratively higher saccharification efficiency than the enzymesfrom brown-rots reported earlier (Table 1).

3.2. Improvement of chick feed

Xylanases for the digestion of antinutritional factors (ANFs)such as non-starch polysaccharides (NSP) are scarce or not presentin monogastric animals. Consequently, the dietary NSPs remainundigested and therefore negatively affect animal performance.The adverse effect is associated with the viscous nature of NSPs,their physiological and morphological effects on digestive tract,interaction with epithelium, mucus and microflora of gut. How-ever, the insoluble NSPs, like pentosans (arabinoxylans andxylans), beneficially decrease transit time, enhance water holdingcapacity and assist in faecal bulking in non-ruminant animals (Sin-ha et al., 2011). Pretreatment of chick feed with xylanase has thepotential to improve feed efficiency, growth rate of the monogas-

tric animals and mitigate environmental pollution (Jozefiak et al.,2007). Moreover, xylanases have been reported to reduce viscosityand increase absorption by breaking down the non-starch polysac-charides (NSPs) in the feeds of monogastric animals (Jozefiak et al.,2007; Kuhad et al., 2011). In the present study an attempt wasmade to study the effect of xylanase on the breakdown of NSPpresent in chick feed and release of reducing sugars in the solution.Among the different enzyme dosage (10–50 U/g dry chick feed)used, xylanase when used at 30 U/g dry chick feed resulted in max-imum release of reducing sugars (11.6 mg/g dry chick feed) andsoluble phosphorous (0.85 mg/g dry chick feed) at 50 �C after 2 hof incubation. The enzyme treatment of chick feed at higherenzyme doses did not increase release of reducing sugars and sol-uble phosphorous (Fig. 2). Xylanase from B. pumilus strain MK001has the potential to improve chick feed, as evident from the releaseof reducing sugars and increase in phosphate availability afterenzyme treatment of poultry feeds.

3.3. Dye decolorization by laccase from Ganoderma sp. rckk-02

In the present study, irrespective of the dyes, 0.1% of HBT, a re-dox mediator, when used in conjunction with 10 U of laccase/ml ofthe dye solution resulted in almost complete decolorization in 12 hof incubation at 30 �C (Table 2). This may be attributed to the factthat redox mediators, the compounds which speed up the rate ofreaction by shuttling electrons from the biological oxidation of pri-mary electron donors or from bulk electron donors to the electron-accepting organic compounds. Our results are well in accordancewith the other reports which showed enhanced laccase activity

Table 1Comparison of enzymatic saccharification of various substrates using celluloses from brown rot-fungi.

Source Substrate Reducing sugar (mg/g) References

Laetiporus sulphureus Pinus dendiflora 70.9 Lee et al. (2008)Fomitopsis pinicola Pinus dendiflora 3.5 Lee et al. (2008)Fomitopsis palustris Avicel 0.1 Yoon et al. (2007)Gloephyllum trabeum Spruce 25.51 Schilling et al. (2009)Gloephyllum trabeum Southern yellow pine 11.07 Schilling et al. (2009)Fomitopsis pinicola Spruce 138.51 Schilling et al. (2009)Fomitopsis pinicola Southern yellow pine 90.63 Schilling et al. (2009)Gloephyllum trabeum Southern yellow pine 46.35 Tewalt and Schilling (2010)Fomitopsis sp. RCK2010 Wheat straw 214.11 Present workFomitopsis sp. RCK2010 Rice straw 151.48 Present work

Fig. 2. Effect of different dosage of xylanase for the removal of ANFs from the chick feed at 50 �C, pH 9.0 after 2 h. Reducing sugars (d) and soluble phosphorous (N).



in the presence of HBT, a redox mediator (Kuhad et al., 2010a).Moreover in conjunction with 0.1% w v�1 of HBT, the effect of dif-ferent dosages of laccase (5–40 U/ml) on the decolorization of boththe dyes (100 mg/l) was also evaluated. The rate of decolorizationof both the dyes increased with increase in the enzyme dosage till30 U/ml, and thereafter it did not increase significantly (Fig. 3a andb). The decolorization pattern of xylidine ponceau and RBBR dye

depicted that when only 5 U/ml laccase dye solution was used,the complete decolorization was observed after 24 and 18 h,respectively (Fig. 3a and b). Interestingly with increased dosageof 30 U/ml laccase of dye solution, the decolorization period forxylidine ponceau and RBBR dyes reduced to 4 and 2 h, respectively(Fig. 3a and b). The rate of decolorization of RBBR was higher thanthe xylidine ponceau, which may be attributed to the structuraldifferences in the dyes (Diwaniyan et al., 2010). Comparatively,the laccase from Ganoderma sp. rckk-02 has shown better decolor-ization rates than the enzymes from other white-rot fungi reportedearlier (Table 3). Enayatzamir et al. (2009) have reported 68%decolorization of xylidine ponceau with laccase from Trametespubescens MB89 in 6 h, while, Baldrian (2004) achieved 74% decol-orization of RBBR (100 mg/l) with laccase from Daedalea quercinain 6 h.

4. Conclusion

In nature there are a wide variety of lignocellulolytic microor-ganisms, which due to there specific properties could be of moreuse in various industrial applications. In the present work, theunique filter paper cellulose producing quality of brown-rot fungusFomitopsis sp. RCK2010, the thermostability of xylanase from B.pumilus MK001 and the high rate of decolorization by laccase fromGanoderma sp. rckk-02 was used for there specific applications. The

Table 2Effect of varied concentrations of redox mediator (HBT) along with laccase on thedecolorization of xylidine ponceau and RBBR dyes.

HBT (% w v�1) Dye decolorization (%)

Xylidine ponceau RBBR

0.00 8.91 15.730.05 72.85 79.940.10 100.00 1000.15 100.00 1000.20 94.64 96.490.25 84.76 83.210.30 75.16 78.53

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a

b

Fig. 3. Decolorization of xylidine ponaceau (a) and RBBR (b) using laccase (alongwith HBT) from Ganoderma sp. rckk-02.

Table 3Comparision of decolorization of various anthraquinonic and azo dyes using laccase.

Enzyme source Dye Decolorization References

% Time(h)

Anthraquinonic dyesDaedaleaquercina

RBBR 74 24 Baldrian (2004)

Pycnoporoussanguineus

RBBR 88 2 Lu et al. (2009)

Trametespubescens

RBBR 90 24 Enayatzamir et al.(2009)

Coriolusversicolor

C.I reactiveviolet 13

74 24 Damle and Shukla(2010)

Coriolusversicolor

C.I reactiveblue 4


Coriolusversicolor

C.I reactiveblue 160


Trametesversicolor

Dispersedblue 3

80 3 Champagne andRamsay (2010)

Ganoderma sp.rckk-02

RBBR 100 2 Present work

Azo dyesTrametes sp.SQ01

Congo red 50 24 Yang et al. (2009)

Trametes sp.SQ01

Acid red 20 24 Yang et al. (2009)

Pycnoporoussanguineus

Congo red 66 18 Lu et al. (2009)

Trametespubescens

Xylidineponceau

68 6 Enayatzamir et al.(2009)

Coriolusversicolor

C.I acid black194


Coriolusversicolor

C.I direct blue54


Coriolusversicolor

C.I directviolet 51


Coriolusversicolor

C.I basic red18


Trametesversicolor

Reactive black5

20 300 Champagne andRamsay (2010)

Ganoderma sp.rckk-02

Xylidineponceau

100 4 Present work

D. Deswal et al. / Bioresource Technology xxx (2011) xxx–xxx 5


exploitation of such enzymes in specific industrial applications canimprove the productivity and cost effectiveness of any product.

Acknowledgements

The authors are grateful to Department of Biotechnology, Min-istry of Science and Technology, Government of India New Delhi,and Council of Scientific and Industrial Research, New Delhi India,for their financial support.

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