Redox balancing in recombinant strains of Saccharomyces cerevisiae

Mikael Anderlund

Department of Applied Microbiology Lund UniversityLund Institute of Technology Sweden 1998 DISCLAIMER Redox balancing in recombinant strains of Saccharomyces cerevisiae

Mikael Anderlund

Department of Applied Microbiology Lund UniversityLund Institute of Technology Sweden 1998

Akademisk avhandling for avlaggande av teknologie doktorsexamen vid tekniska fakulteten vid Lunds universitet. Avhandlingen kommer att forsvaras pH svenska vid en offentlig disputation p%Kemicentrum, Solvegatan 39, Lund, mhdagen den 28 September 1998, kl 10.15.

Fakultetsopponent: Docent Lena Gustafsson, Goteborgs universitet, Sverige . . - ... ._

~cumentname DCTORAL DISSERTATION 1 'teofi~~ September, 1998 3DW: mawring oqdzation Nordic Energy Research Progqme. NUTEK and NFR icchromyces cerevisiae

:xpressing Pichia sripitis XYLl and XY1.2 genes, :DH). respectively. xylitol is excreted as the major Y yields of ethanol are produced. This has been nd the exclusive use of NAD+ by XDH. ormation of glycerol and acetic acid were reduced mited conditions by expressing lower levels of XR were controlled by changing the promoters and

:n Zhermus thermophilus qlA gene was expressed t xylose to ethanol when cultivated on a minimal wo first steps of the xylose metabolism, and 'I XYLI hen the fusion protein. containing a linker of three oxidoreductase. between the two coenzyme systems NADRH) and asformed with a plasma membrane bound AB- anshydrogenase from A. vinelandii. Expression of 3 NADPH/NADp ratio was reduced while the lation of 2-oxoglutarate, glycerol and acetate was

vinelandii. E. coli, xylose. xylitol. glycerol. xylose lide nucleotide transhydrogenas?. _ILanIplpge English

Applied Microbiology. Lurid University. P.O.Box

DVe-mMthned dirratrtion, hereby putto d reference !-mentioned dburtrrtion. CONTENTS

List of papers

1. Introduction 1

2. Glucose and xylose metabolism in yeast 3 2.1 Glucose and xylose metabolism in xylose utilising yeasts 3 2.2 Xylose utilising recombinant S. cerevisiue strains 6

3. Redox balance in recombinant S. cerevisiae strains 9 3.1 The reducing equivalents NADH and NADPH 9 3.2 Glycerol and xylitol in cellular redox balancing 12 3.3 Redox balance in S. cerevisiue strains expressing XYLl and XyL2 genes 14 3.4 Expression of the T. rhemphilis xylA gene in S. cerevisiue 17

4. Artificial bifunctional enzymes 20 4.1 Channelling in metabolic pathways 20 4.2 Metabolite transfer mechanisms 21 4.3 Artificial enzyme systems 22 4.4 Subunit interactions and linker regions in bifunctional enzymes 23 4.5 Proximity studies in vivo 26

5. Nicotinamide nucleotide transhydrogenase 28 5.1 Classes of nicotinamide nucleotide transhydrogenase 28 5.2 AB-transhydrogenases 29 5.3 BB-transhydrogenases 34 5.4 Cloning and expression of AB- and BB-transhydrogenases in S. cerevisiae 36

6. Acknowledgements 42

7. References 43 List of papers

This thesis is based on the following papers:

I Walfridsson, M., Anderlund, M., Bao, X., and Hahn-Hagerdal, B. (1997) Expression of different levels of enzymes from the Pichia stipitis XYLI and XYL2 genes in Saccharomyces cerevisiae and its effect on product formation during xylose utilisation. Appl. Microbiol. Biotechnol. 48:218-224

I1 Walfridsson, M., Bao, X., Anderlund, M., Lilius, G., Bulow, L., and Hahn-Hagerdal, B. (1996) Ethanolic fermentation of xylose with Saccharomyces cerevisiae harboring the Thermus thermophilus xylA gene, which expresses an active xylose (glucose) isomerase. Appl. Microbiol. Biotechnol. 62:4648-465 1

III Anderlund, M., Ridstrom, P., and Hahn-Hiigerdal, B. (1998) Expression of artificial genes encoding bifunctional enzymes with xylose reductase and xylitol dehydrogenase activity in Saccharomyces cerevisiae alter product formation during xylose fermentation. Manuscript.

IV Anderlund, M., Nissen, T., Nielsen, J., Rydstrom, J., Hahn-Hagerdal, B., Villadsen, J., and Kielland-Brandt, M. (1998) Expression of E. coli pntA and pntB genes encoding nicotinamide nucleotide transhydrogenase in Saccharomyces cerevisiae and its effect on product formation during anaerobic glucose fermentation. Submitted for publication.

V Nissen, T., Anderlund, M., Nielsen, J., Villadsen, J., and Kielland-Brandt, M. (1998) Expression of a cytoplasmic transhydrogenase in Saccharomyces cerevisiae results in formation of 2-oxoglutarate due to depletion of the NADPH pool. Submitted for publication. 1. Introduction

Lignocellulosic materials are the most abundant, renewable material found on the earth and can be used as a substrate for microbial fermentation to ethanol. Ethanol derived from lignocellulosic materials can be used as a renewable energy source, replacing fossil fuels. Unlike fossil fuels, the combustion of ethanol produced from lignocellulosic materials does not contribute to the increase in the carbon dioxide level in the atmosphere. Hardwood, softwood and agricultural waste are considered to be suitable for industrial ethanol production (Hayn et aZ., 1993).

Lignocellulosic materials can roughly be divided into three fractions: cellulose, hemicellulose and the lignin fraction. The cellulose fraction is composed of glucose units linked together with f3-1,4-glucosidic bonds forming crystalline polymers. The hemicellulose fraction contains polymers of xylan, glucomannan, galactan and glucan. The ratio of different polysaccharides varies between species and between different parts of the same plant. Lignin is closely associated with cellulose and consists of a complex network of randomly polymerised phenolic monomers.

Prior to fermentation, the lignocellulosic materials must be degraded into a fermentable substrate. The material is pretreated with acids or steam, or with a combination of the two, to make the cellulose fraction accessible to hydrolysis (Saddler et aZ., 1993). The subsequent hydrolysis is achieved with acids or with enzymes, involving endoglucanases, cellobiohydrolases and f3-glucosidase. The hydrolysate is made up mainly of the hexoses glucose, galactose and mannose and the pentoses xylose and arabinose. Between 6 and 28% of the total dry material consist of pentoses where xylose is the dominant one (Ladisch et al., 1983). For economical reasons, all sugars present in the hydrolysate must be fermented into ethanol (von Sivers and Zacchi, 1995). The capability of the microorganism to ferment xylose is especially important since xylose constitutes a substantial part of the sugars.

The performance of ethanol-producing microorganisms in hydrolysates has been extensively investigated (Bjorling and Lindman, 1989; Olsson and Hahn-Hiigerdal, 1993). Saccharomyces cerevisiae, traditionally used for the production of wine and beer, has several properties useful for ethanol production from lignocellulosic hydrolysate: (i) it has the ability to grow and ferment glucose anaerobically, (ii) it shows high ethanol productivity and high ethanol yield from glucose, (E) it has high ethanol tolerance, (iv) it shows high tolerance to inhibitors present in the hydrolysate, (v) and it is also recognised as a GRAS (Genererally Regarded As Safe)

1 microorganism. A major drawback is that S. cerevisiu lacks the ability to metabolise xylose. However, metabolic engineering can be used to extend the substrate range for growth and product formation of an organism. Metabolic engineering can establish new metabolic pathways, remove existing pathways, increase the metabolic flux or alter the distribution of the metabolic flux at a specific branch point.

Genes encoding enzymes required for xylose metabolism have been cloned from xylose utilising yeast and from bacteria, and have been successfully expressed in S. cerevisiae (Kotter et ul., 1990; Tantirungkij er uL, 1993; Walfridsson et al., 1995; Papers I and II). However, adding new metabolic pathways into an organism very often causes disturbances in the native metabolic pathways in the cell due to lack of detailed knowledge about the genetics and biochemistry of the pathway of interest and the physiology of the host organism. In metabolically engineered S. cerevisiue expressing enzymes required for xylose fermentation, xylitol is excreted as the major product during anaerobic xylose fermentation and only low yields of ethanol are produced, indicating a disturbance in the metabolism and the redox balance of the cell.

In the present work, several genes were cloned and expressed in S. cerevisiue in order to construct a recombinant strain able to ferment xylose and glucose efficiently giving an increased ethanol yield and low formation of unwanted by-products such as glycerol and xylitol. The work presented in this thesis covers different approaches to metabolic engineering of S. cerevisiue such as; changes and control of enzyme levels (Paper I), construction of an alternative functional pathway for xylose (Paper II) and the introduction of heterologous redox balancing enzyme systems (Papers IV and V). In order to create a channelled metabolite transfer and thereby prevent byproduct formation e.g. xylitol and glycerol, protein engineering was used to construct artificial bifunctional enzymes with two sequentially operating enzymes (Paper m).

2 : 2. Gluc zose and xylose metabolism in yeasts

2.1 Glue1ose and xylose metabolism in xylose utilising yeasts

Yeasts can be classified into obligate aerobes and facultative anaerobes (Gancedo and Serrano, 19f $9). The obligate aerobes are unable to utilise a carbon source in the absence of oxygen, while the facultative anaerobes are able to utilise a carbon source under both ai erobic and anaerobic conditions. The facultative anaerobes can further be divided into1 fermentative and respiratory yeasts, depending on whether aerobic fermentatior1 is predominant or not. Saccharomyces cerevisiae belongs to the fermentative: group and ferments glucose to ethanol during aerobic conditions. However, a 1particular yeast may be classified as a facultative anaerobe when grown on a specific: sugar, but an obligate aerobe on other sugars. Pichia stipitis has been classified as a facultative anaerobe when grown on glucose or xylose as only carbon source (Ligthelm et aL, 1988), while Candida utilis has been classified as a facultative ai naerobe when grown on glucose but as a strict obligate aerobe on xylose (Bruienberg et al., 1983a). It should be born in mind that this typ of classification may be vagie and it has been reported that most facultative anaerobes need micro- aerobic cond itions for growth and fermentation (Visser et aL, 1990).

Only very few yeasts have been found to ferment xylose anaerobically (Toivola et aL, 1984). This phenomenon, the Kluyver effect, has been defined as the inability of a facultatively fermentative yeast to metabolise a certain sugar anaerobically although the sugar is rapidly metabolised aerobically (Sims and Barnett, 1978). This is a general pher iomenon observed for many sugars in many yeasts and in several cases the metabolic basis for this incapacity is not know. However, the distinct difference between anaerobic fermentation of glucose and of xylose in C. utilis may reflect a disturbed rec lox balance during xylose utilisation (Bruinenberg et al., 1983a). Xylose Glucose COZ 2NADpH 2NADp+ Gluc!se6P t A FNCtOse 6P ulose SP &bose 5p Fructoset 1.6DP

Glyceraldehyde 3P eptulose 7P t Glyceraldehyde \f- 3P t

FNctose 6 Glyeeraldehvd 3P Phosphoeuolpyruvatet I 3- NZ Pyruvate Acetaldehyde Ethanol

Aceticadd OxaloacetateL citrate

Figure 1. Metabolic pathways for xylose and glucose in yeast. XR, XDH, XK, TAL, and TKL are enzymes expressed or overexpressed in S. cerevisia in order to create a functional metabolic pathway for xylose. Abbreviations: XR, xylose reductase; XDH, xylitol dehydrogenase; XK, xylulokinase; TAL, transaldolase; TKL, transketolase. (Modified from Skoog, 1992).

4 The production of ethanol from glucose via glycolysis is redox neutral, since the production of NADH from the conversion of glyceraldehyde 3-phosphate to 1,3 diphosphoglycerateis consumed in the reduction of acetaldehyde to ethanol. Under anaerobic conditions the surplus NADH formed in redox reactions associated with biomass formation is oxidised by dihydroxy acetone phosphate, an intermediate in glycolysis, as endogenous electron acceptor. Reduction of dihydroxy acetone phosphate to glycerol 3-phosphate via glycerol 3-phosphate dehydrogenase and dephosphorylation of glycerol 3-phosphate by glycerol 3-phosphatase lead to glycerol formation (Oura, 1977; Albers et al., 1996). Glycerol in cellular redox balancing is discussed in Chapter 3.

In contrast to anaerobic glucose fermentation, the production of ethanol from xylose is not a redox neutral process in xylose utilising yeasts (Bruinenberg et al., 1984). Xylose is first reduced to xylitol by an NAD(P)H-dependent xylose reductase (XR). Xylitol is then oxidised to xylulose by an NAD+-dependent xylitol dehydrogenase (XDH). However, the cofactor requirements of XR vary between different xylose utilising yeasts and may determine whether xylose fermentation is possible or not. In Pichia stipitis, Pichia segobiensis, Pachysolen tannophilus, Candida shehatae and Candida tenuis, XR can utilise either NADH or NADPH, whereas in C. utilis and other strictly obligate aerobes, XR is NADPH-dependent (Bruinenberg et uL, 1984). The second step in the xylose metabolism, catalysed by XDH, is specific for NAD+ in xylose utilising yeasts. If NADH-coupled XR activity is present, as in P. stipitis, P. segobiensis, P. tannophilus, C. shehatue and C. tenuis, xylose assimilation may proceed with a closed redox balance, since the NADH generated by XDH will be completely reoxidised by XR. On the other hand, if the XR is strictly NADPH- dependent, as in C. utilis, NADH will accumulate and metabolism will stop. This apparent redox imbalance is reflected by the excretion of the intermediate xylitol. If an external electron acceptor is added, xylitol excretion is prevented and anaerobic ethanol production occurs (Bruinenberg et al., 1983a). No transhydrogenase activity has so far been detected in yeast and the redox balance favouring xylose consumption cannot be maintained by this route (Bruinenberg et al., 1983b).

However, most xylose-fermenting yeasts produce some xylitol in addition to ethanol under anaerobic conditions, in spite of XR activity with dual cofactor specificity, which should permit anaerobic ethanol production without xylitol excretion. The addition of an artificial hydrogen acceptor to P. tannophilus under anaerobic and oxygen-limited conditions decreased the xylitol excretion and improved the ethanol production, indicating an overproduction of NADH in xylose-fermenting cells (Ligthelm et al., 1989). However, oxygen has a similar stimulatory effect on xylose,

5 xylitol, xylulose and glucose fermentation by these yeasts (Neirinck et al., 1985; du Preez et al., 1986; Ligthelm et al., 1988; Skoog et al., 1992; Yu et al., 1995), indicating that factors other than the redox imbalance in the conversion of xylose to xylulose retard xylose metabolism under anaerobic conditions. Such factors could be sugar transport or unimpaired mitochondrial function resulting from a lack of oxygen (Slininger et al., 1982; Ligthelm et al., 1988; Skoog et al., 1992; Jeppsson et al., 1995).

Before xylulose enters the pentose phosphate pathway, it is phosphorylated by xylulokinase to xylulose-5-phosphate. The xylulokinase gene has been isolated from P. tannophilus (Stevis et al., 1987) and from S. cerevisiae (Ho and Chang, 1989). The pentose phosphate pathway enzymes, ribulose 5-phosphate epimerase, ribose 5- phosphate ketol-isomerase, transketolase and transaldolase, convert xylulose-5- phosphate to glycolytic fructose-6-phosphate and glyceraldehyde-3-phosphate. The pentose phosphate pathway, which is connected to glycolysis via glucose 6-phosphate dehydrogenase, furnishes the cell with cytosolic NADPH, ribose-5-phosphate, and erythrose-4-phosphate for nucleotide and amino acid biosynthesis. All xylose must be routed by the pentose phosphate pathway, in contrast to glucose where only 1-10% has been estimated to be metabolised via the pentose phosphate pathway (Gancedo and Lagunas, 1973; Nissen et al., 1997).

Fructose-6-phosphate can be metabolised either by cycling via the oxidative part of the pentose phosphate pathway where carbon dioxide and NADPH are produced, or via glycolysis where it ends up as pyruvat. The theoretical yield of ethanol from xylose depends on whether a fraction of the fructose-6-phosphate is cycled via the oxidative pentose phosphate pathway or not. If all the fructose-6-phosphate is metabolised via glycolysis, the yield will be 0.5 1 g g-*,whereas cycling of fructose-6- phosphate, due to high consumption of NADPH, may decrease the yield to as low as 0.3 1 g g-1 (Slininger et aE., 1987). Glyceraldehyde-3-phosphate is metabolised through glycolysis to pyruvate. Pyruvate can either be decarboxylated and reduced to ethanol by pyruvate decarboxylase and alcohol dehydrogenase, or be oxidised to carbon dioxide through the citric acid cycle and respiratory chain.

2.2 Xylose utilising recombinant S. cerevisiae strains

The yeast S. cerevisiae is not able to utilise xylose as the sole carbon source but it can utilise the keto form xylulose both for growth and fermentation to ethanol. The gene for xylulokinase has been deleted in S. cerevisiae and the mutant was unable to grow with xylulose as the carbon source (Ho and Chang, 1989). Obviously, the limiting

6 lose via xylitol by the Amore et al., 1991; :oding XR and XDH, S. cerevisiae. olecular weight of 76 shows homologies to the same aldo-keto :en considered to be a :oh01 dehydrogenase :of P. stipitis xylitol imeric medium-chain ment is important for trameric form of the 1.

2 genes produce both r. However, xylose giderable amounts of Kotter et al., 1993), hway and a disturbed phosphate in xylulose 0), and in a glucose- ierstenschlager et al., rol the xylose flux. in XYLI- and XYL2- : growth on xylose transketolase did not erevisiae strains but ced compared with a aL, 1995; Paper I). lose consumption in pression of the native ition and the ethanol al., 1998). cerevisiue expressing r hexose phosphates, Kotter and Ciriacy, 1993). If hexose phosphates are present at too low levels, the enzymes in the lower part of glycolycis may not be induced (Boles et al., 1993; Boles and Zimmermann, 1993; Muller et al., 1995), leading to slow glycolytic flux during xylose fermentation. The hexose phosphates are also required for inactivation of the gluconeogenic enzyme fructose 1,6-bisphosphatase (Boles et al., 1993), preventing a futile recycling of fructose 6-phosphate and glyceraldehyd 3-phosphate through the oxidative part of the pentose phosphate pathway. However, these problems may be overcome by simultaneous xylose and glucose fermentation. The xylulose consumption rate was 4 times higher during simultaneous xylulose and glucose fermentation than during fermentation on xylulose alone (Jeppsson et al., 1996) indicating a connection between high glycolytic metabolite concentrations and enhanced xylulose consumption.

8 Binant S. cerevisiae strains

DH and NADPH

Jes NAD(H) and NADP(H) play separate and reast. In yeast, NADH may be regarded as a walent, whereas NADPH is mainly involved in concentrations of NAD+, NADH, NADP+ and iaerobic conditions were determined to be 2.87, omass (dry weight) (Papers IV and V). The and NADH in S. cerevisiae have earlier been e per g biomass (dry weight), respectively (De cerevisiae cells grown under aerobic conditions and NADPH values have been published, 1.51, iomass (dry weight), respectively (Saez and [ADP+and NADH/NAD+, in S. cerevisiae cells 1 and 0.15, respectively (Papers IV and V). In a o NADPH/NADP+ was always higher than the >ngbatch fermentation of grape must (Mauricio

nvolves several redox reactions associated with and amino acids (Jones and Fink, 1982). The )und effect upon the redox balance of the cell. :ed by S. cerevisiae are formed in oxidative [thesis the of lipids and nucleic acids involves a actions, leading to a significant net output of metabolic end products, such as acetic acid, ivate, also generates a net surplus of NADH

imilation is even higher than that anticipated levels of sugar and biomass (van Dijken and ed by the fact that NADH produced during the *al metabolism is not the principal reductant for biomass. Instead of NADH, most anabolic which is chemically separated from NADH as a nshydrogenase and NAD(H) kinase activity

9 (Lagunas and Gancedo, 1973; Bruinenberg et al., 1983b). The specific requirement for NADPH in the combination with the absence of transhydrogenase activity, necessitates the conversion of part of the sugar exclusively for the purpose of generating reducing power in the form of NADPH.

Since the concentration of NAD+ is low compared with the amount of intermediates which must be oxidised to maintain cell growth, NADH production must be balanced in order to maintain the cellular redox status and metabolic flow. Under aerobic conditions, yeast may balance any overproduction of reducing equivalents by delivering NADH to the mitochondrial electron transport system via the external NADH dehydrogenase (De Vries and Marres, 1987; Willis, 1990), located on the cytosolic side of the inner mitochondrial membrane, or via the glycerol 3-phosphate shuttle (Larsson et al., 1998). The malate-aspartate shuttle present in higher eucaryotic cells seems to be non-functional and the fatty acid malate-citrate shuttle unimportant in yeast (De Vries and Marres, 1987).

Under conditions of oxygen depletion, the cell must use glucose for the production of endogenous electron acceptors in order to regenerate cytosolic NAD+ (Oura, 1977; Albers et al., 1996). Under these conditions, NADH produced in the mitochondria must be transported to the cytosol in order to maintain the redox balance in this compartment due to lack of transhydrogenase activity. It has been suggested that the diffusion of cytosolically formed acetaldehyde into the mitochondria, followed by reduction by mitochondrial ADHIII, may constitute a functional shuttle (von Jagow and Klingenberg, 1970; Nissen et al., 1997). It has also been suggested that the formation of 2-hydroxyglutarate from 2-oxoglutarate catalysed by 2-hydroxyglutarate dehydrogenase may serve as an additional local sink for NADH produced in the mitochondria when glutamate is used as a nitrogen source (Albers et al., 1998). In this case, NADH produced in the mitochondria is consumed directly in the mitochondria and NADH transport to the cytosol is not necessary. The similar yields of 2-hydroxyglutarate and succinate during anaerobic growth on glucose and glutamate, indicated that the reduction of 2-oxoglutarate takes place in the mitochondria and not in the cytosol (Albers et al., 1998).

While NADH is a reducing equivalent produced and consumed mainly in catabolic reactions, NADPH must be regarded primarily as an anabolic reductant in yeasts. Except for some mitochondrial reduction steps in the biosynthesis of some amino acids, the major NADPH-consuming processes take place in the cytoplasm (Bruinenberg et al., 1983b). Lipid and glutamate synthesis, two major NADPH- consuming reactions, take place in the cytoplasm (Botham and Ratledge, 1979; Roon

10 et al., 1974). Metabolic flux analysis based on continuous cultivations of S. cerevisiae, cultivated in mineral medium containing NH4+ as the sole nitrogen source, has shown that 51% of the cytoplasmic NADPH was consumed in the production of glutamate (Nissen et al., 1997). Glutamate catalysed by the enzyme glutamate dehydrogenase is the donor of nitrogen in many of the biosynthetic pathways of amino acids.

The need for NADPH for biosynthesis is dependent on the carbon source. The metabolism of xylose in the majority of xylose utilising yeasts proceeds via a reduction to xylitol by xylose reductase (XR) which is strictly NADPH dependent (Barnet, 1976) before it enters the pentose phosphate pathway and glycolysis. Therefore, in contrast to glucose, the conversion of xylose to a central intermediate requires NADPH. However, some natural xylose-fermenting yeasts, such as Pichia stipitis, Pachysolen tannophilus, Candida shehatae and Candida tenuis (Bruinenberg et al., 1984) contain a XR which can utilise both NADH and NADPH. In these cases, depending on the Km for the cofactors and the concentration of the cofactors, xylose can be reduced by either NADH or NADPH.

The concentration of NADPH is low in S. cerevisiae cells growing under anaerobic conditions; 1.21 pmole per gram biomass (dry weight) (Papers IV and V). Thus, the NADP+ production must be balanced by conversion to NADPH in order to maintain the cellular redox status and metabolic flow. In yeast, NADPH is considered to be generated primarily by the oxidation of glucose 6-phosphate to ribulose 5-phosphate, using glucose 6-phosphate and 6-phosphogluconate dehydrogenase (Lagunas and Gancedo, 1973; Nissen et al., 1997). However, several other NADPH-generating pathways may exist in yeast. The enzymes isocitrate dehydrogenase IDPl and IDP2, located in the mitochondria and cytosol, respectively, catalyse the reversible conversion of isocitrate into 2-oxoglutarate under the simultaneous formation of NADP(H) (Haselbeck and McAlister-Henn, 1993; Zhao and McAlister-Henn, 1996). At present, it is not known whether these enzymes are indeed functioning for the purpose of NADPH synthesis or whether they only serve to generate 2-oxoglutarate for the biosynthesis of glutamate and glutamine.

Apart from the hexose monophosphate pathway and NADP+-linked isocitrate dehydrogenases, an NADP+ dependent acetaldehyde dehydrogenase may contribute to cytoplasmic NADPH synthesis (Llorente and de Castro, 1977). Recombinant S. cerevisiae strains expressing NADPH-consuming enzymes such as strains expressing XYLl encoding XR from P. stipitis (Meinander et al., 1996) and strains expressing transhydrogenase of different types (Papers IV and V) show increased formation of

11 acetate, indicating an NADPH-producing process connected to the NADP+-dependent acetaldehyde dehydrogenase. A cyclic pathway has been proposed in plants and fungi in which the interconversion of fructose and mannitol is linked to the transfer of reducing equivalents from NADH to NADPH (Hult et al., 1980). This transhydrogenating system, which requires the net input of one ATP per NADPH, has not been detected in yeast, and the operation of this cyclic pathway has been questioned (Singh et al., 1988). A mitochondrial NADP+-linked malic enzyme has recently been demonstrated in S. cerevisiue (Boles el al., 1998). This enzyme catalyses the oxidative decarboxylation of malate to pyruvate and carbon dioxide using NADP+ as an electron acceptor and may contribute to the production of intramitochondrial NADPH under anaerobic growth conditions. A cyclic reaction involving pyruvate carboxylase, NAD+-dependent malate dehydrogenase, a mitochondrial dicarboxylate carrier and an NADP+-linked malic enzyme could act as a transhydrogenase system. Such a system could theoretically convert part of the NADH to NADPH.

A nicotinamide nucleotide transhydrogenase, discussed in Chapter 5, which is present in the mitochondria of mammalian cells and in the plasma membrane and the cytoplasm of some bacteria catalysing the reaction NADH + NADP+ e-> NAD+ + NADPH has so far not been detected in yeast (Lagunas and Gancedo, 1973; Bruinenberg et al., 1983a; Bruinenberg et aL, 1985). In order to analyse the effects of an energy-linked and a non energy linked transhydrogenase in S. cerevisiue, transhydrogenase from E. coli (Paper IV) and A. vinelundii (Paper V) was cloned and transformed into S. cerevisiue. The effect of expressing transhydrogenase of different types in S. cerevisiae is discussed in Chapter 5.

3.2 Glycerol and xylitol in cellular redox balancing

Under both aerobic and anaerobic conditions glycerol synthesis serves as a redox valve to dispose excess reducing power in S. cerevisiue (Oura, 1977; van Dijken and Scheffers, 1986; Albers et ul., 1996). Glycerol is formed during aerobic respiro- fermentative growth conditions which has been considered to be the consequence of NADH/NAD+imbalance (van Dijken and Scheffers, 1986; Verduyn et ul., 1990). The cell requires redox regulation by glycerol synthesis, probably due to glucose repression of the respiratory systems. Substantial overproduction of glycerol can be observed during ethanol-fermentation, when reoxidation of glycolytically formed NADH is restricted by bisulphite (Neuberg and Reinfurth, 1918), indicating that glycerol serves as a redox sink. Recently, a S. cerevisiae mutant was constructed in

12 which both GPDl and GPD2 genes, encoding two isoforms of glycerol 3-phosphate dehydrogenase enzymes, were deleted (Ansell et al., 1997). This double mutant was incapable of glycerol production and could be grown only aerobically. Under anaerobic conditions, metabolism ceased and neither growth nor ethanol production was possible as a consequence of the inability of the cell to regenerate NAD+ (Lid& et al., 1996). Additionally, recombinant S. cerevisiae strains with increased NADH production due to the expression of energy-linked (Paper IV) and non-energy-linked transhydrogenase (Paper V) showed increased formation of glycerol.

In xylose-fermenting yeasts containing a xylose reductase such as Pichia stipitis, Pachysolen tannophilus, Candida shehatae andCandida tenuis (Bruinenberg et al., 1984), xylitol production may serve as a non-energy-demanding redox valve to dispose excess reducing power during oxygen limited conditions. The xylose reductase from these organisms has a dual cofactor specificity, and the reduction of xylose may take place with either NADH or NADPH. Thus, there will be competition for NADH between the glycerol 3-phosphate dehydrogenase and the xylose reductase, giving parallel production of glycerol and xylitol. The production of xylitol is energetically favourable compared with the production of glycerol from glucose which is energetically unfavourable since it occurs at the expense of one ATP. Production of glycerol involves reduction of the ATP phosphorylated dihydroxy acetone phosphate to glycerol 3-phosphate via glycerol 3-phosphate dehydrogenase and dephosphorylation of glycerol 3-phosphate by glycerol 3-phosphatase, while the production of xylitol only involves the reduction of xylose by XR without any ATP- consuming phosphorylation step.

A recombinant strain of S. cerevisiae incapable of glycerol production, due to deletion of the genes GPDl and GPD2, and transformed with XYLI encoding an NAD(P)H-consuming XR from P. stipitis, was found capable of anaerobic glucose conversion in the presence of xylose (Lid& et aL, 1996). This result indicates that the XR can fulfil the role of the glycerol 3-phosphate dehydrogenase reaction as a redox sink. A similar GPDl and GPD2 mutant containing transhydrogenase from E. coli instead of XR from P. stipitis was not capable of anaerobic glucose conversion (unpublished results) indicating a general excess of reducing power during oxygen- limited conditions which not can be alleviated by converting NADH to NADPH.

All xylose utilising yeasts require micro-aerobic conditions for xylose metabolism. In the case of yeast with NADPH-dependent XR and NAD+-dependent xylitol dehydrogenase (XDH), this has been ascribed to a disturbance in the redox balance in the cell caused by inefficient oxidation of NADH produced by XDH (Bruinenberg

13 el al., 1983a). However, oxygen is also essential for efficient xylose utilisation by yeast where XR has a dual cofactor specificity and is able to use both NADH and NADPH. In these yeasts, e.g. Pichia stipits, Pachysolen tannophilus, Candida shehatae and Candida tenuis (Watson et al., 1984; du Preez et al., 1989), XR should be able to reoxidise the NADH produced in the XDH reaction, and the conversion of xylose to xylulose should be a redox neutral process. With decreasing oxygenation, xylitol excretion increased markedly with C. shehatate and C. tenuis, indicating insufficient reoxidation of NADH by XR. In P. tannophilus the addition of hydrogen acceptors decreased xylitol formation and improved ethanol production (Ligthelm et al., 1989), indicating that xylitol formation in P. tannophilus results from the increased NADH formation, due to low consumption of NADH by XR. However, the degree of xylitol production by P. stipitis is much lower than other xylose utilising yeast strains (Skoog et al., 1992). This could be due to the presence of an alternative oxidase in P. stipitis working as an NADH-oxidising redox sink (Jeppsson et al., 1995).

3.3 Redox balance in S. cerevisiae strains expressing XYLl and XYL2 genes

In metabolically engineered S. cerevisiae expressing XYLl and XYL2 genes, encoding XR and XDH, respectively, from P. stipitis, xylitol is excreted as the major product in aerobic and anaerobic xylose fermentation (Kotter and Ciriacy, 1993; Walfridsson et al., 1995). Under aerobic conditions, biomass is the main product accompanied by xylitol formation (Walfridsson et al., 1995). With decreasing oxygenation, the formation of biomass is reduced and the xylitol production is increased. Under anaerobic conditions, all the consumed xylose is converted to xylitol and no growth was observed (Walfridsson et al., 1995). These results may indicate a disturbed redox balance similar to that observed in natural xylose utilising yeasts. This assumption takes into account the fact that XR from P. sripitis preferentially uses NADPH as a cofactor instead of NADH and that the oxidation of xylitol to xylulose by XDH is limited by the availability of NAD+ (Figure 2).

XR has a K~NADPHof 0.0032 mM and a K~NADHof 0.040 mM (Rizzi et al., 1988). The intracellular concentrations of NADPH and NADH under anaerobic conditions have been determined to be 1.21 and 0.44 pmole per g cells (dry weight), respectively, in S. cerevisiae (Papers IV and V). Additionally, the ratio NADPH/NADP+is 5.26 and the ratio NADH/NAD+is only 0.15 (Papers IV and V). Under these conditions, XR preferentially utilises NADPH. Thus, xylitol excretion occurs to prevent the accumulation of NADH since NADH generated by XDH will not be completely reoxidised by XR. However, the addition of acetoin as a hydrogen

I 14 acceptor, which increases the NADH oxidation and enables anaerobic xylose fermentation in C. utilis (Bruinenberg et al., 1983a), did not enhance anaerobic xylose consumption in S. cerevisiae expressing XYLI and XYL2 (Eliasson, personal communication). Xylulose excretion has also been observed under aerobic and anaerobic conditions (Walfridsson and Meinander, personal communication) indicating bottlenecks in the metabolism below XDH. Thus, limitations other than the accumulation of NADH may affect the excretion of xylitol. It has recently been shown that insufficient expression of xylulokinase, the third step in the xylose metabolism, causes a metabolic bottleneck in the xylose utilising S. cerevisiae strains (Ho et al., 1998). Strains overexpressing XYLI and XYL2 from P. stipitis and gene encoding xylulokinase from S. cerevisiae showed high xylose consumption and ethanol production and low xylitol excretion under oxygen-limited conditions, compared with strains expressing only XYLI and XYL2.

Xylose Xylose

NAD(P)+ Xylitol XDH LNm+ NADH Xy Iuiose Xylulose

XK

Xylulose-SP

Figure 2. The initial step(s) in the xylose metabolism of yeasts and bacteria. Abbreviations: XR, xylose reductase; XDH, xylitol dehydrogenase; XK, xylulokinase; XI, xylose isomerase.

The excretion of xylitol was completely stopped and the formation of glycerol and acetic acid were reduced in xylose utilising S. cerevisiae strains cultivated in oxygen- limited conditions by expressing a lower level of XR than of XDH (Paper I). In addition, the xylose consumption rates increased with strains having a low XR:XDH ratio. This result may indicate that there is a correlation between the level of oxygenation and the level of expressed XR and XDH in the cell. The NADH produced by XDH, in the oxidation of xylitol to xylulose, must be reoxidised in the

15 mitochondria before a new xylose molecule can be reduced to xylitol by XR. If the activity of XR is too high, more xylitol will be produced than can be oxidised by XDH due to the limited availability of NAD+. Thus, low XR activity and high XDH activity with a well-balanced oxygenation level will prevent the excretion of xylitol. However, such a system may prevent the formation of xylitol under oxygen-limited conditions, while it will probably not prevent xylitol secretion under strict anaerobic conditions. Xylose utilisation under strict anaerobic conditions without xylitol excretion, will probably only be possible if the conversion of xylose to xylulose via xylitol is a redox neutral process.

The redox problem might be reduced by changing the cofactor preference of XR or XDH by using protein engineering. Site-directed mutagenesis of the conserved lysine-270 (Kostrzynska et al., 1998) and of cysteine residues (Zhang and Lee, 1997) in the P. stipitis xylose reductase has been carried out with varied results. The Lys270Met variant exhibited 4.3-fold higher affinity for NADH over NADPH and showed a 14-fold decrease in Km for xylose. However, the Lys270Met variant exhibited lower enzyme activity than the wild-type enzyme. The introduction of the potential NADP-recognition sequence GSRPVC of the alcohol dehydrogenase from Themoanaerobium brockii into the xylitol dehydrogenase of P. stipitis allowed the mutant enzyme to use both NAD+ and NADP as a cofactor with equal apparent Km values (Metzger and Hollenberg, 1995). This mutant could mediate growth on xylose minimal-medium plates when coexpressed with the XYLl gene but was not able to grow in liquid xylose medium.

Another way of reducing the redox problem is to introduce an artificial bifunctional enzyme with XR and XDH activity (Paper 111). By expressing an artificial bifunctional enzyme with xylose reductase and xylitol dehydrogenase activity in S. cerevisiae, xylitol channelling could be expected due to proximity effects, e.g., shorter transient time. Such an enzyme system would also create a shuttling reaction in which NADH/NAD+ are continuously recycled between the site of oxidation and the site of reduction favouring the consumption of NADH and preventing an accumulation of xylitol. The influence of such a bifunctional enzyme on the xylitol excretion and ethanol production in oxygen-limited conditions is discussed in Chapter 4.

16 3. Expression of the T. thermophilis xylA gene in S. cerevisiae

Xylitol excretion should be eliminated through direct conversion of xylose to xylulose by xylose (glucose) isomerase (Paper 11), which is the initial reaction in bacterial xylose metabolism (Figure 2). Xylose isomerase which catalyses the isomerisation of xylose to xylulose, requires no cofactors for activity. Therefore, heterologous expression of a bacterial xylose isomerase encoding gene in S. cerevisiae has the potential for efficient conversion of xylose to ethanol in a redox neutral process.

Xylose isomerases from different bacteria are tetramers or dimers of similar or identical subunits associated with noncovalent bonds (Chen, 1980). Each subunit has two non-identical binding sites for divalent cations such as Mg2+, Co2+ or Mn2+. One is in direct contact with the active site and the other is responsible for stabilisation of the enzyme by maintaining the ordered conformation, especially the quaternary structure of the enzyme (Whitlow et al., 1991; Cha et al., 1994). Only one eukaryotic xylose isomerase has been described (Kristo et al., 1996). The gene was cloned from barley, Hordeum vulgare.

The xylA genes encoding xylose isomerase from Actinoplanes missouriensis (Amore et al., 1989), Bacillus subtilis (Amore et al., 1989), Clostridium thermosulfirogenes (Moes et al., 1996), Escherichia coli (Sarthy et al., 1987), Luctobacillus pentosus (Hallborn, 1995) Streptomyces rubiginosus (Schriinder et al., 1996) and Thermus thermophilus (Paper II) have been expressed in S. cerevisiae. The xylA genes from A. missouriensis and B. subtilis were isolated by complementation of a xyk4 negative E. coli mutant. The coding regions of the xylA gene from A. missouriensis and B. subtilis were fused to the GAL1 and PDCI promoter and terminator, respectively. The S. cerevisiae transformed with the xylA gene from A. missouriensis produced no protein but showed the presence of xylose-isomerase-specific mRNA. Strains expressing the xyL4 gene from B. subtilis produced xylose isomerase protein but it was catalytically inactive. Similar result was obtained when S. cerevisiae was transformed with the xyL4 gene from E. coli, fused to the yeast ADHl promoter and terminator. Expression of the C. thermosuljkogenes and L. pentosus XyEA genes under the control of A DH2 and PGKI promoter and terminator, respectively, produced no protein or xylose isomerase activity.

Active xylose isomerase was obtained when S. cerevisiae was transformed with the xyL4 genes from S. rubiginosus (Schriinder et al., 1996) and T. thermophilus (Paper

17 11). In contrast to S. rubiginosus, which showed low xylose isomerase activity, the enzyme from T. thermophilus was highly active, showing a similar temperature and pH activity dependency to that of the native enzyme (Paper 11). The strain was able to utilise xylose when cultivated on a minimal medium containing xylose as the only carbon source. Ethanol was produced in addition to acetic acid and xylitol (Paper 11). The xylitol excretion, which is also seen in the reference strain containing plasmid without xylA, was probably caused by unspecific aldo-keto reductases with affinity for xylose. One aldo-keto reductase has been purified from S. cerevisiae with a Kmxyloseof 27.9 mM (Kuhn et al., 1995).

The failure to express xylA genes from bacteria has been ascribed to the difference in codon preference between S. cerevisia and bacteria. However, among the bacterial xylA genes which have been cloned in S. cerevisiae, the T. thermophilus codons are the least preferred by S. cerevisiae, indicating that codon usage is of minor importance for the successful expression of xylA genes in S. cerevisiae.

Table 1. Families or groups of bacterial xylose isomerase based on their amino acid sequences. Asterisks indicate organisms whose xylA genes were cloned in S. cerevisiae.

Family I Family I1

Actinoplanes missouriensis * Bacillus lichenifomis Ampullariella sp Bacillus megaterium Arthrobacter sp Bacillus subtilis* Streptomyces albus Clostridium thermosaccharolyticum* Streptomyces murinus Escherichia coli* Streptomyces olivochromogenes Klebsiella aerogenes Streptomyces rubiginosus* Lactobacillus brevis Streptomyces violaceoniger Lactobacillus pentosus* Thermus aguaticus Staphylococcus xylosus Thermus hemophilus* Themtoga neapolitana Thermoanaerobacter etanolicus Thermoanaerobacter saccharolyticum Thermoanaerobacter thermosulfurogenes The success in producing an active xylose isomerase may depend on which group the :nzyme belongs to (Paper 11). Xylose isomerases are divided into two families Vieille et ai., 1995) or into two clusters within the same family (Meaden et al., 994). The xylose isomerases of A. missouriensis, S. rubiginosus and T. hermophilus belong to the same group (Family I), while the other group (Family E) ncludes the enzymes from B. subtdlis, E. coli, C. thermosulfurogenes and L. pentosus Table 1). The enzymes in the first group are less similar and lack a region of 30 to 40 unino acids at their amino terminus, which are present in the enzymes belonging to he second group. In addition, the G+C content of the DNA is higher in the first group han to the second group. However, in spite of the low homology between the first md second groups, the amino acids involved in the substrate and metal ion binding, LS well as in catalysis, are completely conserved (Bhosale et al., 1996).

The two xylA genes successfully expressed in S. cerevisiae belong to the first group. ill attempts to express xyM genes in the second group have failed, indicating that the :ylose isomerases from the first group are somehow more closely related to ucaryotic enzymes. However, the only known eucaryotic xylose isomerase, Yordeurn vulgare, is more related to the enzymes in the second group, indicating that )ther factors than amino acid similarity are important for successful expression in S. *erevisiae.

19 4. Artificial bifunctional enzymes

4.1 Channelling in metabolic pathways

A living cell is a very complex entity in which most of the biochemical reactions are organised into multistep pathways constituted by a sequence of enzymes through which a substrate is converted into an end product, e.g. the pyruvate dehydrogenase complex (Reed, 1974) and the pentafunctional AROM protein (shikimata pathway) (Lamb et al., 1992). The enzymes often operate sequentially in a reaction pathway; the product of the first enzyme serving as the substrate for the next enzyme, whose product will serve as the substrate for the subsequent enzyme. In eukaryotic cells many of the soluble enzymes of a metabolic pathway are compartmentalised in organelles such as the nucleus, the mitochondrion, the endoplasmatic reticulum, the Golgi apparatus or the chloroplast. The enzymes may be membrane bound (Harris and Winzor, 1990) or attached to the cytoskeleton F-actin proteins (Walsh and Knull, 1988) in an organised form operating in an ordered, non-random way (Srere, 1987; Knull and Walsh, 1992).

The transfer of the intermediates in a reaction pathway occurs either by free diffusion (Gutfreund, 1972) or by channelling (Keleti et al., 1989; Srivastava and Bemhard, 1986). If the enzymes of a metabolic pathway can not bind to each other in solution, then the intermediates must be transferred through solution via diffusion. If the producing and consuming enzymes do have the capacity to bind to each other or localise each other by protein-protein, protein-membrane, protein-DNA or protein- polysaccharide interaction (Srere and Ovadi, 1990), then a channelled metabolite transfer mechanism may be operative. The enzyme complexes formed might have properties such that the intermediately formed metabolite can be directly transferred from one enzyme to another, without prior release into solution. Such a direct transfer is usually referred to as channelling and has been defined as the transfer of metabolites between two enzymes without complete equilibrium with the surrounding bulk solution (Ovadi, 1991).

The arrangement of sequentially operating enzymes into multi-enzyme complexes and multifunctional enzymes could offer several catalytic advantages (Ovadi, 199 1). (i) Channelling prevents loss of intermediates by diffusion. (ii) Channelling decreases the transit time required for an intermediate to reach the active site of the next enzyme. (iii) Channelling protects chemically labile intermediates.

20 (iv) channelling circumvents unfavourable equilibria by creating a higher local concentration of the intermediate around the enzyme complex. (v) Channelling segregates the intermediates of competing enzymatic reactions. (vi) Channelling reduces the transient time for the system to reach the new steady state. The transient time is the time required to attain a steady state. A shorter transient time means that the intermediate metabolite does not equilibrate with the bulk solution so locally higher concentration of the intermediate around the next enzyme is expected. (vii) Channelling may lower the concentration of an intermediate due to sequestering of the intermediate between enzymes.

4.2 Metabolite transfer mechanisms

There has been, and still is, much discussion about the molecular mechanism of channelling but the processes belived to be behind have been divided into two groups: direct transfer processes and proximity effects (Spivey and Merz, 1989). In the "direct transfer process", the intermediate metabolites are passed directly from the first enzyme to the second enzyme during transiently formed enzyme-enzyme interactions. Classical examples of such complexes are the mammalian fatty acid synthases (Wakil, 1989) and 2-oxoacid dehydrogenase (Perham, 1991). In these cases, the intermediates are covalently bound to the complex. Another classical example of direct transfer is tryptophan synthase (Anderson et al., 1991). In this case, there is no covalent interaction of the intermediate with the complex, but crystallographic analysis of the structure has revealed the presence of a tunnel through which the intermediate can be transferred (Hyde et al., 1988).

Evidence of electrostatic channelling has also been reported for bifunctional dihydrofolate reductase-thymidylate synthase (Knighton et al., 1994; Trujillo et al., 1996) and for an artificial bifunctional malate dehydrogenase-citrate synthase (Elcock and McCammon, 1996). The X-ray structure of dihydrofolate reductase-thymidylate synthase indicates that the transfer of dihydrofolate between the active sites does not occur by transient binding at both sites, but rather by movement of dihydrofolate across the surface of the protein. The enzyme has an unusual surface charge distribution which could account for channelling between the active sites (Knighton et al., 1994).

Direct transfer between soluble enzymes in glycolysis has been reported in systems involving pairs of enzymes, such as glycerol-3-phosphate dehydrogenase, lactate dehydrogenase, alcohol dehydrogenase and glyceraldehyde-3-phosphate

21 dehydrogenase (Keleti et al., 1989; Neuzil et al., 1990; Weber and Bernhard, 1982; Srivastava and Bernhard, 1985). The design of the experiments and interpretation of the data have been questioned (Chock and Gutfreund, 1988; Gutfreund and Chock, 1991; Wu etal., 1991; Arias, 1997).

The ”proximity effect” is proposed to exist for any coupled enzyme reaction when the second enzyme is locally concentrated near the first enzyme, creating a favourable micro-environment in which the local concentration of the intermediate is higher than in the surrounding bulk medium. This situation is believed to take place when enzymes are aggregated together in complexes, e.g. pyruvate dehydrogenase complex (Reed, 1974) and when enzymes are closely bound to cellular structures, e.g. glycolytic enzymes associated with cytoskeletal proteins (Knull and Walsh, 1992). The local increase in enzyme concentration leads to an increased “kinetic power”, kat[E]~/Km,where [E]T is the total enzyme concentration, and therefore results in a more efficient pathway (Keleti and Welch, 1984). Most attention has been focused on the tricarboxylic acid cycle in the mitochondrial matrix. It has been suggested that the enzymes in the Krebs cycle behave as a multi-enzyme entity, rather than free enzymes in solution (Srere, 1987; Robinson et al., 1989). NMR studies on yeast cells (Sumegi et al., 1990; Sumegi et al., 1993) and mutagenesis of citrate synthase (Kispal et al., 1989) have provided evidence that proximity effects do exist in the Krebs cycle. Another example where proximity effects have been demonstrated is in the pentafunctional AROM protein which catalyses the first five steps in the biosynthesis of aromatic amino acids of fungi and yeast (shikimata pathway) (Lamb et aZ., 1992).

4.3 Artificial enzyme systems

Proximity between two or more enzymes can be artificially generated by making an in-frame gene fusion of the corresponding structural genes. The translational stop codon of the first structural gene is removed, followed by in-frame ligation with the second structural gene. This procedure, which results in a novel chimaeric gene encodes a single polypeptide chain carrying two or more active centres.

The first example of the successful artificial fusion of two enzymes at the genetical level was the fusion of two enzymes which are involved in the biosynthesis of histidine (Yourno et al., 1970). In another example, the lac repressor from the p- galactosidase operon was fused in frame with &galactosidase, without altering the individual activities of the proteins (Muller-Hill and Kania, 1974). Two sequentially operating enzymes, P-galactosidase and galactokinase, were structurally linked to study whether the proximity of the enzymes in vitro affected the reaction kinetics in

22 the conversion of lactose to galactose-I-phosphate (Biilow et al., 1985). A small proximity effect was detected which was more pronounced when polyethylene glycol was added to the solution. The galactose dehydrogenase-bacterial luciferase enzyme system exhibited both galactose dehydrogenase activity and twofold higher bioluminescence in vitro, than a corresponding reference system with separate native enzymes (Lindbladh et al., 1992). Similar in vitro results were obtained when galactose dehydrogenase was fused in frame with P-galactosidase (Ljungcrantz et al., 1989). This complex exhibited a markedly decreased transient time and a higher steady state rate than a corresponding reference system of native enzymes, indicating substrate channelling.

To create channelled metabolite transfer in the two first sequential steps of the xylose metabolism in S. cerevisiae, two genes of Pichia stipitis, XYLl and XYL2 encoding xylose reductase (XR) and xylitol dehydrogenase (XDH), were structurally linked in frame. These fusion proteins exhibited both XR and XDH activities in cell-free extracts from S. cerevisiae, and strains expressing the proteins were observed to excreted less xylitol during anaerobic xylose fermentation than strains expressing native XR and XDH, indicating proximity effects (Paper III).

4.4 Subunit interactions and linker regions in bifunctional enzymes

The properties of artificial bifunctional enzymes depend primarily on the gene fusion accuracy. If the entire primary sequences of the native enzymes are maintained in the fusion enzyme, the enzymes should retain their native specific activities. However, low or no enzyme activities of artificial fusion proteins consisting of oligomeric enzymes with several subunits have been reported. The fusion between the tetrameric enzymes glyceraldehyde-3-phosphate dehydrogenase from E. coli and lactate dehydrogenase from Bacilus stearotthermophilus did not result in functional complexes with both enzyme activities (Carlsson, 1993). Similarly, when the intact cDNA of mouse pancreatic a-amylase was fused to Bacilus subtilis glucoamylase in both possible orders, no glucoamylase activity was found (de Moraes et al., 1995). However, low levels of a-amylase activity were found when the fusion had this domain at its amino terminus. Similarly, the specific activities of XR and XDH, in strains expressing fusion proteins composed of the enzymes XR and XDH, were also dependent on the order of the enzymes in the hybrid polypeptide chain (Paper In). Constructs with XDH at the N-terminus and XR at the C-terminus of the final gene product (Figure 3 B,C,D), had XR and XDH activities while a construct with XR at the N-terminus and XDH at the C-terminus lack XR activity (Figure 3 A). The

23 juxtaposition of XR and XDH in this fusion protein may result in missfolding of the catalytic domain of XR or unfavourable interactions between the chimeric subunits.

A. XR Linker XDH --1 --1 Phe Val Gln Ser Arg Met H 317 318 12Thr B. XDH Linker XR --1 --, Y Pro Glu Gly Ser Arg Met Pro 362 363 12 C.

XDH rl Linker XR a bl Pro Glu Gly Ser Arg Asn Gln Met Pro 362 363 Thr 12 D.

XDH d Linker -1 XR 4 Pro Glu Gly Ser Arg Pro Ser Pro Thr Ala Ser Thr Asn Gln Met Pro 362 363 12 Figure 3. Scheme of linker region in bifunctional enzymes with XR and XDH activity. A, B, C, and D show the order of the enzymes (XR and XDH) in the hybrid polypeptide chain and the length and composition of the connecting region in different bifunctional enzymes.

It is likely that enzymes composed of several subunits can not form optimal conjugates in terms of subunit interactions when the chimeric subunits associate to form an active conjugate. In addition, intracellular expression of fusion proteins composed of two oligomeric proteins might generate extensive polypeptide chain interactions, resulting in inactive, soluble three-dimensional protein networks or insoluble inclusion bodies. Expression of protein A-0-galactosidase fusion protein in E. coli resulted in intracellular inclusion bodies (Hellebust et aE., 1989). However, the amount of inclusion bodies depended on the cultivation conditions.

One way to circumvent the constraints of proper subunit interactions in fusion proteins is to vary the length of the linker in the hybrid polypeptide chain. The linker was important for the specific activity of the XDH part of the xylitol dehydrogenase- xulose reductase fusion protein complexes (Paper III) and the introduction of a longer linker between XR and XDH (Figure 3) appeared to relax the intramolecular strain in the fusion protein, as demonstrated by an increase in the specific activity of XDH. The highest activity was obtained with a linker of 12 amino acids (Figure 3 D). The activity of the XR part of the enzyme was not affected to the same extent as the XDH

24 part. Similar results have been obtained in an E-2 and IL-6 lymphokine fusion protein (Rock et al., 1992), in a human granulocyte/macrophage colony-stimulating factor and interleukin 3 fusion protein (Curtis et al., 1991) and in a P-galactosidase- galactose dehydrogenase fusion protein (Carlsson et aL, 1996). The opposite effect has been reported for the fusion protein metallothionein (Rhee et al., 1990) where the level of activity decreased with the length of the linker.

The amino acid composition of the linker region has proven to be important for the stability and for the function of the artificial fusion protein. Linkers consisting of multiple proline and glycine residues have been introduced between P-galactosidase and galactokinase enzymes at the gene level (Carlsson et al., 1992) to give the connecting region a rigid or a flexible character, respectively. These proteins with poly-proline and poly-glycine linker regions were not expressed to the same degree as fusion proteins with linkers consisting of randomly chosen amino acids. In addition, proteolytic degradation has been observed in linker regions consisting of multiple glycine and proline residues (Carlsson, 1993). Degradation of the linker region in the fusion protein, protein A-P-galactosidase, has also been reported (Hellebust et al., 1988), whereas proteolytic degradation of linker regions of different lengths and compositions in fusion proteins composed of XR and XDH (Paper JII) expressed in S. cerevisiae was not observed, probably due to high contents of non-hydrophobic amino acids (Figure 3). It has been shown that non-hydrophobic amino acids in the linker region prevent proteolytic degradation of the protein (Argos, 1990).

Enzyme functions can be further improved by expressing native subunits and the corresponding chimeric subunit in the same cell (Lindblad, 1992). The native subunits may then exchange with chimeric subunits to form more favourable interactions between the monomers, generating a ”second generation” fusion protein complex consisting of chimeric and native subunits. This approach has been used to create P-galactosidase/galactosedehydrogenase enzyme complexes with 30% higher galactose dehydrogenase activity than the fusion enzyme (Lindblad, 1992). By using this technique, the relative XR and XDH activities of the fusion protein complexes, composed of the fusion protein and native monomers of XR and XDH, were increased 10 and 9 times, respectively, compared with the original fusion protein complexes produced in strains overexpressing fusion protein only (Paper In). In addition, the expression of XR and XDH monomers prevented formation of higher order aggregates of the fusion protein complexes so that smaller complexes (190 and 230 kDa) were formed than when only the artificial fusion protein was expressed (300,450 and 600 ma).

25 4.5 Proximity studies in vivo

The use of artificial fusion proteins is not only restricted to studies directed towards the understanding of the enzyme organisation and proximity effects in the cell. The technique can also be used for metabolic engineering. By fusing enzymes at metabolic branch points, the intermediate can be efficiently transferred within the artificial enzyme and thereby be guided to the desired pathway. Proximity effects in vivo have been reported for E. coli cells with a scavenger enzyme, galactose dehydrogenase, competing with the galactokinase part of the p-galactosidase- galactokinase fusion protein for galactose formed by p-galactosidase (Carlsson, et aZ., 1992). A similar in vivo effect has been observed in the TCA cycle when fused citrate synthase and malate dehydrogenase was expressed in mutants of S. cerevisiae in which the genes for citrate synthase and malate dehydrogenase had been deleted (Lindbladh et al., 1994a,b; Elcock and McCammon, 1996). The malate dehydrogenase-citrate synthase fusion protein exhibited a shorter transient time for the coupled reaction sequence than for the free enzymes. Bovine cytochrome P-450 and yeast NADPH cytochrome P-450 reductase have been fused and introduced into yeast cells (Shibata et al., 1990). This strain showed increased hydroxylation activity towards progesterone as a result of more efficient electron transfer between the enzymes. An a-amylase-glucoamylase enzyme system showed higher levels of activity than mixtures of the two individual enzymes (Shibuya et al., 1992; de Moraes et al., 1995).

26 ...... : JCVLtlSE : : : :xvLuc6sE: : ...... ::: : :::: ...... :) ...... -...... ::: -:::: ::::::: (b......

Figure 4. Schematic representation of the artificial bifunctional fusion protein with xylose reductase and xylitol dehydrogenase activities. The dotted part represent region with higher local concentration of xylitol, NAD'and NADH than the bulk solution. i. cerevisiae was metabolically engineered by expressing fusion proteins composed )f XR and XDH (Figure 4) to create a channelled metabolite transfer in the first two iequential steps of xylose metabolism (paper In). The formation of the intermediate cylitol during anaerobic xylose fermentation decreased, indicating a more efficient )athway. Xylitol formation is ascribed to a redox imbalance created by the fact that he first enzyme, XR, prefers NADPH as cofacter whereas the second enzyme, XDH, :xclusively uses NAD+. In addition, less acetic acid and more ethanol were formed. The decreased xylitol formation indicates proximity effects created by the fusion )rotein. The proximity effects probably circumvent the unfavourable equilibria in the irst two steps of the xylose metabolism by creating a favourable micro-environment where the local concentrations of xylitol and NADH around XDH and XR are higher han in the surrounding bulk medium. The higher local concentration of xylitol and VADH leads to an increased flux through XDH and increased consumption of NADH )y XR. The enzyme system creates a shuttling reaction in which NADH/NAD+ is :ontinuously recycled between the site of oxidation and the site of reduction Bvouring the consumption of NADH instead of the preferred cofactor NADPH and ireventing the accumulation of xylitol.

27 5. Nicotinamide nucleotide transhydrogenase

5.1 Classes of nicotinamide nucleotide transhydrogenase

Nicotinamide nucleotide transhydrogenase (EC 1.6.1.1) was discovered in Pseudomonas jluorescens (Colowich et al., 1952). Investigations on isocitrate dehydrogenase in this bacteria led to the discovery that NAD+ could be reduced by isocitrate provided a catalytic amount of NADP+ was added to the cell extract. It was proposed that an enzyme, nicotinamide nucleotide transhydrogenase, catalysed the formation of NADP+ and NADH from NAD+ and NADPH without the mediation of any other substrates. Determination of the stoichiometry of oxidised and reduced nicotinamide nucleotides, respectively, indicated the following reaction;

NADH + NADP+ <-> NAD+ + NADPH

Indirect transfer of the hydrogen and transfer of the phosphate or nicotinamide moieties could be eliminated, suggesting that the reduction of NAD+ by NADPH was direct and stereospecific. It was later demonstrated that a similar activity was present in animal tissue (Kaplan et al., 1953). In contrast to the soluble enzyme from Pseudomonas jluorescens, the enzyme was insoluble and bound to the mitochondrial membrane. Differences between the soluble and insoluble transhydrogenases, with respect to kinetic and regulatory properties, indicated that the enzymes were unrelated.

Based on their difference with respect to the stereospecificity of the hydrogen transfer at position C4 of the nicotinamide ring of NAD(P)H, the two enzyme groups were later denoted BB- and AB- transhydrogenase. The nicotinamide ring of NAD(P)H is asymmetric because of the carboxamide substituent at its C3 atom. The two hydrogen atoms at C4 in the reduced form of NAD(P)+ are therefore not equivalent. The 4-A hydrogen is located above the plane of the ring while 4-B is located below the ring. The BB-transhydrogenases transfer the 4-B hydrogen of NADH and NADPH to NADP+ and NAD+, respectively, whereas the AB-transhydrogenases transfer the 4-A hydrogen of NADH and the 4-B hydrogen of NADPH to NADP+ and NAD+, respectively. The BB-transhydrogenases are exclusively bacterial enzymes. They are easy to solubilise, are flavoproteins containing FAD, and have a single catalytic site for both NAD(H) and NADP(H). They react according to a binary ping-pong mechanism. The AB-transhydrogenases occur in the mitochondria of eucaryotic organisms and certain bacteria, including respiring and photosynthetic bacteria. They

28 are membrane: bound and are not flavoproteins, have separate catalytic sites for NAD(H) and 1NADP(H), and react according to a ternary complex mechanism. AB- transhydrogenases (energy linked transhydrogenase) are functionally linked to the energy -transfer system of the bacterid and mitochondrial membrane (Danielsson and Ernster, 1963)I, while BB-transhydrogenases (non-energy linked transhydrogenase) are not related to energy generated by respiration, ATP or light.

5.2 AB-traIns hydrogenases

AB-transhydrc lgenases are found in the inner membranes of animal mitochondria (Danielsson arId Ernster, 1963) and plant mitochondria (Carlenor et al., 1988) and in the cytoplasmic membranes of many bacteria (Fisher and Earle, 1982). They catalyse the reversible transfer of a hydride ion equivalent between NAD(H) and NADP(H) and are coupled to the proton motive force. The reaction can be summarised as;

NADH + NADP + nH+out <-> NAD+ + NADPH + nH+in (2)

where "n " deriotes the number of protons pumped across the membrane and "in" and "out" denotes the matrix and intermembrane space, respectively, of mitochondria, or cytoplasm and periplasmic space, respectively, of bacteria (Figure 5). The number of protons pump:d across the membrane has been determined to be close to unity (Earle and Fisher, 1980; Eytan et aE., 1987a). The reaction from left to right is the predominant dlirection of reaction (2) in the intact mitochondrion and bacterial cell, driven by the existing electrochemical proton potential (Ap) (Eytan et al., 1987a,b; Hu et al., 199:5). In the absence of a proton motive force, reaction (2) proceeds to an equilibrium pc )int with Keq = 0.79, in agreement with the difference in the reduction potentials of 1JADH/NAD+ (E'F -320 mV) and NADPH/NAPD+(E'o= -325 mV) (Olson and An Ifinsen, 1953; Kaplan et aL, 1953).

The transhydrogenase reaction may also be reversed so as to generate, for instance ATP, but onljI at very high [NADPH] [NAD+]/[NADP+] [NADH] ratios (Van de Stadt et d.,1 97 1). However, despite repeated attempts, reconstituted transhydro- genase-ATPai se vesicles have not been found to generate ATP under similar conditions (01ausson et al., 1995).

29 CYTOPLASM t

Plasma membrane

nH PERIPLASM

Figure 5. Ilustration of E. coli transhydrogenase showing the forward reaction. The a-subunit contains the NAD(H) site (dotted part) and transmembrane helices 1-4 (grey part), and the P-subunit the NADP(H) site (dotted part) and transmembrane helices 5-10 (grey part).

Transhydrogenase has been purified from bovine mitochondria (Anderson and Fisher, 1978), E. coli (Clarke and Bragg, 1985) and Rhodobacter capsulatus (Lever et al., 1991), and amino acid sequences based on the DNA sequences of those from man and mouse (Lagberg Arkblad et al., 1996), ox (Yamaguchi et al., 1988), E. coli (Clarke et al., 1986), Rhodospirillum rubrum (Yamaguchi and Hatefi, 1994; Williams et al., 1994), Eimeria tenelZa (Kramer et al., 1993; Vermeulen et al., 1993), and Entamoeba histolytica (Yu and Samuelson, 1994) have been published. The E. coli (Clarke and Bragg, 1985) and R. rubnun (Diggle et al., 1995) transhydrogenase genes have been overexpressed in E. coli.

Saccharomyces cerevisiae was transformed with the gene encoding transhydro- genases from E. coli (Paper IV) and showed high transhydrogenase activity. This is the first time a transhydrogenase has been successful expressed in an eucaryotic system. An attempt to express the transhydrogenase gene from humans in S. cerevisiae was also made but no activity was observed in cellfree extracts (unpublished results).

30 The human and mouse transhydrogenases are identical to the bovine protein in length (Lagberg-Arkblad et al., 1996). All three mammalian proteins have a single gene product in contrast to the bacterial enzymes which have 2-3 gene products (Olausson et aL, 1995) (Figure 6). Based on the high similarity between bovine, human and mouse transhydrogenase it has been concluded that all mammalian transhydrogenases have a similar structure (Lagerberg Arkblad et al., 1996). The mammalian transhydrogenases are homodimers with a monomer molecular mass of 110 kDa. The monomer is composed of three domains (Olausson et al., 1995); a 433 residue hydrophilic N-terminal domain I, which is extramembranous and binds NAD(H); a 407 residue hydrophobic central domain 11, which is largely integrated in the membrane and harbours the enzyme's proton channel; and a 204 residue hydrophilic C-terminal domain 111, which is extramembranous and binds NADP(H) (Figure 6). Domains I and Ill extend into the mitochondrial matrix, where they presumably come together to form the catalytic site of the transhydrogenase.

NAD(H)-binding Transmembrane NADP(H)-binding domain domain domain 1 432 839 1043 Human

a1 a404 a510 pl p260 E. coli

Figure 6. Linear representation of the transhydrogenase from humans and E. coli. The blocks represent the various domains and gene products. Transhydrogenase from humans consists of 1043 amino acids in a single gene product. Transhydrogenase from E. coli has two gene products, a and p, which consist of 404 and 462 amino acids, respectively. (Modified from Olausson et al., 1995).

The transhydrogenase of E. coli has the same overall tridomain structure as the mammalian transhydrogenases (Figure 6) and similar to mammalian transhydro- genases, domains I and 111 extend into the cytoplasm where the form the catalytic site of the transhydrogenase (Figure 5). However, the E. coli transhydrogenase is composed of two subunits; a, with a molecular mass of 54 ma, and p, with a molecular mass of 49 kDa, and has been shown to be an heterotetramer (Hou et al., 1990). The 3-dimensional structure of the NAD(H) binding site of the E. coli transhydrogenase, which is located in domain I of the a subunits, has been predicted (Fjellstrom et aZ., 1995). Hydropathy analyses suggest that domain I1 could be composed of a maximum of 14 a helices, but prediction algorithms (Holmberg et al.,

31 several times. The explanation for this is that the binding and release of NADP(H) from transhydrogenase are intimately involved in the proton translocation reaction. The binding of NADP+ is accompanied by the uptake of a proton at the outside of the bacterium or mitochondrion. After subsequent transfer of H- from NADH to NADP+, the release of NADPH is accompanied by the release of the proton to the inside of the bacterium or mitochondrion. The effect of pH on NADP(H) binding, suggests that a single acid or basic residue is responsible for proton binding and proton release. This single amino acid residue has been identified to be J3 His-91 in E. coli (Hu et uL, 1995; Glavas et al., 1995; Bragg and Hou, 1996).

// 'NAD+

'NADH 'NADH

Figure 8. A model of the intermediate conformational states of the transhydrogenase reaction. "Out" denotes the periplasmic space, "In" denotes the cytosol and "His" denotes p His-91. The shift of the histidine residue should be regarded as accessibility to either side of the membrane, rather than a physical movement. The sequence of steps in proton-translocating transhydrogenation are indicated by numbers 1 to 6. (Modified from Hu et ul., 1995).

In the model (Figure S), NADH and NADP+ or NAD+ and NADPH, determine on which side of the membrane J3 His-91 is exposed and also the pKa of J3 His-91. When NAD+ and NADPH are bound to the protein it is proposed that J3 His-91 is directed towards the cytosolic side and buried in the hydrophobic phase of the membrane, thus increasing its pKa. When NADH and NADP+are bound, it is proposed the p His-91 is directed to the periplasmic space side and becomes freely accessible to the bulk solute, leading to a normal pKa. In agreement with this model, the pH dependency for the reduction of AcPyAD+ by NADH supported by NADP(H) is a consequence of the assumed lack of deprotonation at low pH of p His-91. At low pH NADP(H) will remain bound to the enzyme and the kinetics of the reduction of AcPyAD+ by NADH

33 supported by NADP(H) will therefore be a ping-pong mechanism. The kinetics of the normal transhydrogenase reaction (2) at neutral pH is that of a random bi-bi mechanism.

5.3 BB-transhydrogenases

The second class of transhydrogenases constitute water-soluble B B -specific transhydrogenases. These enzymes are flavoproteins and have been isolated in pure form from several bacterial sources. The enzymes isolated from Pseudomonas aeruginosa (Cohen and Kaplan, 1970; Hojeberg et al., 1976) and Azotobacter vinilandii (Middleditch et aL, 1972) have been studied most extensively. Both consist of a single polypeptide chain of molecular weight 54 kDa (Wermuth and Kaplan, 1976; Hojeberg et al., 1976).

The purified enzyme of A. vinelandii is present in solution at neutral pH as a polydisperse mixture of threads or rods with a width of 11- 12 nm and a maximum length of 1000-2000 nm (Voordouw et al., 1979; Voordouw et al., 1980a). Similar polymers are present in purified preparations of P. aeruginosa transhydrogenase with lengths ranging from 20 nm to 500 nm and a reported diameter of 8-10 nm (Louie et al., 1972; Hojberg et al., 1976). These polymers dissociate at alkaline pH to an octamer consisting of eight subunits of relative molecular mass 54 kDa (Voordouw et al., 1982). This octamer is believed to be the smallest functional unit. It is not known whether the octamer/polymer interconvertion fulfils a role in vivo. Studies on the enzyme in cell-free extracts indicate a limited degree of polymerisation (Voordouw et al., 1979) or even absence of polymers (Voordouw et al., 1980b). Amino acid analysis (Wermuth and Kaplan, 1976; Middleditch et al., 1972) reveals a similar amino acid composition for both enzymes.

The gene encoding transhydrogenase from Azotobacter vinilandii was cloned and the nucleotide sequence was determined (Paper V). The deduced amino acid sequence shows high similarity to the transhydrogenase from Pseudomonas fluorescens (French et al., 1997) and to a dehydrogenase with unknown function from E. coli (Gustafsson and Warne, 1992). The enzyme shows high similarity to the flavoproteins in the disulfide oxidoreductase family (Schinner and Schulz, 1987), particularly dihydrolipoamide dehydrogenase, mercuric ion reductase, glutathione reductase, and trypanothione reductase. Both the transhydrogenases from A. vinelandii (Paper V) and P. fZuorescens (French et al., 1997) contain consensus patterns typical for this family but lack one of the conserved redox-active cysteine residues in their active sites (PaperV). The enzymes of the disulfide oxidoreductase

34 family consist of three domains; an N-terminal flavin-binding domain containing the redox-active disulphide bond characteristic of this family, a central NAD(P)-binding domain and a C-terminal dimerisation domain. The enzymes have a high degree of sequence similarity in the FAD- and NAD(P)H-binding regions of the amino-acid chain (McKie and Douglas, 1991).

Kinetic and fluorometric studies (Rydstrom et aZ., 1987) have shown that transhydrogenases from A. vinilandii and P. aeruginosa undergo marked allosteric changes in the presence of AMP, NADPH, Ca2+ and NADP+. Whereas AMP NADPH and Ca2+ stimulate the rate of reduction of NADP by NADH, NADP is strongly inhibitory and NAD(H) is without effect. The rate of reduction of NAD+ by NADPH is maximal in the absence of effectors and is only marginally inhibited by NADP+ and AMP.The mechanism by which the enzyme is regulated by the effectors is unknown. Obvious possibilities are that the effects of NADP(H) and AMP on the activity are mediated through binding either to the catalytic site or to second separate regulatory site(s) (Hojeberg et aL, 1976; van den Broek et aL, 197 1). The existence of a second separate NADP+ site with a markedly different dissociation constant, in A. vinelandii transhydrogenase, is strongly supported by the results of spectral titration analysis and changes in the absorption spectrum at higher NADP+ concentrations (van den Broek et aL, 1971). Circular dichroism measurements also indicated that the two NADP+-enzyme complexes had different conformations.

This regulatory separate NADP(H) binding site was not found in the sequences of either P. fluorescens (French et al., 1997) or A. vinelandii (Paper V) However, certain of the binding sites in the native enzyme of A. vinelandii , consisting of eight subunits, may act as regulatory sites whereas others function as active sites. If this is true, then there could be two types of NAD(P) binding sites which are identical in amino acid sequence but which differ in secondary, tertiary or quaternary structure. The two structurally different binding sites may have different function and binding constant for NADP+. Such a mechanism was proposed to account for incomplete reduction of flavin in transhydrogenase from P. aeruginosa (Cohen and Kaplan, 1970).

35 5.4 Cloning and expression of AB- and BB-transhydrogenasesin S. cerevisiae

In mammals, an important role of AB-transhydrogenase is to produce NADPH for utilisation by the combined actions of intramitochondrial glutathione reductase and glutathione peroxidase to reduce H202, which results from the dismutation of 02 radicals produced by electron leak to oxygen along the respiratory chain (Chance et al., 1979). Via the actions of intra- and extramitochondrial isocitrate dehydrogenase, NADPH is exported and utilised in cytosolic biosynthetic reactions (Hoek and Rydstrom, 1988). Transhydrogenase, together with NAD(H)- and NADP(H)-linked isocitrate dehydrogenase may contribute to fine tune the tricarboxylic acid cycle activity in mitochondria (Sazanov and Jackson, 1994). In addition, it is thought that by producing NADPH, the transhydrogenase provides a protective buffer against the dissipation of cellular redox power and mitochondrial energy supply (Hoek and Rydstrom, 1988).

The function of AB-transhydrogenase in E. coli is to provide NADPH for the synthesis of glutamate (Liang and Houghton, 1981). It was found that high concentrations of NH4Cl induced transhydrogenase synthesis and that glutamate strongly repressed the synthesis of both transhydrogenase and glutamate dehydrogenase.

Bacterial BE-transhydrogenases have generally been believed to be involved primarily in NADPH oxidation, which is consistent with the preferential generation of NADPH rather than NADH in catabolic processes in bacteria containing BE- transhydrogenases (Voordouw, et al., 1983). For instance in A. vinelandii several catabolic steps are exclusively NADP+ linked e.g. isocitrate dehydrogenase, whereas others are linked to both NAD+ and NADP+ e.g. the enzymes of the Entner-Doud- oroff pathway. The Km values of the latter enzymes for NADP+ are generally about 10-fold lower than for NAD+ (Voordouw, et al., 1983). Thus, a catabolic over- production of NADPH could be foreseen in the absence of transhydrogenase.

Enzymatically catalysed pyridine nucleotide transhydrogenation has not been observed in yeast (Lagunas and Gancedo, 1973; Bruinenberg et al., 1983b; Bruinenberg et al., 1985, Papers IV and V). The lack of pyridine nucleotide transhydrogenation has considerable consequences for the redox balances of both coenzyme systems NAD(H) and NADP(H) and hence for sugar and amino acid metabolism in yeast (van Dijken and Scheffers, 1986; Albers et al., 1996). Each coenzyme system has to maintain a delicate balance between formation and consumption of reducing equivalents. The formation of NADPH occurs via central

36 metabolic routes, primarily in the pentose phosphate pathway (Lagunas and Gancedo, 1973; Nissen et al., 1997).

Most anabolic reductive reactions require NADPH rather then NADH. Since transhydrogenase is absent in yeast, the overall process of assimilation leads to the production of a considerable surplus of NADH (Oura, 1977; Albers et al., 1996). During aerobic growth this is balanced by the oxidation of NADH with oxygen in the mitochondria. In the absence of oxygen as an electron acceptor, glycerol is formed from dihydroxy acetone phosphate in glycolosis at the expense of one mole of NADH per mole glycerol.

To analyse the physiological effect of transhydrogenation between the two coenzyme systems, NADP(H) and NAD(H), during anaerobic growth, S. cerevisiae was transformed with a mitochondrial and a plasma-membrane-bound A B-transhydro- genase from a human (unpublished results) and E. coli (Paper IV), respectively, to study the effect of energy-linked transhydrogenase present in the mitochondria and in the cytoplasma. S. cerevisiae was also transformed with a cytoplasmic BB- transhydrogenase from A. vinelandii (Paper V) to study the effect of non-energy linked-transhydrogenase in the cytoplasma.

Our hypothesis was, that since NADH can be consumed and NADPH produced by a transhydrogenase, expression of the gene encoding transhydrogenase would result in a decrease in both glycerol formation and in the carbon flux through the pentose phosphate pathway, where there is a loss of carbon in the form of carbon dioxide.

The cDNA sequence encoding A B-transhydrogenase from human was subcloned from three plasmids containing three overlapping cDNA sequences. After PCR amplification, the complete structural gene was subcloned into a YEp24 plasmid containing the PGK promoter and terminator. The sequences of the PCR-amplified fragments were verified by DNA sequencing. S. cercevisiae was transformed with the YEp24 plasmid containing the cDNA sequence encoding the transhydrogenase under the control of the PGK promoter-terminator. Cell-free extracts from transformants showed no transhydrogenase activity. Western blot analysis, using transhydrogenase- specific polyclonal antibodies, showed no transhydrogenase in cell-free extracts from transformants containing the cDNA sequence encoding the transhydrogenase, indicating an unstable enzyme or mRNA (unpublished results).

S. cerevisiae transformed with the expression plasmid containing the gene encoding AB-transhydrogenase from E. coli showed transhydrogenation activity (Paper IV).

37 However, the protein was not transported to the plasma membrane but seemed to be accumulated in internal membrane systems, mostly in rough endoplasmic reticulum (ER) (Paper IV). It is known that missfolded or unassembled proteins tend to accumulate in ER and degrade rapidly (Schauer et al., 1985; Haguenauer-Tsapis et al., 1986; Hurtley and Helenius, 1989) but our work indicates a correctly folded and catalytically active recombinant protein.

Integral membrane proteins enter the secretory and endocytic pathways via insertion into the ER membrane (Sanders and Schekman, 1992). The pathway taken by yeast integral membrane proteins to the plasma membrane appears to at least partially coincide with that of soluble secretory proteins since sec mutations (Pryer et al., 1992), which block membrane traffic at various stages of the secretory pathway, prevent trafficking of both types of passenger proteins. New evidence indicates that the transport of integral membrane proteins to the yeast vacuole occurs by default and yeast plasma membrane proteins contain positive sorting information necessary for their localisation (Rayner and Pelham, 1997). Previous work has also established that proteins can be prevented from moving from the thin phospholipid-rich ER- Golgi- membranes to the thick sterol- and sphingolipid-rich plasma membranes by the length of their transmembrane domains (Rayner and Pelham, 1997; Munro, 1995). It is possible that a certain sequence necessary for exit from rough ER is missing in the sequence of E. coli transhydrogenase (Nothwehr and Stevens, 1994) oriand the iength of the transmembrane domains do not correspond to the thickness of yeast plasma membranes.

S. cerevisiae transformed with the gene encoding BB-transhydrogenase from A. vinelandii also showed transhydrogenase activity, demonstrating a functional enzyme (Paper V). To analyse the effect of transhydrogenation on the ratio of the two cofactor systems, NADP(H) and NAD(H), the intracellular concentrations of the four nucleotides were measured in S. cerevisiae strains expressing AB- and BB- transhydrogenase (Table 2). Expression of energy-linked A B-transhydrogenase changed the intracellular nucleotide levels (Paper N). The NADPH/NADP+ ratio was reduced from 4.96 to 1.95, while the NADH/NAD+ ratio was almost constant (Table 2). A similar effect was observed in strains expressing non-energy-linked BB- transhydrogenase (Paper IV). The NADPH/NADP+ratio remained reduced to 2.96 (Table 2). The different levels of NADPH and NADP+ in strains expressing either AB-transhydrogenase or BB-transhydrogenase could be an effect of different regulation mechanism of the proteins. The forward reaction (2) in AB-transhydro- genase is known to be strongly product inhibited by NADPH and NAD+ in a manner that varies with Ap (Persson et al., 1986), and BB-transhydrogenase undergoes

38 marked allosteric changes in the presence of AMP, NADPH, Ca2+ and NADP+ (Rydstrom et ul., 1987).

Table 2. The intracellular concentrations of NAD(H) and NADP(H) in pol per gram biomass (dry weight) in the cells sampled during exponential growth in anaerobic, glucose-limited batch cultivations.

Strain NAD+ NADP+ NADH NADPH NADW NADPW NAD+ NADP+

TN3 2.85k0.11 0.24k0.01 0.43k0.01 1.19kO.07 O.lSfl.01 4.96k0.52 TN24 3.45k0.07 0.2420.02 0.58H.04 0.47H.10 0.17H.01 1.95k0.50 AB-type TN4 3.17k0.07 0.27k0.02 0.54k0.02 O.SOkO.10 0.17H.01 2.96k0.60 BB-type

TN3, control strain containing expression vector without gene. TN24, strain expressing transhydrogenase from E. coli. TN4,strain expressing transhydrogenasefrom A. vinehdii.

Increased excretion of 2-oxoglutarate was observed during anaerobic glucose fermentation in recombinant S. cerevisiae strains overexpressing AB-transhydro- genase from E. coli (Paper IV) and BB-transhydrogenase from A. vinelundii (Paper V), compared with wild type (TN3) (Table 3). During growth with ammonium as the nitrogen source, ammonium and 2-oxoglutarate are converted into glutamate by glutamate dehydrogenase under oxidation of NADPH to NADP+ (Moye et aL, 1985). Conversion of NADPH and NAD+ into NADP+ and NADH in strains expressing transhydrogenase leads to a significant decrease in the intracellular pool of NADPH (Table 2) and could result in a reduced conversion rate of the reaction catalysed by the NADPH-dependent glutamate dehydrogenase. The reduced consumption of 2- oxoglutarate by glutamate dehydrogenase results in secretion of this compound (Table 3).

An increasing formation of both glycerol and acetate was observed in strains expressing transhydrogenase (Papers IV and V; Table 3). The increase in acetate formation probably occurs to compensate for the consumption of NADPH by the transhydrogenase. A similar effect has been observed in recombinant S. cerevisiae strains overexpressing XYLZ encoding an NADPH-consuming xylose reductase (Meinander et uL, 1996).

39 Table 3. Product yields in anaerobic, glucose-limited cultivations. Unit: c-mole per c-mole glucose

Strain

Product TN3 TN24 TN4 AB-typ BB-tyw

Ethanol 0.493 0.455 0.488 Glycerol 0.093 0.118 0.110 0.27 1 0.253 0.270

Succinate 0.005 0.006 0.005 Pyruvate 0.005 0.005 0.005

Acetate 0.005 0.014 0.004 Biomass 0.112 0.101 0.111

2-oxoglutarate 0.007 0.033 0.004

Total 0.991 0.985 0.997

TN3, control strain containing expression vector without gene. TN24, strain expressing transhydrogenase from E. coli. TN4, strain expressing transhydro- genase from A. vinehndii.

It was not possible to redirect the formation of NADH to NADPH by overexpressing transhydrogenase from E. coli or A. vinelandii in S. cerevisiae. However, by using protein engineering, e.g. the addition of sorting signals present in integral membrane proteins from yeast and the use of genetic screens to identify genes encoding proteins that facilitate the sorting events, it should be possible to redirect the membrane-bound AB-transhydrogenase to the plasma or to the vacuolar membranes where a Ap is present. In the presence of a Ap the forward reaction (2) should be stimulated and the [NADPH] [NAD+]/[NADP+] [NADH] ratio should increase.

The coupling of transhydrogenase to an existing Ap, has implications for the physiological function of the enzyme. The rate of the forward reaction (2) increased while the reverse reaction was simultaneously inhibited in reconstituted vesicles of bovine transhydrogenase (Rydstrom et al., 1971). Furthermore, the productlsubstrate ratio of [NADPH] [NAD+]/[NADP+] [NADH], increased by a factor of up to 500 (Lee and Emster, 1964; Rydstrom et al., 1970). However, this latter ratio is normally not reached in the intact mitochondrion or bacteria cells because of competing reactions which will reoxidise NADPH and rereduce NAD+ (Lee and Ernster, 1964;

40 Rydstrom el al., 1970). The affinity for NADP increased 5-fold while that for NAD+ decreased 5-fold for in the presence of Ap (Rydstrom et aZ., 1971). Less pronounced affinity changes have been observed for NADH and NADPH (Rydstrom et aL, 1971). Similar changes in dissociation constants have been reported for transhydrogenase from E. coli (Hu et aZ., 1995).

Non-energy-linked BB-transhydrogenase can also be used as an instrument to redirect the formation of NADH to NADPH in metabolically engineered S. cerevisiae strains where the consumption of NADPH and the production of NADH is unnaturally high. Such a system could be anaerobic xylose-fermenting strains of S. cerevisiae (Paper I), expressing NADPH, consuming XR and NADH producing XDH enzymes.

41 6. Acknowledgements

I would like to express my sincere gratitude to my supervisor, Professor Biirbel Hahn- Hagerdal, and to Professor Morten Kielland-Brandt, who gave me the opportunity to work in a stimulating and creative atmosphere.

I would also like to thank all my colleagues at the Department of Applied Microbiology, Lund, and at the Department of Yeast Genetics, Carlsberg Laboratory, Copenhagen, especially my former colleagues Torben Nissen, Mats Walfridsson and Xiaoming Bao.

Finally, I wish to thank my wife Christine and my son Markus, for their tremendous support and love.

This work was supported by the Nordic Energy Research Programme, the Swedish National Board for Technical and Industrial Development (NUTEK), and the Swedish National Science Research Council (NFR). 7. References

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