To compare the production of recombinant cellulase enzymes in two Saccharomyces species so as to ascertain the most suitable heterologous host for the degradation of cellulose-based biomass and its conversion into bioethanol.
To compare the production of recombinant cellulase enzymes in two Saccharomyces species so as to ascertain the most suitable heterologous host for the degradation of cellulose-based biomass and its conversion into bioethanol.
cDNA copies of genes representing the three major classes of cellulases (Endoglucanases, Cellobiohydrolases and β-glucosidases) from Trichoderma reesei were expressed in Saccharomyces pastorianus and Saccharomyces cerevisiae. The recombinant enzymes were secreted by the yeast hosts into the medium and were shown to act in synergy to hydrolyse cellulose. The conditions required to achieve maximum release of glucose from cellulose by the recombinant enzymes were defined and the activity of the recombinant enzymes was compared to a commercial cocktail of T. reesei cellulases.
We demonstrate that significantly higher levels of cellulase activity were achieved by expression of the genes in S. pastorianus compared to S. cerevisiae. Hydrolysis of cellulose by the combined activity of the recombinant enzymes was significantly better at 50°C than at 30°C, the temperature used for mesophilic yeast fermentations, reflecting the known temperature profiles of the native enzymes.
The results demonstrate that host choice is important for the heterologous production of cellulases. On the basis of the low activity of the T. reesei recombinant enzymes at fermentation temperatures, we propose a two-step process for the hydrolysis of cellulose and its fermentation into alcohol using cellulases produced in situ.
With dwindling fossil fuel resources and the necessity to reduce greenhouse gas emissions, there is a growing need to identify alternative environmentally sustainable energy sources. One potential green energy source is that derived from biomass, which can be converted to useable energy such as bioethanol (Goldemberg 2007). While first generation biofuels relied on biomass from agricultural crops such as corn and sugarcane, ethical issues raised over the use of ‘food for fuel’ has moved the field in the direction of extracting energy from nonfood sources such as woody plants, grasses, decaying or discarded biological waste, paper and municipal waste (Carere et al. 2008).
A challenge for the industry is to extract useable sugars from biomass. Lignocellulose accounts for 50% of world biomass (50 billion ton per annum) and is composed of cellulose, haemicellulose and lignin. The most abundant component, cellulose, is a linear polymer of repeating units of cellobiose, linked by β-1,4-glycosidic bonds (Guo et al. 2012), which exists as two distinct forms, a tightly packed insoluble crystalline form and nonorganized soluble amorphous regions. Haemicellulose is a heteropolysaccharide polymer composed of hexose and pentose sugars such as xylose, mannose, galactose, rhamnose and arabinose. Lignin links both haemicelluloses and cellulose together forming a recalcitrant physical barrier in the plant cell wall. However, lignin, which is composed of phenylpropanoids does not contribute to the carbohydrate pool in plants (Guo et al. 2012).
Cellulose can be hydrolysed to glucose by the concerted action of three classes of o-glycoside hydrolases (cellulases), namely endoglucanases (EGs), cellobiohydrolases (CBHs) and β-glucosidases (BGLs) (Elkins et al. 2010). EGs cleave randomly at amorphous sites along the cellulose fibre leading to a rapid depolymerization of cellulose, exposing new chain ends which are acted upon by CBHs to release cellobiose. Finally, BGLs hydrolyse cellobiose to release d-glucose (Asztalos et al. 2012). Several novel classes of enzymes such as the copper-requiring polysaccharide mono-oxygenases contribute to cellulose degradation by acting in synergy with the exo- and endoglucanases (Leggio et al. 2012; Žifčáková and Baldrian 2012).
A current goal in the biofuel industry is to generate micro-organisms that can degrade cellulose-based biomass and convert the released sugars into alcohol without the need to add cellulase enzymes. Cellulases are produced by saprophytic organisms such as filamentous fungi and anaerobic bacteria. Fungi, such as Trichoderma reesei, synthesize an array of cellulases (Martinez et al. 2008), which are mostly secreted from the cell (Foreman et al. 2003; Kubicek 2013). However, T. reesei is not an efficient fermenter of sugars to alcohol (Xu et al. 2009). Therefore, a suitable approach is to introduce cellulase genes into micro-organisms with known fermentative capacity such as yeasts of the Saccharomyces genus (Lynd et al. 2005). Recombinant cellulase genes mainly from T. reesei, Saccharomycopsis fibuligera or from bacterial sources have been expressed in yeasts either individually or in various combinations (Penttila et al. 1987, 1988; Cummings and Fowler 1996; Fujita et al. 2004; Den Haan et al. 2007b). The genes have been expressed from different promoters (Fujita et al. 2004; Gundllapalli et al. 2007; Marin-Navarro et al. 2011) and secretion of the proteins from the cell has been directed by secretory signals from heterologous genes (Van Rooyen et al. 2005). Cellulases have also been tethered to the yeast cell surface with anchors such as the S. cerevisiae α-agglutinin gene or secreted into the medium (Fujita et al. 2002; Yamada et al. 2010; Hasunuma and Kondo 2012).
The heterologous expression of cellulase genes has mainly been confined to strains of S. cerevisiae, the baker's yeast (Den Haan et al. 2007a; Du Plessis et al. 2010; Ilmen et al. 2011; Yamada et al. 2011) although a few studies have examined the expression of individual genes in other yeast species (Zurbriggen et al. 1991; Gurgu et al. 2011). To expand the potential host range for heterologous genes expression, we compared the production and secretion of the three major classes of T. reesei cellulase genes in a standard S. cerevisiae strain and in the polyploid lager yeast S. pastorianus. Our results indicate that S. pastorianus strains produce substantially higher levels of all three classes of cellulases. The released sugars from cellulose hydrolysis were sufficient to support fermentations using cellulose-based substrates as a sole carbon source and yielded substantial levels of alcohol.
Saccharomyces cerevisiae strain S150-2B (Mata, leu2-3, 112 ura3-52, trp1-289, his3) is a haploid laboratory yeast strain (Campbell et al. 2002). The industrial S. cerevisiae strains Y2699 and K103 were kindly supplied by Guinness Brewery, Dublin, Ireland. Saccharomyces pastorianus strain CMBS-33 was obtained from the Centre for Malting and Brewing Science, Leuven, Belgium (Bond et al. 2004; James et al. 2008). Strains CM51, C10 and C1051 are stress tolerant derivatives of the strain CMBS-33 and were generated by an adaptive evolution protocol to identify variants with tolerance to high temperatures and high-specific gravity (James et al. 2008). Trichoderma reesei strain QM9123 was purchased from the Commonwealth Agricultural Bureaux International, CABI, (Surrey, UK). The S. cerevisiae and S. pastorianus strains, unless otherwise stated, were cultivated at 30°C in yeast extract peptone dextrose (YPD; 10 g l−1yeast extract, 20 g l−1peptone, 20 g l−1 dextrose) or synthetic complete (SC) medium (1·7 g l−1 yeast nitrogen base (Foremedium, Norwich, England), supplemented with 5 g l−1 (NH4)2SO4, 20 g l−1of casamino acids and 20 g l−1glucose or sucrose (Den Haan et al. 2007b). For growth comparisons, yeast strains were inoculated into 0·1 ml medium in a 96-well plate (Sarstedt, Numbrecht, Germany) using a 96-micropin inoculator. Absorbance was measured every hour at 620 nm on a Multiskan Ascent (Thermo Labsystems, Milford, MA). Trichoderma reesei was cultured at 25°C in SC medium containing 30 g l−1lactose. Strains harbouring plasmids were grown in YPD or SC supplemented with either 0·2 g l−1Geneticin (G418; Melford, Ipswich, UK) or 0·3 g l−1hygromycin (Foremedium, Hunstanton, UK).
Cellulosic substrates were purchased from Sigma-Aldrich, Dublin, Ireland and were prepared as follows: Carboxymethyl cellulose [CMC;1% (w/v)] and Avicel PH-101[10% (w/v)] were suspended in 50 mmol l−1sodium acetate, pH 5. Phosphoric acid swollen cellulose (PASC) was generated from Avicel PH-101 as previously described (Zhang et al. 2006). To determine the concentration of cellulose (g l−1) in PASC, varying amounts of Avicel were pretreated with phosphoric acid. The weight of the swollen cellulose (wet weight) was plotted against the starting dry weight of Avicel. A linear correlation between the dry weight of Avicel and the final wet weight of PASC was observed (R2 = 0·997). Using the conversion factor, calculated from the slope of the line, 1 g of PASC was deemed to contain 0·025 g cellulose. Based on this conversion factor, the theoretical yield of glucose from PASC was calculated.
With the exception of egl2, cDNAs for each of the cellulase genes were generated from T. reesei mRNA using the primers listed in Table S1. The mRNA was generated as previously described (Wilkins and Smart 1996), cDNA was synthesized using the one-step High-capacity cDNA Reverse Transcription (RT) Kit from ABI (Applied Biosystems, Foster City, CA) as per the manufacturer's instructions. Full-length copies of each cellulase gene were generated by amplifying a series of DNA fragments with overlapping sequence homology of between 92 and 245 nts using gene specific primers (Table S1). Details of the cloning strategy are outlined in the supplementary Data (S1). The DNA fragments were transformed into S. cerevisiae S150-2B strain together with the low copy number plasmid pGREG586 and the genes were reconstituted and inserted into the plasmid by homologous recombination in vivo, placing the genes downstream of the GAL1 promoter and upstream of the CYC termination cassette, which directs 3′ end mRNA processing and transcription termination (Jansen et al. 2005). The endogenous GAL1 promoter in pGREG586 was then replaced with various S. cerevisiae promoters such as PGK1, TEF1 and HXT7 [Table S1, Fig. 2 and supplementary Data (S1)]. Reconstitution of the full-length cDNAs in the context of the promoter and terminator sequences was verified by DNA sequencing at GATC Biotech (Konstanz, Germany) using the same gene-specific primers as listed in Table S1. The gene cassettes were subsequently cloned into the high copy number plasmid pRSH42 (Taxis and Knop 2006) by homologous recombination in vivo as described in the supplementary Data (S1). The egl2 gene was amplified from T. reesei genomic DNA and cloned directly into the pRSH42 plasmid, referred herein as pRSH, as described in the supplementary Data (S1).
The copy number of plasmid pRSH was determined by Real-time PCR. Briefly genomic DNA was isolated from cultures of S. cerevisiae and S. pastorianus, harbouring the pRSH plasmid as previously described (Bond et al. 2004). A known amount the plasmid pGREG586 was added to the genomic DNA preparations. Real-time PCR was carried out as previously described (Bond et al. 2004) using primers specific to the Hygromycin- and Kanamycin-phosphotransferase genes (Table S1) that are present on the plasmids pRSH and pGREG586 respectively. The efficiency of amplification (Eff) for each primer set was determined from a plot of Ct values of serial dilutions of the template DNA using the formula Eff = 10(−1/slope)–1. The copy number of the pRSH plasmid in both species was determined using the standard curve method and/or the ΔΔCt method using the formula Hygro 1·5Ct(S.c)-Ct(S.p)/Kan 1·8Ct(S.c)-Ct(S.p) where S.c denotes S. cerevisiae and S.p denotes S. pastorianus and the efficiency of amplification of the Hygro gene was 1·5, whereas that for the Kan gene was 1·8.
Yeast strains were inoculated into 0·2 ml SC-sucrose in 96-well microtitre plates using a 96-micropin inoculator. Cultures were grown for 48 h at 30°C. The spent medium supernatants (0·2 ml) were incubated in a 1 : 20 volumetric ratio with a reaction buffer [45 mmol l−1 sodium acetate, pH 5, 1 mmol l−1 p-nitrophenylglucopyranoside (p-NPG)] at 50°C. The reaction was stopped after 30 min by the addition of an equal volume of 50 mmol l−1 sodium carbonate. Enzyme activity was quantified by measuring the absorbance at 414 nm.
CBHII activity was monitored as previously described (Den Haan et al. 2007a) with some modifications. Briefly, yeast strains expressing cbh2 were cultured in 20 ml YP-sucrose (50 g l−1) for 24 h. The supernatants of the cultures were filtered through a PES membrane of pore size 0·2 μm (Corning, NY). CBHII, present in culture supernatant, was adsorbed onto phosphoric acid-swollen cellulose (PASC, 20 g l−1, wet weight) by incubation at 4°C overnight. The cellulose was washed with sodium acetate buffer (50 mmol l−1, pH 5) and then resuspended in 5 ml of the same buffer and incubated at 50°C for 48 h. Total sugars released were assayed using the phenol sulphuric acid method (Wood and Bhat 1988). Alternatively, enzyme activity was determined using an indirect glucose release assay. Culture supernatants were mixed in a 1 : 1 volumetric ratio with supernatants from cultures expressing the gene cassette TEFbgl1. The mixture was then incubated in a 1 : 1 ratio with PASC; 200 g l−1wet weight in sodium acetate (50 mmol l−1, pH 5·0) and incubated at 50°C. Glucose released by the action of EGI and BGLI was quantified after 24 h using the glucose HK assay kit (Sigma, Dublin, Ireland) as per manufacturer's instructions.
Yeast strains expressing endoglucanase gene cassettes egl1, egl2 and egl2-egl1 were inoculated into 0·2 ml SC-sucrose in a 96-well plate using a 96-micropin inoculator. Cultures were grown for 48 h at 30°C. The cultures were centrifuged and the supernatants were filtered through a PES membrane of pore size 0·2 μm (Corning, NY). The culture supernatants were mixed in a 1 : 1 volumetric ratio with 1% CMC in 50 mmol l−1 sodium acetate, pH 5, and incubated at 50°C for 3 h. The released reduced sugars were measured using a dinitrosalicylic acid (DNS) assay as described previously (Wood and Bhat 1988).
A standard unit of cellulase activity was derived for each cellulase enzyme. The measured absorbance value from each of the enzyme assays described above was compared to absorbance values generated from a standard curve using commercial T. reesei cellulase cocktail (0–60 mU, Sigma; C8546). The exact composition of the commercial cellulase was not divulged by the manufacturer, however, it was derived from culture supernatants of T. reesei strain ATCC 26921, which contains the major three classes of cellulases used here. The derived mU value was divided by the total number of cells in the original yeast culture (mU cell−1) and then multiplied by 1 × 108 to give a value of mU 108−1cells. The resultant units are referred to as UEG, UCBHII and UBGLI respectively.
The release of glucose from PASC by individual or combinations of recombinant cellulases was assayed as follows. Recombinant yeasts were cultured in SC-sucrose in volumes ranging from 5 to 15 ml for 72–120 h. The spent culture supernatants, either individually, in pair-wise combinations or combined in 1 : 1 : 1 volumetric ratios were incubated with PASC (100–1000 g l−1 in 50 mmol l−1 sodium acetate buffer, pH 5) at 50°C in volumes ranging from 0·25 to 10 ml for 48–72 h. The temperature used for hydrolysis reflects that used in similar studies (Fujita et al. 2004; Den Haan et al. 2007a; Du Plessis et al. 2010; Yamada et al. 2010) and is within the optimum temperature range for cellulase activity (Andrade et al. 2011). Alternatively, culture supernatants were concentrated (1·5X–60X) by filtration through a 0·2 μmol l−1 PES membrane filter, followed by application to a Spin-X UF column (molecular weight cut off, 10 kDa; Corning, NY), prior to incubation with PASC. Glucose release was determined using a Glucose HK assay kit (Sigma) as per manufacturer's instructions.
Yeast strains expressing the gene cassettes PGKegl2-PGKegl1, TEFcbh2 and TEFbgl1 (referred to as Cel 2.0) were individually cultured in a 170 ml of SC-sucrose for 48 h using a starting inoculum of 1 × 105 cells ml−1. The supernatants were mixed in a 1 : 1 : 1 volumetric ratio and then concentrated (17X) as described above. PASC (1000 g l−1, wet weight) in 1 × YP was added to the concentrate. The solution was incubated at 50°C for 24 h and then cooled to room temperature. Freshly grown yeast strains expressing the gene cassettes PGKegl2-PGKegl1, TEFcbh2 and TEFbgl1 (5 × 106 cells ml−1 of each) were added to yield a final total cell concentration of 1·5 × 107 cells ml−1. Two independent fermentations were carried out at 30°C in a reaction volume of 10–20 ml. Alternatively, yeast strains expressing the gene cassettes were cultured either separately (5 ml) or together in a co-culture (15 ml) in YPD for 48 h with a starting inoculum of 1 × 105 cells ml−1. The supernatants from the separately grown cultures were pooled together. PASC (1000 g l−1, wet weight) in 1X YP was added to the supernatants. The pretreatment and fermentations were carried out as described above. Alcohol levels during and at the end of the fermentation were determined using an alcohol dehydrogenase assay kit (Sigma) and using a standard curve of commercial ethanol at various dilutions. The value at time 0 was subtracted from subsequent readings.
The data sets for enzyme activities and glucose release were tested for statistical significance using a Student's t-test. P-values <0·05 were deemed significant.
To establish a baseline for comparison of cellulase expression, the growth rates of the yeast strains chosen for analysis were examined in rich medium supplemented with either glucose or sucrose (Fig. 1a,b). The latter was examined as subsequent analysis of enzyme activity was carried out in this medium. The S. cerevisiae strain displayed a faster doubling time when grown in either glucose or sucrose at 30°C (Fig. 1a,b) and reached a higher final cell density than any of the S. pastorianus strains. The stress-tolerant S. pastorianus strains showed similar growth kinetics as the parental strain CMBS-33 when grown in either medium, with the exception of C10, which showed an apparent slower growth rate in glucose, however, this growth difference was not statistically significant.
Initially, a single gene representing each of the three major classes of cellulases, namely endoglucanases, cellobiohydrohydrolyses and beta-glucosideases, was chosen for expression in the heterologous hosts. The genes cbh2, bgl1 and egl1, (Fig. 2a–c), encoding CBHII, BGLI and EGI, respectively, were chosen for expression based on previous data relating to their known enzymatic activity and final cellular location (Penttila et al. 1987; Saloheimo et al. 2002; Ilmen et al. 2011). Subsequently, the egl2 gene was cloned separately or was co-expressed along with egl1 (Fig. 2d–e). The genes were cloned as cDNA copies under the control of several promoters [Phosphoglycerate Kinase 1 (PGK), Translation Initiation Factor 1 (TEF) or Hexokinase 7 (HXT), Fig. 2a–e] and contained the secretory signal sequence native to each gene (black boxes Fig. 2a–e).
To determine if the recombinant enzymes were secreted from the cells into the medium, yeast cells expressing each cellulase gene individually were cultured and the spent culture supernatants were assayed for each individual enzyme activity as described in the Methods section. The enzymatic activity for all three enzymes was significantly higher in the S. pastorianus strains compared to S. cerevisiae strain S150-2B (Fig. 3a–c). The levels of enzymes produced were dependent on the gene copy number as significantly more activity was detected in the supernatants from cells containing the high copy number plasmids. All S. pastorianus strains produced relatively similar levels of enzymes. To ensure that the low levels of enzymatic activity observed in S. cerevisiae S150-2B was not an anomaly particular to that strain, the activity from the bgl1 gene was examined in two further S. cerevisiae strains. As shown in Table 1, BGLI activity was 8–10-fold lower in the S. cerevisiae strains compared to the S. pastorianus strains. The enzyme activity was not influenced by the ploidy of the cells as isogenic 1n-4n strains of S. cerevisiae showed similar levels of BGLI activity (data not shown). To rule out that the observed enzyme activity resulted from differences in the rate of plasmid replication in the two yeast species, the copy number of the pRSH plasmid in both species was determined. Using two independent quantification methods, no more than a twofold difference in the copy number of pRSH was observed, therefore this parameter cannot account for the observed 8–10-fold difference in enzyme activity between the two yeast species. Likewise, expression from the PGK promoter does not significantly differ between the two species (James et al. 2002).
|Saccharomyces cerevisiae S150-2B||5·01 (0·13)|
|Saccharomyces cerevisiae Y2699||8·12 (1·01)|
|Saccharomyces cerevisiae K103||9·29 (1·55)|
|Saccharomyces pastorianus CMBS||42·30 (1·29)|
|Saccharomyces pastorianus CM51||57·41 (0·90)|
|Saccharomyces pastorianus C10||75·45 (12·10)|
|Saccharomyces pastorianus C1051||58·13 (17·2)|
The ability of the recombinant strains to hydrolyse cellulose-based substrates to release sugars for subsequent fermentation was next examined. As significantly more enzyme activity was produced in the S. pastorianus strains and there was no significant difference in enzyme production between the S. pastorianus strains, just a single S. pastorianus strain, the stress-tolerant CM51, containing the cellulose genes expressed on the high copy plasmid, was carried forward for further analysis. To determine if synergy exists between the three classes of recombinant enzymes, hydrolysis of the cellulose substrate PASC by spent medium supernatants from cells expressing each of the genes individually, in pair-wise combinations or when combined all together was examined.
CM51 expressing PGKcbh2 produced negligible levels as glucose, whereas low yet discernible levels of glucose were detected in supernatants from CM51 expressing PGKegl1 and PGKbgl1 individually, reflecting the known activities of the individual enzymes against PASC (Fig. 4). Low levels of glucose were also observed when supernatants from strains expressing PGKcbh2 and PGKegl1 were combined, whereas combining supernatants from strains expressing PGKbgl1 and PGKcbh2 produced higher glucose levels. Significantly higher levels of glucose were produced from a combination of supernatants from cells expressing PGKbgl1 and PGKegl1 and the levels of released glucose almost doubled when all three supernatants (PGKbgl1, PGKcbh2, PGKegl1, herein referred to as Cel 1.0), were combined. On the basis of the maximum theoretical yield of glucose that could be released from the amount of PASC used, we deduced that the combination of supernatants from all three cellulase-expressing strains generated a 7·3% yield. Taken together, the data suggest that the minimum combination of BGLI and EGI was sufficient to release glucose from PASC but that the addition of CBHII significantly increases glucose release. We also examined the hydrolysis of other cellulose-based substrates with the recombinant enzymes. Similar yields were obtained using the substrate CMC, whereas crystalline cellulose (Avicel) produced substantially lower yields (data not shown).
To increase the levels of recombinant enzymes, expression of the genes from different promoters as well as co-expression of two endoglucanase genes (egl2 and egl1) were examined. Use of the TEF promoter increased bgl1activity c. 1·7-fold, whereas HXT7 did not significantly alter bgl1 expression (Fig. 5a). Likewise, an c. 1·7-fold increase in cbh2 activity was achieved by switching to the TEF promoter (Fig. 5b).
To improve endoglucanase production, a second endoglucanase gene egl2 was cloned under the control of a PGK promoter (Fig. 2d, PGKegl2). Secondly, both egl1 and egl2 were co-expressed in strain CM51 under the control of PGK (Fig. 2e, PGKegl2-PGKegl1). Enzyme activity of PGKegl2 was 1·7-fold greater than PGKegl1 and furthermore, co-expression of both egl genes (PGKegl2-PGKegl1) increased activity by 2·7-fold, indicating that activity from egl1 and egl2 genes is additive (Fig. 5c). As the highest levels of enzyme activity were achieved by the improved gene cassettes, TEFbgl1, TEFcbh2 and PGKegl2-PGKegl1, (herein referred to as Cel 2.0), strains carrying these plasmids were used in all subsequent analysis.
The hydrolysis of PASC by the Cel 2.0 gene products was examined. The spent medium supernatants of CM51 cells expressing the Cel 2.0 gene cassettes individually were combined and concentrated (1·5-fold). The hydrolysis of PASC by the recombinant enzymes was compared to hydrolysis by a commercial cocktail of T. reesei cellulases. The combined spent medium supernatants released c. 2·5 times more glucose than 0·1 unit ml−1 of the commercial cocktail (Fig. 6a). Taking the dilution factor into account, we estimate that the concentration of cellulase activity in the combined supernatants to be equivalent to 0·17 units ml−1 of the commercial cellulase. The amount of reduced sugars released from PASC hydrolysis was also examined. Interestingly, the commercial cellulase cocktail produced more reduced sugars than the recombinant enzymes (Fig. 6b) indicating the accumulation of reduced intermediates. Analysis of the PASC hydrolysate by Capillary Electrophoresis-Laser Induced Fluorescence (CE-LIF) confirmed that substantially more di-saccharide (cellobiose) accumulated in the samples treated with the commercial cellulases (mono:di ratio; 1 : 1), whereas the recombinant cellulases produced substantially more monosaccharides (mono:di ratio 3 : 1, data not shown).
The optimum conditions for PASC hydrolysis by the recombinant enzymes was determined by varying the enzyme and substrate concentrations as well as the temperature and time of incubation. Concentrating the enzyme led to a concomitant increase in yield of glucose from PASC as well as in the rate of hydrolysis (Fig. 6c and Table 2). Hydrolysis at 30°C, the temperature at which mesophilic yeast fermentations are conducted, was substantially lower than that achieved at the 50°C (Fig. 6c). This requirement for a very high reaction temperature reflects the known activities of the native T. reesei enzymes. The enzyme activity was also influenced by the starting substrate concentration (data not shown) as well as the length of hydrolysis (Table 2). We observed that complete hydrolysis (>95%) is achieved by concentrating the recombinant enzymes 60-fold, however, the rate of hydrolysis is greatest in the first 24 h when a yield of 55·5% was achieved using the same concentration of substrate (Table 2, mg l−1 h−1 g). It should be noted that enzyme production was limited by the fact that cells were cultured in minimal medium to facilitate enzyme assays and higher levels were produced when cells were cultured in rich medium.
|Strains||Glucose g l−1 (±SD)||% yielda(±SD)||Time (h)||Glucose mg l−1 h−1 (±SD)||Glucose yield mg l−1 h−1 g PASC−1 (±SD)|
|Cel 2.0 (1X)||3·32 (0·05)||13·3 (0·19)||120||28 (0·39)||1·11|
|Cel 2.0 (6·7X)||10·36 (1·68)||41·4 (6·73)||72||143·8 (23)||5·76|
|Cel 2.0 (17X)||13·80 (0·14)||55·2 (0·56)||72||192 (1·93)||7·67|
|Cel 2.0 (33X)||15·96 (1·67)||63·9 (6·67)||72||222 (23)||8·87|
|Cel 2.0 (60X)||13·87 (0·14)||55·5 (2·43)||24||578 (25)||23·12|
|Cel 2.0 (60X)||23·76 (1·35)||95·1 (5·39)||240||99 (5·6)||3·96|
With the knowledge that recombinant enzyme activity was better at 50°C than at 30°C, the normal temperature required for mesophilic yeast fermentations, we devised a two-step hydrolysis and fermentation approach to generate alcohol from cellulose as a sole carbohydrate source. CM51 strains, expressing the Cel 2.0 gene cassettes, were cultured in SC-sucrose medium. The spent medium supernatant containing the secreted cellulases were pooled, concentrated and then added to PASC. Following an initial hydrolysis step at 50°C to generate glucose from PASC, a fresh inoculum of CM51 cells expressing the Cel 2.0 gene cassettes or empty vector (pRSH) was added to the cooled PASC mixture and fermentations were carried out at 30°C. Substantial levels of alcohol (12–13 g l−1) were produced after 24 h from cultures containing either the Cel. 2·0 gene cassettes or the empty vector (Fig. 7a). This burst of alcohol production results from glucose generated from PASC in the hydrolysis step. The yield of alcohol corresponds to a 27% conversion of the available glucose in the cellulose substrate. Alcohol production increased over the course of the fermentation in cells expressing the Cel 2.0 gene cassettes reaching a concentration of 16·5 g l−1 after 7 days, indicating that PASC hydrolysis continued during the fermentation. This corresponds to an additional conversion of 6·7% of the available glucose in the cellulose substrate, resulting in a total yield of 33·7%. In the absence of the Cel 2.0 gene cassettes, alcohol levels remained relatively constant.
Recombinant enzyme production was limited by the growth of cells in synthetic minimal medium. To obviating the need to concentrate the enzymes in the spent medium supernatant, we examined the possibility of increasing enzyme production by culturing Cel 2.0-expressing cells in rich YPD medium. We also queried if co-culturing together the three CM51 strains expressing each of the Cel 2.0 gene cassettes rather than culturing them separately would influence enzyme production. The two-step hydrolysis and fermentation of PASC was repeated using the supernatant from the co-culture or supernatants combined after culturing, without prior concentration. Up to 5 g l−1 alcohol was generated after 48 h of fermentation of the PASC hydrolysate from both the co-culture or the mixed spent medium supernatants as a sole carbohydrate source, with slightly lower levels produced at 24 h (Fig. 7b). Thus, the two-step process can be used to generate alcohol from PASC without the need to concentrate the enzymes.
A goal of research into utilizing cellulosic biomass for the production of bioethanol is the generation of micro-organisms capable of degrading cellulose into simple sugars that can be fermented into alcohol at a high yield (Lynd et al. 2005; Olson et al. 2012). Here, we compared the heterologous expression of genes encoding the three major classes of T. reesei cellulases in two different Saccharomyces species. The cellulase genes were individually expressed rather than co-expressed in a single cell as previous studies have indicated that co-expression of recombinant cellulase genes often leads to reduced activity for one or both enzymes (Den Haan et al. 2007b). While several other studies have co-expressed cellulase genes from different fungal species in S. cerevisiae (Fujita et al. 2004), here we choose a single donor organism, T. reesei, to preserve the natural synergistic cellulose-degradation activity of the cellulase enzymes. The expressed genes also preserved the native secretory signal of the proteins as previous studies have demonstrated that the use of nonnative secretory signals can result in the misdirection and/or periplasmic retention of cellulases during the secretory process (Van Rooyen et al. 2005).
Our data show that all three classes of T. reesei cellulases were expressed in both S. cerevisiae and S. pastorianus. However, up to 10-fold higher levels of enzyme activity were observed in S. pastorianus strains for all three recombinant enzymes. The stress-tolerant strains of S. pastorianus did not appear to confer any selective advantage on cellulase production over the parental strain CMBS-33 under the experimental conditions employed in this study. It is possible, however, that under different environmental conditions these strains may have a selective advantage as they produce higher alcohol levels under industrial fermentation conditions (U. Bond, unpublished data). Under the experimental conditions employed here, the S. cerevisiae strain displayed a much faster growth rate and higher final cell density than the S. pastorianus strains, yet in each case cellulase expression was higher in the S. pastorianus strains suggesting that yeast biomass may be counterproductive to heterologous enzyme production.
The differential cellulase activity was not restricted to the haploid S. cerevisiae strain chosen for analysis here as lower levels of enzymatic activity were also detected when bgl1 was expressed in several other S. cerevisiae industrial strains. The increased expression could not be accounted for by differences in plasmid copy number or cell ploidy or by differential promoter expression in the two species. Therefore, differential enzyme activity most likely results from differences in post-translational processing, protein stability and/or export. Previous analysis of the expression of the S. fibuligera bgl1 gene in a number of yeast species (Gurgu et al. 2011) indicated that the genetic background of the host can influence expression of the heterologous gene, however, in that analysis, only 2–3-fold differences in expression were noted. Thus, species choice may be critical to ensure maximum recombinant cellulase production.
Several approaches were taken to optimize for enzyme production. Enzymatic activity was only marginally increased by expression of the genes from several different promoters. This suggests that promoter choice, in particular when expression is driven from constitutive promoters, does not significantly influence enzyme production. Co-expression of additional egl genes did also not significantly improve enzyme activity.
As a starting point to assess the activity of the recombinant enzymes, we used equivalent volumetric ratios of the three classes of cellulases to hydrolyse the cellulose substrate PASC. Under inducing conditions, CBHs account for up to 80% of secreted protein from cultures of T. reesei (Takashima et al. 1998), therefore, it is possible that recombinant cellulase activity could be further increased by altering the ratio of the three enzymes. Like the native cellulases, the recombinant cellulases acted in synergy to hydrolyse cellulose. The rate of enzyme activity was maximum in the first 24 h and at 50°C, indicating that maximum hydrolysis of cellulose-based substrates may best be achieved with consecutive short reaction times and at higher temperatures than normally used for mesophilic fermentations. Interestingly, the recombinant cellulases produced more glucose per gram of PASC than the commercial cellulases as the latter appeared to accumulate higher levels of intermediates such as cellobiose, a known inhibitor of cellulase activity.
As cellulose hydrolysis was significantly reduced at 30°C, the temperature used for most fermentations, we choose a two-step approach for the production of alcohol from cellulose. Hydrolysis of PASC using concentrated supernatants from cultures grown in SC minimal medium produced up to 16·5 g l−1alcohol in subsequent fermentations. However, sufficient enzymes were produced from recombinant strains grown in rich medium and up to 5 g l−1 of alcohol was generated in subsequent PASC fermentations, without the need to concentrate the enzymes.
Several other studies have examined alcohol yields from the hydrolysis of cellulose with recombinant cellulase genes expressed in yeasts (Fujita et al. 2004; Den Haan et al. 2007b; Yamada et al. 2010; Yanase et al. 2010). The most widely employed host for expression of recombinant cellulases has been S. cerevisiae. It is difficult to compare alcohol production from PASC hydrolysis by these recombinant strains due to the variations in experimental parameters, such as starting substrate concentrations, incubation times and the combination of cellulase genes used but in general alcohol yields in all studies are in the order of 1–7 g l−1. One common parameter, however, in these studies, is the use of very high cell densities (200 g l−1wet cell weight or 50 ODs) to achieve these yields. Here, we used at least 2 Logs less cells but achieved greater alcohol yields (13 g l−1) after 24 h of fermentation. Our results also show that the recombinant yeast strain continue to degrade cellulose, albeit at a slower rate, as fermentations proceeded at 30°C, adding to the final yield of alcohol. This rate reflects the slower hydrolysis by enzymes at 30°C. The increased yields achieved here are reflective of the increase in enzyme production in S. pastorianus and the use of a pre-fermentation hydrolysis step.
For the industrial application of cellulose degradation by recombinant cellulases, even higher enzyme yields could be achieved through the use of a fed batch system for propagation of the yeast strains in rich medium. The pursuit of improvements in the efficiency of cellulose hydrolysis, coupled with selection of yeast strains showing increased enzyme production, may contribute to the knowledge required for the on-going development of consolidated bioprocessing of cellulosic biomass and inform further attempts to determine the optimal process operating window.
This research was funded by a grant from the Environmental Protection Agency (2006-PhD-ET-5) to U.B. W.K. is supported by an Innovation Technology grant from Trinity College Dublin, awarded to U.B. We thank Dr. Simone Albrect and Dr. Pauline Rudd, University College Dublin for carrying out the CE-LIF analysis.
No conflict of interest is declared.