Challenges for the Production of Bioethanol from Biomass Using Recombinant Yeasts

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Challenges for the Production of Bioethanol from Biomass Using Recombinant Yeasts William Kricka, James Fitzpatrick and Ursula Bond1 Microbiology Department, School of Genetics and Microbiology, Trinity College Dublin, Dublin, Ireland 1 Corresponding author: E-mail: [email protected]

Contents 1. Introduction 2. Lignocellulosic Biomass Structure 2.1 Pretreatment of Lignocellulose Biomass 3. Saccharomyces Species as Microbial Factories for Conversion of Biomass to Bioethanol 3.1 Engineering Yeasts for Cellulose Metabolism 3.2 Challenges for Cellulose Hydrolysis in Yeasts 4. Utilizing Hemicellulose Pentose Sugars by Yeasts 4.1 Engineering Yeasts for Xylose and Arabinose Metabolism 4.2 Challenges to Pentose Metabolism in Yeast 4.2.1 Transport of Pentose Sugars into Saccharomyces Species 4.2.2 Cofactor Imbalances 4.2.3 Improvement in Metabolic Fluxes in Xylose Assimilation

5. Hexose and Pentose Sugar Coutilization 6. Challenges to Using Real Biomass for Bioethanol Production by Recombinant Yeasts 7. Conclusions References

2 4 4 6 6 13 15 18 19 19 19 20

22 23 25 27

Abstract Lignocellulose biomass, one of the most abundant renewable resources on the planet, is an alternative sustainable energy source for the production of second-generation biofuels. Energy in the form of simple or complex carbohydrates can be extracted from lignocellulose biomass and fermented by microorganisms to produce bioethanol. Despite 40 years of active and cutting-edge research invested into the development of technologies to produce bioethanol from lignocellulosic biomass, the process remains commercially unviable. This review describes the achievements that have been made in generating microorganisms capable of utilizing both simple and complex sugars from lignocellulose biomass and the fermentation of these sugars into ethanol. We also Advances in Applied Microbiology, Volume 92 ISSN 0065-2164 http://dx.doi.org/10.1016/bs.aambs.2015.02.003

© 2015 Elsevier Inc. All rights reserved.

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provide a discussion on the current “roadblocks” standing in the way of making secondgeneration bioethanol a commercially viable alternative to fossil fuels.

1. INTRODUCTION At the start of 2015 as we write this perspective, prices of Brent crude oil have more than halved in the past year from a high of $112 to $46 per barrel. The revolutionary technologies of hydraulic fracturing and horizontal drilling (fracking) of underground shale formations have opened up new reservoirs of oil and gas and are reshaping the discussion on the availability and limits of fossil fuel resources. In the past 5 years, the net import of oil into the USA dropped by 44% and fracking technology promises to make the USA energy self-sufficient by 2030. This current hiatus from “peakoil” fears may have temporarily eased the pressure on identifying and developing alternative sources of renewable and environment friendly energy sources. However, growing concerns regarding the contribution of CO2 emissions from fossil fuels to climate change, coupled with the vagaries of oil production, geopolitical and energy economics, ensure that the development of renewable energy technology will remain a major priority for society in this century and beyond. Biomass, organic material derived mainly from plant matter, has been identified as a possible alternative sustainable energy source for the production of biofuels due to its vast abundance and renewable nature. The global production of plant biomass amounts to approximately 2  1011 Mt per annum, of which between 8 and 20  109 Mt is potentially accessible for processing (Lin & Tanaka, 2006). It is estimated that replacing fossils fuels with biofuels could decrease CO2 emissions by 60e90% (Wang, Wu, & Huo, 2007). Biomass can be derived from agricultural, forestry or industrial waste, or from dedicated energy crops. By far, the most successfully utilized commercial biofuel is bioethanol. Interestingly, the original Ford’s model T car was designed to run on 100% ethyl alcohol and Henry Ford was a major supporter of bioethanol. In 1925, he told a New York Times reporter that ethyl alcohol was “the fuel of the future. It (sic) is going to come from fruit like that sumach out by the road, or from apples, weeds, sawdustdalmost anything. There is fuel in every bit of vegetable matter that can be fermented.” It was only later in the twentieth century that ethanol was replaced with oil-derived products such as petroleum. Worldwide production of bioethanol has increased from 17 to 86  1010 L in the last decade (Lennartsson,

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Erlandsson, & Taherzadeh, 2014) and today bioethanol is widely used in the USA and Brazil, although mainly as a blending component with petroleum. Commercial bioethanol is classified by the source of biomass used for its production. First-generation bioethanol is produced from food crops such as corn or sugarcane. Sugars are easily extracted directly from the crops and used as a carbohydrate source for fermentation and ethanol production. Although commercially successful, the cost of bioethanol production from edible raw material is dictated by the international commodities market. There is also an ongoing ethical debate as to whether arable land should be used for the cultivation of food or fuel crops. This has led to a switch from the use of edible to nonedible lignocellulosic biomass, one of the most abundant renewable resources on the planet, for biofuel production. Bioethanol generated from nonedible lignocellulose biomass is termed second generation and has the benefit of reduced biomass cost and better environmental performance (Farrell et al., 2006; Granda, Zhu, & Holtzapple, 2007; Lennartsson et al., 2014), however generally the starting material is much more complex than that used in first-generation biofuel production, and requires thermochemical and/or biological pretreatments. Thermochemical pretreatments utilizes heat and oxidizing agents to generate three main products, biochar, bio oil, and syngas (synthetic gas; a mixture of CO, CO2, and H2). Biochar can be utilized as a solid fuel for burning, while bio oil and syngas, through further processing, can be converted into transportation fuels (Latif, Zeidan, Nielsen, & Zengler, 2014; Lee & Lavoie, 2013). Alternatively sugars can be extracted from lignocellulose biomass following a chemical pretreatment step and fermented to produce ethanol. Bioethanol can also be generated from sugars released from algal biomass (often referred to as third-generation biofuel). The latter offers several benefits over standard plant lignocellulosic biomass, such as a short harvesting cycle (Ho et al., 2013). Currently, over 90% of the commercial bioethanol is produced from edible biomass (Carere, Sparling, Cicek, & Levin, 2008). Despite 40 years of active and cutting-edge research invested into the generation of lignocellulosic bioethanol, second-generation bioethanol remains commercially unviable. This review focuses on the challenges of developing strategies for the production of bioethanol from lignocellulose biomass. We discuss the achievements that have been made in generating microorganisms capable of utilizing sugars from lignocellulose biomass and the fermentation of these sugars into ethanol. We also provide a discussion on the current “roadblocks” standing in the way of making second-generation bioethanol a commercially viable alternative to fossil fuels.

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2. LIGNOCELLULOSIC BIOMASS STRUCTURE Lignocellulosic biomass is comprised of three main components, cellulose, hemicellulose, and lignin. The percentage of each varies between different types of biomass, although cellulose is generally the most abundant fraction (Saha, 2003). Cellulose, a linear polysaccharide with reducing and nonreducing ends, is composed of up to 15,000 repeating units of the disaccharide cellobiose, which contains two glucose molecules, linked by a b 1e4 glycosidic bond. Intermolecular hydrogen bonding and van der Waals forces allow cellulose chains to stack in parallel to form microfibrils. Microfibrils contain both highly ordered (crystalline regions) and disordered regions (amorphous regions) (Fernandes et al., 2011; Ruel, Nishiyama, & Joseleau, 2012) and are bound together by hemicellulose and lignin to form larger structures called macrofibrils. The second major fraction of lignocellulosic biomass is hemicellulose, a heteropolymer of pentose (xylose and arabinose) and hexose (glucose, mannose, and galactose) sugars. Generally hemicellulose is classified into four groups (xylan, xyloglucan, mannan, and glucomannan) based on the sugar moieties making up the polysaccharide. Xylan, a major component of hardwood trees, is a homopolymer of xylose linked by b 1e4 glycosidic bonds, while xyloglucan contains a b 1e4-linked glucose backbone with mainly b 1e6-linked xylose side chains although other sugar moieties can also be present as side chains. Xyloglucans have an important role in linking adjacent cellulose microfibrils, via hydrogen bonding between xyloglucans and cellulose. The final types of hemicellulose mannans and glucomannans, the principal components of softwoods, are composed of mannose homopolymers and heterpolymers of mannose and glucose respectively. Lignin, the third major component of lignocellulosic biomass is responsible for the structural rigidity of plants (Galbe & Zacchi, 2007). It is a heteropolymer comprised of p-hydroxyphenyl, guaiacyl, and syringyl monolignol units that form a complex branched network around cellulose microfibrils (Phitsuwan, Laohakunjit, Kerdchoechuen, Kyu, & Ratanakhanokchai, 2013; Weng, Li, Bonawitz, & Chapple, 2008).

2.1 Pretreatment of Lignocellulose Biomass Pretreatment is the first and most important step in conversion of lignocellulose to bioethanol. The pretreatment step is designed to effectively increase cellulose accessibility, generally through the alteration of the physical

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structure of cellulose and the solubilization of the hemicellulose and lignin fractions of biomass (Agbor, Cicek, Sparling, Berlin, & Levin, 2011). Pretreatments can be physical, chemical, or biological or combinations of the three. Any pretreatment method should ideally fulfill certain requirements such as high recovery levels of the desired carbohydrates, low levels of degradation by-products and inhibitors, low operation costs and be applicable to an industrial setting (Galbe & Zacchi, 2007). The first step in any pretreatment is the physical size reduction of the lignocellulose substrate to increase the accessible surface area and decrease the crystallinity of the substrate. Size reduction is typically performed by chipping, grinding, and milling. The average size of the lignocellulose material should be 0.2e2 mm (Chiaramonti et al., 2012). The most environment friendly and cost-effective pretreatment methods involve steam explosion and hot water extraction. Steam explosion significantly solubilizes the hemicellulose fraction, allowing for increased cellulose accessibility, however, lignin extraction is limited by this method (Liu et al., 2013). Hemicellulose is extracted by steam explosion through the action of naturally occurring acetic and other acids released from biomass during treatment (Chiaramonti et al., 2012; Mosier et al., 2005). Steam explosion also presents disadvantages such as the release of inhibitors due to the conversion of xylan to volatile organic compounds (Chiaramonti et al., 2012). The yield of cellulose and hemicellulose can be improved by the addition of sulfur dioxide to the steam explosion. Hemicellulose can also be extracted by treatment with dilute acid (DA) solutions at high temperatures, which hydrolyze the polysaccharides into monomers of xylose, glucose, arabinose, and galactose. The release of sugars using DA pretreatment is dependent on factors such as acid concentration, pretreatment time, solid loading, and temperature (Ahmed et al., 2013; Kim, Kreke, & Ladisch, 2013; Lim & Lee, 2013; Sindhu et al., 2011). Pretreatment of biomass by ammonia fiber expansion (AFEX) significantly alters the lignin structure within lignocellulose. The anhydrous ammonia cleaves the ligninecarbohydrate linkage, increasing accessibility to cellulose (Mosier et al., 2005). Ionic liquid (IL) pretreatments interact with the hydrogen-bonding network in cellulose, disrupting the 3-D structure leading to amorphous cellulose generation (Moulthrop, Swatloski, Moyna, & Rogers, 2005). A comparison of three of the pretreatments (DA, IL, and AFEX) reveals that significant levels of lignin (89.9%) and xylan (23.4%) are extracted from biomass by IL pretreatment, while much higher levels of xylan (87%) but significantly lower levels of lignin (2.8%) are extracted

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by DA, and finally very low levels of xylan (0.8%) are extracted by AFEX (Gao et al., 2014). When combined with biological treatments, the greatest release of glucose was achieved by IL pretreatment (Gao et al., 2014). Additionally, IL pretreatments appear to produce fewer inhibitory compounds than other pretreatment methods (Galbe & Zacchi, 2007).

3. SACCHAROMYCES SPECIES AS MICROBIAL FACTORIES FOR CONVERSION OF BIOMASS TO BIOETHANOL By far the most successful strategy for the production of bioethanol from biomass has been the metabolic engineering of natural ethanologenic yeast species to ferment sugars extracted from lignocellulose. While naturally capable of fermenting sugars such as glucose, yeasts do not possess the enzymes necessary for the hydrolysis of cellulose nor for the utilization of pentose sugars such as xylose and arabinose. Over the past 25 years great strides have been made to introduce the necessary genes into yeasts to allow for this metabolism. The culmination of these efforts is outlined in Tables 1e3 and is discussed below.

3.1 Engineering Yeasts for Cellulose Metabolism The extraction of fermentable sugars from cellulose requires both chemical and physical pretreatments and subsequent enzymatic hydrolysis by the synergistic action of three major classes of cellulases, namely endoglucanases (EG), exoglucanases or cellobiohydrolases (CBH), and b-glucosidases (BGL) (Figure 1). Endoglucanases cleave the cellulose backbone randomly at amorphous sites along the cellulose fiber exposed by the pretreatments, leading to a rapid decrease in the degree of polymerization of the fiber and exposing new chain ends. CBH act processively on reducing and nonreducing chain ends to release mainly the disaccharide cellobiose. The processive action of CBH enzymes is intrinsically slow and a critical bottleneck in cellulose hydrolysis (Horn, Vaaje-Kolstad, Westereng, & Eijsink, 2012; Ilmen et al., 2011). Finally, b-glucosidases hydrolyze the b-1,4 glycosidic bond of cellobiose and cello-oligosaccharides to release glucose. Cellulase enzymes have been identified in a diverse range of fungal and bacterial species and are either defined as complexed or noncomplexed (Figure 1). Complexed cellulase enzymes, often referred to as cellulosomes, are tethered to the cell wall while organisms with noncomplexed cellulase systems

Saccharomyces cerevisiae EBY100

S. cerevisiae MT8-1/ cocdBEC3 S. cerevisiae MT8-1

Saccharomyces pastorianus CM-51

S. cerevisiae Y294

Tethered

Saccharomycopsis fibuligera BGLI

Secreted

10

2.12

0.21

Baek et al. (2012)

Cellulosome 10

1.80

0.18

Kim, Baek, Lee, and Hahn (2013)

20

7.60

0.38

Yamada et al. (2011)

10

2.10

0.21

Yanase et al. (2010)

25

5.00

0.20

Fitzpatrick et al. (2014)

10

1.00

0.10

Den Haan, Rose, et al. (2007)

Tethered Tethered

Cellulosome Tethered Tethered Tethered Tethered Tethered Tethered Tethered Secreted Secreted Secreted Secreted Secreted

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(Continued)

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S. cerevisiae EBY100

Thermoascus aurantiacus EGI Trichoderma reesei CBHII Aspergillus aculeatus BGLI Clostridium thermocellum CelA (EG) T. reesei CBHII A. aculeatus BGLI T. reesei EGII T. reesei CBHII A. aculeatus BGLI T. reesei EGII T. reesei CBHII A. aculeatus BGLI T. reesei EGI T. reesei EGII T. reesei CBHII T. reesei BGLI T. reesei EGI

References

Challenges for the Production of Bioethanol from Biomass Using Recombinant Yeasts

Table 1 Ethanol production from cellulose hydrolysis and fermentation using recombinant Saccharomyces sp. Tethered or Host Cellulase enzyme Secreted PASC (g/L) Ethanol (g/L) Yield (g/g)

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Table 1 Ethanol production from cellulose hydrolysis and fermentation using recombinant Saccharomyces sp.dcont'd Tethered or Host Cellulase enzyme Secreted PASC (g/L) Ethanol (g/L) Yield (g/g) References

S. cerevisiae BY4742

S. cerevisiae MT8-1

S. cerevisiae EBY100

C. thermocellum CelA (EG) T. reesei CBHII T. aurantiacus BGLI C. thermocellum CelA (EG) T. reesei CBHII T. aurantiacus BGLI T. reesei EGII T. reesei CBHII A. aculeatus BGLI T. reesei EGII T. reesei CBHII A. aculeatus BGLI

Cellulosome 10

1.25

0.12

Goyal et al. (2011)

Cellulosome Cellulosome Secreted 10

0.43

0.04

Goyal et al. (2011)

2.90

0.29

Fujita et al. (2004)

1.80

0.18

Wen et al. (2010)

Secreted Secreted Tethered 10 Tethered Tethered Cellulosome 10 Cellulosome Cellulosome

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PASC, phosphoric acid swollen cellulose; CBH, cellobiohydrolases; BGL, b-glucosidases; EG, endoglucanases.

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S. cerevisiae BY4742

Pichia stipitis

P. stipitis S. cerevisiae e

27.50

0.40

Neurospora crassa Candida parapsilosis P. stipitis Candida guilliermondii P. stipitis

P. P. P. P. P.

e e þ þ e

27.50 7.50 12.30 6.10 12.60

0.40 0.23 0.24 0.33 0.35

S. cerevisiae YPH499XU Scheffersomyces stipitis S. cerevisiae 1-dX-70 S. stipitis S. cerevisiae TMB 3057 P. stipitis

S. stipitis S. cerevisiae e S. stipitis S. cerevisiae e P. stipitis S. cerevisiae þ

e e 13.30

0.24 0.36 0.33

S. cerevisiae TMB 3400 P. stipitis S. cerevisiae BY4741X S. stipitis S. cerevisiae W303-1ATe S. stipitis

P. stipitis S. cerevisiae e S. stipitis S. cerevisiae e S. stipitis S. cerevisiae e

12.10 25.40 12.20

0.34 0.28 0.27

S. cerevisiae sun048T S. cerevisiae YRH388

S. stipitis P. stipitis

S. stipitis S. cerevisiae e P. stipitis S. cerevisiae e

16.60 5.60

0.34 0.23

S. cerevisiae YRH396

P. stipitis

P. stipitis S. cerevisiae e

7.80

0.27

stipitis stipitis stipitis stipitis stipitis

Xiong et al. (2011) Ma et al. (2012) Matsushika, Inoue, Murakami, Takimura, and Sawayama (2009) Kato et al. (2013) Karhumaa, Garcia Sanchez, et al. (2007) Fujitomi et al. (2012) Ismail, Sakamoto, Hatanaka, Hasunuma, and Kondo (2013) Hector, Dien, Cotta, and Qureshi (2011)

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PPP: þ indicates the overexpression of pentose phosphate pathway genes.

S. cerevisiae S. cerevisiae S. cerevisiae P. stipitis S. cerevisiae

Bera et al. (2011)

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Saccharomyces cerevisiae 424A S. cerevisiae 424A S. cerevisiae 424A S. cerevisiae F106X S. cerevisiae YY5A S. cerevisiae MA-N4

Challenges for the Production of Bioethanol from Biomass Using Recombinant Yeasts

Table 2 Ethanol production from xylose using recombinant Saccharomyces sp. strains expressing the xylose reductase (XR)/xylitol dehydrogenase (XDH) pathway Enzyme Theoretical Host XR XDH XK PPP Ethanol (g/L) yield (g/g) References

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Table 3 Ethanol production from xylose using recombinant Saccharomyces sp. strains expressing the xylose isomerase (XI) pathway Enzyme Ethanol Theoretical Host XI XK PPP (g/L) yield (g/g) References

7.30

0.43

e e S. cerevisiae e

e 4.10

0.23 0.35

S. S. S. S. S. S. S. S. S.

P. ruminicola Piromyces sp. E2 Clostridium phytofermentans C. phytofermentans Orpinomyces sp. Piromyces sp. E2 Piromyces sp. E2 Thermus thermophilus C. phytofermentans

S. S. S. S. S. S. S. e e

e e e e e þ þ e e

13.60 5.30 e e 6.93 17.50 7.00 1.30 8.00

0.42 0.35 0.23 0.46 0.32 0.45 0.42 0.13 0.43

S. cerevisiae RWB202

Piromyces sp. E2

e

e

8.05

0.42

S. cerevisiae INVSc1

Orpinomyces sp.

S. cerevisiae e

4.06

0.39

cerevisiae cerevisiae cerevisiae cerevisiae cerevisiae cerevisiae cerevisiae cerevisiae cerevisiae

YRH1114 Y7092 HDY.GUF5 GS1.11-26 MT8-1 CIBTS0735 BY4741-S2A3K H158 BarraGrande

PPP: þ indicates the overexpression of pentose phosphate pathway genes.

cerevisiae cerevisiae cerevisiae cerevisiae cerevisiae cerevisiae cerevisiae

Karhumaa, Garcia Snachez, et al. (2007) De Figueiredo Vilela et al. (2013) Hector, Dien, Cotta, and Mertens (2013)

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S. cerevisiae þ

Usher et al. (2011) Demeke et al. (2013) Tanino et al. (2010) Diao et al. (2013) Lee, Jellison, and Alper (2012) Walfridsson et al. (1996) Brat, Boles, and Wiedemann (2009) Kuyper, Winkler, Van Dijken, and Pronk (2004) Madhavan et al. (2009)

William Kricka et al.

Saccharomyces cerevisiae TMB Piromyces sp. E2 3066 S. cerevisiae BY4741 Burkholderia cenocepacia S. cerevisiae YRH631 Prevotella ruminicola

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Figure 1 Expression of recombinant cellulase systems in yeasts. The concerted action of cellulases, endoglucanases (EG), cellobiohydrolyases (CBH), and b-glucosidases (BGL) is required for hydrolysis of cellulose. Due to the polymeric nature of cellulose, hydrolysis must occur external to the yeast cell. Cellulases are secreted from the cell and can be assembled into cellulosomes, be tethered to the cell surface through cell wall anchors or secreted into the surrounding medium. Adapted from Kricka et al. (2014).

produce enzymes that are secreted from the cell into the surrounding environment (Kricka, Fitzpatrick, & Bond, 2014; Peterson & Nevalainen, 2012). For the production of recombinant yeasts capable of hydrolyzing and metabolizing cellulose from biomass, the vast majority of studies have focused on the coexpression of genes encoding at least one enzyme from each of the three major classes of cellulases. Genes from several fungal and bacterial species, including Aspergillus niger, Clostridium thermocellum, Aspergillus aculeatus, Saccharomycopsis fibuligera, Thermoascus aurantiacus, and Trichoderma reesei have been heterologously expressed in yeasts. Generally, the cellulase machinery has been reconstructed by combining cellulase genes from different species, rather than by using genes from a single organism and both complexed and noncomplexed systems have been reconstituted (Table 1). A comparison of the two systems indicated that the complexed

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cellulosome system was more efficient, producing up to threefold more ethanol than the secreted enzyme system (Goyal, Tsai, Madan, Dasilva, & Chen, 2011; Wen, Sun, & Zhao, 2010), however, a comparison of all reconstituted cellulase systems indicates that in general, similar levels of ethanol are produced by both complexed and noncomplexed systems (Table 1). A comparison of reconstituted cellulase systems is hindered by differences in the choice of genes used for expression, differences in cell growth conditions used in each study, and by variations in the units used to express ethanol yields and enzyme activities. The activities of recombinant cellulase enzymes have been compared in an effort to identify the enzymes with maximum activity. There appears to be no consensus donor species that out performs others. A comparison of BGL1 genes expressed in Saccharomyces cerevisiae indicated that the gene isolated from S. fibuligera displayed the highest activity (Tang et al., 2013; Van Rooyen, Hahn-Hagerdal, La Grange, & Van Zyl, 2005). A similar comparison of recombinant CBH demonstrated that CBHII displayed higher specific activities than CBHI, with the highest activities obtained from a CBHII gene from Chrysosporium lucknowense (Den Haan, Mcbride, La Grange, Lynd, & Van Zyl, 2007; Ilmen et al., 2011). The comparison of recombinant endoglucanase activity has mainly been limited to genes from T. reesei (Du Plessis, Rose, & Van Zyl, 2010), although endoglucanase enzymes from various protists isolated from termites have also been expressed in S. cerevisiae (Todaka et al., 2011). The ratio of the three cellulases also affects cellulose hydrolysis. In T. reesei, CBH represent up to 80% of the secreted cellulases while endoglucanase and b-glucosidase make up the remainder. To examine the effects of different ratios of the three cellulases on cellulose hydrolysis by recombinant yeasts, different copy numbers of the three different cellulase genes were randomly integrated into the multiloci transposable element (Ty delta) sites in S. cerevisiae (Yamada et al., 2010). The highest yield of glucose from the pure cellulose substrate, phosphoric acid swollen cellulose (PASC), was obtained in a strain containing EGL1, BGL1, and CBH2 genes in a ratio of 16:2:6. Hydrolysis was slightly lower (1.4-fold) when the ratio was 5:9: 6 and the lowest activity was observed when the genes were present in single copies in a 1:1:1 ratio. Many different promoters have been used to increase recombinant gene expression (Nacken, Achstetter, & Degryse, 1996; Sun et al., 2012). Constitutive promoters such as the PGK (Yamada et al., 2011), TEF1 (Fitzpatrick, Kricka, James, & Bond, 2014), SED1 (Inokuma, Hasunuma, & Kondo, 2014), and ENO (Den Haan, Rose, Lynd, & Van Zyl, 2007) have been

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utilized for continuous expression of cellulase genes. Inducible promoters such as GAL1/10 have also been used (Jeon et al., 2009). These promoters drive vastly higher gene expression, however, they are repressed by glucose, the end product of cellulose hydrolysis and require the addition of an expensive starting substrate (galactose) to the medium. Efficient secretion is required for high cellulase enzyme activity. In several studies, the native secretory signals of cellulases have been exchanged for either S. cerevisiae secretory signals (mating factor-a) (Zhu, Yao, & Wang, 2010) or the xyn2 secretory signal from T. reesei (Den Haan, Rose, et al., 2007; Van Rooyen et al., 2005). Comparison of the activity of BGL1 (S. fibuligera), secreted from the cell by various secretory signals, including its native secretory signal, revealed little difference between the efficiencies of the secretory signals (Tang et al., 2013). Modifications of disulfide bond formation, glycosylation, protein folding and trafficking have been shown to increase protein secretion and activity of cellulase enzymes, however, such modifications only resulted in modest (1.3- to 6-fold) increases (Xu et al., 2014). Recent research has suggested that other nonhydrolytic auxiliary proteins play a crucial role in enzymatic degradation of cellulose (Harris, Xu, Kreel, Kang, & Fukuyama, 2014). The copper-dependent monooxygenases such as the bacterial CBM33 and the fungal AA9 (formerly GH61) disrupt the packing of crystalline cellulose, thereby increasing accessibility for the hydrolytic enzymes (Horn et al., 2012). Elastin-like proteins, such as swollenin, also contribute to the hydrolysis of cellulose by increasing access for the cellulase enzymes to the cellulose chains ends. Therefore, in order to increase the efficiency of enzymatic hydrolysis of cellulose by recombinant organisms, the heterologous expression of these nonhydrolytic “enhancer” enzymes might be considered.

3.2 Challenges for Cellulose Hydrolysis in Yeasts Presently, first-generation bioethanol is commercially produced by a process referred to as separate hydrolysis and fermentation in which the cellulose hydrolysis and the fermentation are performed in separate reaction vessels, with cellulases supplied ex vivo (Kricka et al., 2014). The ultimate goal of the industry is to produce a single microorganism capable of both producing biomass-hydrolyzing enzymes and fermenting the resultant released sugars to ethanol, a process referred to as consolidated bioprocessing (CBP). There is an inherent problem in developing yeast strains for the metabolism of cellulose as a sole carbohydrate source. As mentioned above,

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cellulose is a large polysaccharide that must be hydrolyzed in the extracellular medium to release glucose, which is then transported into the cell as a fermentable sugar. This creates a classic chicken and egg conundrum as in order to produce the cellulases for cellulose hydrolysis, cell growth is required, which is limited by the lack of fermentable sugars (glucose) at the start of the fermentation. This problem might be alleviated by the addition of glucose in low quantities into the medium at the start of fermentation; however, cellulase genes are repressed by glucose, thus compounding the problem. The use of complexed cellulosomes tethered to the yeast cell surface provides a partial solution to this problem yet even with these systems, cellulose hydrolysis is not substantially improved when compared with the secreted enzyme systems (Table 1) (Baek, Kim, Lee, Lee, & Hahn, 2012; Fitzpatrick et al., 2014; Fujita, Ito, Ueda, Fukuda, & Kondo, 2004; Nakatani, Yamada, Ogino, & Kondo, 2013; Yamada et al., 2011). The choice of host for heterologous expression of cellulase genes for CBP is an important factor for consideration. The most commonly used host is the bakers yeast, S. cerevisiae. Saccharomyces cerevisiae is generally regarded as safe and has been extensively characterized biochemically and genetically. It is a suitable host for the high-level production and secretion of recombinant proteins in their correctly folded forms. Saccharomyces cerevisiae has several advantages over other ethanologenic microbes, the most important from an industrial point of view being its high fermentative capacity and robust tolerance to ethanol. Saccharomyces cerevisiae strains can produce up to 200 g/L ethanol in classic industrial fermentations (Lin & Tanaka, 2006). As a heterologous host, S. cerevisiae can produce recombinant protein levels of up to 1 g/L (Ilmen et al., 2011). However, this pales in comparison to the high protein levels that natural cellulolytic organisms are known to secrete. Indeed the hypersecretory T. reesei mutant RUTC30 has been shown to secrete between 20 and 100 g/L of cellulase enzymes (Lambertz et al., 2014). Interestingly, significantly higher recombinant protein levels and enzyme activities can be obtained by expressing cellulases in the brewer’s yeasts Saccharomyces pastorianus (Fitzpatrick et al., 2014). The increased enzymatic activity in S. pastorianus appears to result from increased protein stability in this tetraploid species. In general, increased enzymatic activities appear to correlate with the DNA content of yeast cells with higher levels observed in diploid compared with haploid cells, and even greater levels produced in tetraploid cells (Fitzpatrick et al., 2014). Other robust S. cerevisiae industrial strains such as Ethanol Red, which has been reported to produce up to 250 g/L ethanol, are also being

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used for heterologous gene expression for bioethanol production (Demeke et al., 2013). This strain has desired qualities such as high fermentative capacity and is tolerant to dehydration. One major problem associated with cellulose hydrolysis by recombinant yeasts relates to the optimum temperature for cellulase activities. All three classes of cellulases show optimum activities between 50 and 70  C (Johnson, Sakajoh, Halliwell, Madia, & Demain, 1982; Kupski, Pagnussatt, Buffon, & Furlong, 2014) and activities are substantially reduced at the optimum growth temperatures (30  C) of mesophilic yeasts. Several studies have focused on identifying cold-adapted cellulase enzymes (Ueda et al., 2014; Yang & Dang, 2011) to tackle this problem. Rational protein design, adaptive or targeted mutagenic approaches might be used in the future to increase the activities of these enzymes at the lower temperatures.

4. UTILIZING HEMICELLULOSE PENTOSE SUGARS BY YEASTS Pretreatment of lignocellulose biomass releases both hexose and pentose sugars. Glucose is by far the most abundant sugar in lignocellulose biomass accounting for approximately 30e50% of the biomass dry weight. Mannose and galactose are present in much lower amounts of approximately 1e3% and 1e2% respectively. The xylose content of lignocellulose biomass varies from source to source and on average accounts for approximately 18% of the dry weight while arabinose accounts for approximately 3% (Madhavan, Srivastava, Kondo, & Bisaria, 2012). The hexose sugars, glucose, mannose, and galactose can be readily fermented to ethanol by most yeasts species, on the other hand, only a small number of yeast species that naturally metabolize xylose or arabinose have been identified. These species are often associated directly with lignocellulose or indirectly through association with another organism. Xylose-utilizing yeast species (Candida tropicalis, Candida parapsilosis, Geotrichum sp., Candida mengyuniae, Sporopachydermia lactativora, Trichosporon asahii, and Spathaspora passalidarum) have been isolated from sources such as the wood-boring beetle Odontotaenius disjunctus, the wood roach Cryptocercus sp., and from buffalo feces (Hou, 2012; Lorliam, Akaracharanya, Suzuki, Ohkuma, & Tanasupawat, 2013; Urbina, Frank, & Blackwell, 2013). Other xylose-metabolizing yeasts include isolates of Pachysolen tannophilus and the well-characterized Pichia stipitis. Arabinose is naturally metabolized by several fungal species including Candida and Pichia sp., Arxula adeninivorans, Debaryomyces hansenii, and

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Figure 2 Metabolism of pentose sugars by engineered yeast. (a) Genes required for the metabolism of xylose and arabinose expressed in Saccharomyces cerevisiae. The pentose sugars are transported into the cell by native hexose transporters. Both sugars are metabolized by either isomerase or cofactor requiring oxidoreductase pathways. The endogenous xylulose kinase in S. cerevisiae, converts the product of these pathways, xylulose, to xylulose-5-phosphate, which then enters the pentose phosphate pathway (PPP) and is directed toward ethanol production. (The key enzymes in the pathways are shown: XRdxylose reductase, XDHdxylitol dehydrogenase, XKSdxylulose kinase, XIdxylose isomerase, ARdaldose reductase, LADdL-arabitol

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:

Kluveromyces marxianus and several of the genes required for uptake and metabolism have been characterized in these species (Madhavan et al., 2012). Despite their potential for pentose sugar metabolism, these species have not been advanced for development as cell factories for bioethanol production due to low ethanol yields, low tolerance to high ethanol concentrations, and limited genetic characterization, however, several studies have focused on improving ethanol yields in P. stipitis through genome shuffling (Bajwa, Pinel, Martin, Trevors, & Lee, 2010; Shi et al., 2014) or mutagenesis (Grabek-Lejko, Ryabova, Oklejewicz, Voronovsky, & Sibirny, 2006; Watanabe, Watanabe, Yamamoto, Ando, & Nakamura, 2011). Despite being the most well-characterized species for fermentation and ethanol production from hexose sugars, in general, Saccharomyces species lack several essential enzymes required for the metabolism of the pentose sugars xylose and arabinose. Natural Saccharomyces isolates have been identified that grow slowly in medium containing xylose as a sole carbohydrate source (Wenger, Schwartz, & Sherlock, 2010) and several genes encoding putative xylose-utilizing enzymes have been identified in the genomes of Saccharomyces species, however, xylose metabolism by Saccharomyces species is inefficient and unsuitable for industrial processes and most likely serves as a fail-safe mechanism for survival under conditions when nutrients are limiting. Much of the effort in developing a cell factory for the conversion of biomass to bioethanol has focused on developing recombinant yeast strains capable of cofermenting hexose and pentose sugars through the heterologous expression of genes required for xylose and arabinose metabolism. In nature, xylose can be metabolized by microorganisms in two different pathways (Figure 2). In most bacterial species, some fungi (Piromyces, Orpinomyces) and plants, xylose is converted to xylulose in a one-step process by the action of xylose isomerase (Figure 2(a)). The second pathway, utilized by most fungi, requires sequential oxidoreduction reactions by the enzymes xylose reductase (XR) and xylitol dehydrogenase (XDH), which require the cofactors NADPH and NADþ respectively in the forward reactions. The product of both pathways, xylulose, is then phosphorylated by the dehydrogenase, ALXdL-xylulose reductase, RKdribulose kinase, R5PE-ribulose-5-Phosphate epimerase, AIdL-arabinose isomerase). (b) Xylulose-5-phosphate produced from xylose and arabinose is metabolized by the PPP to produce fructose-6-phosphate and glyceraldehyde-3-phosphate (underlined). These metabolites enter the glycolytic and fermentative pathways to produce ethanol. (The key enzymes in the pathways are shown: RPE1: D-ribulose-5-phosphate 3-epimerase, RKL1: ribose-5-phosphate ketolisomerase, TAL: transaldolase, TKL: transketolase).

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enzyme xylulose kinase to generate xylulose-5-P, which is metabolized via the pentose phosphate pathway (PPP; Figure 2(b)). Intermediate metabolites such as glyceraldehyde-3-P and fructose-5-P generated by the PPP are directed to the glycolytic and fermentation pathways to yield ethanol (Figure 2(b)). Arabinose can also be metabolized by two distinct pathways. In the first pathway, mainly identified in bacterial species, the enzyme arabinose isomerase (AI) converts arabinose to ribulose, which is then phosphorylated by ribulose kinase to yield ribulose-5-P. Finally ribulose-5-P epimerase converts ribulose-5-P to xylulose-5-P. Most fungal species utilize a second pathway involving sequential redox reactions involving NADPHdependent reductases and NADþ-dependent dehydrogenases to convert arabinose to xylulose. Following phosphorylation by the same xylulose kinase as required for xylose metabolism, xylulose generated from arabinose is fed into the PPP and subsequently into the glycolytic and fermentation pathways (Figure 2(a) and (b)).

4.1 Engineering Yeasts for Xylose and Arabinose Metabolism Since xylose represents the most abundant pentose sugar in hemicellulose, it follows that most efforts have focused engineering S. cerevisiae strains to express the xylose metabolic pathways. The genes required for both the xylose isomerase (XI) and XR/XDH pathways have been introduced into strains of S. cerevisiae (Tables 2 and 3). Genes encoding XR and XDH enzymes are most commonly sourced from P. stipitis, while a more diverse range of XI genes from different organisms have been examined (Tables 2 and 3). While S. cerevisiae contains a functional copy of the gene XKS encoding xylulose kinase, conversion of xylulose to xylulose-5-P appears to be a rate-limiting step in the utilization of xylose in S. cerevisiae, therefore to increase the flux through the pathway, most studies have additionally overexpressed the endogenous XKS gene (Tables 2 and 3) allowing for increased ethanol production and reduced by-product formation (Johansson, Christensson, Hobley, & Hahn-Hagerdal, 2001; Toivari, Aristidou, Ruohonen, & Penttila, 2001). Genes required for arabinose utilization via the two pathways described in Figure 2 have also been introduced into yeasts with some limited success (Bera, Sedlak, Khan, & Ho, 2010; Bettiga, Bengtsson, Hahn-Hagerdal, & Gorwa-Grauslund, 2009; Wisselink et al., 2007). Ethanol production from strains expressing the bacterial arabinose pathway appears to be more successful than from the fungal pathway. The lower yields obtained in yeast expressing the fungal arabinose-utilizing genes have been attributed

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to redox imbalances in the cell due the accumulation of reduced cofactors under anaerobic conditions. Saccharomyces cerevisiae strains expressing both arabinose and xylose metabolic pathways have also been generated (Demeke et al., 2013; Karhumaa, Wiedemann, Hahn-Hagerdal, Boles, & GorwaGrauslund, 2006; Wisselink, Toirkens, Wu, Pronk, & Van Maris, 2009).

4.2 Challenges to Pentose Metabolism in Yeast 4.2.1 Transport of Pentose Sugars into Saccharomyces Species Native xylose-metabolizing yeasts possess both xylose-specific and nonspecific transport systems. The xylose-specific transporters utilize a proton symport system while nonspecific transport is mediated by facilitated diffusion through other low-affinity sugar transporters (Leandro, Goncalves, & Spencer-Martins, 2006). While lacking xylose-specific transporters, Saccharomyces species can uptake xylose via glucose transporters encoded by the HXT gene family (Hamacher, Becker, Gardonyi, Hahn-Hagerdal, & Boles, 2002), however, in mixed sugar medium, glucose is preferentially transported into cell due to a 100-fold lower affinity of xylose for the transporters. This problem has been addressed by the introduction of genes encoding xylose transporters into S. cerevisiae. The overexpression of genes such as GXF1/GXS1 from Candida intermedia, Trxlt1 from T. reesei, SUT1 from P. stipitis, and At5g17010 from Arabidopsis thaliana have met with mixed results, with some improvements of xylose uptake reported in mixed sugar fermentations (Leandro et al., 2006; Runquist, Hahn-Hagerdal, & Radstrom, 2010; Saloheimo et al., 2007). Saccharomyces cerevisiae also lacks arabinose-specific transporters, however, arabinose can be transported in the cells via the galactose transporter, Gal2p, albeit at a slow rate. Sugar transporters that mediate arabinose uptake such as AraT from Scheffersomyces stipitis, Stp2 from the plant A. thaliana, and LAT1 and LAT2 from Ambrosiozyma monospora facilitate increased arabinose uptake when overexpressed in S. cerevisiae (Subtil & Boles, 2011; Verho, Penttila, & Richard, 2011). These transporters appear to be specific for arabinose and did not facilitate glucose or other hexose sugar uptake (Wang et al., 2013). The overexpression of various native hexose transporters (HXT1, HXT7, HXT13, and GAL1) can also increase xylose transport into the cell, however, ethanol production was not significantly increased (Tanino et al., 2012; Young, Poucher, Comer, Bailey, & Alper, 2011). 4.2.2 Cofactor Imbalances Redox imbalances arise when yeasts are grown under strict anaerobic conditions due to the absence of transhydrogenase activity, which is used for the

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interconversion of NADPH and NADH. This is particularly problematic in xylose metabolism carried out under anaerobic conditions, as NADPþ generated in the conversion of xylose to xylitol cannot be used for the NADþ-linked oxidation of xylitol to xylulose (Jeffries & Jin, 2004; Van Dijken, Van Den Bosch, Hermans, De Miranda, & Scheffers, 1986). The net effect of these redox imbalances is the accumulation of xylitol in the cell. Altering cofactor specificity of the oxidoreductase enzymes is one possible approach to reduce the redox imbalance and to increase ethanol yields (Ghosh, Zhao, & Price, 2011). The alteration of XDH cofactor specificity from NADþ to NADPþ has been reported to increase ethanol yields and decrease xylitol excretion, although ethanol levels were not increased greatly (Matsushika et al., 2008; Watanabe et al., 2007b) and no distinct correlation between XDH (NADPþ) activity and alcohol production was observed (Watanabe et al., 2007b). The alteration of XR cofactor specificity from NADPH to NADH resulted in increased ethanol production and reduced xylitol formation (Bengtsson, Hahn-H€agerdal, & GorwaGrauslund, 2009; Krahulec, Klimacek, & Nidetzky, 2012; Petschacher & Nidetzky, 2008; Watanabe et al., 2007a; Xiong, Chen, & Barford, 2011). While expression of the single enzyme, XI avoids redox imbalances, a comparison of the XI and XR/XDH expression systems revealed that the XR/ XDH pathway produced significantly higher ethanol levels from xylose than the XI pathway under semianaerobic fermentation conditions (Karhumaa, Garcia Sanchez, Hahn-Hagerdal, & Gorwa-Grauslund, 2007). This appears to be a consequence of the low activity of recombinant XI coupled with its inhibition by xylitol, generated in the cell by reduction of xylose by endogenous reductases such as GRE3. The additional overexpression of the endogenous gene XKS, which encodes for xylulose kinase, partially alleviates the deficiencies in the early steps of xylose metabolism by providing a forward flux “pull” of metabolites in the direction of the PPP. In general, redox imbalances can be avoided by growth of yeasts under semianaerobic conditions, which are sufficient for ethanol production. 4.2.3 Improvement in Metabolic Fluxes in Xylose Assimilation To improve metabolic fluxes toward ethanol production in S. cerevisiae, manipulation of specific enzymes in the PPP and in other metabolic pathways has been extensively examined. Deletions and overexpression of genes encoding key PPP enzymes have shown mixed effects on xylose metabolism (Matsushika, Inoue, Kodaki, & Sawayama, 2009). Overexpression of genes

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encoding the PPP enzymes such as transaldolase (TAL1), transketolase (TKL1), ribose-5-phosphate ketol-isomerase (RKL1), and D-ribulose-5phosphate 3-epimerase (RPE1) (Figure 2(a) and (b)) increased growth rates on xylose, although ethanol levels were only increased fractionally (Bera, Ho, Khan, & Sedlak, 2011; Karhumaa, Fromanger, Hahn-Hagerdal, & Gorwa-Grauslund, 2007). Deletion of either TAL1 or TKL1 led to decreased growth and ethanol production from xylose, demonstrating the essential nature of these enzymes in xylose fermentation (Matsushika et al., 2012). Interestingly when the secondary TAL (NQM1) and TKL2 genes were deleted, growth and ethanol production increased compared with the control (Matsushika et al., 2012). Transcriptome analysis has proved useful in identifying not only PPP gene targets, but also other genes for optimizing xylose utilization. Transcriptome analysis of cells grown in xylose identified increased mRNA levels from genes such as TAL1, TKL1, SOL3, and GND1. Interestingly non-PPP genes such as those involved in galactose metabolism were also upregulated (Bengtsson et al., 2008; Wahlbom, Cordero Otero, Van Zyl, Hahn-Hagerdal, & Jonsson, 2003). Identification of genes downregulated in xylose fermentations was also used as a guide to identify possible gene targets for improving xylose metabolism. Generating deletion mutants in the identified genes validated the transcriptome analysis: strains carrying deletions in YLR042C, MNI1, and RPA49, showed a vast improvement in growth rates on xylose compared to the control strain (Bengtsson et al., 2008). The deletion of YLR042C along with the overexpression of the PPP genes (TAL1, TKL1, RKL1, and RPE1) increased ethanol yields twofold (Parachin, Bengtsson, Hahn-Hagerdal, & Gorwa-Grauslund, 2010). Other gene deletions leading to increased ethanol yields from xylose include PHO13, ALP1, ISC1, RPL20B, and BUD21 (Usher et al., 2011; Van Vleet, Jeffries, & Olsson, 2008). The mechanism of how these deletions increase xylose fermentation is currently unknown. It is clear that improved pentose metabolism is controlled by the expression of a variety of genes and involves complex genetic and biochemical interactions. To mimic this phenotype through overexpression or deletion of identified target genes may prove complex and unpredictable. The identification of key genes required for optimum PPP metabolism has been invaluable, however, an alternative strategy might be to develop an evolutionary approach to identify natural mutations that lead to increased xylose metabolism in S. cerevisiae.

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5. HEXOSE AND PENTOSE SUGAR COUTILIZATION For efficient and complete CBP of lignocellulose to ethanol, both the hexose and pentose sugars from the cellulose and hemicellulose fractions must be utilized. This necessitates the development of CBP from mixed sugar hydrolysates as well as from semisolid biomass slurries. Fermentations of mixed sugar solutions pose several problems. In general, glucose is preferentially fermented due to its preferred uptake by glucose transporters even in the presence of xylose and arabinose-specific transporters (Subtil & Boles, 2012). Glucose catabolism has also been shown to affect xylose utilization through alterations in metabolic fluxes in xylose metabolism (Pitkanen, Aristidou, Salusjarvi, Ruohonen, & Penttila, 2003). In of itself, this is not necessarily problematic as initial glucose fermentation allows for cell growth and the concomitant expression of genes required for pentose sugar fermentation. Since sequential sugar utilization is the norm in yeasts, this should not be considered a drawback to CBP. The coutilization of hexose and pentose sugars from hemicellulose with sugars extracted from the cellulose fraction of biomass is more problematic. The polymeric structure of cellulose precludes its transport into the yeast cell and cellulose hydrolysis cannot begin until sufficient cellulases have been produced and secreted from the cell. As a first step to generating recombinant yeasts with a capacity to ferment cellulosic and hemicellulosic sugars, yeast strains have been engineered for the metabolism of xylose and cellobiose, the repeating disaccharide found in cellulose. Two different approaches for cellobiose and xylose coutilization have been tested. Firstly the genes encoding xylose-utilizing enzymes (XR/XDH/XKS1) and an intracellular BGL1 were coexpressed along with a cellodextran transporter to allow for intracellular transport of cellobiose (Chomvong et al., 2014; Ha et al., 2011, 2013). The second approach expressed genes encoding xylose-utilizing enzymes and a secreted form of b-glucosidase to allow for extracellular cellobiose hydrolysis followed by glucose and xylose transport into the cell (Saitoh, Hasunuma, Tanaka, & Kondo, 2010). Both strategies supported coutilization of xylose and cellobiose with similar ethanol yields. To develop a comprehensive cellulose and hemicellulose coutilization system, Kricka, James, and Bond (unpublished) coexpressed the three major cellulase genes EGL2, CBH2, and BGL1 along with the XR/XDH genes from T. reesei and XKS from S. cerevisiae in a stress-tolerant strain of Saccharomyces pastorianus (James, Usher, Campbell, & Bond, 2008). In fermentations

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carried out at 30  C, hydrolysis of the cellulose substrate PASC was ineffectual, confirming the poor activity of cellulase enzymes at fermentation temperatures. Xylose metabolism was extremely efficient with an observed 73% conversion of xylose to ethanol. Cofermentation of both xylose and cellulose (in a semisolid state) using the recombinant strain generated an ethanol yield to 82%. In the mixed fermentation, xylose was initially metabolized allowing for cellulase enzyme production. Thus, the initial metabolism of xylose facilitates cellulase production resulting in sequential pentose and hexose sugar utilization. The slow release of glucose from cellulose at 30  C appears to be beneficial toward the coutilization of xylose and cellulose as the controlled low-level release of glucose from complex sugars minimizes inhibitory effects of glucose on xylose utilization and fluxes within the cell.

6. CHALLENGES TO USING REAL BIOMASS FOR BIOETHANOL PRODUCTION BY RECOMBINANT YEASTS A major disadvantage to all pretreatment methods is the production and release of various undesirable by-products such as acetic acid, formic and levulinic acids resulting from the hydrolysis of sugar molecules. These weak acids can affect cellular growth and ethanol yield through diffusion across the plasma membrane and altering cytosolic pH (Palmqvist & Hahn-Hagerdal, 2000). The resulting increase in intracellular [Hþ] concentration affects cellular ATP concentrations and causes DNA damage. Other inhibitors such as furfurals and 5-hydroxymethyl furfural (HMF) are formed from the degradation of pentose and hexose sugars at high temperature and pressure. Furfurals have been shown to affect cellular growth, enzyme activity (Modig, Liden, & Taherzadeh, 2002), and cellular redox balance (Ask, Bettiga, Mapelli, & Olsson, 2013), although interestingly glycolytic activity is maintained (Horvath, Taherzadeh, Niklasson, & Liden, 2001; Sarvari Horvath, Franzen, Taherzadeh, Niklasson, & Liden, 2003; Taherzadeh, Gustafsson, Niklasson, & Liden, 2000). Phenolic compounds, which are a by-product of lignin degradation, have also been shown to greatly affect ethanol production and yield (Klinke, Olsson, Thomsen, & Ahring, 2003). In addition to inhibiting cellular growth, the main by-products of pretreatment have a specific inhibitory effect on xylose metabolism in recombinant yeast expressing xylose-utilizing genes (Wang et al., 2014) (Kricka and Bond, unpublished). Metabolome analysis of xylose fermentations in the presence of inhibitors revealed the accumulation of

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various nonoxidative PPP intermediates (Hasunuma et al., 2011), indicating a bottleneck in PPP metabolism. This inhibition can be partially relieved by overexpression of the PPP TAL1 (Hasunuma, Ismail, Nambu, & Kondo, 2014; Hasunuma et al., 2011). A transcriptome analysis of cells grown in the presence of inhibitors revealed reduced levels of transcripts coding for proteins required not only for carbohydrate metabolism but also for transcriptional and translational control, indicating the pleotrophic effect of inhibitors on cell metabolism (Bajwa et al., 2013; Li & Yuan, 2010). To improve fermentation efficiency from biomass, it is clear that the presence of inhibitors must be addressed. One approach has been to remove inhibitors chemically or by filtration (Chandel, Kapoor, Singh, & Kuhad, 2007; Grzenia, Wickramasinghe, & Schell, 2012; Sasaki et al., 2014; Zhuang, Liu, Wu, Sun, & Lin, 2009), although this adds substantial costs to bioethanol production. Other strategies involve engineering yeasts to metabolize inhibitory compounds present in biomass hydrolysates. Furfurals and HMF can be naturally metabolized by yeasts into less toxic alcohols. The overexpression of genes associated with furfural and HMF metabolism has been examined as a solution to overcome the effects of inhibitors. In particular the overexpression of the alcohol dehydrogenase genes ADH6, ADH7, and a mutated ADH1 led to increased growth, ethanol yields, and productivity in the presence of inhibitors (Ishii, Yoshimura, Hasunuma, & Kondo, 2013; Laadan, Almeida, Radstrom, Hahn-Hagerdal, & Gorwa-Grauslund, 2008; Liu, Moon, Andersh, Slininger, & Weber, 2008). The increased metabolism of furfural and HMF through overexpression of ADH genes comes at a cost due to the cofactor requirements of both Adh6 and Adh7 (NADH and NADPH respectively), which may lead to redox imbalances (Ask et al., 2013; Petersson et al., 2006). While furfurals and HMF, can be metabolized by yeasts, it is the weak acids present in biomass hydrolysates that specifically appear to pose a major problem (Hasunuma et al., 2011). In a recent study, yeast strains were engineered to metabolize acetic acid through expression of acetylating acetaldehyde dehydrogenase, leading to a reduction in acetate in the medium and improved growth and ethanol production (Wei, Quarterman, Kim, Cate, & Jin, 2013). The overexpression of genes such as MSN2 (Sasano et al., 2012), GSH1 (Ask, Mapelli, Hock, Olsson, & Bettiga, 2013), and FLO1 (Westman, Mapelli, Taherzadeh, & Franzen, 2014), which are not specifically associated with furfural or acetic acid detoxification, has also proved beneficial to inhibitor tolerance. The increased ethanol production and inhibitor tolerance associated with an increased flocculent phenotype is interesting. A similar study using cell

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encapsulation suggested that the higher cell density found in encapsulated cultures not only allowed higher inhibitor tolerance but also improved simultaneous mixed sugar utilization (Westman, Bonander, Taherzadeh, & Franzen, 2014; Westman, Manikondu, Franzen, & Taherzadeh, 2012). The replacement of encapsulation by highly flocculent strains may be a more cost-effective approach to increasing inhibitor tolerance. Deletion of genes has also been shown to increase tolerance to inhibitors. The deletion of PHO13 improved ethanol production from lignocellulose hydrolysate (Fujitomi, Sanda, Hasunuma, & Kondo, 2012), while the deletions of genes identified to be downregulated in the presence of inhibitors (PCL1, RPS22A, TOS6, RPL8B, RPL22A, OPI11, RPS17B, and RPL13A) also demonstrated improved acetic acid tolerance (An et al., 2014). The exact mechanism of how these deletions increased inhibitor tolerance is not clearly understood. Evolutionary engineering and mutagenesis have been carried out to adapt yeast for growth in the presence of inhibitors (Demeke et al., 2013; Koppram, Albers, & Olsson, 2012). Adaptive evolution is achieved by culturing of yeast cells for many generations (40e100) in the presence of inhibitors (Sato et al., 2014; Smith, Van Rensburg, & Gorgens, 2014). Increased inhibitor tolerance has been linked to an increase in fitness and cell viability (Almario, Reyes, & Kao, 2013; Heer & Sauer, 2008; Koppram et al., 2012; Wallace-Salinas & Gorwa-Grauslund, 2013), however significant increases in ethanol production from tolerant strains have been limited (Wallace-Salinas & Gorwa-Grauslund, 2013). High-throughput screening of natural robust industrial yeasts has been used to identify suitable species for bioethanol production for biomass. A comparison of lab and industrial S. cerevisiae isolates has highlighted the limitations of lab strains: peak ethanol levels using biomass hydrolysate from lab strains were up to 25-fold less than that produced by industrial isolates (Pereira, Romani, Ruiz, Teixeira, & Domingues, 2014). Isolates from the environment have also been shown to have natural high tolerance to inhibitor compounds, with isolates from grape marc showing increased ethanol production from sugarcane hydrolysate (Favaro et al., 2013).

7. CONCLUSIONS More than four decades of research have provided us with the background scientific knowledge of the genes required for cellulose and

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hemicellulose metabolism and the mechanisms to introduce and express these genes in suitable Saccharomyces host species. Knowledge of the host biochemistry and genetics has furthered our understanding of the metabolic pathways required for pentose and hexose conversion to ethanol. Armed with this knowledge and taking into account recent advances in synthetic biology, we are not far from generating the ultimate microorganism that is capable of efficiently utilizing all of the sugars that can be extracted from lignocellulose biomass in a CBP fermentation. We have learned that despite our best efforts, the complexities of cellular metabolism and unforeseen subtle interactions between cellular metabolites may limit our ability to bioengineer yeasts for CBP and lower conversion efficiencies of sugars to ethanol may have to be accepted. Roadblocks, such as reduced enzyme activities at fermentation temperatures, sugar uptake, and effects of inhibitors on yeast cell growth and metabolism, continue to provide challenges for the bioengineer into the future. The art of converting sugars to ethanol has been perfected for millennia and industrialized for several centuries. The lessons learned over the centuries can guide the quest for a commercially viable industrialized process for converting biomass to bioethanol. We propose a step-wise fermentation process, based on current practices in industrial breweries, as a model for CBP (Figure 3). Firstly, biomass, such as spent grains, a waste product from industrial brewing, is milled and pretreated to generate (1) a liquor (hydrolysate) containing extracted pentose and hexose sugars from the hemicellulose fraction of biomass and (2) an amorphous form of cellulose (solids). Secondly, a microorganism engineered to utilize cellulose, xylose, and arabinose is grown in batch in rich media. The cells generated are used for the subsequent fermentation of the hemicellulose liquor while the supernatant, containing recombinant cellulase enzymes, is incubated with the remaining insoluble biomass (mashing stage) at an optimal temperature for cellulase activity. The biomass cellulase slurry is then separated (lautering stage) into soluble (containing glucose release from cellulose) and insoluble fractions, with the soluble fraction being combined with hemicellulose liquor for fermentation. The remaining insoluble fraction can be used as solid fuel for energy production. This process could easily be integrated into present-day commercial brewery practice where a similar elevated temperature mashing stage is used for extraction of sugars from malted barley by amylases, however, success is heavily dependable on improved tolerance toward inhibitors, improved xylose utilization, and increased recombinant cellulase yields.

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Figure 3 A step-wise industrial fermentation process for biomass. Biomass such as spent grains (SG) are first pretreated to extract sugars from the hemicellulose fraction and to expose amorphous cellulose. The SG liquor and insoluble material (here referred to as spent spent grains; SSG) are then separated. Next recombinant yeast strains expressing xylose, arabinose, and cellulose metabolizing enzymes are grown to high cell densities. The supernatant from the cultures, containing secreted cellulases is used for hydrolysis at high temperatures of the SSGs (mashing). The SSG and soluble sugars are then separated (lautering). The released sugars (glucose) are cooled, combined with the liquor and added to the cells for fermentation. The remaining SSGs can be utilized as solid fuel.

REFERENCES Agbor, V. B., Cicek, N., Sparling, R., Berlin, A., & Levin, D. B. (2011). Biomass pretreatment: fundamentals toward application. Biotechnology Advances, 29, 675e685. Ahmed, I. N., Santoso, S. P., Tran-Nguyen, P. L., Huynh, L. H., Ismadji, S., & Ju, Y. H. (2013). Impact of pretreatments on morphology and enzymatic saccharification of shedding bark of Melaleuca leucadendron. Bioresource Technology, 139, 410e414. Almario, M. P., Reyes, L. H., & Kao, K. C. (2013). Evolutionary engineering of Saccharomyces cerevisiae for enhanced tolerance to hydrolysates of lignocellulosic biomass. Biotechnology and Bioengineering, 110, 2616e2623. An, J., Kwon, H., Kim, E., Lee, Y. M., Ko, H. J., Park, H., et al. (2014). Tolerance to acetic acid is improved by mutations of the TATA-binding protein gene. Environmental Microbiology. http://dx.doi.org/10.1111/1462-2920.12489. Ask, M., Bettiga, M., Mapelli, V., & Olsson, L. (2013). The influence of HMF and furfural on redox-balance and energy-state of xylose-utilizing Saccharomyces cerevisiae. Biotechnology for Biofuels, 6, 22. Ask, M., Mapelli, V., Hock, H., Olsson, L., & Bettiga, M. (2013). Engineering glutathione biosynthesis of Saccharomyces cerevisiae increases robustness to inhibitors in pretreated lignocellulosic materials. Microbial Cell Factories, 12, 87. Baek, S. H., Kim, S., Lee, K., Lee, J. K., & Hahn, J. S. (2012). Cellulosic ethanol production by combination of cellulase-displaying yeast cells. Enzyme and Microbial Technology, 51, 366e372.

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