FUNCTIONAL GENOMICS OF GABA METABOLISM IN YEAST THERMOTOLERANCE Except where reference is made to the work of others, the work described in this dissertation is my own or was done in collaboration with my advisory committee. This dissertation does not include proprietary or classified information. ________________________________________ Juxiang Cao Certificate of Approval: ________________________ ________________________ Joe H. Cherry Robert D. Locy, Chair Professor Professor Biological Sciences Biological Sciences _______________________ ________________________ Narendra K. Singh Fenny Dane Professor Professor Biological Sciences Horticulture _______________________ Joe F. Pittman Interim Dean Graduate School FUNCTIONAL GENOMICS OF GABA METABOLISM IN YEAST THERMOTOLERANCE Juxiang Cao A Dissertation Submitted to the Graduate Faculty of Auburn University in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy Auburn, Alabama May 10, 2008 iii FUNCTIONAL GENOMICS OF GABA METABOLISM IN YEAST THERMOTOLERANCE Juxiang Cao Permission is granted to Auburn University to make copies of this dissertation at its discretion, upon request of individuals or institutions and at their expense. The author reserves all publication rights ___________________________________ Signature of Author ___________________________________ Date of Graduation iv VITA Juxiang Cao, daughter of Zhenbin, Cao and Yinxiu, Xiao, was born in Septerber 18, 1972 in Hubei, China. She graduated from number 1 high school in Gong'an, Hubei, China in 1992. In July of 1992, she entered Hubei Agricultural University and graduated in 1995 with a Bachelor's degree in the department of Plant Protection. She worked in industry for three years and in September 1998, she entered the department of Plant Pathology, college of natural resources and environmental sciences of South China Agricultural University and received her Master's degree in July 2001. In January of 2002, she joined the department of Biological Sciences to pursue her PhD degree in biochemistry and molecular biology. She is married to Shaoqiang, Dong in 2003. v DISSERTATION ABSTRACT FUNCTIONAL GENOMICS OF GABA METABOLISM IN YEAST THERMOTOLERANCE Juxiang Cao Doctor of Philosophy, May 10, 2008 (M.S., South China Agricultural University, Guangzhou, China, 2001) (B.S., Hubei Agricultural University, Jinzhou, China, 1995) 198 Typed pages Directed by Robert D. Locy ?-Aminobutyric acid (GABA) is a ubiquitous non-protein amino acid which accumulates rapidly in response to diverse environmental stresses. The GABA shunt is a pathway involving three enzymes, glutamate decarboxylase (GAD, encoded by GAD1), GABA aminotransferase (GABA-T, encoded by UGA1), and succinate semialdehyde dehydrogenase (SSADH, encoded by UGA2). These three enzymes acting in concert convert glutamate to succinate. GABA specific permease (encoded by UGA4) mediates the transportation of GABA into cells. The GABA specific transcription factor (encoded by UGA3) regulates the expression of GABA genes (UGA1, UGA2 and UGA4). We have constructed deletion mutants of each of these genes in yeast and have found that vi mutants of GAD, GABA-T and SSADH are more susceptible to stress induced by lethal temperature (45?C) than wild type yeast cells. Additionally, set of the combinations of double and triple mutants were examined. With a pretreatment at 40?C (a non-lethal temperature) for 30 min, the mutants retained susceptibility to stress at 50?C compared to the wild type. The levels of accumulated ROS were correlated to the susceptibility of heat stress. In addition, in the uga1 and uga2 mutants, GABA and ?-ketoglutarate accumulated markedly higher compared to the wild-type, while glutamate accumulated at higher levels in gad1 mutant. Deletion mutations of UGA3 and UGA4 grown showed heat tolerant to 45?C with overexpression of antioxidant genes superoxide dismutase compared to the wild-type. However, ?uga3 mutant strain grown in minimal-GABA medium showed heat sensitive phenotype while ?uga4 maintained heat tolerance. RT-PCR analysis showed that the expression of all GABA shunt genes and UGA3 and UGA4 genes were GABA inducible and were also up-regulated by lethal heat at 45?C. In addition, acidic pH in the growth medium induced the expression of UGA1, UGA2 and UGA4 but not that of GAD1 and UGA3. Under heat stress, deletion of UGA3 suppressed the expression of UGA1 and UGA4 but not on GAD1 and UGA2, while deletion of UGA4 did not affect the expression of all GABA shunt genes and UGA3. Additionally, the antioxidant genes superoxide dismutase (encoded by SOD1 and SOD2) were found to be gradually induced by heat in the wild-type strain but overexpressed in the ?uga3 and ?uga4 mutant strains. Bioinformatic programme "TargetP and pSORT' predicts that GABA transaminase from Arabidopsis is localized in mitochondria with a 54 nucleotide mitochondrial transit peptide sequence, yeast GABA transaminase is localized in cytosol. We constructed vectors expressing ScGABA-TKG and AtGABA-TP in both yeast cytosol and vii mitochondria to complement yeast GABA transaminase mutant ?uga1 and succinate semialdehyde dehydrogenase mutant ?uga2 phenotypes: GABA growth defect, thermo sensitivity and heat induced production of reactive oxygen species (ROS). Our studies revealed that plant AtGABA-TP is functionally interchangeable with yeast ScGABA- TKG for GABA growth, thermotolerance and limiting production of ROS whether located in mitochondria or cytosol in yeast. However, yeast GABA-TKG, whether located in mitochondria or in the cytosol, displayed much stronger effect. The yeast succinic semialdehyde dehydrogenase gene (SSADH; EC 1.2.1.16) was cloned and overexpressed in E. coli., and kinetically characterized. It has a molecular mass of subunit around 54 kDa, the purified enzyme has a tetramer molecular mass of 200 kDa. The recombinant protein was highly specific for succinate semialdehyde, can use both NAD+ and NADP+ as a cofactor but with higher affinity to NAD+. The enzyme activity can be inhibited by substrate SSA, product NADH and adenine nucleotides AMP, ADP and ATP. viii ACKNOWLEDGEMENTS The author would like to thank her advisor Dr. Robert D. Locy and also Dr. Narendra K. Singh for their continued support and encouragement during her work. Their work ethic and motivation have provided her with the guidance needed to be successful in this challenging field. She is forever indebted. Special thanks given to her committee members Dr. Joe H. Cherry and Dr. Fenny Dane for guidance, helpful comments, and discussions. Without their help, this dissertation would never have been completed. Thanks to Dr. Floyd Woods for agreeing to be my outsider reader and his valuable comments on her work. She also thank Dr. Cathy McVay, her TA supervisor for providing her part of the material for expressing recombinant protein, She learned a lot from her while she was assisting her with Cell Biology Lab teaching. ix Style manual of journal used: Journal of Applied and Environmental Microbiology Computer software used: Microsoft Word, Microsoft Excel, SigmaPlot 2000, Adobe Photoshop 7.0 x TABLE OF CONTENTS LIST OF TABLES.............................................................................................................xii LIST OF FIGURES ......................................................................................................... xiii I. LITERATURE REVIEW.................................................................................................1 Literature cited..........................................................................................................25 II. THE GABA SHUNT MEDIATES BASAL THERMOTOLERANCE IN SACCHAROMYCES CEREVISIAE BY REDUCING THE PRODUCTION OF REACTIVE OXYGEN SPECIES Abstract.....................................................................................................................43 Introduction...............................................................................................................45 Material and Methods ...............................................................................................46 Results.......................................................................................................................54 Discussion.................................................................................................................60 References.................................................................................................................64 III. GABA SHUNT GENE EXPRESSION ANALYSIS AND HEAT STRESS RESPONSE OF THE GABA SPECIFIC TRANSCRIPTION FACTOR AND TRANSPORT GENES IN SACCHAROMYCCEREVISIAE Abstract.....................................................................................................................83 Introduction...............................................................................................................84 Material and Methods ...............................................................................................86 Results.......................................................................................................................89 Discussion.................................................................................................................95 References.................................................................................................................99 IV. GABA TRANSAMINASE: COMPLEMENTATION ANALYSIS AND INTRACELLULAR LOCALIZATION IN SACCHAROMYCES CEREVISIAE Abstract...................................................................................................................108 Introduction.............................................................................................................109 Material and Methods .............................................................................................111 Results.....................................................................................................................118 Discussion...............................................................................................................124 References...............................................................................................................127 V. MOLECULAR CLONING, EXPRESSION AND CHARACTERIZATION OF THE RECOMBINANT YEAST SUCCINIC SEMIALDEHYDE DEHYDROGENASE xi Abstract...................................................................................................................136 Introduction.............................................................................................................137 Material and Methods .............................................................................................138 Results.....................................................................................................................141 Discussion...............................................................................................................145 References...............................................................................................................150 VI. IN GEL STAINNING METHOD FOR DETECTING GABA TRANSAMINASE ACTIVITY Abstract...................................................................................................................168 Introduction.............................................................................................................169 Material and Methods .............................................................................................171 Results and Discussion ...........................................................................................173 Conclusions.............................................................................................................176 References...............................................................................................................177 APPENDIX I ...................................................................................................................181 APPENDIX II ..................................................................................................................182 xii LIST OF TABLES Table II.1.Yeast strains used in this study .........................................................................67 Table II.2. Primers used for GAD1, UGA1 and UGA2 gene cloning, deletions and verifications........................................................................................................................68 Table III.1. Yeast strains used in this study.....................................................................102 Table III.2. Primers used for UGA3 and UGA4 gene deletions and verifications...........104 Table IV.1.Yeast strains used in this study......................................................................129 Table IV.2. Primers used for yeast cytosolic and mitochondrial forms of GABA-TKG and plant GABA-TP expression .............................................................................................129 Table IV.3. Enzyme activities in yeast wild type, ?uga1 deletion mutant strain and ?uga1 mutant transformants .......................................................................................................130 Table V.1. Single step purification of recombinant yeast SSADH..................................153 Table V.2. Kinetic parameters of recombinant yeast SSADH.........................................153 Table V.3. Inhibition of recombinant yeast SSADH by the products and nucleotides.................................................................................................................154 xiii LIST OF FIGURES II.1. Viability of wild-type and mutant strains after lethal heat treatment at 45?C................................................................................................................................69 II.2. Viability of wild-type and ?gad1mutant with overexpression of yeast GAD1 (A), ?uga1 mutant with overexpression of yeast UGA1 (B) and ?uga2 mutant with overexpression of yeast UGA2 (C) after lethal heat stress at 45oC...................................70 II.3. Viability of wild-type and mutants at lethal heat stress50?C.....................................71 II.4. Intracellular ROS levels induced by lethal heat stress at 45?C in wild-type and mutant strains.....................................................................................................................72 II.5. Intracellular ROS levels induced by lethal heat stress at 45?C in wild-type and ?gad1mutant transformed with plasmid P425-GPD- GAD1 ............................................73 II.6. Intracellular ROS levels induced by lethal heat stress at 50?C after a sub-lethal heat stress at 40 ?C for 30min in wild-type and mutant.............................................................74 II.7. Effect of PBN on lethal heat induced cell death under 45?C for wild-type and all mutant strains.....................................................................................................................75 II.8. Effect of PBN on production of ROS under 45?C for wild-type and all mutant strains .........................................................................................................76 II.9. Effect of PBN on heat induced cell death under 50?C with preheat treatment at 40?C for 30min for wild-type and all mutant strains ..................................................................77 II.10. Effect of PBN on production of ROS under 50?C with pre-heat treatment at 40 ?C for 30min for wild-type and all mutant strains ..................................................................78 II.11. Changes in the levels of GABA under lethal heat stress at 45oC in wild-type and mutant strains.....................................................................................................................79 II.12. Changes in the levels of glutamate under lethal heat stress of 45oC in wild-type and mutant strains.....................................................................................................................80 II.13. Changes in the levels of a-ketoglutarate under lethal heat stress at 45oC in wild-type and mutant strains ..............................................................................................................81 xiv II.14. Changes in the levels of succinate semialdehyde under lethal heat stress at 45oC in wild-type and mutant strains..............................................................................................82 III.1. Transcript levels of GABA shunt genes (GAD1, UGA1, UGA2), the GABA specific transcription factor (UGA3) and GABA permease (UGA4) in response to minimal media with different nitrogen sources and acidic pH .................................................................104 III.2. Viability of wild-type, ?uga3, and ?uga4 mutant strains after lethal heat treatment at 45oC..............................................................................................................................105 III.3. Transcript levels of GABA shunt genes (GAD1, UGA1 and UGA2) and the GABA specific regulatory and transport genes (UGA3 and UGA4 respectively) in response to lethal heat stress at 45?C ..................................................................................................106 III.4. Transcript levels of the cytosolic and mitochondrial superoxide dismutases (SOD1 and SOD2 respectively) in response to lethal heat stress at 45?C....................................107 IV.1. (A) Growth of wild type and mutant strains on YNB medium with NH4+ as the sole nitrogen source; (B) YNB medium with GBAB as the sole nitrogen source ..................131 IV.2. Viability of wild-type, mutant strains and mutant strains transformed with (A) cytosolic and (B) mitochondrial forms of yeast endogenous GABA-TKG after lethal heat treatment at 45oC..............................................................................................................132 IV.3. Intracellular ROS levels induced by lethal heat stress at 45?C in wild type, mutant strains and mutant strains transformed with cytosolic (A) and mitochondrial (B) forms of yeast endogenous GABA TKG........................................................................................133 IV.4. Viability of wild-type, mutant strains and mutant strains transformed with (A) cytosolic and (B) mitochondrial forms of plant GABA-TP after lethal heat treatment at 45oC..................................................................................................................................134 IV.5. Intracellular ROS levels induced by lethal heat stress at 45?C in wild type, mutant strains and mutant strains transformed with (A) cytosolic and (B) mitochondrial forms of plant GABA-TP ...............................................................................................................135 V.1. Expression and purification of recombinant yeast SSADH. 12% of SDS-PAGE analysis of crude cell extracts of BL21 transformed with expression vector pET16b containing the SSADH coding sequence .........................................................................155 V.2. Molecular mass determination of native SSADH by FPLC gel filtration on Sephacryl S-300-HR .........................................................................................................................156 V.3. Double reciprocal plot from initial velocity experiments of yeast recombinant xv succinate semialdehyde dehydrogenase with varied concentrations of SSA......................................................................................................157 V.4. Double reciprocal plot from initial velocity experiments of yeast recombinant succinate semialdehyde dehydrogenase with varied concentrations of NAD+........................................................................................158 V.5. Double reciprocal plot from initial velocity experiments of yeast recombinant succinate semialdehyde dehydrogenase with varied concentrations of NADP+ ...............................................................................................159 V.6. Inhibition of yeast recombinant SSADH by NADH. Double-reciprocal plots of the rate of NADH formation vs NAD+ concentration at different NADH concentrations...................................................................................160 V.7. Inhibition of yeast recombinant SSADH by NADH. Double-reciprocal plots of the rate of NADH formation vs SSA concentration at different NADH concentrations...................................................................................161 V.8. Inhibition of yeast recombinant SSADH by AMP. Double-reciprocal plots of the rate of NADH formation vs NAD+ concentration at different AMP concentrations......................................................................................162 V.9. Inhibition of yeast recombinant SSADH by AMP. Double-reciprocal plots of the rate of NADH formation vs SSA concentration at different AMP concentrations......................................................................................163 V.10. Inhibition of yeast recombinant SSADH by ADP. Double-reciprocal plots of the rate of NADH formation vs NAD+ concentration at different ADP concentrations.......................................................................................164 V.11. Inhibition of yeast recombinant SSADH by ADP. Double-reciprocal plots of the rate of NADH formation vs SSA concentration at different ADP concentrations................................................................165 V.12. Inhibition of yeast recombinant SSADH by ATP. Double-reciprocal plots of the rate of NADH formation vs NAD+ concentration at different ATP concentrations ..................................................................................................................166 V.13. Inhibition of yeast recombinant SSADH by ATP. Double-reciprocal plots of the rate of NADH formation vs SSA concentration at different ATP concentrations.......................................................................................167 VI.1. Zymogram staining analysis of GABA transaminas activities from various yeast sources ...............................................................................179 xvi VI.1. Zymogram staining analysis for the substrate identification of Arabidopsis GABA transaminase.....................................................................................................................180 1 I. LITERATURE REVIEW GABA is a ubiquitous non protein amino acid which is widely found from prokaryotic to eukaryotic organisms. It was first discovered by Steward et al. (1949). in potato tubers . Later on, it was found in rat brain and many parts of vertebrates, invertebrates, plants, bacteria, and fungi (Santosh et al., 1997). Interest in GABA shifted to animals when it was found in large quantity in brain. It is well known that GABA functions as a negative neurotransmitter in animals (Erdo and Kiss, 1986). In plants, GABA accumulates in response to various biotic and abiotic stresses including cold shock, heat shock, mechanical damage, and water stress (Streeter and Thompson, 1972; Wallace et al., 1984; Reggiani et al., 1988; Craw ford et al., 1994). However, in most organisms, the role of GABA is not well known. GABA is produced in the cytosol through the decarboxylation of L-glutamate by glutamate decarboxylase (GAD). GABA is then transaminated to ?-ketoglutarate or pyruvate by GABA transaminase (GABA-T) producing glutamate or alanine and succinate semialdehyde (SSA), which is an irreversible reaction. Finally, SSA is converted to succinate (SUCC), a metabolite of the tricarboxylic acid cycle (TCA cycle) by succinate semialdehyde dehydrogenase (SSADH). These three enzymes (GAD, GABA-T and SSADH) work in concert to convert glutamate to succinate, and along with glutamate dehydrogenase or several ?-ketoglutarate-dependent aminotransferases to 2 bypass 2 enzymes in the TCA cycle (Appendix). Therefore, these reactions constitute the GABA shunt. In plants, Bouch?, N. et al. (2003) demonstrated that the disfunction of SSADH leads to hyper sensitivity to environmental stresses, and accumulation of higher levels of reactive oxygen species (ROS). In addition, in yeast, Coleman et al. (2001) found that the GABA shunt is involved in oxidative stress. However, the role of the GABA shunt in response to other environmental stress conditions is uncertain. Of particular interest in this dissertation, is defining the role of GABA shunt in thermotolerance. This review will discuss the role of GABA in different organisms, the role of the three enzymes of GABA shunt, and the role of these enzymes and the GABA shunt in heat stress and its relation to the production of ROS, and finally the expression of the three enzymes and the related regulatory proteins under heat stress. GABA Shunt enzymes The GABA shunt is a metabolic pathway that links nitrogen and carbon metabolism using glutamate as a substrate in the first catalytic step while producing succinate as its end product. The reactions of the GABA shunt are catalyzed by three enzymes, glutamate decarboxylase, GABA transaminase and succinate semialdehyde dehydrogenase. Understanding the properties of each individual enzyme in the metabolic pathway helps to understand the specific regulation of GABA shunt under stress conditions and the possible physiological roles of GABA in different organisms. 3 Glutamate decarboxylase In mammals, the cDNAs of GAD have been isolated and sequenced, it is encoded by two different genes of GAD1 and GAD2 which are located on different chromosomes (Erlander et al., 1991), the two isoforms are GAD67 and GAD65 which have molecular weights of 67 and 65 kDa respectively. GAD1 is only expressed in brain tissue, and GAD2 is expressed in brain but also in pancreas (Erlander et al., 1991). It is widely thought that the two GAD isoforms in brain are localized within different subcellular compartments; GAD67 is related mainly to cytoplasmic pools of GABA, while GAD65 is associated with vesicular pools of GABA (Soghomonian and Martin, 1998). It is possible that GAD67 regulates GABA synthesis for metabolic functions of the cell, while GAD65 regulates GABA synthesis for synaptic release (Soghomonian and Martin, 1998). Mammalian GAD plays an important role in the regulation of GABA synthesis through interaction with its co-factor pyridoxal-phosphate (pyridoxal-P) (Martin and Rimvall, 1993).The two GAD isoforms in brain differ in binding with the co-factor, recombinant GAD data showed that GAD65 is more preferable to co-factor pyridoxal-P than GAD67 (Erlander et al., 1991), but characterization of the differences is not yet available. In most regions of the rat brain, GAD65 appears to be the major GAD isotype, however, research from knockout mice and rats treated with vigabatrin, an irreversible inhibitor of GABA transaminase, indicated that most of GABA in the brain was synthesized by GAD67 (Soghomonia and Martin, 1998). In neuronal cell cultures or brain slices, it was found that GABA can be released through both Ca2+-dependent and -independent mechanisms (Pin and Brockaert, 1989, Attwell et al., 1993, Belhage et al., 1993). The mammalian GADs have a pH optimum of approximately 7 (Wu et al., 1974). 4 In plants, GAD genes from various sources such as Petunia (Baum et al., 1993), tomato (Gallego et al., 1995), tobacco (Yu and Oh, 1998), and Arabidopsis (Turano and Fang, 1998, Zik et al., 1998) have been identified. The activity of plant GAD was found to be associated with senescence, seed germination and ripening (Gallego et al., 1995). Plant GAD utilizes L-glutamate as a substrate and has pyridoxal 5'-phosphate as a cofactor (Satya and Nair, 1986). Plant GADs demonstrate Km's between 3 and 25 mM depending on the plant species, and tissue sources (Satya and Nair, 1985; Snedden et al., 1995; 1996) and from 22 to 25 mM respectively (Chen et al., 1960). GAD appears to be localized in cytosol of plant cells and possesses a calmodulin-binding domain (Breitkreuz and Shelp, 1995). Unlike mammalian GAD, plant GADs have sharp acidic pH optima around pH 5.8 (Satya and Nair, 1985; Snedden et al., 1995). The in vitro activity assay of recombinant Petunia GAD shows that it has little activity in pH 7.0 in the absence of calcium ion and calmodulin (Snedden et al., 1996). Environmental stresses such as oxygen deficiency or heat shock ((Roberts et al., 1984; 1992; Locy et al., 2000) and acid or ammonium treatments (L.A. Crawford et al., 1994) induce GABA synthesis resulting from a reduced cytosolic pH, presumably as a result of stimulated GAD activity at lowered cytosolic pH. Other environmental stresses such as cold shock or touch are known to increase cytosolic Ca2+ levels (Knight et al., 1991). GAD activity in vitro in a lot of species and plant tissues is shown to be stimulated by Ca2+/calmodulin at neutral pH 7.0 ~ 7.5, instead of acidic pH lower than 6.5 (Snedden et al., 1995; 1996; Ling et al., 1994). In vivo experiments with 1 hour pretreatment by Ca2+ channel-blockers and calmodulin antagonists in aerobic conditions for Ca2+/calmodulin activation of GAD in 5 the rice roots did not result in GABA accumulation under 3 hour of anoxia (Aurisano et al., 1995). Hence, both in vivo and in vitro experiments suggest that the stimulation of GABA synthesis results from the stimulated GAD which is regulated by increased Ca2+ levels and calmodulin. Furthermore, in isolated Asparagus mesophyll cells, Chung et al. (1992) found that GAD activity is also stimulated by elevated glutamate levels. The catalytic activity of plant GAD is inhibited by reagents which can react with sulfhydryl groups. GAD activity is both transcriptionally and translationally regulated in different Petunia organs (Chen et al., 1994). In Arabidopsis, there are at least two GAD isoforms: GAD1 is root specific and GAD2 is distributed in all organs (Zik et al., 1998; Turano and Fang, 1998). Additionally, with the whole genome sequenced in Arabidopsis, three more putative GAD isoforms have been revealed by sequence comparisons, GADs 2 ? 5 are 75 ~ 82% identical to GAD1 for their protein sequences (Shelp et al., 1999). In fungi, the GAD enzyme was first reported in yeast in 1953 (Krishnaswamy and Giri, 1953; 1956). Fungal GAD has been purified and characterized in several species such as R. glutinis (Krishnaswamy and Giri, 1953), N. crassa (Schmit and Brody, 1975), A. niger ((Kubicek et al., 1979), and best studied in A. bisporus (Baldy, 1975). Like mammalian and plant GADs, fungal GAD is exclusively localized in the cytosol , the optimum pH is between 4.0 an 6.0, the Km for L-glutamate is in the lower millimolar range and requires pyridoxal phosphate as a cofactor. Partially purified R. glutinis GAD is inhibited by hydroxylamine and the addition of excess amount of pyridoxal phosphate can reverse the inhibition (Krishnaswamy and Giri, 1953; 1956). N. crassa GAD has been purified from conidia, it is a 30 to 33 kDa monomer and the Km for pyridoxal phosphate of 40 nM (Schmit and Brody, 1975). A. bisporus GAD is similar to N. crassa GAD. In A. 6 niger, only the conidiating mycelia, but not vegetative cells, show GAD activity (Baldy, 1975). Blast searches show that GAD from yeast shares sequences with GADs from Petunia hybrida and A. thaliana with identities of 38% and 39% respectively. Recombinant GAD studies demonstrated that yeast GAD can also bind to CaM like plant GAD does, but no activities were found with the CaM bound form, the functional GAD inside cells is required for oxidative stress tolerance (Coleman et al., 2001). In bacteria, GAD is best characterized in Escherichia coli, it is easily induced in the growth media by glutamate (Fonda, 1985). Two isoforms of gadA and gadB were cloned in E. coli, (Smith et al., 1992), both isozymes showed identical kinetic and physico- chemical properties (Biase et al., 1996). Purified GAD from E. coli is a 53 kDa hexamer, with one pyridoxal 5'-phosphate (PLP) site on each subunit (Strausbauch and Fischer, 1970). The optimum pH is between 4.0 and 4.5, and it has broad ranges of substrates such as ?-methylene -glutamate, threo-?-hydroxy- glutamate, and homocysteine sulfinate, but -glutamate was shown to be the best substrate (Fonda, 1972). The activity of Escherichia coli GAD was inhibited by suicide substrates such as serine-O-sulfate, R-(?)-4-aminohex- 5-ynoic acid, and ?-fluoromethyl-glutamate (Kuo and Rando, 1981; Sukhareva and Braunshtein, 1971; Likos et al., 1982), the inhibition mechanism was found to be the same as aspartate aminotransferase (Likos et al., 1982). GAD has also been purified and characterized from other bacteria such as Streptococcus pneumoniae which resembles mammalian GAD (Garcia and Lopez, 1995) and Lactobacillus brevis which resembles E. coli GAD (Ueno et al., 1997). 7 GABA transaminase In mammals, GABA is present in many mammalian tissues and is a major inhibitory neurotransmitter in the central nervous system (Fonnum, 1987). The concentration of GABA is controlled by GAD and GABA transaminase (GABA-T). After its synthesis by GAD, GABA is catabolized by GABA-T to succinic semialdehyde for subsequent conversion into succinate. In mammals, GABA-T has been purified and characterized mostly from brain tissues such as mouse, rat, rabbit and human brains (Maitre et al., 1975; John and Fower, 1976; White and Sato, 1978). The basal level of GABA-T activity is low. Recombinant human GABA-T has an apparent Km values for substrates ?-ketoglutarate and GABA of 0.11 mm and 1.27 mm respectively (Seong et al., 2000). These values are similar to those from bovine and pig brains (Choi et al., 1993). The activity of GABA-T is inhibited by vigabatrin. Mammalian GABA-T requires pyridoxal-5-phosphate as co- factor. It catalyzes the transamination of GABA with a Km of 1.1 mM and catalyzes the transamination of ?-alanine to the same extent of GABA (Schousboe et al.,1973; Buzenet et al., 1978). In plants, the catabolism of GABA takes places in mitochondria (Breitkreuz and Shelp, 1995). It was shown that tobacco crude extracts apparently contain two GABA transaminases. One enzyme utilizes GABA, together with pyruvate while the other enzyme utilizes GABA and ?-ketoglutarate. The two enzymes form succinate semialdehyde and alanine or glutamate respectively (Shelp et al., 1999; Satya and Nair, 1990; Van Cauwenberghe et al., 1999). However, the ?-ketoglutarate-dependent GABA- T remains to be further elucidated, the activity can only be detected in tobacco crude extract and is undetectable subsequently during purification (Van Cauwenberghe et al., 8 1999). Additionally, it is noteworthy that there are no ?-ketoglutarate-dependent GABA- T sequences found in any plant genome utilizing the BLAST search engine at NCBI to search the GenBank database. Plant GABA-T has not been purified from any plant. However, biochemical studies from plant crude preparations show the optimum pH for plant GABA-T is between 8.6 to 9.0 (Streeter and Thompson, 1972; Satya and Nair, 1986), and requires pyridoxal- phosphate (PLP) as a cofactor ( Satya and Nair,1986; 1990, Van Cauwenberghe et al., 1999). Plant pyruvate dependent GABA-T has been cloned from Arabidopsis (Van Cauwenberghe et al., 2002) and rice (Ansari et al., 2005). These enzymes show putative mitochondrial targeting sequences and conserved PLP binding domains. In addition, the GABA-T knockout in Arabidopsis provides evidence that GABA accumulates greatly in flowers up to 113- fold compared with wild type suggesting GABA-T functions in vivo (Palanivelu et al., 2003). In yeast and other fungi, Roberts et al. (1953) discovered that ?-ketoglutarate- dependent GABA-T is involved in nitrogen utilization. Subsequently, Yonaha et al. (1983) found that GABA-T exists in a variety of micro-organisms including yeast and other molds. GABA-T was induced in yeast species like Saccharomyces, Hansenula and Candida on medium with GABA but not on ?-alanine, while in molds and other genera such as Rhizopus, Aspergillus, Penicillium and Neurospora, GABA-T was detected on both GABA and ?-alanine-containing media (Yonaha et al., 1983). Characterization of GABA-Ts purified from Candida (Der et al., 1986) or A. bisporus carpophore (Baldy, 1976) indicated that they share some catalytic features: pyridoxal phosphate-dependent, optimum pH of 8.0, ?-ketoglutarate specificity and inhibition by short chain fatty acids 9 such as propionate and butyrate. GABA-T has been cloned from yeast Saccharomyces cerevisiae (Andre and Jauniaux, 1990), and it is encoded by UGA1 gene. Previous studies from UGA1 knockout demonstrated that yeast GABA-T, like other genes in GABA shunt pathway, plays an essential role in oxidative stress tolerance (Coleman et al., 2001). Bacterial GABA-T, is most similar to yeast GABA-T, but different from the GABA-T of other fungi and mammals. It is specific for GABA, does not utilize ?-alanine, and requires ?-ketoglutarate as an amino acceptor (Yonaha et al., 1983). GABA-T was found to be abundantly distributed in most strains of bacteria grown on media with ?-alanine or GABA as nitrogen sources (Yonaha et al., 1983). Studies from cowpea Rhizobium also show that GABA-T requires ?-ketoglutarate. The activity of GABA-T is induced on media containing GABA as the sole carbon and nitrogen source (Jin et al., 1990). Recently, a GABA-T gene (gabT) from Rhizobium leguminosarum bv. Viciae was identified, cloned, and characterized (Prell et al., 2002). It is induced by GABA and highly expressed in bacteroids. Mutants of gabT lost the ?-ketoglutarate-dependent GABA-T activity, but they are still able to grow on GABA as the sole carbon and nitrogen source, suggesting the existence of multiple GAB-Ts which have different substrate specificities in this organism. According to Tunnicliff (1993), the activity of GABA-T from Pseudomonas fluorescens is inhibited by ATP, and its analogues ADP, CTP and XTP in a competitive manner, the inhibition effect can be antagonized by GABA, indicating ATP competes with GABA for the enzyme binding site. Voellym and Leisinger (1976) purified GABA-T from Pseudomonas aeruginosa. In addition to GABA, the purified enzyme catalyzed the transamination of N2-acetyl-L-ornithine, L-ornithine, 10 putrescine, L-lysine, and cadaverine in order of decreasing activity. The enzyme is induced by GABA, guanidinobutyrate, or putrescine as well. Succinate semialdehyde dehydrogenase In mammals, SSADH has been purified to apparent homogeneity from rat, pig and human brains (Chambliss and Gibson, 1992; Ryzlak and Pietruszko, 1988; Lee et al., 1995). The purified human brain SSADH is localized in mitochondria, has SSA as the best substrate, but also has activities for the substrates of glutaric semialdehyde, nitrobenzaldehyde, and short chain aliphatic aldehydes. The enzyme can only use NAD+ as a cofactor (Ryzlak and Pietruszko, 1988). The enzyme from pig brain is substrate inhibited by SSA and by the product, NADH (Duncan and Tipton, 1971). In plants, SSADH has been purified from wheat embryos (Galleschi et al., l983), barley seeds (Yamura et al., 1988) and potato tubers (Satya and Nair, 1989). The first cloned SSADH was from Arabidopsis (Bu et al., 1999). Biochemical analysis from previous studies show plant SSADH is localized in mitochondria (Breitkreuz and Shelp, 1995). Mitochondrial localization is common among most organisms except yeast. The plant SSADH has an optimum pH of 9.0, is highly specific for substrate SSA, and can only use NAD+ as a cofactor, the catalytic activity is inhibited by the substrate SSA at higher concentrations, the product NADH and also adenine nucleotides such as AMP, ADP and ATP. A disruption of the unique SSADH gene in Arabidopsis has been shown to cause necrotic lesions and programmed cell death during periods of heat stress, UV light stress, or hydrogen peroxide stress as result of accumulation of toxic reactive oxygen species (Bouch? et al., 2003). 11 In yeast and other fungi, SSADH has received relatively little attention. Most studies have been limited to enzyme activities from crude cell preparations (Pietruszko & Fowden, 196; Ramos et al., 1985). Ramos et al. (1985) found that SSADH from Saccharomyces has a pH optimum of 8.4. It is highly specific for the substrate succinate semialdehyde with a Km in the ?M range. It can use both NAD+ and NADP+ as a cofactor but has higher affinity for NAD+. Its activity is substrate inhibited by succinate semialdehyde. These kinetic properties are also shared by SSADH from niger mycelia and plants (Kumar and Punekar, 1994; Pietruszko and Fowden, 1961; Rating et al., 1984; Satya and Nair, 1989). Ramos et al. (1985) found a thio requirement is not necessary for yeast SSADH activity. Some exception was also observed in fungi, for example, the T. utilis extracts were shown to catalyze the reverse reaction by reducing SSA to 4- hydroxybutyric acid (Pietruszko and Fowden, 1961). In Saccharomyces cerevisiae, SSADH is encoded by the UGA2 gene (Coleman et al., 2001). It is localized in the cytosol (Huh et al., 2003). Together with UGA1, UGA2 is involved in the degradation of GABA to succinate. Mutation of either of UGA1 or UGA2, makes yeast cells more sensitive to oxidative stress (Coleman et al., 2001), and prevent them from growing on GABA as the sole nitrogen source cells (Ramos et al., 1985). Like GAD1 and UGA1, UGA2 expression is also induced by GABA (Ramos et al., 1985). The expression levels of UGA1 and UGA2 are under the control of transcription activator UGA3p (Ramos et al., 1985; Vissers et al., 1989; Talibi, et al., 1995; Coleman et al., 2001). Under oxidative stress, levels of UGA2 transcript were found to be up-regulated (Coleman et al., 2001). Detailed regulation of the expression of GABA shunt genes and the genes which control their expression will be reviewed later. 12 In bacteria, two proteins are involved in the activity of succinate semialdehyde dehydrogenase: NAD+-dependent and NADP+-dependent succinate semialdehyde dehydrogenases. Cozzani et al. (1980) separated these two proteins from E. coli. The partially purified enzymes differ in their co-factors (NAD+ or NADP+). The NADP+- dependent SSADH is very specific for the substrate SSA while the NAD+-dependent SSADH can utilize n-butyraldehyde in addition to SSA. In E. coli. B, both NAD+-and NADP+-dependent SSADH were induced by GABA, but the NADP+-dependent enzyme has higher activity of GABA-T (Donnelly and Cooper,1981). SSADH has also been cloned from E. coli.. It is part of the gene cluster which encodes the enzymes for GABA degradation. (Metzer and Halpern, 1990; Bartsch et al., 1990; Niegemann et al., 1993). In E. coli, a second sad-encoded SSADH was identified in addition to the gabD-encoded SSADH (Donnelly and Cooper, 1981; Marek and Hensen,1988), and in Pseudomonas sp., Klebsiella pneumoniae, and Ralstonia eutropha also an NADP+- and/ or an NAD+-dependent SSA-DH were detected (Nirenberg and Jacoby, 1960; Sanchez et al., 1989) The role of GABA in different organisms In animal systems, GABA has been well known as an inhibitory neurotransmitter mainly found in the central nervous systems (CNS). In vertebrates, GABA acts at inhibitory synapses in the CNS by binding to specific transmembrane receptors, which causes the opening of ion channels to allow negatively-charged chloride ions to flow into the cell or positively-charged potassium ions to flow out of the cell (Roth et al., 2003). Three general classes of GABA receptor have been found: GABAA and GABAC 13 ionotropic receptors, which are ion channels themselves, and GABAB metabotropic receptors, which are G protein-coupled receptors. (Dzitoyeva et al., 2003; Mihic et al., 1997; Boehm et al., 2006; Dimitrijevic et al., 2005). In plants, for many years, GABA has been shown from numerous reports to accumulate rapidly upon various stress factors including heat, cold, drought, acidosis, anoxia, mechanical damage. For example, in heat-stressed cowpea cells, GABA accumulated 1,800-fold in 24 hours as compared to the non stressed control (Mayer et al., 1990). In soybeans subjected to cold stress and mechanical damage, GABA level in soybean leaves increased 2,000- and 2,700- fold respectively in 5 min (Wallace et al., 1984). However, the physiological role GABA plays in plants is not well understood yet. Early studies suggested different roles of GABA in response to stress conditions. These roles are generally related to: contributing to the C: N balance (Rolin et al., 2000, Breitkreuz et al., 1999, Snedden and Fromm, 1999), regulation of cytosolic pH (Snedden et al., 1995, 1996), protection against oxidative stress (Bouch? et al., 2003), defense against insects (Ramputh and Brown, 1996), or GABA as an osmoregulator (Rentsch et al., 1996) More recently, scientists are trying to find evidences to show that GABA is a signaling molecule by analogy to such a clearly defined role in animals. In Arabidopsis, the pollen?pistil-interaction2 (pop2) gene encodes GABA-T. The pop2 mutant accumulated GABA in flowers, and the guidance and growth of the pollen tube was disturbed. In the wild type, a GABA gradient was created in the pistil, that was required for proper pollen tube guidance to the ovule (Palanivelu et al., 2003; Bouch? et al., 2004). Thus, this finding indicates GABA may play an important role in signaling. However, 14 there is a strong need in plants to elucidate the GABA signaling pathway by showing the presence of GABA receptors. Genes which share high sequence similarities with animal ionotropic glutamate receptors have been identified in Arabidopsis as putative glutamate receptors (designated as ATGLRs) (Lacombe et al., 2001), these ATGLRs posses domains which are structurally highly homologous to GABAB receptors, thus, it is reasonable to assume GABA interacts with ATGLRs to play its signaling role by binding to these domains. In bacteria, GABA was first found to play a role in carbon and nitrogen metabolism, wild type laboratory strains of E. coli K-12 are unable to utilize ?-aminobutyrate (GABA) as the sole carbon and nitrogen source. However, Dover and Halpern (1972) found UV induced mutants are able to grow on GABA as a sole carbon and nitrogen source, and also the activities of GABA aminotransferase and SSADH were both increased six- to nine-fold. Foester and Foester (1973) reported that GABA plays a role in the germination of the Bacillus megaterium spores. GAD activity increased dramatically during germination leading to accumulation of GABA. In addition, Castaniethere et al. (1999) reported that the production of GABA was involved in acidic pH resistance in E. coli. The GAD enzyme protects the cell from extreme acid conditions during transit through the host stomach since GAD consumes protons by catalyzing decarboxylation of glutamate. The product GABA is finally exported from the cells. In yeast and other fungi, GABA was first isolated from acid treated yeast extracts (Reed, 1950). It was subsequently found in various other fungi. GABA accumulated unusually in A. niger during acidogenesis coupled with the increase of citric acid (Kubicek et al., 1979). In understanding the role of GABA in fungi, the following have 15 been summarized from previous studies: Firstly, GABA metabolism is involved in the catabolism of nitrogen compounds. In S. cerevisiae (Ramos et al., 1985) and A. nidulans (Yonaha et al., 1983), GABA in the growth media as a sole nitrogen source induces the expression of the enzymes which catabolize GABA such as GABA-T and SSADH, the GABA transporter gene (UGA4) is also induced (Ramos et al., 1985). GABA metabolism is involved in conidiation and germination of conidia. This role was established from the work of Schmit et al (Schmit and Brody, 1975; Hao and Schmit, 1991; 1993), Schmit and Brody (1975) who found GAD can only be detected at conidiation stage but GAD is not found in mycelia. The levels of GAD increase at conidia as they mature and decline in the early phase of conidial germination. Therefore, this work enabled Schmit et al. (1975) to first clone GAD as a conidiation stage-specific marker enzyme. Furthermore, GABA metabolism seems to be involved in oxidative stress tolerance. Coleman and co-workers (Coleman et al., 2001) showed the three genes including GAD1, UGA1 and UGA2 in the GABA metabolic pathway are all required to tolerate oxidative stress. Strains bearing single mutations in these genes are more sensitive to oxidative stress compared to wild type, and the single mutants of gaba-t and ssadh can no longer grow on GABA as their sole nitrogen source. However, a definitive physiological role of GABA in yeast, other fungi and most of other organisms remains understood for other stress conditions. Heat shock and yeast thermotolerance Induced thermotolerance, or the increased resistance of cells and tissues to severe 16 (or lethal) heat shock following a prior exposure to mild (sublethal) heat shock has been described in all living organisms from prokaryotes (Morozov et al., 1997) to eukaryotes (Mager and Moradas, 1993). Direct exposure to elevated lethal temperature can damage major components of cells such as proteins and membranes. Prior exposure to a mild heat shock can induce heat shock proteins (HSP) and other cellular changes, thereby making cells more resistant to a subsequent, even more severe heat shock. For yeast Saccharomyces cerevisiae, the optimal growth temperature is within the range of 25 to 35?C, at temperature of 35 to 37?C, yeast cells continue to grow, but are moderately stressed, developing a protective tolerance against elevated lethal temperature. At temperatures above 45?C, yeast cells are severely stressed, and 99% of growing non- adapted aerobic yeast cells die after a 5 min exposure at 50?C (Davidson et al., 2001a, 2001b). Thermotolerant yeasts are considered to have an optimum growth temperature above 40?C (Walker, 1998). Heat-shock is one of the best studied stress-inducible responses, not only of yeast but also of virtually all living organisms. The heat shock response system is understood very well at the DNA level (Craig, 1986; Lindquist and Craig, 1988; Parsell and Lindquist, 1993). For yeasts, the induced thermotolerance acquired from a pre-exposure to a mild heat treatment is related to heat shock factors and stress response element pathways that regulate the synthesis of heat shock proteins (HSPs) (Mager and Moradas, 1993). Miller et al. (1982) have shown a temperature shift from 23 to 37?C in Saccharomyces cerevisiae transiently induces approximately 80 proteins. Twenty of these induced proteins belong to major HSP families. These proteins play important roles in helping cells cope with the toxic effects of high temperatures, but they have different 17 expression patterns and molecular functions. Some of the HSPs have been functionally characterized, but the function of most of the HSPs remains to be understood (Mager and Moradas, 1993). In the yeast Saccharomyces cerevisiae, heat shock proteins of Hsp90 and Hsp70 are important at all temperatures and are required for growth at higher temperature (Borkovich et al., 1989). Sanchez and Lindquist (1990) have shown Hspl04 is induced by heat but expressed at a low level under non stressful temperatures. Hsp104 apparently plays a crucial role in helping cells tolerate short exposures to extreme temperatures. Sanchez and Lindquist (1990) show that with a preheat treatment (30 min at 37?C) followed by stress at 50?C, both wild type and cells bearing a deletion in Hsp 104 acquired thermotolerance, but the hsp104 deletion mutant died 100-1000 fold faster than the wild type cells. In general, heat shock proteins are molecular chaperones required for survival under stress conditions (Brosnan et al., 2000). HSPs confer thermotolerance in organisms by preventing the denaturation of proteins during the heat stress and by facilitating the refolding of already damaged proteins. HSPs are induced in yeast cells under stresses other than heat shock. Parsell et al. (1994) have shown that Hspl04 functions to promote the resolubilization and reactivation of proteins that have unfolded and aggregated after exposure to high temperatures. Hsp70 and Hsp90 prevent aggregation by binding to unfolded proteins, maintaining them in a native, functional state (Parsell and Lindquist, 1993). Other Hsps help to usher unfolded proteins along the degradation pathway. In addition to HSPs, it has been suggested that acquired thermotolerance is mediated by other factors. For example, in heat shock factor (HSF) mutant hsfl-m3 yeast cells, the 18 induction of thermotolerance was not impaired (Smith and Yaffe, 1991). Furthermore, recent work by De Virgilio et al. (1994) has shown that trehalose plays an important role in acquired thermotolerance in S. cerevisiae, and the mutations that block trehalose synthesis sharply reduce thermotolerance. Trehalose is a disaccharide that changes the solvent properties of the fluid phase and reduces the denaturation of proteins at high temperatures (Yancy et al., 1982). Trehalose acts synergistically with HSP104 to protect cells of S. cerevisiae from heat damage (Estruch, 2000). Heat shock and reactive oxygen species The production of reactive oxygen species (ROS) is an unavoidable consequence of life in an aerobic environment. ROS are characterized by their high chemical reactivity and include both free radicals (that is, species with one or more unpaired electrons, such as superoxide (O2 .?) and hydroxyl radicals (OH.)), and non-radical species such as hydrogen peroxide (H2O2) (Shah and Channon, 2004). Indeed, organisms keep a balance between ROS generation and antioxidant systems that scavenge or reduce ROS concentrations. The imbalance caused by over production of ROS and / or reduced antioxidant capabilities generates oxidative stress. However, during episodes of environmental stresses such as high temperature or during the fermentation process, ROS levels increase dramatically, which can result in significant damage to cellular components including DNA fragmentation, protein or enzyme inactivation, the modification of carbohydrate compounds, and changes in membrane fluidity via lipid peroxidation (Elisa et al., 2000). Cells are normally able to defend themselves against ROS damage through the use of detoxifying enzymes such as 19 superoxide dismutase, catalase, and glutathione peroxidase, small molecule antioxidants such as ascorbic acid (vitamin C), uric acid, and glutathione, polyphenol antioxidants assist in preventing ROS damage by scavenging free radicals (Arrigo et al., 2002). Heat stress has been documented to produce oxidative stress (Davidson et al., 1996; Davidson and R. H. Schiestl, 2001a; 2001b), and to some extent, heat stress is equivalent to oxidative stress ( Sugiya et al., 2000). For example, Collinson and Dawes (1992) have shown heat stress induces HSPs as well as a set of antioxidant enzyme genes such as catalase (CTT1), thioredoxin peroxidase (TPX1 and TPX2), and cytochrome c peroxidase (CCP1). Mutant cells with deletion of protein antioxidant enzymes including catalase, cytochrome c peroxidase, superoxide dismutase, and thioredoxin peroxidase are more sensitive to heat stress compared to the wild type ( Davidson et al., 1996). Bouch? et al. (2003) show disfunction of mitochondrial succinate semialdehyde dehydrogenase in plants caused cells to be very sensitive to heat stress and over- accumulation of ROS, suggesting a role of GABA shunt in thermotolerance by restricting the production of ROS. Krahe et al. (1996) found aerobic exposure of yeast cells to higher temperatures increases the concentration of intracellular oxygen, consequently increasing the production and reactivity of ROS (Issels et al., 1986). In addition, it has been found that H2O2 pretreatment strongly induces many of HSPs (Godon et al., 1998). Thus, some overlap must exist between mechanisms of heat and oxidative stress supporting the concept that both HSPs and antioxidants contribute to the thermotolerance of yeast cells (Jamieson, 1995). In respiring yeast cells, the main source of ROS production is the mitochondrial electron transport chain, the principal site being proximal to the cytochrome c oxidase 20 complex (Guidot et al., 1993). In the cytosol, ROS such as H2O2 and superoxide may be formed by P450 cytochromes and dioxygenases (Dalton et al., 1999), or generated during glycolytic fermentative metabolism (Hamm-Kunzelmann et al., 1997). In mammalian cells, Halliwell and Gutteridge (1989) indicated the major source of ROS is mitochondria, other potential ROS sources include, xanthine oxidase, cytochrome P450 based enzymes, NADPH oxidases, dysfunctional NO synthases, peroxisomes and infiltrating inflammatory cells. However, a major ROS source for redox signalling is NADPH oxidases (Shah and Channon, 2004). In plant cells, ROS are generated at a number of cellular sites, predominantly in mitochondria, chloroplasts, and peroxisomes. Another source includes the extracellular side of the plasma membrane (Laoli et al., 2004). Heat shock and the expression of yeast GABA shunt genes The three genes GAD1, UGA1 and UGA2 in the GABA shunt pathway play important role in the metabolism of GABA. To cope with the damaging effects from the environmental and physiological stresses such as heat stress, osmotic stress, or oxidative stress, yeast cells have developed rapid molecular responses by changing their gene expression patterns (Estruch, F.. 2000). In response to heat stress, heat shock proteins may not be the major factor regulating thermotolerance. Expression analysis upon environmental changes in yeast has been investigated by several scientists using the technique of DNA microarray. Gasch et al. (2000) show in response to mild heat stress 37?C shifting from various temperatures from 17?C to 33?C, all the three enzymes in the GABA shunt pathway were similarly up-regulated at least 3 fold upon heat stress at 37?C in a time dependent manner. A genome wide gene profiling conducted by Sakaki et al. 21 (2003) indicated that 104 genes were up-regulated and 287 genes were down-regulated under mild heat stress by shifting cells from 25?C to 37?C, among the down-regulated or repressed genes are mitochondrial related, among the up-regulated genes are heat shock proteins and antioxidant genes, the three stress responsive genes (GAD1, UGA1 and UGA2) in GABA shunt pathway, and also genes involved in production of oxaloacetic acid (OAA) and acetyl-coenzyme A (CoA) were induced, therefore an increased synthesis of ?-ketoglutarate may occur through TCA cycle leading to an elevated level of glutamate, which may subsequently stimulate GABA synthesis. These results further indicated that cells adapted to heat stress by down-regulation of mitochondrial genes to avoid heat induced ROS and also by up-regulation of genes to activate metabolic pathways. However, the effect of lethal temperature on the gene expression pattern and the specific response of GABA shunt remain to be investigated. Under other environmental stress conditions, Coleman et al (2003) demonstrated UGA1p and UGA2p together with GAD1p are oxidative stress responsive proteins. By screening among different nitrogen sources, Ramos et al. (1985), Andre et al. (1993) Talibi et al.(1995) and Patrice Godard et al. (2007) found the expression of UGA1 and UGA2 genes were induced 30 - 240 fold by the presence of GABA as sole nitrogen source instead of ammonium sulfate; According to Coleman et al. (2001), the expression of UGA2 was induced 3- fold when yeast cells were exposed to 1 mM H2O2 stress, however, information obtained on the expression of these three genes (GAD1, UGA1 and UGA2) under stress conditions is limited. 22 The expression of genes regulating GABA metabolism related genes In addition to the environmental changes, the expression of GABA shunt genes is also under the control of two other regulatory genes: UGA3 and UGA4. UGA3 gene is a transcription factor which regulates the expression of GABA metabolism related UGA genes such as UGA1, UGA2 and UGA4 (Coornaert et al., 1991), UGA35 or DAL81 gene is a general positive regulator of genes involved in nitrogen utilization related to metabolisms of GABA, urea, arginine and allatoin (Vissers et al., 1990). Together with UGA35p/DAL81p, UGA3p controls the gene expression by recognizing the promoter elements centered around a GATAA sequence involving in the metabolism of poor nitrogen source such as GABA (Cunningham et al., 1994). In yeast, UGA3p is required for the transcriptional activation of UGA1 and UGA4 (Vissers et al., 1989), deletion of either UGA3 or UGA35 impairs the expression of UGA1 and UGA4 (Vissers et al., 1990). Gasch et al. (2000) has found the expression of UGA3 was strongly induced by nitrogen depletion but unaffected by mild heat stress at 37?C. In yeast, GABA is imported into cells by three proteins: the general amino acid permease (GAP1), proline specific permease (PUT4p) and the GABA permease (UGA4p) (Grenson et al., 1987). The expression of the UGA4 gene was induced by GABA as sole nitrogen source (Ramos et al., 1985), but also dependent on the cell growth conditions (Moretti et al., 1998). As reviewed above, the constitutive expression of UGA4 requires two positive-acting proteins, the specific UGA3p and the pleiotropic UGA35p/DAL81p (Andre et al., 1995; Garc?a et al., 2000). By measuring ?-galactosidase activity in the cells carrying a UGA4::lacZ fusion gene, Moretti et al. (2001) have shown the expression of UGA4 was induced in response to acid pH (4.0) medium in the condition without the 23 repression by UGA43 repressor factor or induction by GABA. Previous studies have demonstrated cells with the mutation of UGA1 or UGA2 are unable to grow on GABA as the sole nitrogen source (Ramos et al., 1985; Coleman et al., 2001), Andre et al. (1993) found mutation of UGA4 has the same phenotype if with the mutation of the other GABA transporter genes GAP1 and PUT4. From Gash et al.'s (2000) expression investigation, in response to mild heat stress at 37?C, slight induction (~1.5 fold) was observed after 5 min, but this induction was only temporary. In contrast, the expression of UGA4 was induced after the temperature was shifted from 37?C to 25?C for 45 min. Yeast is powerful genetic tool "Yeast" is often taken as a synonym for S. cerevisiae (Kurtzman, 1994). It is unicellular, and is a proven model eukaryote for molecular and cellular biology studies. It is well known that yeast has a small genome, grows fast, and is easy to manipulate which makes S. cerevisiae a popular tool for genome wide studies. The genome has been sequenced, and the corresponding databases are available online. The most comprehensive database about yeast genes and research references is provided by Saccharomyces Genome Database. By the strategy of homologous recombination, knockouts of almost every yeast gene have been constructed (as heterozygous diploids for essential genes; as homozygous diploids and haploids for the others), and are available from American Type Culture Collection (ATCC). These collections have been widely used for numerous genome wide studies, or for competition studies using the specific ?bar codes? of each disruption for identification (Fr?hlich et al., 2007). Yeast has also been widely used for protein-protein interactions by two-hybrid or 24 synthetic lethality studies. Because of its simple genome, yeast provides a "clean background" for expressing genes from other eukaryotic systems such as humans or plants. Yeast also provides a convenience for knocking out genes which have only single copies in yeast but exist in homologues of multiple isoforms in other organisms such as plants, which is still lack of an effective gene knockout strategy for functional characterization under stress conditions. Because of these advantageous features, yeast has become the model organism for medical, applied, and fundamental research (Mager and Winderickx, 2005). 25 LITERATURE CITED Andre, B. and J. C. Jauniaux. 1990. Nucleotide sequence of the yeast UGA1 gene encoding GABA transaminase. Nucleic Acids Res. 18(10): 3049. Andre, B., C. Hein, M. Grenson, and J. C. Jauniaux. 1993. Cloning and expression of the UGA4 gene coding for the inducible GABA-specific transport protein of Saccharomyces cerevisiae. Mol. Gen. Genet. 237:17-25. Ansari, M. I., R. H. Lee, and S. G. Chen. 2005. A novel senescence-associated gene encoding ?-aminobutyric acid (GABA):pyruvate transaminase is upregulated during rice leaf senescence. Physiologia Plantarum. 123(1): 1?8. Apel, K., and H. Hirt. 2004. REACTIVE OXYGEN SPECIES: Metabolism, Oxidative Stress, and Signal Transduction Annual Review of Plant Biology.Vol. 55: 373-399. Arazi, T., G. Baum, W. A. Snedden, B. J. Shelp, and H. Fromm. 1995. Molecular and biochemical analysis of calmodulin interactions with the calmodulin-binding domain of plant glutamate decarboxylase. Plant Physiol. 108: 551?561. Arrigo, A. P., C. Paul, C. Ducasse, O. Sauvageot, and C. Kretz-Remy. 2002. Small stress proteins: modulation of intracellular redox state and protection against oxidative stress. In A.-P. Arrigo and W.E.G. Muller (eds.), Small stress proteins. Springer-Verlag Berlin, Germany. p171-184. Attwell, D., B. Barbour, and M. Szatkowski. 1993. Nonvesicular release of neurotransmitter. Neuron. 11: 401?407. Aurisano, N., A. Bertani and R. Regianni. 1995. Involvement of calcium and calmodulin in protein and amino acid metabolism in rice roots under anoxia. Plant Cell Physiol. 36: 1525?1529. Baldy, P. 1975. Metabolisme du c-aminobutyrate chez Agaricus bisporus. I. La l- glutamate-carboxy-lyase. Physiologia Plantarum. 34: 365?372. Baldy, P. 1977. Metabolisme du c-aminobutyrate chez Agaricus bisporus. III. La succinate-semialdehyde: NAD(P)+ oxydoreductase. Physiologia Plantarum 40: 91?97. Baldy, P.. 1976. Metabolism of ?-aminobutyrate in Agaricus bisporus Lge. II. c- Aminobutyrate: a-ketoglutarate aminotransferase. Planta. 130: 275?281. Balzan, R., K. Sapienza, D. R. Galea, N. Vassallo, H. Frey, and W. H. Bannister. 2004. Aspirin commits yeast cells to apoptosis depending on carbon source, Microbiology. 150: 109?115. 26 Bartsch, K., A. Johnn-Marteville, and A. Schulz. 1990. Molecular analysis of two genes of the Escherichia coli gab cluster: nucleotide sequence of the glutamate:succinic semialdehyde transaminase gene (gabT) and characterization of the succinic semialdehyde dehydrogenase gene (gabD). J. Bacteriol. 172: 7035?7042. Baum, G., Y. Chen, T. Arazi , H. Takatsuji, and H. Fromm. 1993. A plant glutamate decarboxylase containing a calmodulin-binding domain. J. Biol. Chem. 268: 19610? 19617. Belhage B., G. H. Hansen, and A. Schousboe. 1993. Depolarization by K+ and glutamate activates different neurotransmitter release mechanisms in GABAergic neurons: Vesicular versus non-vesicular release of GABA. Neuroscience 54: 1019?1034. Berm?dez, M., S. Moretti, C. Garc?a, and A. Batlle. 1998. UGA4 gene expression in Saccharomyces cerevisiae depends on cell growth conditions. Cell. Mol. Biol. 44: 585? 590. Biase, D. D., A. Tramonti, R. A. John, and F. Bossa. 1996. Isolation, overexpression, and biochemical characterization of the two isoforms of glutamic acid decarboxylase from Escherichia coliProtein Expression Purif. 8: 430-438. Boehm, S. L., I. Ponomarev, Y. A. Blednov, and R. A. Harris. 2006. "From gene to behavior and back again: new perspectives on GABAA receptor subunit selectivity of alcohol actions". Adv. Pharmacol. 54: 171-203 Borkovich, K. A., F. W. Farrelly, D. B. Finkelstein, J. Taulien, and S. Lindquist. 1989. Mo. Cell. Bio. 9: 3919-3930. Bouch? N, A. Fait, D. Bouchez, S. G. M?ller, and H. Fromm. 2003. Mitochondrial succinic-semialdehyde dehydrogenase of the ?-aminobutyrate shunt is required to restrict levels of reactive oxygen intermediates in plants. Proc. Natl. Acad. Sci. U.S.A 100: 6843? 6848. Bouch?, N., and H. Fromm. 2004. GABA in plants: just a metabolite? Trends in Plant Sci. 9(3): 110-115. Bouch?, N., B. Lacombe, and H. Fromm. 2003. GABA signaling: a conserved and ubiquitous mechanism. Trends Cell Biol. 13: 607?610. Bown, A. W., and Shelp B. J. 1997. The metabolism and function of y-aminobutyric acid. Plant Physiol. 115: 1-5. Bown, A., and Shelp B. J.. 1989. The metabolism and physiological roles of ?-amino butyric acid. Biochem (Life Sci Adv). 8: 21-2. 27 Brace J. L., W. D. J. Van der, and C. M. Rudin. 2005. SvfI inhibits reactive oxygen species generation and promotes survival under conditions of oxidative stress in Saccharomyces cerevisiae. Yeast 22: 641-652. Breitkreuz K. E., and B. J. Shelp. 1995. Subcellular compartmentation of the 4- aminobutyrate shunt in protoplasts from developing soybean cotyledons. Plant Physiol. 108: 99?103. Breitkreuza, K. E., B. J. Shelp, W. N. Fischera, R. Schwackea, and D. Rentsch. 1999. Identification and characterization of GABA, proline and quaternary ammonium compound transporters from Arabidopsis thaliana. FEBS Lett. 450: 280?2. Brosnan, M. P., D. Donnelly, T. C. James, and U. Bond. 2000.The stress response is repressed during fermentation in brewery strains of yeast. Appl. Microbiol. 88: 746-755. Bu, D. F., M. G. Erlander, B. C. Hitz, N. J. Tillakaratne, D. L. Kaufman, C. B. Busch, K. B., and H. Fromm. 1999. Plant succinic semialdehyde dehydrogenase. Cloning, purification, localization in mitochondria, and regulation by adenine nucleotides. Plant Physiol. 121: 589?597. Buzenet A. M., C. Fages, T. M. Bloch, and P. Gonnard. 1978. Purification and properties of 4-aminobutyrate 2-ketoglutarate aminotransferase from pig liver. Biochem. Biophys. Acta. 522: 400-411. Carratore, R. D., C. Della Croce, M. Simili, E. Taccini, M. Scavuzzo, and S. Sbrana, 2002. Cell cycle and morphological alterations as indicative of apoptosis prompted by UV irradiation in S. cerevisiae. Mutat. Res. 513: 183-191. Castanie-Cornet, M. P., T. A. Penfound, D.Smith, J. F. Elliott, and J. W. Foster. 1999. Control of acid resistance in Escherichia coli. J. Bacteriol. 181: 3525?3535. Chambliss, K. L., D. L. Caudle, D. Hinson, C. R. Moomaw, C. A. Slaaughter, C. Jacobs, K. M. Gibson. 1995. Molecular Cloning of the mature NAD-dependent Succinic Semialdehyde Dehydrogenase from Rat and Human cDNA ISOLATION, EVOLUTIONARY HOMOLOGY, AND TISSUE EXPRESSION. Journal of biological chemistry. 270: 461-467. Chambliss, K. L., and K. M. Gibson. 1992. Succinic semialdehyde dehydrogenase from mammalian brain: subunit analysis using polyclonal antiserum.Int. J. Biochem. 24: 1493- 1499. Chen, W., P. Linko, and M. Milner. 1960. On the nature of glutamic acid decarboxylase in wheat embryos. Plant Physio. l35: 68-71. 28 Chen, Y., G. Baum, and H. Fromm. 1994. The 58-kD calmodulin-binding glutamate decarboxylase is a ubiquitous protein in petunia organs and its expression is developmentally regulated. Plant Physiol. 106: 1381?1387. Choi, S. Y., I. Kim, S. H. Jang, S. J. Lee, M. S. Song, Y. S. Lee, and S. W. Cho. 1993. Purification and properties of GABA transaminase from bovine brain. Mol. Cells. 3: 397?401. Chung I., A. W. Bown, and B. J. Shelp. 1992. The production and efflux of 4- aminobutyrate in isolated mesophyll cells. Plant Physiol. 99: 659?664. Coleman S. T., T. K Fang ., S. A. Rovinsky, F. J. Turano, and W. S. Moye-Rowley. 2001. Expression of a glutamate decarboxylase homologue is required for normal oxidative stress tolerance in Saccharomyces cerevisiae. J Biol Chem. 276(1): 244-50. Collinson, L. P., and I. W. Dawes. 1992. Inducibility of the response of yeast cells to peroxide stress. J. Gen. Microbiol. 138: 329-335. Coornaert, D., S. Vissers, and B. Andre. 1991. The pleiotropic UGA35(DURL) regulatory gene of Saccharomyces cerevisiae: cloning, sequence and identity with the DAL81 gene.Gene (Amst.) 97: 163-171. Cozzani I., Fazio A. M., Felici E., and G. Barletta. 1980. Separation and characterization of NAD- and NADP-specific succinate-semialdehyde dehydrogenase from Escherichia coli K-12 3300. Biochim Biophys Acta. 613(2): 309-317. Craig, E. A. 1986. The heat-shock response. Crit Rev Biochem 38: 239-280. Crawford L., A. Bown, K. Breitkreuz, and F. Buinel. 1994. The synthesis of gamma- arninobutyric acid in response to treatments reducing cytosolic pH. Plant Physiol 104: 865-871. Cunningham, T. S., R. A. Dorrington, and T. G. Cooper. 1994. The UGA4 UASNTR site required for GLN3-dependent transcriptional activation also mediates DAL80- responsive regulation and DAL80 protein binding in Saccharomyces cerevisiae. J. Bacteriol. 176: 4718-4725. Dalton, T. P., H. G. Shertzer, and A. Puga. 1999. Regulation of gene expression by reactive oxygen. Annu Rev Pharmacol Toxicol. 39: 67?101. Danial, N. N., and S. J. Korsmeyer. 2004. Cell death: critical control points. Cell. 116: 205?219. Davidson, J. F., and R. H. Schiestl. 2001a. Cytotoxic and genotoxic consequences of heat stress are dependent on the presence of oxygen in Saccharomyces cerevisiae. J. Bacteriol. 183: 4580-4587. 29 Davidson, J. F., and R. H. Schiestl. 2001b. Mitochondrial respiratory electron carriers are involved in oxidative stress during heat stress in Saccharomyces cerevisiae. Mol. Cell. Biol. 21: 8483-8489. Davidson, J. F., B. Whyte, P. H. Bissinger, and R. H. Schiestl. 1996. Oxidative stress is involved in heat-induced cell death in Saccharomyces cerevisiae. Proc. Natl. Acad. Sci. USA. 93: 5116-5121. De Virgilio, C., T. Hottiger, J. Dominguez, T. Boller, and A. Wiemken. 1994. The role of trehalose synthesis for the acquisition of thermotolerance in yeast. II. Physiological concentrations of trehalose increase the thermal stability of proteins in vitro. Eur. J. Biochem. 219: 179-186. Der Garabedian, P. A., L. Lotti, and J. J. Vermeersch. 1986. 4-Aminobutyrate:2- oxoglutarate aminotransferase from Candida: purification and properties. European Journal of Biochemistry. 156: 589?596. Dimitrijevic, N., S. Dzitoyeva, R. Satta, M. Imbesi, S. Yildiz, and H. Manev. 2005. "Drosophila GABAB receptors are involved in behavioral effects of gamma hydroxybutyric acid (GHB)". Eur. J. Pharmacol. 519 (3): 246-52. Donnelly M. I., and Cooper R. A. 1981. Two succinic semialdehyde dehydrogenases are induced when Escherichia coli K-12 Is grown on gamma-aminobutyrate. J Bacteriol. 145(3): 1425-1427. Donnelly, M.I., and R. A. Cooper. 1981. Succinic semialdehyde dehydrogenases of Escherichia coli. Their role in the degradation of p-hydroxyphenylacetate and ?- aminobutyrate. Eur. J. Biochem. 113: 555?561. Dover, S., and Y. S. Halpern. 1972. Control of the Pathway of ?-Aminobutyrate Breakdown in Escherichia coli K-12. J Bacteriol. 110(1): 165?170. Duncan, R. J. S. and K. F. Tipton. 1971.The kinetics of pig brain aldehyde dehydrogenase, Eur. J. Biochem. 22: 538?543. Dzitoyeva, S., N. Dimitrijevic, and H. Manev. 2003. "Gamma-aminobutyric acid B receptor 1 mediates behavior-impairing actions of alcohol in Drosophila: adult RNA interference and pharmacological evidence". Proc. Natl. Acad. Sci. U.S.A. 100(9): 5485- 90. Elisa, C., P. Eva, E. Pedro, H. Enrique, and R. Joaquim. 2000. Oxidative damage stress promotes specific protein damage in Saccharomyces cerevisiae. J. Biol. Chem. 275: 27393-27398. 30 Erdo, S. L. and B. Kiss. 1986. Presence of GABA, glutamate decarboxylase, and GABA transaminase in peripheral tissues: a collection of quantitative data. p. 5-17. In S. L. Erdo, and N. G. Bowery (ed.) GABAergic Mechanisms in the Mammalian Periphery. Raven Press, New York. Erlander M. G., N. J. K. Tillakaratne, S. Feldblum, N. Patel, and A. J. Tobin. 1991. Two genes encode distinct glutamate decarboxylases. Neuron 7: 91-100. Estruch, F. 2000. Stress-controlled transcription factors, stress-induced genes and stress tolerance in budding yeast. FEMS Microbiol. Rev. 24: 469-486. Fleury, C., B. Mignotte, and J. L. Vayssiere. 2002. Mitochondrial reactive oxygen species in cell death signaling. Biochimie. 84 (2-3): 131?141. Foerster, C. W., and H. F. Foerster. 1973. Glutamic acid decarboxylase in spores of Bacillus megaterium and its possible involvement in spore germination. Journal of Bacteriology 114: 1090?1098. Fonda, M. L. 1972. Glutamate decarboxylase. Substrate specificity and inhibition by carboxylic acids Biochemistry. 11(7): 1304-1309. Fonda, M. L. 1985. L-Glutamate decarboxylase from bacteria. Methods Enzymol. 113: 11-16. Fonnum, F.. 1987. Biochemistry, anatomy, and pharmacology of GABA neurons. In Psychopharmacology: the Third Generation of Progress (Metzler, H.Y., ed.). 173?182. Raven Press, New York. Fr?hlich, K. U., H. Fussi, and C. Ruckenstuhl. 2007. Yeast apoptosis?From genes to pathways. Seminars in Cancer Biology. 17(2): 112-121. Fr?hlich, K.-U., and F. Madeo. 2000. Apoptosis in yeast - a monocellular organism exhibits altruistic behaviour. FEBS Lett. 473: 6-9. Gallego P.P., L. Whotton, S. Picton, D. Grierson, and J. E. Gray. 1995. A role for glutamate decarboxylase during tomato ripening: The characterisation of a cDNA encoding a putative glutamate decarboxylase with a calmodulin-binding site. Plant molecular biology. 27(6): 1143-1151. Galleschi L., C. Nocchi, C. Floris, G. Bedini, M. C. Anguillesi, and 1. Griiii. 1983. Succinic semialdehyde dehydrogenase in higher plants: purification and properties of the enzyme from Triticum dunrm embryos. Biochem Physiol Pflanzen. 178: 645-651. Garcia, E. and R. Lopez. 1995. Streptococcus pneumoniae type 3 encodes a protein highly similar to the human glutamate decarboxylase (GAD65). FEMS Microbiol. Lett. 133: 113-118. 31 Garc?a, S. C., M. B. Moretti, and A. Batlle. 2000. Constitutive expression of the UGA4 gene in Saccharomyces cerevisiae depends on two positive-acting proteins, Uga3p and Uga35p. FEMS Microbiol. Lett. 184: 219?224. Gasch, A. P., P. T. Spellman, C. M. Kao, O. Carmel-Harel, M. B. Eisen, G. Storz, D. Botstein, and P. O. Brown. 2000. Genomic expression programs in the response of yeast cells to environmental changes. Mol Biol Cell. 11(12): 4241-57. Gibson, K. M., E. Chistensen, C. Jakobs, B. Fowler, M. A. Clarke, G. Hammersen, K. Raab, J. Kobori, A. Moosa, D. Vollmer, E. Rossier, A. K. Iafolla, D. Matern, O. F. Brouwer, J. Finkelstein, F. Aksu, H. P. Weber, J. A. J. M. Bakkeren, F. J. M. Gabreels, D. Bluestone, T. F. Barron, P. Beauvais, D. Rabier, C. Santos, R. Umansky, and W. Lehnert. 1997. The clinical phenotype of succinic semialdehyde dehydrogenase deficiency (4-hydroxybutyric aciduria): case reports of 23 new patients, Pediatrics. 99: 567?574. Gibson, K. M., G. F. Hoffmann, A. K. Hodson, T. Bottiglieri, and C. Jakobs. 1998. 4-Hydroxybutyric acid and the clinical phenotype of succinic semialdehyde dehydrogenase deficiency, an inborn error of GABA metabolism, Neuropediatrics 29: 14?22. Godon, C., G. Lagniel, J. Lee, J. M. Buhler, S. Kieffer, M. Perrot, H. Boucherie, M. B. Toledano, and J. Labarre. 1998. The H2O2 stimulon in Saccharomyces cerevisiae. J Biol Chem 273(35): 22480?22489. Godard, P., A. Urrestarazu, S. Vissers, K. Kontos, G. Bontempi, J. V. Helden, and B. Andr?1. 2007. Effect of 21 Different Nitrogen Sources on Global Gene Expression in the Yeast Saccharomyces cerevisiae. Mol Cell Biol. 27(8):3065-86 Guidot, D. M., J. M. McCord, R. M. Wright, and J. E. Repine. 1993. Absence of electron transport (Rho 0 state) restores growth of a manganese superoxide dismutase- deficient Saccharomyces. J. Biol. Chem.. 268(35): 26699-26703. Grenson, M., F. Muyldermans, K. Broman, and S. Vissers. 1987. 4-Aminobutyric acid (GABA) uptake in baker?s yeast Saccharomyces cerevisiae is mediated by the general amino acid permease, the proline permease and a specific permease integrated into the GABA-catabolic pathway. Biochemistry Life Sci Adv 6: 35?39. Halliwell, B., and J. M. C. Gutteridge. 1989. Free Radicals in Biology and Medicine, 2nd ed., Oxford: Clarendon. Hamm-Kunzelmann, B., D., Schafer, C. Weigert, and K. Brand. 1997. Redox- regulated expression of glycolytic enzymes in resting and proliferating rat thymocytes. FEBS Lett. 403(1): 87?90. 32 Hao, R. and J. C. Schmit. 1991. Purification and characterisation of glutamate decarboxylase from Neurospora crassa conidia. Journal of Biological Chemistry. 266: 5135?5139. Hao, R. and J. C. Schmit. 1993. Cloning of the gene for glutamate decarboxylase and its expression during conidiation in Neurospora crassa. Biochemical Journal 293: 735?738. Huh, G. H., B. Damsz, T. K. Matsumoto, M. P. Reddy, A. M. Rus, J. I. Ibeas, M. L. Narasimhan, R. A. Bressan, and P. M. Hasegawa. 2002. Salt causes ion disequilibrium-induced programmed cell death in yeast and plants. Plant J. 29: 649-659. Huh, W. K., J. V. Falvo, L. C. Gerke, A. S. Carroll, R. W. Howson, J. S. Weissman, and E. K. O'Shea. 2003. Global analysis of protein localization in budding yeast. Nature. 425(6959): 686-691. Inatomi K., and J. C. Slaughter. 1975. Glutamate decarboxylase from barley embryos and roots. General properties and the occurrence of three enzymic forms. Biochem. J. 147: 479-484. Intaomi, K., and J. C. Slaughter. 1975. Glutamate decarboxylase from barley embryos and roots. Biochem J. 147: 479-484. Issels, R. D., R. M. Fink, and E. Lengfelder. 1986. Effects of hyperthermic conditions on the reactivity of oxygen radicals. Free Rad Res Comm. 2(1?2): 7?18. Jamieson, D. J. 1995. The effect of oxidative stress on Saccharomyces cerevisiae. Redox Rep 1: 89?95. Jin, H. N., M. J. Dilworth, and A. R. Glenn. 1990. 4-aminobutyrate is not available to bacteroids of cowpea Rhizobium MNF2030 in snake bean nodules Arch Microbiol.. 153: 455-462. John, R. A. and L. J. Fowler. 1976. Kinetics and spectral properties of rabbit brain 4- aminobutyrate aminotransferase. Biochem. J. 155: 645?651. Jung, M. J., B. W. Metcalf, B. Lippert, and P. Casara. 1978. Mechanism of the stereospecific irreversible inhibition of bacterial glutamic acid decarboxylase by (R)-(-)- 4-aminohex-5-ynoic acid, an analog of 4-aminobutyric acid. Biochemistry. 17(13): 2628 - 2632. Kaufmann, S. H., and Hengartner M. O. 2001. Programmed cell death: alive and well in the new millennium.Trends Cell Biol. 11: 526- 534. King, D.A., D. M. Hannum, J. S. Qi, and J. K. Hurst. 2004. HOCl-mediated cell death and metabolic dysfunction in the yeast Saccharomyces cerevisiae, Arch Biochem Biophys. 423: 170?181. 33 Kinnersley, A. M., and F. J. Turano. 2000. Gamma Aminobutyric Acid (GABA) and Plant Responses to Stress. Critical Reviews in Plant Sciences. 19(6): 479-509. Kishnaswamy, P. R., and K. V. Giri. 1956. Glutamic acid decarboxylase in Rhodotorula glutinis. Biochemical Journal. 62: 301?303. Knight M. R., A. K. Campbell, S. M. Smith, and A. J. Trewavas. 1991. Transgenic plant aequorin reports the effect of touch and cold shock and elicitors on cytoplasmic calcium. Nature 352: 524?526. Krahe, M., G. Antranikian, and H. M?arkl. 1996. Fermentation of extremophilic micro-organisms. FEMS Microbiol Rev 18: 271?285. Krishnaswamy, P. R., and K. V. Giri. 1953. The occurrence of 4-aminobutyric acid and glutamic acid decarboxylase in Red yeast (Rhodotorula glutinis). Current Science. 22: 143?144. Kroemer, G., and S. J. Martin. 2005. Caspase-independent cell death. Nat Med. 11(7): 725-30. Kubicek, C. P., W. Hampel, and M. Rohr. 1979. Manganese deficiency leads to elevated amino acid pools in critic acid accumulating Aspergillus niger. Archives of Microbiology 123: 73?79. Kumar, S. and N. S. Punekar. 1994. Succinicsemialdehyde dehydrogenase from Aspergillus niger. A kinetic enquiry. Abstract P7-343, XVI IUBMB. International Congress, New Delhi. Kuo, D. and R. R. Rando. 1981. Irreversible inhibition of glutamate decarboxylase by alpha.-(fluoromethyl)glutamic acid. Biochemistry. 20(3): 506-511. Kurtzman, C.P.. 1994. Molecular taxonomy of the yeasts. Yeast. 10(13): 1727-1740 . Lacombe B., D. Becker, R. Hedrich, R. DeSalle, M. Hollmann, J.M. Kwak, J.I. Schroeder, N. Le Novere, H.G. Nam, E.P. Spalding, M. Tester, F.J. Turano, J. Chiu, and G. Coruzzi. 2001. The identity of plant glutamate receptors. Science 292: 1486-1487. Laloi, C., K. Apel and A. Danon. 2004. Reactive oxygen signalling: the latest news. Current Opinion in Plant Biology. 7(3): 323-328. Laun, P., A. Pichova, F. Madeo, J. Fuchs, A. Ellinger, S. Kohlwein, I. Dawes, K. U. Fr?hlich, and M. Breitenbach. 2001. Aged mother cells of Saccharomyces cerevisiae show markers of oxidative stress and apoptosis. Mol Microbiol. 39: 1166-1173. 34 Lee, B. R., J. W. Hong, B. K. Yoo, S. J. Lee, S. W. Cho and S. Y. Choi. 1995. Bovine brain succinic semialdehyde dehydrogenase: purification, kinetics and reactivity of lysyl residues connected with catalytic activity. Mol. Cells. 5: 611?617. Ligr, M., F. Madeo, E. Frohlich, W. Hilt, K. U. Frohlich, and D. H. Wolf. 1998. Mammalian Bax triggers apoptotic changes in yeast. FEBS Lett. 438: 61-65. Likos, J. J., H. Ueno, R. W. Feldhaus and D. E. Metzler. 1982. A novel reaction of the coenzyme of glutamate decarboxylase with L-serine O-sulfate Biochemistry. 21: 4377- 4386. Lindquist, S., and E. A. Craig. 1988.The heat-shock proteins. Annu Rev Genet 55: 1151-1191. Lindquist, S., and G. Kim. 1996. Heat-shock protein 104 expression is sufficient for thermotolerance in yeast. Proc Nat Acad Sci 93: 5301?5306. Ling V., W. A. Snedden, B. J. Shelp, and S. M. Assmann. 1994. Analysis of a soluble calmodulin-binding protein from fava bean roots: identification of glutamate decarboxylase as a calmodulin-activated enzyme. Plant Cell. 6: 1135?1143 Lockshin, R. A., and C. M. Williams. 1964. "Programmed cell death?II. Endocrine potentiation of the breakdown of the intersegmental muscles of silkmoths". Journal of Insect Physiology. 10(4): 643-649. Locy R. D., S. J. Wu, J. Bisnette, T. W. Barger, D. McNabb, M. Zik , H. Fromm, M. R. Knight , J. H. Cherry, N. K. Singh . 2000. The regulation of GABA accumulation by heat stress induced increases in cytoplasmic calcium. In: NATO ARW Series: Plant Tolerance to Abiotoc Stresses in Agriculture: Role of Genetic Engineering. JH Cherry, ed. Longo, V. D., J. Mitteldorf, and V. P. Skulachev. 2005. Programmed and altruistic ageing. Nat. Rev. Genet. 6: 866?872. Longo, V. D., L. Ellerby, D. Bredesen, J. Valentine, and E. Gralla. 1997. Human Bcl- 2 reverses survival defects in yeast lacking superoxide dismutase and delays death of wild-type yeast. J Cell Biol. 137: 1581-1588. Ludovico, P., M. J. Sousa, M.T . Silva, C. Leao, and M. Corte-Real. 2001. Saccharomyces cerevisiae commits to a programmed cell death process in response to acetic acid, Microbiology 147: 2409?2415. Lutke-Eversloh T., and A. Steinbuchel. 1999. Biochemical and molecular characterization of a succinate semialdehyde dehydrogenase involved in the catabolism of 4-hydroxybutyric acid in Ralstonia eutropha.. FEMS Microbiol Lett. 181(1): 63-71. 35 Madeo, F., E. Fr?hlich, and K. U. Fr?hlich. 1997. A yeast mutant showing diagnostic markers of early and late apoptosis. J Cell Biol. 139: 729-734. Madeo, F., E. Fr?hlich, M. Ligr, M. Grey, S. J. Sigrist, D. H. Wolf, and K. U. Fr?hlich. 1999. Oxygen stress: a regulator of apoptosis in yeast. J Cell Biol. 145: 757? 767. Madeo, F., E. Herker, C. Maldener, S. Wissing, S. Lachelt, M. Herlan, M. Fehr, K. Lauber, S. J. Sigrist, S. Wesselborg, and K. U. Frohlich. 2002. A caspase-related protease regulates apoptosis in yeast. Mol Cell. 9(4): 911-917. Madeo, F., E. Herker, S. Wissing, H. Jungwirth, T. Eisenberg, and K. U. Frohlich. 2004. Apoptosis in yeast. Curr. Opin. Microbiol. 7: 655?660. Madeo, F., S. Engelhardt, E. Herker, N. Lehmann, C. Maldener, A. Proksch, S. Wissing, and K. U. Frohlich. 2002. Apoptosis in yeast: a new model system with applications in cell biology and medicine. Curr. Genet. 41: 208?216. Mager, W. H., and F. P. Moradas. 1993. Stress response of yeast. Biochem J. 290: 1? 13. Mager, W. H., and J. Winderickx. 2005. Yeast as a model for medical and medicinal research, Trends Pharmacol Sci 26: 265?273. Mager, W. H., and P. M. Ferreira. 1993. Stress response of yeast. Biochem. J. 290: 1- 13. Maitre, M., L. Ciesielski, C. Cash, and P. Mandel. 1975. Purification and studies on some properties of the 4-aminobutyrate: 2-oxoglutarate transaminase from rat brain. Eur. J. Biochem. 52: 157?169. Marek L. E., and J. M. Henson. 1988. Cloning and expression of the Escherichia coli K-12 sad gene. J. Bacteriol. 170: 991?994. Martin D. L., and K. Rimvall. 1993. Regulation of ?-Aminobutyric Acid Synthesis in the Brain. J. Neurochem. 60: 395?407. Martin D. L., S. B. Martin, S. J. Wu, and N. Espina. 1991. Regulatory properties of brain glutamate decarboxylase (GAD): The apoenzyme of GAD is present principally as the smaller of two molecular forms of GAD in brain. J. Neurosci. 11: 2725?2731. Matsumoto, T., 1. Yamaura, and M. Funastu. 1986. Purification and properties of glutamate decarboxylase from squash. Agric Biol Chem. 50: 1413-1417. 36 Mayer, R. R., J. L. Cherry, and D. Rhodes. 1990. Effects of heat shock on amino acid metabolism of cowpea cells. Phytochemistry 94: 796?810. Medina-Kauwe, L. K., N. J. K.Tillakaratne, J. Y. Wu, and A. J. Tobin. 1994. A rat brain cDNA encodes enzymatically active GABA transaminase and provides a molecular probe for GABA-catabolizing cells. J. Neurochem. 62: 1267-1275. Metzer E., and Y. S. Halpern. 1990. In vivo cloning and characterization of the gabCTDP gene cluster of Escherichia coli K-12. J Bacteriol. 172(6): 3250-6. Mihic, S. J., Q. Ye, M. J. Wick, V. V. Koltchine, M. D. Krasowski, S. E. Finn, M. P. Mascia, C. F. Valenzuela, K. K. Hanson, E. P. Greenblatt, R. A. Harris, and N. L. Harrison. 1997. "Sites of alcohol and volatile anaesthetic action on GABAA and glycine receptors". Nature 389 (6649): 385-9. Miller, M. J., N. H. Xuong, and E. P. Geiduschek. 1982. Quantitative analysis of the heat shock response of Saccharomyces cerevisiae. J. Bacteriol. 151: 311-327. Morozov, II., V. G. Petin, and B. V. Dubovick. 1997. Influence of tonicity and chloramphenicol on hyperthermic cytotoxicity and cell permeability under various heating rates. Int J Hyperther. 13: 49?57. Moretti, M. B., A. Batlle, and S. C. Garcia. 2001. UGA4 gene encoding the ?- aminobutyric acid permease in Saccharomyces cerevisiae is an acid-expressed gene. The International Journal of Biochemistry & Cell Biology. 33(12): 1202-1207. Niegemann, E., A. Schulz, and K. Bartsch. 1993. Molecular organization of the Escherichia coli gab cluster: nucleotide sequence of the structural genes gabD and gabP and expression of the GABA permease gene. Arch. Microbiol. 160: 454?460. Nirenberg, M. W., and W. B. Jakoby. 1960. Enzymatic utilization of ?-hydroxybutyric acid. J. Biol. Chem. 235: 954?960. Oh, S. H., W. G. Choi, I. T. Lee, S. J. Yun. 2005. Cloning and characterization of a rice cDNA encoding glutamate decarboxylase. JOURNAL OF BIOCHEMISTRY AND MOLECULAR BIOLOGY. 38(5): 595-601. Palanivelu, R., L. Brass, A. Edlund, and D. Preuss. 2003. Pollen tube growth and guidance is regulated by POP2, an Arabidopsis gene that controls GABA levels. Cell. 114: 47?59. Palanivelu, R., L. Brass, A. F. Edlund, and D. Preuss. 2003. Pollen tube growth and guidance is regulated by POP2, an Arabidopsis gene that controls GABA levels. Cell. 114: 47?59. 37 Parsell, D. A., A. S. Kowal, M. A. Singer, and S. Lindquist. 1994. Protein disaggregation mediated by heat-shock protein Hsp104.Nature (London) 372: 475-478. Parsell, D. A., and S. Lindquist. 1993.The function of heat-shock proteins in stress tolerance: degradation and reactivation of damaged proteins. Annu Rev Genet 27: 437- 496. Petruszko, R., and L. Fowden. 1961. 4-Aminobutyric acid metabolism in plants. I. Metabolism in yeasts. Annals of Botany 25: 491?511. Pin J. P., and J. Bockaert. 1989. Two distinct mechanisms, differentially affected by excitatory amino acids, trigger GABA release from fetal mouse striatal neurons in primary culture. J. Neurosci. 9: 648?656. Pozniakovsky, A. I., D. A. Knorre, O. V. Markova, A. A. Hyman, V. P. Skulachev, and F. F. Severin. 2005. Role of mitochondria in the pheromone- and amiodarone- induced programmed death of yeast. J Cell Biol 168: 257?269. Prell J., B. Boesten, P. Poole, U. B. Priefer. 2002. The Rhizobium leguminosarum bv. viciae VF39 gamma-aminobutyrate (GABA) aminotransferase gene (gabT) is induced by GABA and highly expressed in bacteroids.Microbiology. 148: 615-623. Ramos F, M. Guezzar, M. Grenson, and J. M. Wiame. 1985. Mutations affecting the enzymes involved in the utilization of 4-aminobutyric acid as nitrogen source by the yeast Saccharomyces cerevisiae. Eur J Biochem 149: 401?404. Ramputh, A. I., and A. W. Bown. 1996. Rapid gama-aminobutyric acid synthesis and the inhibition of the growth and development of oblique-banded leaf-roller larvae. Plant Physiol. 111: 1349-1352. Rating, D., F. Hanefeld, H. Siemes, and J. Kneer. 1984. 4-Hydroxybutyric acidurea : a new inborn error of metabolism. Journal of Inherited Metabolic Research. 7 (Supplement 1): 90?96. Reed, L. J.. 1950. The occurrence of c-aminobutyric acid in yeast extract: its isolation and identification. Journal of Biological Chemistry 183: 451?458. Reggiani, R., N. Aurisano, M. Mattana, and A. Bertani. 1988. Accumulation and interconversion of amino acids in rice roots under anoxia. Plant Cell Physio. l29: 981-987 Rentsch, D., B. Hirner, E. Schmelzer, and W. B. Frommer. 1996. Salt stress-induced proline transporters and salt stress-repressed broad specificity amino acid permeases identified by suppression of a yeast amino acid permease-targeting mutant. Plant Cell. 8: 1437?1446. 38 Roberts, E., P. Ayengar, and I. Posner. 1953. Transamination of ?-aminobutyric acid and ?-alanine in microorganisms. Journal of Biological Chemistry. 203: 195?204. Roberts, J. K. M., J. Calfis, D. Wemmer, and V. Walbot. 1984. Mechanism of cytoplasmic pH regulation in hypoxic maize root tips and its role in survival under hypoxia. Proc Nat Acad Sci USA. 8: 3379-3383. Rolin, D., P. Baldet, D. Just, C. Chevalier, M. Biran, and P. Raymond. 2000. NMR study of low subcellular pH during the development of cherry tomato fruit. Aust. J. Plant Physiol. 27: 61?69. Roth, Robert J., J. R. Cooper, and F. E., Bloom. 2003. The Biochemical Basis of Neuropharmacology. Oxford [Oxfordshire]: Oxford University Press, 416 pages. Ryzlak, M. T., and R. Pietruszko. 1988. Human brain ?high Km? aldehyde dehydrogenase: Purification, characterization, and identification as NAD+-dependent succinic semialdehyde dehydrogenase Arch. Biochem. Biophys. 266: 386-396. Sakaki, K., K. Tashiro, S. Kuhara, and K. Mihara. 2003. Response of genes associated with mitochondrial function to mild heat stress in yeast Saccharomyces cerevisiae. J Biochem (Tokyo). 134(3): 373-84. Sanchez, M., J. Fernandez, M. Martin, A. Gibello, and A. Garrido-Pertierra. 1989. Purification and properties of two succinic semialdehyde dehydrogenases from Klebsiella pneumoniae. Biochim. Biophys. Acta. 990: 225?231. Santosh K., and S. P. Naryan. 1997. The metabolism of 4-aminobutyrate (GABA) in fungi. Mycol. Res. 101(4): 403?409. SatyaNarayan V., and P. M. Nair. 1989. Potato tuber succinic semialdehyde dehydrogenase: Purification and characterization. Arch Biochem Biophys. 275: 469-477. SatyaNarayan V., and P.M. Nair. 1990. Metabolism, enymology and possible roles of 4- aminobutyrate in higher plants. Phytochem 29: 367-375. SatyaNarayan, V., and P. M. Nair. 1986. Enhanced operation of 4-aminobutyrate shunt in y-irradiated potato tuben. Phytochem. 25: 1801?1805. SatyaNarayan, V.,and P. M. Nair. 1985. Purification and characterization of glutamate Schmit, J. C. and S. Brody. 1975. Neurospora crassa conidial germination: role of endogenous amino acid pools. Journal of Bacteriology. 124: 232?242. Schousboe, A., J.Y. Wu, and E. Roberts. 1973. Purification and. characterization. of. the. 4-aminobutyrate-2-ketoglutarate. transaminase from mouse brain. Biochemistry. 12: 2868-2873. 39 Seong G. J., H. B. Jae, S. J., Joong, P. Jinseu, S. K., Oh, W. C., Sung, and Y. C., Soo 2000. Human brain GABA transaminase. Eur. J. Biochem. 267: 5601?5607. Shah, A. M., and K. M. Channon. 2004. Free radicals and redox signalling in cardiovascular disease. Heart. 90(5): 486?487. Shelp B. J., A. W. Bown and M. D. McLean. 1999. Metabolism and functions of gamma-aminobutyric acid. Trends Plant Sci. 4: 446?452. Shelp, B. J., A. W. Bown, and M. D. McLean. 1999. Metabolism and functions of gamma-aminobutyric acid. Trends Plant Sci. 4: 446?452. Shelp, B. J., C. S. Walton, W. A. Snedden, L. G. Tuin, I. J. Oresnik, D. B. Layzell. 1995. GABA shunt in developing soybean seeds is associated with hypoxia. Physiol Plant 94: 219-228 Shmit, J. C. and S. Brody. 1975. Neurospora crassa conidial germination: role of endogenous amino acid pools. Journal of Bacteriology 124: 232?242. Smith D. K., T. Kassam, B. Singh, and J. F. Elliott. 1992. Escherichia coli has two homologous glutamate decarboxylase genes that map to distinct loci. J. Bacteriol. 174: 5820-5826. Smith, B. J., and M. P. Yaffe. 1991. Uncoupling thermotolerance from the induction of heat shock proteins. Proc. Natl. Acad. Sci. USA. 88: 11091-11094. Snchez, Y., and Lindquist, S. L. 1990. HSP104 required for induced thermotolerance. Science 248: 1112-1115. Snedden, W. A., T. Arazi, H. Fromm, and B. J. Shelp. 1995. Calcium/calmodulin activation of soybean glutamate decarboxylase. Plant Physiol. 108: 543?549. Snedden, W. A., K. Nataly, B. Gideon, and H. Fromm. 1996. Activation of a recombinant petunia glutamate decarboxylase by calcium/calmodulin or by a monoclonal antibody which recognizes the calmodulin-binding domain. J. Biol. Chem. 271: 4148? 4153. Snedden, W.A., and H. Fromm. 1999. Regulation of the ?-aminobutyrate-synthesizing enzyme, glutamate decarboxylase, by calcium?calmodulin: a mechanism for rapid activation in response to stress. In: H.R. Lerner, Editor, Plant Responses to Environmental Stresses: From Phytohormones to Genome Reorganization, Marcel Dekker. 549?574. Soghomonia J. J., and D. L. Martin. 1998. Two isoforms of glutamate decarboxylase: why? Trends in Pharmacological Sciences. 19(12): 500-505. 40 Steller, H. 1995. Mechanisms and genes of cellular suicide. Science 267: 1445-1449. Steward F.C., J. F. Thosmpson, and C. E. Dent. 1949. ?-Aminobutyric acid: a constituent of the potato tuber? Science 110 (2861): 439?440. Strasser, A., L. O?Connor, and V. M. Dixit. 2000. Apoptosis signaling. Annu Rev Biochem. 69: 217-245. Strausbauch, P. H. and E. H. Fischer. 1970. Chemical and physical properties of Escherichia coli glutamate decarboxylase. Biochemistry. 9(2): 226- 233. Streeter J. G., and J. F. Thompson. 1972. Anaerobic accumulation of gamma- aminobutyric acid and alanine in radish leaves (Raphonur sativus L.). Plant Physio. l49: 572-578. Sugiyama, K., S. Izawa, and Y. Inoue. 2000. The Yap1p-dependent induction of glutathione synthesis in the heat-shock response of Saccharomyces cerevisiae. J Biol Chem. 275(20): 15535?15540. Sukhareva, V. S., and A. E. Braunshtein. 1971. Investigation of the nature of the interactions of glutamate decarboxylase from Escherichia coli with the substrate and its analogs. Mol. Biol. (Moscow). 5: 241-252. Sweetman, K. M. L., W. L. Nyhan, C. Jakobs, D. Rating, H. Siemes, and F. Henefeld. 1983. Succinic semialdehyde dehydrogenase deficiency: an inborn error of gamma- aminobutyric acid metabolism, Clin. Chim. Acta 133: 33?42. Talibi D., M. Grenson, and B. Andr?. 1995. Cis- and trans-acting elements determining induction of the genes of the gamma-aminobutyrate (GABA) utilization pathway in Saccharomyces cerevisiae. Nucleic Acids Res. 23(4): 550-557. Tnnicliff G. 1993. Inhibition of 4-aminobutyrate aminotransferase from Pseudomonas fluorescens by ATP. Biochem Mol Biol Int. 31(1): 41-47. Tsushida, T., and T. Murai. 1987. Conversion of glutamic acid to gamma-aminobutyric acid in tea leaves under anaerobic conditions. Agric Biol Chem. 51: 2805-2871. Turano, F. J., and T. K. Fang. 1998. Characterization of two glutamate decarboxylase cDNA clones from Arabidopsis. Plant Physiol. 117: 1411?1421. Ueno, Y., K. Hayakawa, S. Takahashi and K. Oda. 1997. Purification and Characterization of Glutamate Decarboxylase from Lactobacillus brevis IFO 12005. Biosci., Biotechnol., Biochem. 61: 1168-1171. 41 Vacca, R. A., D. Valenti, A. Bobba, R. S. Merafina, S. Passarella, and E. Marra, 2006. Cytochrome c is released in a reactive oxygen speciesdependent manner and is degraded via caspase-like proteases in tobacco BY-2 cells en route to heat shock-induced cell death.Plant Physiol. 141: 208-219. Van Cauwenberghe O. R., and B. J. Shelp. 1999. Biochemical characterization of partially purified GABA:pyruvate transaminase from Nicotiana tabacum. Phytochemistry 52: 575-581. VanCauwenberghe, O. R., A. Makhmoudova, M. D. McLean, S. M. Clark, and B. J. Shelp. 2002. Plant pyruvate-dependent gamma-aminobutyrate transaminase identification of an Arabidopsis cDNA and its expression in Escherichia coli. Can. J. Bot. 80: 933?941. Vanovska, I., and J. M. Hardwick. 2005. Viruses activate a genetically conserved cell death pathway in a unicellular organism. J Cell Biol. 170: 391-399. Vissers, S., B. Andre, F. Muyldermans, and M. Grenson. 1989. Positive and negative regulatory elements control the expression of the UGA4 gene coding for the inducible 4- aminobutyric acid-specific permease in Saccharomyces cerevisiae. European Journal of Biochemistry. 181: 357?361. Vissers, S., B. Andre, F. Muyldermans, and M. Grenson. 1990. Induction of the 4- aminobutyrate and urea-catabolic pathways in Saccharomyces cerevisiae. Specific and common transcriptional regulators. Eur. J. Biochem. 187: 611-616. Voellym R, T. Leisinger. 1976. Role of 4-aminobutyrate aminotransferase in the arginine metabolism of Pseudomonas aeruginosa. J Bacteriol. 128(3): 722-729. Wagner-McPherson, G. A. Evans, and A. J. Tobin. 1992. Two human glutamate decarboxylases, 65-kDa GAD and 67-kDa GAD, are each encoded by a single gene. Proc Natl Acad Sci U S A. 89(6): 2115?2119. Walker, G. M. 1998. Yeast physiology and biotechnology. John Wiley & Sons Ltd. Chichester, England. Wallace W., J. Secor, and L. E. Schrader. 1984. Rapid accumulation of gamma- aminobutyric acid and alanine in soybean leaves in response to an abrupt transfer to lower temperature, darkness, or mechanical manipulation. Plant Physio. l75: 170-175. Wallace, W., J. Secor, and L. E. Schrader. 1984. Rapid accumulation of ?- aminobutyric acid and alanine in soybean leaves in response to an abrupt transfer to lower temperature, darkness, or mechanical manipulation. Plant Physiol. 75: 170?175. Weinberger, M., L. Ramachandran, and W. C. Burhans. 2003. Apoptosis in yeasts. IUBMB Life. 55: 467- 472. 42 White, H. E., and T. L. Sato. 1978. GABA transaminase of human brain and peripheral tissues: kinetics and molecular properties. J. Neurochem. 31: 41?47. Wu, J.Y., T. Matsuda, and E. Roberts. 1974. Purification and characterization of glutamate decarboxylase from mouse brain. J Biol Chem. 245: 3029-3034 Yamaura, 1., T. Matsumoto, M. Funatsu, T. Shinohara. 1988. Purification and some properties of succinic sernialdehyde dehydrogenase fiom barley seeds. Agric Biol Chem 52: 2929-2930 Yancey, P. H., M. E. Clark, S. C. Hand, R. D. Bowlus, and G. N. Somero. 1982. Living with water stress: evolution of osmolyte systems. Science 217: 1214-1222. Yonaha, K., K. Suzuki, H. Minei, and S. Toyama. 1983. Distribution of o-amino acid: pyruvate transaminase and aminobutyrate: a-ketoglutarate transaminase in microorganisms. Agricultural and Biological Chemistry. 47: 2257?2265. Yu, S.J., and S-H. Oh. 1998. Cloning and characterization of a tobacco cDNA encoding calcium/calmodulin-dependent glutamate decarboxylase. Mol. Cell. 8: 125?129. Zik, M., T. Arazi, W.A. Snedden, and H. Fromm. 1998. Two isoforms of glutamate decarboxylase in Arabidopsis are regulated by calcium/calmodulin and differ in organ distribution. Plant Mol. Biol. 37: 967?975. Zuppini, A., C. Andreoli, and B. Baldan. 2007. Heat Stress: an Inducer of Programmed Cell Death in Chlorella saccharophila. Plant Cell Physiol. 48(7): 1000?1009. Zuppini, A., V. Bugno, and B. Baldan. 2006. Monitoring programmed cell death triggered by mild heat shock in soybean-cultured cells. Funct. Plant Biol. 33: 617?627. 43 II. THE GABA SHUNT MEDIATES BASAL THERMOTOLERANCE IN SACCHAROMYCES CEREVISIAE BY REDUCING THE PRODUCTION OF REACTIVE OXYGEN SPECIES Abstract The GABA shunt pathway involves three enzymes: glutamate decarboxylase (GAD), GABA aminotransferase (GABA-TA), and succinate semialdehyde dehydrogenase (SSADH). These enzymes act in concert to convert ?-ketoglutarate (through glutamate) to succinate. Deletion mutations in each of these genes in Saccharomyces cerevisiae resulted in growth defects at 45?C or at 50?C following an induction period at 40?C. Double and triple mutation constructs were compared for thermotolerance with the wild type and single mutant strains. Although wild type and all mutant strains were highly susceptible to even brief heat stress at 50?C, a 30 min at 40?C (a non-lethal temperature) induced tolerance of wild type and all of the mutants to the 50?C (lethal temperature). The mutant strains collectively exhibited similar susceptibility at both 45?C and the induced 50?C treatments. Intracellular reactive oxygen species (ROS) accumulation was measured in wild type and each of the mutant strains. ROS accumulation in each of the mutants and under various stress conditions was correlated to heat susceptibility of the mutant strains. The addition of ROS scavenger N-tert-butyl-?- phenylnitrone (PBN) enhanced the mutant growth defect and strongly inhibited the 44 accumulation of ROS, but did not have significant effect on the wild-type. Measurement of intracellular GABA, glutamate and ?-ketoglutarate during lethal heat exposure at 45?C showed higher level of accumulation of GABA and ?-ketoglutarate in the uga1 and uga2 mutants, while glutamate accumulated at higher level in gad1 mutant. These results suggest that GABA shunt pathway plays a crucial role in protecting yeast cells from heat damage by restricting reactive oxygen species production involving the flux of carbon from ?-ketoglutarate in yeast cells to succinate during heat stress in mutant cells lacking a functional GABA shunt pathway. 45 Introduction Virtually all living organisms contain the GABA shunt pathway consisting of 3 enzymes: glutamate decarboxylase (GAD) catalyzing the conversion of glutamate (Glu) to GABA, GABA aminotransferase (GABA-AT) catalyzing the conversion of GABA and ?-ketoglutarate (?-KG) into succinate semialdehyde (SSA) and Glu, and succinate semialdehyde dehydrogenase (SSADH) catalyzing the NAD(P)-dependent conversion of SSA into succinate. The pathway serves to move ?-KG to succinate bypassing two reactions of the tricarboxylic acid cycle. The biological function of the GABA shunt is varied depending on the organism in which it exists. In bacteria and fungi, this pathway is responsible for the assimilation of exogenously supplied and excess glutamate (Jacob, 1962; Piquemal et al., 1961). In animals, it is well established that GABA and associated metabolism is associated with inhibitory neurotransmission (Petroff, 2002; Wang and Joseph, 1999). The disfunction of each enzyme in the pathway is related to human neurological genetic disorders (Jakobs et al., 1993). In plants, the GABA shunt appears to contribute to the control of cytosolic pH, balance between carbon and nitrogen metabolism, and adaptation to stress (Bouche and Fromm, 2004). GABA has been shown to play a potential signaling role in pollen tube guidance (Palanivelu et al., 2003). Disruption of the SSADH gene in Arabidopsis results in the accumulation of reactive oxygen species (ROS), necrotic leaf lesions, dwarfism, and hypersensitivity to environmental stresses (Bouche et al., 2003). In baker?s yeast, GAD1 is required for tolerance to oxidative stress and the loss of UGA1 and UGA2 reduced the oxidative stress tolerance (Coleman et al., 2001). These studies suggest that the GABA shunt pathway or at least an enzyme or enzymes in the pathway play a role in 46 limiting the production or lethality of ROS produced during environmental stress episodes. However, the molecular mechanism via which such actions are mediated remains largely unknown. In fungi and higher plants, GAD is known to contain a calmodulin binding domain, and the activity of the GABA shunt as well as GABA accumulation appears to be regulated by the binding of calcium and calmodulin to the GAD enzyme (Baum et al., 1993; Coleman et al., 2001). Thus, the accumulation of GABA during episodes of environmental stresses including heat stress in plants and fungi appears to be modulated by calmodulin and stress modulated intracellular calcium pools (Baum et al., 1996, Bouche et al., 2005; Coleman et al., 2001). Although the mechanism of modulating GABA shunt activity appears to be elucidated, the role of the enzymes of the GABA shunt in abating the deleterious effects of stress in fungi and plants remains to be elucidated. Here, we report the role of the three GABA shunt enzymes during heat stress in Saccharomyces cerevisiae. Deletion mutants of ?gad1, ?uga1, and ?uga2 and the double and triple mutation constructs of these three genes were made. Viability and thermotolerance of cells, intracellular levels of ROS, GABA shunt metabolite levels were measured in the wild type and various deletion strains during the mid-log phase of growth during heat stress. Materials and Methods Yeast strains and growth media. Yeast strains used in this study were all derived from stain W303-1A (Mat a leu2-3,112 ura3-1 trp1-1 his3-11,15 ade2-1, lys2, met) and the precise genotypes of the mutants are listed in Table 1. 47 In various experiments as detailed in text, table and figure legends, the strains were grown on either YPD medium or YNB medium. YPD medium contains: 2% glucose, 1% yeast extract, 2% yeast bactopeptone, and was used for growth and cell survival assay.YNB medium contains: 0.67% [wt/vol] yeast nitrogen base, 2% glucose, supplemented with essential amino acids (Sherman et al., 1979). Disruption of GAD1, UGA1 and UGA2 genes. The GAD1 disruption was generated by PCR amplification / homologous recombination strategy (1) that replaced the entire open reading frame of GAD1 with TRP1 gene. The TRP1 gene was amplified from plasmid pRS414 template DNA using a forward primer (named GAD1UP45TRP1A, Table 1) that consisted of the 45 nucleotides immediately upstream of the GAD1 gene fastened upstream of the 5?most portion of the gene. The reverse primer (named GAD1DN45TRP1D, Table 2) consisted of the 45 nucleotides immediately downstream of the GAD1 gene ligated to the 22 nucleotides at the 3? end of the TRP1 gene contained in pRS414. The PCR product was then transformed into yeast strain W303-1A by lithium acetate as previously described (Geitz and Woods, 2003), and TRP+ colonies were selected for genomic DNA isolation. The correct integration was verified by isolating genomic DNA (Hoffman and Winston, 1987) from each selected strain and conducting a series of PCR reactions using primers that land 300-500 nucleotides upstream and downstream of GAD1 open reading frame and within the TRP1 gene (Table 2). The UGA1 gene was disrupted by replacing a fragment of the gene between bases +293 to +911 with the HIS3 gene. The UGA1 coding sequence was amplified by PCR 48 from yeast genomic DNA utilizing PCR primers UGA1HIS3FOR and UGA1HIS3REV (Table 2). The amplified UGA1-orf was digested using KpnI and SacI according to manufacturer?s procedures and ligated into pBluescript, which had been linearized by restriction with the same two enzymes to create pBS-UGA1-orf. This pBluescript -UGA1- orf construct was then double digested with SalI and AgeI according to manufacturer's procedures generating a 618 nucleotide deletion from the UGA1-orf. The HIS3 gene was PCR amplified from the pRS413 plasmid using forward and reverse primers that had a SalI restriction site added 5' to the 5'-25 nucleotides of the HIS3 gene and an AgeI restriction site added 3' to the 3' end of the HIS3 gene sequence. This PCR product was then ligated into pGEM-T easy vector, and the plasmid was amplified in E. coli strain DH5-?, the insert was recovered from this amplified plasmid by digestion with SalI and AgeI restriction endonucleases. The SalI / AgeI insert was subsequently ligated into the linearized, digested pBS?UGA1-orf to replace the deleted 618 base-pair sequence of UGA1-orf generating pBS?uga1-orf::HIS3. The disrupted uga1-orf::HIS3 sequence was PCR amplified from pBS?uga1-orf::HIS3 and this PCR product (uga1::HIS3) was used as the disruption cassette to transform and eventually disrupt the endogenous W303-1A UGA1 gene. The UGA2 disruption was generated by PCR amplification / homologous recombination strategy (Bauldin et al., 1993) that replaced the entire open reading frame of UGA2 with URA3 gene. The URA3 gene was PCR amplified from plasmid pRS416 template DNA using a forward primer (named UGA2UP45URA3A, Table2) that consisted of the 45 nucleotides immediately upstream of the UGA2 gene fastened upstream of the 5?most portion of the gene. The reverse primer (named 49 UGA2DN45URA3D, Table 2) consisted of the 45 nucleotides immediately downstream of the UGA2 gene ligated to the 22 nucleotides at the 3? end of the URA3 gene contained in pRS416. The PCR product was then transformed into yeast strain W303-1A (Geitz and Woods, 2003), and URA3+ colonies were selected for genomic DNA isolation. The correct integration was verified by isolating genomic DNA from each selected strain (Hoffman and Winston, 1987) and conducting a series of PCR reactions using primers that land 300-500 nucleotides upstream and downstream of UGA2 open reading frame and within the URA3 gene (Table 2). To make the double deletion construct, gad1::TRP1 cassette (see above) was used to transform the single uga1::HIS3 deletion strain and the uga2::URA3 deletion strain to make ?gad1?uga1 and ?gad1?uga2 strains respectively. The uga1::HIS3 cassette was also transformed into the uga2::URA3 deletion strain to make ?uga1?uga2 deletion strain, and finally ?uga1?uga2 double deletion was transformed by gad1::TRP1 cassette to make the triple deletion construct of ?gad1?uga1?uga2. In addition to the chromosomal PCR verification, all disruptions of GAD1, UGA1, UGA2 in the single, double and triple deletion strains were verified by appropriate PCR for each of the single mutants as described above, and by failure to growth on YNB minimal medium containing 0.1% GABA or 0.2% ammonium sulfate as the sole nitrogen source. All deletion mutants except ?gad1 failed to grow on GABA-containing plates. Plasmid Construction. The S. cerevisiae glutamate decarboxylase gene (GAD1) was PCR amplified from genomic DNA isolated from wild type strain W303-1A using a forward primer GAD1FOR (Table 2) and a reverse primer GAD1REV (Table 2). To facilitate plasmid construction, two restriction sites for SmaI and XhoI (underlined, Table 50 2) were introduced at each end of the GAD1 specific primers. Following PCR amplification, the 1.778-kb SmaI / XhoI fragment containing the entire gene was digested from the PCR product by SmaI and XhoI (Biolabs, New England) according to manufacturer's procedures and ligated into the unique SmaI / XhoI sites of yeast shuttle vector p425 GPD (American Type Culture Collection) using DNA ligase (Promega) according to manufacturer's procedure to create plasmid p425 GPD-GAD1. The S. cerevisiae GABA aminotransferase gene (UGA1) was PCR amplified as described above using forward primer UGA1FOR (Table 2) and reverse primer UGA1REV (Table 2). The 1.436-kb BamHI / HindIII fragment (sites underlined in primers in Table 2) containing the entire gene digested from the PCR product and ligated into the unique BamHI / HindIII sites of p425 GPD as described above to create plasmid p425 GPD-UGA1.The S. cerevisiae succinate semialdehyde dehydrogenase gene (UGA2) was similarly PCR amplified using forward primer UGA2FOR (Table 2) and reverse primer UGA2REV (Table 2). The 1.504-kb SmaI / XhoI fragment (restriction sites underlined in the primers above) containing the entire gene was digested from the PCR product and ligated into unique SmaI / XhoI restriction sites of plasmid p425 GPD to create a plasmid p425 GPD- UGA2. The constructed plasmids p425 GPD-GAD1, p425 GPD-UGA1 and p425 GPD- UGA2 were used for overexpression in wild-type and single deletion strains of ?gad1, ?uga1 and ?uga2 by yeast transformation as described (Geitz and Woods, 2002). Lethal heat stress survival assays. A modification of the procedure of Davidson and Schiestl (2001) was used. Briefly, yeast cells were grown to mid log phase (5 ? 106 cells/ml) in YPD medium. To assess a possible protective effect of free spin trap reagent 51 N-tert-butyl-?-phenylnitrone (PBN, Sigma Chemical Co.), a final concentration of 5 mM PBN was added directly to the YPD medium for 4 ? 5 h as described (Ren et al., 2005) before heat stress. Cells were harvested by centrifugation (15,000 g), washed in 0.87% NaCl, and concentrated to 2 ? 108 cells/ml in fresh YPD medium. Aliquots of 100 ?l were taken into 0.6-ml PCR tubes for each time point and heated at 45oC in a thermocycler. At set time points, tubes were removed and placed on ice for 1 minute. Cells were then diluted so that colonies were countable and spread onto YPD plates. Colonies were counted after plates were incubated at 30oC for 2 days, the survival rate was calculated against unheated cells. The effect of PBN on cell survival was determined by 60 min lethal heating, cell survival without PBN treatment served as control. Induced thermotolerance assays. Exponentially growing cultures with and without PBN pretreatment were washed and concentrated to 2 ? 108 cells/ml as described above. Aliquots of 100 ?l in 0.6ml PCR tubes were preheated at 40?C in thermocycler for 30 min followed by quickly raising temperature to 50?C. Aliquots of cells were taken out at different time and placed on ice for 1 minute. Suitable dilutions of cells were plated on YPD plates and incubated at 30?C for two days to determine percent of survival as described earlier. The effect of PBN on induced cell survival was determined by 45 min heating under 50?C after preheating at 40?C, cell survival without PBN treatment served as control. Measurement of ROS production. Intracellular ROS generated during heat stress were measured using 2?, 7?-dichlorofluoroscin diacetate (DCFH-DA) essentially as described by Davidson (2001). With uptake of the probe, the intracellular esterase 52 removes the acetate groups, resulting in the formation of the non fluorescent substrate 2', 7'-dichlorofluoroscin (DCFH). Subsequent oxidation of DCFH produces highly fluorescent 2?, 7?-dichlorofluoroscein (DCF) which absorbs at a wavelength of 504 nm and emits at a wavelength of 524 nm (Wang and Joseph, 1999). Fluorescence was measured for cells both with and without PBN pretreatment, under lethal heat directly, and cells heated after preheat treatment. Overnight culture growing in SC or selective media were diluted into 10 ml of fresh YPD media to 0.2 OD600 and allowed to grow at 30?C on a shaker until an OD600 of 1.0 was achieved. Cells were then washed twice and concentrated to 2 ? 108 cells/ml in phosphate-buffered saline (pH7.4) (PBS). A 5 mM stock solution of dichlorofluorescin diacetate dissolved in ethanol was added to each culture to reach a final concentration of 10 ?M and incubated at 30?C for 15 min to allow entry of the probe into the cells. Five hundred ?l of each culture with DCFH-DA included was taken for heating under different temperature conditions as described in the text, table and figure legends. Immediately after heating, cells were cooled on ice for 1 min, washed twice in ice-cold distilled water and resuspended in 100 ?l of ice cold distilled water. Cells were disrupted by agitation on a vortex mixer with an equal volume of glass beads at maximum speed five times for 1 min each. Between agitation, samples were cooled on ice for 1 min. The clear supernatant was collected after 2.5 min centrifugation in a microcentrifuge at maximum speed in a cold room (4?C) and kept on ice. Thirty ?l of each supernatant was mixed with 2.5 ml of water and the fluorescence was recorded at 25?C at an excitation wavelength (Ex) of 504 nm and an emission (Em) wavelength of 524 nm, using a Hitachi F-2000 53 spectrofluorimeter. Background fluorescence was estimated using cells heated without DCFH-DA and using DCFH-DA without any cells. Measurement of intracellular GABA, glutamate, ?-ketoglutarate and succinate semialdehyde. Exponentially growing cells were harvested, washed and suspended in fresh YPD to 2 ? 108 cells/ml. Aliquots of 500 ?l were submitted to lethal heat stress at 45?C for varying periods of time as indicated in text, table and figure legends. Immediately after stress, cells were cooled on ice for 1 minute, washed twice with distilled water, and quickly flash frozen in liquid nitrogen and resuspended in 500 ?l distilled water. Cell dry weight was determined by incubation at 105?C overnight until there is no change of weight. Intracellular glutamate and GABA were extracted by boiling cells for 10 min. Intracellular succinate semialdehyde was extracted by breaking the cells with glass beads. After centrifugation for 5 min (15,000 g), the supernatant was collected for measurement of intracellular amino acids. Cell extract for ?-ketoglutarate was prepared by boiling ethanol method as described in Gonzalez et al. (1997). The amount of GABA was determined in a 200 ?l reaction mixture which contained 86 mM potassium phosphate buffer (pH 8.6), 3.3 mM 2-mercaptoethanol, 1.2 mM ?- NADP, and 0.004 unit of GABAase (Sigma-Aldrich Fine chemicals, St. Louis, MO), and an appropriate amount of the cell free extract prepared as above. The reaction was initiated by the addition of 5 mM ?-ketoglutarate after pre-incubation of components at 30oC for 5 min. 54 Glutamate assays were performed in a 200 ?l reaction mixture which contained 90 mM triethanolamine hydrochloride (pH 7.3), 60 ?M NAD+, 250 ?M EDTA, 0.06 units of L-glutamic dehydrogenase (sigma-Aldrich Fine chemicals, St. Louis, MO) and an appropriate amount of the cell free extract prepared as above. ?-ketoglutarate was determined in a 200 ?l reaction mixture which contained 90 mM triethanolamine hydrochloride (pH 7.3), 53 mM ammonium acetate, 60 ?M NADH, 250 ?M EDTA, 0.06 units of L-glutamic dehydrogenase, and an appropriate amount of cell free extract prepared as described above. Succinate semialdehyde was determined 200 ?l reaction mixture which contained 87 mM potassium phosphate buffer (pH8.4), 3 mM 2-mercaptoethanol, 1.3 mM ?-NAD, 0.83% glycerol and 0.025-0.05 unit of yeast succinate semialdehyde dehydrogenase (Ramos et al., 1985). The enzymatic determination of GABA, glutamate and succinate semialdehyde is based on the change in absorbance at 340 nm caused by the reduction of NAD(P)+ to NAD(P)H and the enzymatic determination of ?-ketoglutarate is based on the change in absorbance at 340 nm caused by the oxidation of NADH to NAD+. All enzymatic reactions are performed at 30oC for 60 min and the change of absorbance at 340 nm was monitored using microplate spectrophotometer (Power Wave? XS, BioTek instruments, Inc, Winooski, VT, USA). Concentrations of glutamate, ?-ketoglutarate, GABA and succinate semiladehyde were expressed as ?mol/g cell dry weight and calculated from the calibration curve of the standard solutions. All assays were repeated at least three times and data represented the mean ? SD of the results from at least three experiments. 55 Results Gene disruption of GABA shunt enzymes and heat sensitivity. A GAD1, UGA1 and UGA2 wild type strain, and the isogenic deletion strains ?gad1, ?uga1, and ?uga2 were subjected to lethal heat stress at 45oC for 15, 30, 45 and 60 min (Fig. 1). After 60 min, the wild type cells showed 65% survival compared to wild- type without heat treatment. The ?gad1, ?uga1, and ?uga2 deletions showed 30%, 19%, and 25% survival respectively. The double deletions of ?gad1?uga1, ?gad1?uga2 and ?uga1?uga2 showed an enhanced heat sensitivity and survival of 10%, 18% and 1%, respectively after 60 min at 45oC. While the viability of the triple deletion strain (?gad1?uga1?uga2) was 0% . Induced thermotolerance in GABA shunt mutant strains. Induction of thermotolerance by pretreatment at a sublethal temperature of 40oC for 30 min before exposure to a lethal temperature of 50oC in wild-type and the isogenic deletion mutants of GABA shunt enzymes was investigated. The survival rate at 50oC for 5 min without pretreatment in wild-type was 14% and below 10% in all deletion strains (Fig. 3A and Fig. 3B). Survival of all mutant and wild-type strains increased at 50oC with preheating at sublethal temperature. Wild-type showed survival above 50% and mutant strains showed survival up to 40% after 15 min at 50oC. After longer exposure at 50oC, the survival rate of the deletion strains declined more rapidly than in the wild-type. ROS production during heat stress in wild-type and GABA shunt mutants It has been shown that lethal heat stress damages cells by production of reactive oxygen species (ROS) including hydroxyl radicals (OH?), hydrogen peroxide (H2O2) and 56 the super oxide anion (O2?-), which can damage macromolecules and other cellular components (Cryer et al., 1975; Davidson, and Sehiestl, 2001; Wang and Joseph, 1999). The level of intracellular oxidants produced during lethal heating at both 45?C, and 50?C after preheating at 40?C for 30 min was measured by monitoring the oxidation of DCFH- DA. Fig. 4 shows when cells of the deletion strains were exposed to lethal heating at 45?C, DCFH oxidation increased 2- to 5-fold compared to unheated controls, while in wild-type, the increase was less than 1-fold. Generation of higher level of intracellular ROS in mutants is consistent with survival of the yeast cells at higher temperature. With longer exposure to heat, significantly higher levels of DCFH oxidation were observed in single, double and triple mutants than in wild-type (Fig. 4). After preheating at 40oC for 30 min (Fig. 6), the level of ROS increased 3- to 9- fold in deletion strains after exposure at 50oC, while in wild-type this increase was about 1.2- fold. These findings are consistent with the results of the survival experiments. The viability of deletion mutant cells was correlated with the level of DCFH oxidation observed. This correlation of heat stress tolerance with oxidation status of DCFH is further substantiated by results summarized in Fig. 5 (compared to Fig. 2) where overexpression of the three enzymes of the GABA shunt in each of the three respective single mutant strains led to levels of DCFH oxidation similar to wild type. Overexpression of GAD1, UGA1, or UGA2 in wild type resulted in either wild type levels of DCFH oxidation (GAD1), or reduction in ROS production below the levels found in heat treated wild type cells (UGA1 or UGA2). These data further support the hypothesis that all the three 57 enzymes in the GABA shunt pathway are involved in restricting the intracellular levels of ROS during lethal heat stress. To determine whether the increased level of ROS is essential for heat induced cell death, or ROS is just a byproduct of the heat stress, oxygen radicals were scavenged with free radical spin trap, PBN. Wild-type and all mutant strains were cultured at 30?C. The cultures were pretreated with PBN at a final concentration of 5 mM for 5 h. Cells were then submitted to heat stress for the determination of cell survival and production of ROS. As shown in Fig. 7, under 45 ?C lethal heating for 60 min, PBN treatment had no significant effect on cell survival for wild-type, however, the cell survival was greatly enhanced for all mutants from 1-fold (?gad1 mutant) up to more than 10-fold (?gad1?uga1?uga2), comparatively, in Fig. 8, the accumulation of ROS in the control decreased from 1- to 2-fold for mutant strains by the pretreatment of PBN, while the accumulation of ROS had no significant change. Similar effects had been found for the cells stressed under 50?C with preheating treatment (Fig. 9 and Fig. 10). These results suggest by scavenging ROS production under heat stress, cells were more able to tolerate heat stress, the ROS accumulation played an essential role in heat induced cell death. Effect of GABA shunt deficiency on intracellular metabolite levels. GABA is produced from decarboxylation of L-glutamate by glutamate decarboxylase (GAD1). GABA and ?-ketoglutarate is then converted to succinate semialdehyde and glutamate by GABA transaminase (UGA1) which is then used to make GABA. Changes in the levels of GABA, glutamate, ?-ketoglutarate and succinate semialdehyde during lethal heat stress in wild-type and mutant strains were investigated. 58 Under normal growth conditions, GABA level was very low in wild type and all strains bearing deletion mutants. Under lethal heat stress at 45oC, an 80-fold increase in GABA occurred in the wild-type in 15 min and remained at that level for up to 60 minutes of stress (Fig. 11). However, in the ?gad1 strain and in all double and triple mutant strains containing ?gad1, GABA levels increased less than 3-fold during 15 minutes of heat stress at 45oC and remained comparatively lower at all time points up to 60 minutes (Fig. 11). In strains bearing a uga1 deletion (?uga1 and ?uga1?uga2) the intracellular GABA pool increased over 160-fold during 15 minutes of heat stress at 45oC. This value increased to over 200-fold after 60 minutes of heat stress. Slightly lower increases in GABA levels were observed in the uga2 deletion mutants at all time points examined. These results suggest GABA accumulation during heat stress requires a functional GAD gene, and that the highest levels of GABA accumulation occur in strains with impaired GABA degradation (?uga1, ?uga2, and ?uga1?uga2). In unstressed wild type cells glutamate levels were substantially higher than GABA levels (39.2 ?mole/g dry weight versus 0.065 ?mole/g dry weight), and glutamate levels increased 3.2-fold and 4.6- fold during 15 minutes and 60 minutes respectively at 45oC ( Fig. 12). The same general pattern of glutamate accumulation was observed in all mutant strains examined, but the absolute levels accumulated in all strains bearing a gad1 mutation (?gad1, ?gad1?uga1, ?gad1?uga2, and ?gad1?uga1?uga2) were approximately doubled at each time point compared to the wild-type strain. Glutamate levels in ?uga1, ?uga2, ?uga1?uga2 were approximately the same as or slightly lower than wild type. These results confirm that GAD1p is the crucial enzyme involved in glutamate consumption in yeast cells. 59 In unstressed wild type W303-1A cells, ?-ketoglutarate concentrations were approximately 1/3 the glutamate levels (13 versus 39 ?mole/g dry weight). Following heat stress at 45oC, ?-ketoglutarate accumulated linearly for at least 60 minutes reaching 61 ?mole/g dry weight by 60 minutes of heat stress (Fig. 13). Similar linear patterns of ?- ketoglutarate accumulation were observed for all deletion strains, but accumulation rates were significantly higher in ?uga1 and ?uga2 strains (6.6-fold and 6.1-fold respectively after 60 minutes) compared to wild type (Fig. 13). The ?uga1?uga2 double mutant strain demonstrated the higher rate of ?-ketoglutarate accumulation (7.3-fold after 60 minutes) compared to wild-type (Fig. 9). In all strains carrying a gad1 deletion (?gad1, ?gad1?uga1, ?gad1 ?uga2 and ?gad1?uga1?uga2), the accumulation of ?- ketoglutarate showed a slow linear pattern of accumulation reaching only 2-fold after 60 minutes of heating at 45oC. Since ?-ketoglutarate is not a product of the reaction catalyzed by GAD1p, it is not clear why strains bearing a gad1 mutation accumulate ?- ketoglutarate. By comparison,???-ketoglutarate is a substrate consumed in the reaction catalyzed by UGA1p. Thus, strains bearing a uga1 mutation would logically accumulate greater levels of ?-ketoglutarate. Since succinate semialdehyde is a metabolite unique to the GABA shunt, accumulation of this metabolite was investigated. Unstressed wild type cells produced 2.95 ?mol/g dry weight SSA while the ?uga2 strain produced 4.37 ?mol/g dry weight SSA. All of the other mutants produced less than 2.5 ?mol/g dry weight SSA. Heat stress at 45oC produced a linear accumulation of SSA only in wild type and in the ?uga2 strain while no SSA accumulation occurred in any of the other deletion mutants (Fig. 14). SSA accumulation levels in ?uga2 were nearly double the levels accumulated in wild type. 60 Mutation of gad1 or uga1 reduces accumulation of SSA while mutation of uga2 leads to high levels of SSA accumulation. Discussion Mutation of each of the 3 genes of the GABA shunt resulted in a reduction in tolerance of yeast cells to a lethal heat stress at 45?C and to heat stress at 50?C following an inductive period at 40?C (Fig. 1 and Fig. 3). Deletion of GAD1 resulted in the least reduction in stress tolerance, while deletion of UGA1 resulted in the greatest effect. All double mutants were at least as sensitive to heat stress as ?uga1 (e.g. ?gad1 ?uga2) or more heat sensitive (?gad1 ?uga1 and ?uga1 ?uga2) than the single mutants, and the triple mutant was most sensitive. Essentially identical results were obtained whether the cells were grown on YPD medium (Fig. 1) or YNB minimal medium with ammonium sulfate as nitrogen source (data not shown) indicating that the rich source of amino acids present in YPD medium does not contribute to the heat stress tolerance phenotype of wild type or the mutants. Tolerance of lethal heat stress essentially reverted to wild type levels when each of the single genes was transformed into the respectively comparable mutant driven by a strong promoter (Fig. 2), and overexpression of each gene in wild type resulted in only small increases in heat tolerance. This further demonstrates a role for the enzymes of the GABA shunt in abating some aspect of heat stress. Tolerance to even more severe heat stress (50oC) can be induced by pretreatment of cells at sublethal stress of 35-40oC (Davidson and Schiestl, 2001). The sensitivity of the GABA shunt mutants used above to 50oC heat stress following a 40oC adaptation period was proportionately similar to the 45oC lethal heat stress above (Fig. 3 compared to Fig. 1). Thus, the basic 61 mechanism(s) involved in abating heat stress by function of the enzymes of the GABA shunt are critical to basal thermotolerance but appear not to be involved in induced thermotolerance (Lindquist and Kim, 1996). The enzymes of the GABA shunt have been shown to play a role in protecting yeast cells against damage from exogenous oxidative stress generated by H2O2 (Coleman et al., 2001). Similarly in Arabidopsis thaliana SSADH appears to play a role in heat and UV stress tolerance by restricting ROS accumulation during episodes of stress (Bouche et al., 2003). Thus, the production of ROS was investigated in mutants of the GABA shunt. Either during lethal heat stress at 45oC (Fig. 4 and Fig. 5) or at 50oC following an inductive period at 40oC (Fig. 6) in all single, double, and triple mutants and in cells expressing various genes on plasmids, the level of heat-stress-induced ROS produced relative to wild type was proportional to the lethality of the mutant(s) relative to wild type. The presence of 5 mM free radical scavenger PBN strongly inhibited the ROS accumulation in the mutant strains (Fig. 8 and Fig. 10) at both stress conditions and enhanced the cell survival. Furthermore, in the single mutants, the heat-stress-induced ROS production was reversed when the respective gene was overexpressed on a plasmid. These observations are consistent with previously published results that the production of ROS is a critical component of basal heat-stress-induced damage to yeast cells (Davidson and Schiestl, 2001) and with the conclusion that heat-stress-induced-damage to yeast cells is prevented by the function of the enzymes of the GABA shunt through a mechanism that involves abatement of the accumulation of ROS during heat stress. However, the role of the enzymes of the GABA shunt pathway in protecting cells from heat stress damage involving heat-induced ROS accumulation is not clear. In yeast 62 the GABA shunt pathway is known to be involved in the assimilation of supplied glutamate, and to feed the carbon from this metabolite into the respiratory chain within the mitochondria with succinate (Pietruszko and Fowden, 1961; Ramos et al., 1985). As expected, all strains bearing a GAD1 deletion demonstrated a reduction in the intracellular pool of GABA produced during heat stress (Fig. 11), while strains bearing a deletion of UGA1 accumulated the highest heat-stress-induced levels of GABA. UGA2 deletion strains produced intermediate GABA levels during heat stress. The effect of deletion of GAD1 was epistatic to deletion of UGA1 or UGA2 which is consistent with the hypothesis that GAD1p is the primary if not exclusive source of GABA derived from glutamate. Since the ?gad1 strain, the least thermosensitive strain and the strain producing lowest ROS levels during heat stress, produced essentially identical heat-stress-induced levels of GABA with the triple mutant strain, the most thermosensitive and highest heat-stress- induced ROS producing strain, yet neither strain produced even wild type levels of GABA, it would appear that GABA itself is not a metabolite directly or indirectly involved in the protection of yeast cells from thermo-damage. Specifically, the accumulation of GABA in these mutants does not support a role for GABA as a signaling molecule during heat stress even though GABA is known to be a powerful signaling molecule in animal systems (Calver et al., 2000; Gamel-Didelon et al., 2003, Geigerseder et al., 2003). The highest levels of heat-stress-induced glutamate were observed in the triple mutant strain, but all other strains bearing a GAD1 deletion produced accumulated glutamate pools that were larger than wild type. Strains bearing a UGA1 or UGA2 deletion (without a GAD1 deletion) produced lower levels of glutamate during heat stress 63 relative to wild type. The opposite patterns were observed for ?-ketoglutarate, i.e. in strains bearing GAD1 deletions, ?-ketoglutarate levels were lower than wild type during heat stress while in strains bearing UGA1 or UGA2 deletions, ?-ketoglutarate was elevated relative to wild type. The fact that glutamate and ?-ketoglutarate levels are altered dramatically during heat stress in GABA shunt mutants is consistent with the hypothesis that the GABA shunt is a major pathway for the assimilation of glutamate during episodes of stress. However, the specific pattern of glutamate and ?-ketoglutarate accumulation in the mutants does not suggest a specific mechanism for limiting ROS production, but rather that flux through the pathway is the critical feature for heat-stress tolerance and limiting ROS production. This conclusion is further substantiated by examination of succinate semialdehyde (SSA) accumulation (Fig. 14). All of the mutant strains except ?uga2 accumulated little or no SSA during heat stress beyond the level found in unstressed cells. However, unexpectedly, wild type accumulated substantial SSA during heat stress, and the ?uga2 strain accumulated even higher levels of SSA. This accumulation was dependent on functional GAD1 and UGA1 genes, and thus the function of the complete pathway is required for the production of SSA. However, the accumulation of SSA in the various mutants does not suggest that this metabolite plays an exclusive role in mediating ROS production during heat stress. Taken together, it appears that carbon flux through the entire GABA shunt is required to abate ROS production during heat stress. This suggests that moving carbon from ?-ketoglutarate to succinate may be the function most critical to abating ROS production and consequent damage during heat stress. Clearly, this function is only 64 efficiently accomplished by the 3 enzymes of the GABA shunt working in concert, and disturbance of any one of these enzymes can have a variable effect on the pathway based on alternative competing mechanisms for moving carbon past various steps in the pathway. Because it is likely that the greatest flux of carbon from ?-ketoglutarate to succinate occurs inside the mitochondria as part of the tricarboxylic acid cycle and UGA1p and UGA2p both appear to be cytosolic proteins in Saccharomyces cerevisiae (Chapter IV), it is a reasonable working hypothesis that the compartmentation of metabolites into the cytosol and/or the vacuole may be a critical aspect of the function of the GABA shunt. Such a movement of these metabolites out of the mitochondria would clearly have an effect on the redox status of the mitochondria during stress and thus could affect ROS accumulation. References 1. Bauldin A., O. Ozier-Kalogeropoulos, A. Danouel, F. Ladcroute, and C. Cullin. 1993. A simple and efficient method for direct gene deletion in Saccharomyces cerevisae. Nucleic Acids Research 21:3329-3330. 2. Baum, G., S. Lev-Yadun, Y. Fridmann, T. Arazi, H. Katsnelson, M. Zik, and H. Fromm. 1996. Calmodulin binding to glutamate decarboxylase is required for regulation of glutamate and GABA metabolism and normal development in plants. EMBO J. 15:2988-2996. 3. Baum, G., Y. Chen, T. Arazi, H. Takatsuji, and H. Fromm. 1993. A plant glutamate decaroxylase containing a calmodulin binding domain. Cloning, sequence, and functional analysis. J. Biol. Chem. 268:19610?19617. 4. Bouch? N, A. Fait, D. Bouchez, S. G. M?ller, and H. Fromm. 2003. Mitochondrial succinic-semialdehyde dehydrogenase of the ?-aminobutyrate shunt is required to restrict levels of reactive oxygen intermediates in plants. Proc. Natl. Acad. Sci. U.S.A 100:6843?6848. 5. Bouch? N., and H. Fromm. 2004. GABA in plants: just a metabolite? Trends in Plant Science 9:3110-11. 65 6. Bouch?, N., A. Yellin, W. A. Snedden, and H. Fromm. 2005. Plant-specific calmodulin-binding proteins. Annu. Rev. Plant Biol. 56:435?466 7. Calver, A.R., A. D. Medhurst, M. J. Robbins, K. J. Charles, M. L. Evans, D. C. Harrison, M. Stammers, S. A. Hughes, G. Hervieu, A. Couve, S. J. Moss, D. N. Middlemiss, and M. N. Pangalos. 2000. The expression of GABA(B1) and GABA(B2) receptor subunits in the CNS differs from that in peripheral tissues. Neuroscience 100:155?170 8. Coleman S.T., T. K. Fang , S. A. Rovinsky, F. J. Turano, and W. S. Moye- Rowley. 2001. Expression of a glutamate decarboxylase homologue is required for normal oxidative stress tolerance in Saccharomyces cerevisiae. J Biol Chem. 276:244-50. 9. Cryer, D. R., R. Eccleshal, and J. Marmur. 1975. Isolation of yeast DNA. Meth. Cell Biol. 12:39-44. 10. Davidson, J. F., and R. H. Schiestl. 2001. Cytotoxic and genotoxic consequences of heat stress are dependent on the presence of oxygen in Saccharomyces cerevisiae. Journal of Bacteriology. 183:4580-4587. 11. Gamel-Didelon, K., L Kunz, K. J F?hr, M. Gratzl, and A. Mayerhofer. 2003. Molecular and physiological evidence for functional gamma-aminobutyric acid (GABA)-C receptors in growth hormone-secreting cells. J. Biol. Chem. 278:20192?20195. 12. Geigerseder C., R. Doepner, A.Thalhammer, M. B. Frungieri, K. Gamel- Didelon, R. S. Calandra, F. M. K?hn, and A. Mayerhofer. 2003. Evidence for a GABAergic system in rodent and human testis: local GABA production and GABA receptors. Neuroendocrinology 77:314?323. 13. Geitz, R. D. and R. A. Woods. 2002. Transformation of yeast lithium acetate/single-stranded carrier DNA / polyethylene glycol method. Methods Enzymol. 350:87-96. 14. Gonzalez B, J. Fran?ois and M. Renaud. 1997. A rapid and reliable method for metabolite extraction in yeast using boiling buffered ethanol. Yeast 13:1347-1356. 15. Hoffman, C. S., and R. Winston. 1987. A ten-minute DNA preparation from yeast efficiently releases autonomous plasmids for transformation of Escherichia coli. Gene (Amst.) 57:267-272. 16. Jacoby, W. B. 1962. Enzymes of 4-aminobutyrate metabolism. Methods in Enzymology 5:765-779. 66 17. Jakobs, C., J. Jaeken, and K. M. Gibson. 1993. Inherited disorders of GABA metabolism. J. Inherit. Metab. Dis. 16:704-715. 18. Lindquist, S., and G. Kim. 1996. Heat-shock protein 104 expression is sufficient for thermotolerance in yeast. Proc. Natl. Acad. Sci. USA 93:5301?5306. 19. Palanivelu R, L. Brass, A. F. Edlund, and D. Preuss. 2003. Pollen tube growth and guidance is regulated by POP2, an Arabidopsis gene that controls GABA levels. Cell 114:47?59. 20. Petroff O.A. 2002. GABA and glutamate in the human brain. Neuroscientist 8:562?573. 21. Pietruszko, R. and L. Fowden. 1961. 4-Aminobutyric acid metabolism in plants. I. Metabolism in yeasts. Annals of Botany 25: 491-511. 22. Piquemal, M., J. C. Latche, and P. Bald. 1972. Importance de l'aide glutamique dans la nutrition et le metabolisme d'Agaricus bisporus Lge. Mushroom. Science 8:413-424. 23. Ramos F, M. Guezzar, M. Grenson, and J. M. Wiame. 1985. Mutations affecting the enzymes involved in the utilization of 4-aminobutyric acid as nitrogen source by the yeast Saccharomyces cerevisiae. Eur J Biochem 149:401 24. Ren, Q., H. Yang, M. Rosinski, M. N. Conrad, M. E. Dresser, V. Guacci and Z. J., Zhang. 2005. Mutation of the cohesin related gene PDS5 causes cell death with predominant apoptotic features in Saccharomyces cerevisiae during early meiosis Mutation Research 570:163-173. 25. Sanchez, Y., and S. L. Lindquist. 1990. HSP104 required for induced thermotolerance. Science 248:1112?1115. 26. Sherman, F., G.R. Fink, and C.W. Lawrence. 1979. Methods in Yeast Genetics. Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y. 27. Varju, P., Z. Katarova, E. Madarasz, and G. Szabo. 2001. GABA signalling during development: new data and old questions. Cell Tissue Res. 305:239-246. 28. Wang H., and J. A. Joseph. 1999. Quantifying cellular oxidative stress by dichlorofluorescin assay using microplate reader. Free Rad Biol Med. 27:612-616. 67 Table 1. Yeast strains used in this study Strain Genotype Source and description W303-1A Mat a leu2-3,112 ura3-1 trp1-1 his3-11,15 ade2-1, lys2, met ATCCa, WT ?gad1 Mat a leu2-3112 ura3-1 trp1-1 his3-11,15 ade2-1, lys2, met gad1::TRP1 This study, gad1 disruptant ?uga1 Mat a leu2-3112 ura3-1 trp1-1 his3-11,15 ade2-1, lys2, met uga1::HIS3 This study, uga1 disruptant ?uga2 Mat a leu2-3112 ura3-1 trp1-1 his3-11,15 ade2-1, lys2, met uga2::URA3 This study, uga2 disruptant ?gad1?uga1 Mat a leu2-3112 ura3-1 trp1-1 his3-11,15 ade2-1, lys2, met gad1::TRP1 uga1::HIS3 This study, gad1 uga1 double disruptant ?gad?1uga2 Mat a leu2-3112 ura3-1 trp1-1 his3-11,15 ade2-1, lys2, met gad1::TRP1 uga2::URA3 This study, gad1 uga2 double disruptant ?uga1?uga2 Mat a leu2-3112 ura3-1 trp1-1 his3-11,15 ade2-1, lys2, met uga1::HIS3 uga2::URA3 This study, uga1 uga2 double disruptant ?gad1?uga1?uga2 Mat a leu2-3112 ura3-1 trp1-1 his3-11,15 ade2-1, lys2, met gad1::TRP1 uga1::HIS3 uga2::URA3 This study, gad1 uga1 uga2 triple disruptant aATCC, American Type Culture Collection 68 Table 2. Primers used for GAD1, UGA1 and UGA2 gene cloning, deletions and verifications Primer Name Sequence (5? to 3?) GAD1FOR GAGACCCGGGATGTTACACAGGCACGGTTCTAAG GAD1REV GTGTCTCGAGTCAACATGTTCCTCTATAGTTTCTCG UGA1FOR GAGAGGATCCATGTCTATTTGTGAACAATACTACCCA UGA1REV GTGTAAGCTTTCATAATTCATTAACTGATTTGGCTAA UGA2FOR GAGACCCGGGATGACTTTGAGTAAGTATTCTAAACCAAC UGA2REV GTGTCTCGAG TTAAATGCTGTTTGGCAAATTCC GAD1UP45TRP1A CACGTCGCTCTTAACAATCCAGGCTGAACAAAACAAGGA ATAATGGGAAGCATTTAATAGACAGCATCGT GAD1DN45TRP1D TACATACATATAGGGGGCGGTATATTGGATGACCTTTTC AACTCAAGGCAAGTGCACAAACAATACT GAD1/TRP1KO_A TGCGTTTATAAATAATCTTTCTGGC GAD1/TRP1KO_B CATCTTTACCAGCATTCTTCATTCT GAD1/TRP1KO_C ATTCAGGGTATAGAACACAATTCCA GAD1/TRP1KO_D TATTCCCGCATAACTTTTCTATCAC UGA1HIS3FOR GGGGTACCCCAGAACAGACAAGAAACCGTCA UGA1HIS3REV GAT GAG CTC GCG GCC TCG CTA ATA TAC AAT UGA2UP45URA3A CCAGCTACATTAAAAGCAAATTTTACAAACTACTATTTCA ACATGCGGTTTCTTTGAAATTTTTTTGA UGA2DN45URA3D ACATGAAACCATACCAGTTTCCAAAGCTTCAGACACAGT GTATTAGGGTAATAACTGATATAATTAAATTGAAGC. UGA2/URA3KO_A CGGTCGTTGAAGTGCTATAGTTTAT UGA2/URA3KO_B AGGTTGCCTAATTGTGAATACTCTG UGA2/URA3KO_C ACTTTTACTGGTTCTACAAACGTCG UGA2/URA3KO_D GTGAAAAACTTCAAAACTCCGTAAA 69 ?gad1W303-1A (WT) ?uga1 ?uga2 ?gad1?uga1 ?gad1?uga2 ?uga1?uga2 ?gad1?uga1?uga2 Time at 45oC (min) 0 10 20 30 40 50 60 Su rvi va l (% ) 0 20 40 60 80 100 Su rvi va l (% ) Fig. 1. Viability of wild-type and mutant strains after lethal heat treatment at 45oC. Strains were grown to mid log phase (5 ? 106 cells/ml) in YPD, washed in 0.87% NaCl, and concentrated to 2 ? 108 cells/ml in fresh YPD. Aliquots of 100 ?l were heated at 45oC in a thermocycler for 0, 15, 30, 45 and 60 min. Viability was determined as described. Each point represents the mean ? SD of the results from at least three experiments, some error bars are smaller than the symbol sizes. 70 A Time at 45oC (min) 0 10 20 30 40 50 60 Ce ll s ur viv al (% ) 0 20 40 60 80 100 B Time at 45oC (min) 0 10 20 30 40 50 60 Ce ll s ur viv al (% ) 20 40 60 80 100 C Time at 45oC (min) 0 10 20 30 40 50 60 Ce ll s ur viv al (% ) 0 20 40 60 80 100 W303-1A (WT) ?gad1 ?gad1-P425 GPD-GAD1 W303-1A-P425 GPD-GAD1 ?uga1 ?uga1-P425 GPD-UGA1 W303-1A-P425 GPD-UGA1 ?uga2-P425 GPD-UGA2 W303-1A-P425 GPD-UGA2?uga2 Ce ll s ur viv al (% ) Ce ll s ur viv al (% ) Ce ll s ur viv al (% ) Ce ll s ur viv al (% ) Fig. 2. Viability of wild-type and ?gad1mutant with overexpression of yeast GAD1 (A), ?uga1 mutant with overexpression of yeast UGA1 (B) and ?uga2 mutant with overexpression of yeast UGA2 (C) after lethal heat stress at 45oC. Heat stress was performed and viability was determined as the same in Fig. 1. Each point represents the mean ? SD of the results from at least three experiments, some error bars are smaller than the symbol sizes. Data for empty vector were not shown. 71 0 2 4 6 8 10 12 14 16 W30 3-1A (WT ) ?ga d1 ?uga1 ?ug a2 ?ga d1? uga 1?u ga2 ?ga d1? uga 2 ?ga d1? uga 1 ?ug a1? uga 20 10 20 30 40 0 20 40 60 80 100 ?gad1W303-1A (WT) ?uga1 ?uga2 ?gad1?uga1 ?gad1?uga2 ?uga1?uga2 ?gad1?uga1?uga2 A B Heat stress at 50?C for 5 min Time (min) at 50?C after 30 min preheated at 40?C Su rvi va l (% ) Su rvi va l (% ) Su rvi va l (% ) Su rvi va l (% ) Fig. 3. Viability of wild-type and mutants at lethal heat stress 50?C. Mid log phase cells grown in YPD were collected and washed in 0.87% NaCl, aliquots of 100 ?l concentrate in fresh YPD were heated at 50oC preceded by sub-lethal heat stress at 40?C for 30 minutes (A), as a control, cells were heated at 50?C for 5 min without preheat treatment (B). Cell survival was determined as described. Each point represents the mean ? SD of the results from at least three experiments, some error bars are smaller than the symbol sizes. 72 Time at 45oC (min) 0 10 20 30 40 50 60 He at ind uc ed flu ore sc en ce in cre as e 0 200 400 600 800 1000 ?gad1W303-1A (WT) ?uga1 ?uga2 ?gad1?uga1 ?gad1?uga2 ?uga1?uga2 ?gad1?uga1?uga2 He at ind uc ed flu ore sc en ce in cre as e Fig. 4. Intracellular ROS levels induced by lethal heat stress at 45?C in wild-type and mutant strains. Fluorescence was measured at an excitation wavelength of 504nm and an emission wavelength of 524nm in crude cell extracts as described. As a control, fluorescence was recorded during heat stress without cells and cells without DCFH-DA (date not shown). Data shown represent the mean ? SD of the results from at least three experiments, some error bars are smaller than the symbol sizes. 73 A Time at 45oC (min) 0 10 20 30 40 50 60 He at ind uc ed flu ore sc en ce in cre as e 0 100 200 300 400 500 600 B Time at 45oC (min) 0 10 20 30 40 50 60 He at ind uc ed flu ore sc en ce in cre as e 0 100 200 300 400 500 600 C Tim e at 45oC (m in) 0 10 20 30 40 50 60 He at ind uc ed flu or es ce nc e i nc rea se 0 100 200 300 400 500 600 W303-1A (WT) ?gad1 ?gad1-P425 GPD-GAD1 W303-1A-P425 GPD-GAD1 ?uga1 ?uga1-P425 GPD-UGA1 W303-1A-P425 GPD-UGA1 ?uga2-P425 GPD-UGA2 W303-1A-P425 GPD-UGA2?uga2 He at ind uc ed flu ore sc en ce in cre as e He at ind uc ed flu ore sc en ce in cre as e He at ind uc ed flu or es ce nc e i nc rea se Fig. 5. Intracellular ROS levels induced by lethal heat stress at 45?C in wild-type and ?gad1mutant transformed with plasmid P425 GPD- GAD1 (A), ?uga1 mutant transformed with plasmid P425 GPD- UGA1 (B), ?uga2 mutant transformed with plasmid P425 GPD- UGA2 (C). Fluorescence was recorded the same as described in Fig. 4. Data shown represent the mean ? SD of the results from at least three experiments, some error bars are smaller than the symbol sizes. 74 Time at 50oC (min) 0 10 20 30 40 50 He at ind uc ed flu ore sc en ce in cre as e 0 500 1000 1500 ?gad1W303-1A (WT) ?uga1 ?uga2 ?gad1?uga1 ?gad1?uga2 ?uga1?uga2 ?gad1?uga1?uga2 He at ind uc ed flu ore sc en ce in cre as e Fig. 6. Intracellular ROS levels induced by lethal heat stress at 50?C after a sub-lethal heat stress at 40 ?C for 30min in wild-type and mutant strains. Fluorescence was recorded the same as described in Fig. 4. Data shown represent the mean ? SD of the results from at least three experiments, some error bars are smaller than the symbol sizes. 75 Ce ll s urv iva l (% ) 0 20 40 60 80 Control PBN W3 03- 1A (WT ) ?ug a2?uga1?gad1 ?ga d1? uga 1?u ga2 ?ga d1? uga 2 ?ug a1? uga 2 ?ga d1? uga 1 Ce ll s urv iva l (% ) Fig. 7. Effect of PBN on lethal heat induced cell death under 45?C for wild-type and all mutant strains. Cells of wild-type and mutants were cultured at YPD medium at 30?C. The free spin trap reagents PBN (5 mM) was added directly to the cell cultures for 5 h before heat stress. Cell survival was determined as described for a 60 min point. Cells without PBN treatment server as control. Data shown represent the mean ? SD of the results from at least three experiments. 76 He at ind uc ed flu ore sc en ce in cre as e 0 200 400 600 800 1000 Control PBN W30 3-1A (W T) ?ga d1 ?uga1 ?ug a2 ?ga d1? uga 1 ?ga d1? uga 2 ?ug a1? uga 2 ?ga d1? uga 1?u ga2 He at ind uc ed flu ore sc en ce in cre as e Fig. 8. Effect of PBN on production of ROS under 45?C for wild-type and all mutant strains. Cells of wild-type and mutants were cultured at YPD medium at 30?C. The free spin trap reagents PBN (5 mM) was added directly to the cell cultures for 5 h before heat stress. Fluorescence was recorded the same as described in Fig. 4 for a 60 min point. Cells without PBN treatment server as control. Data shown represent the mean ? SD of the results from at least three experiments. 77 Ce ll s urv iva l (% ) 0 20 40 60 80 Control PBN W3 03-1 A (W T) ?ug a2 ?ug a1 ?ga d1 ?ga d1u ga1 uga 2 ?ga d1u ga2 ?ug a1u ga2 ?ga d1u ga1 Ce ll s urv iva l (% ) Fig. 9. Effect of PBN on heat induced cell death under 50?C with preheat treatment at 40 ?C for 30min for wild-type and all mutant strains. Cells of wild-type and mutants were cultured at YPD medium at 30?C. The free spin trap reagents PBN (5 mM) was added directly to the cell cultures for 5 h before heat stress. Cell survival was determined as described for a 45 min point. Cells without PBN treatment server as control. Data shown represent the mean ? SD of the results from at least three experiments. 78 He at ind uc ed flu ore sc en ce in cre as e 0 500 1000 1500 2000 Control PBN W3 03-1 A (W T) ?ug a2 ?ug a1 ?ga d1 ?ga d1u ga1 uga 2 ?ga d1u ga2 ?ug a1u ga2 ?ga d1u ga1 He at ind uc ed flu ore sc en ce in cre as e Fig. 10. Effect of PBN on production of ROS under 50?C with preheat treatment at 40 ?C for 30min for wild-type and all mutant strains. Cells of wild-type and mutants were cultured at YPD medium at 30?C. The free spin trap reagents PBN (5 mM) was added directly to the cell cultures for 5 h before heat stress. Fluorescence was recorded the same as described in Fig. 4 for a 45 min point. Cells without PBN treatment server as control. Data shown represent the mean ? SD of the results from at least three experiments. 79 Time at 450C (min) 0 10 20 30 40 50 60 GA BA co nc en tra tio n ( um ol/ g o f d ry we igh t) 0.0 2.0 4.0 6.0 8.0 10.0 12.0 14.0 ?gad1W303-1A (WT) ?uga1 ?uga2 ?gad1?uga1 ?gad1?uga2 ?uga1?uga2 ?gad1?uga1?uga2 GA BA co nc en tra tio n ( um ol/ g o f d ry we igh t) Fig. 11. Changes in the levels of GABA under lethal heat stress at 45oC in wild-type and mutant strains. Intracellular GABA was extracted from heat stressed cells by boiling cells for 10 min. After centrifugation for 5 min (15,000g), supernatant was taken for determination of GABA by GABase as described. Data shown represent the mean ? SD of the results from at least three experiments, some error bars are smaller than the symbol sizes. 80 Time at 45oC (min) 0 10 20 30 40 50 60 Gl uta ma te co nc en tra tio n ( um ol/ g o f d ry we igh t) 0 50 100 150 200 250 300 ?gad1W303-1A (WT) ?uga1 ?uga2 ?gad1?uga1 ?gad1?uga2 ?uga1?uga2 ?gad1?uga1?uga2 Gl uta ma te co nc en tra tio n ( um ol/ g o f d ry we igh t) Fig. 12. Changes in the levels of Glutamate under lethal heat stress of 45oC in wild-type and mutant strains. Intracellular Glutamate was extracted from heat stressed cells by boiling cells for 10 min. After centrifugation for 5 min (15,000g), supernatant was taken for determination of Glu by glutamate dehydrogenase as described. Data shown represent the mean ? SD of the results from at least three experiments, some error bars are smaller than the symbol sizes. 81 Time at 45oC (min) 0 10 20 30 40 50 602 -ke tog lut ara te co nc en tra tio n ( um ol/ g o f d ry we igh t) 0 20 40 60 80 100 ?gad1W303-1A (WT) ?uga1 ?uga2 ?gad1?uga1 ?gad1?uga2 ?uga1?uga2 ?gad1?uga1?uga2 2-k eto glu tar ate co nc en tra tio n ( um ol/ g o f d ry we igh t) Fig. 13. Changes in the levels of a-ketoglutarate under lethal heat stress at 45oC in wild- type and mutant strains. Cell extracts for intracellular a-ketoglutarate from heat stressed cells were prepared by boiling ethanol method as described (12). ?-ketoglutarate was determined by glutamate dehydrogenase assay as described. Data shown represent the mean ? SD of the results from at least three experiments, some error bars are smaller than the symbol sizes. 82 Time at 45oC (min) 0 10 20 30 40 50 60 SS A co nc en tra tio n ( um ol/ g d ry we igh t) 0 2 4 6 8 10 12 ?gad1W303-1A (WT) ?uga1 ?uga2 ?gad1?uga1 ?gad1?uga2 ?uga1?uga2 ?gad1?uga1?uga2 SS A co nc en tra tio n ( um ol/ g d ry we igh t) Fig. 14. Changes in the levels of succinate semialdehyde under lethal heat stress at 45oC in wild-type and mutant strains. Intracellular succinate semialdehyde was extracted by breaking the cells with glass beads. After centrifugation for 5 min (15,000g), supernatant was taken for determination of SSA by succinate semialdehyde dehydrogenase as described. Data shown represent the mean ? SD of the results from at least three experiments, some error bars are smaller than the symbol sizes. 83 III. GABA SHUNT GENE EXPRESSION ANALYSIS AND HEAT STRESS RESPONSE OF THE GABA SPECIFIC TRANSCRIPTION FACTOR AND TRANSPORT GENES IN SACCHAROMYCES CEREVISIAE Abstract The GABA shunt pathway is composed of three enzymes: glutamate decarboxylase (GAD, coded by GAD1), GABA aminotransferase (GABA-TA, coded for UGA1), and succinate semialdehyde dehydrogenase (SSADH, coded by UGA2). In Saccharomyces cerevisiae, the expression of UGA1, UGA2, and a GABA permease gene, UGA4 is regulated by the GABA specific transcription factor, UGA3. GABA permease UGA4 mediates the transportation of GABA into vacuoles. The deletion mutation of UGA3 grown in minimal medium with GABA as sole nitrogen source resulted in heat sensitivity, while ?uga3 conferred heat tolerance if grown on YPD or minimal medium with non GABA nitrogen sources. While a deletion mutant of ?uga4 was more heat tolerant than wild type on YPD medium is equivalent to wild type on all other media. The increased heat tolerance of the deletion mutant strains was not correlated with the expression of the enzymes of the GABA shunt, but appears to be at least partially mediated by the expression of ROS scavenging superoxide dismutases. The transcripts of GABA shunt genes, UGA3 and UGA4 were induced by GABA, and the expression of 84 UGA1, UGA2 and UGA4 were also induced by acidic pH. Under lethal heat stress at 45?C, each of the GABA shunt genes together with the regulatory genes of UGA3 and UGA4 were up-regulated in wild-type strain. Deletion mutation of the transcription factor UGA3 repressed the transcription activation of GABA transaminase (UGA1) and GABA permease (UGA4), but did not result in the change of the induction pattern for glutamate decarboxylase (GAD1) compared to the wild-type. Deletion mutation of UGA4 did not have significant effect on the expression pattern of GAD1, UGA1, UGA2, or UGA3 under lethal heat stress at 45?C. These results suggest that GABA shunt pathway and the regulatory genes of UGA3 and UGA4 play an important role in utilizing GABA as a nitrogen source and the GABA shunt protects yeast cells from heat damage through transcriptional up-regulation, which is partly regulated by the transcription factor UGA3. Introduction GABA is a four carbon non protein amino acid which is widely found in all prokaryotic and eukaryotic organisms (Kumar and Puneker, 1997). The GABA shunt is a metabolic pathway consisting of three enzymes: glutamate decarboxylase (GAD, encoded by the GAD1 gene) catalyzing the conversion of glutamate (Glu) to GABA, GABA aminotransferase (GABA-AT, encoded by the UGA1 gene) catalyzing the conversion of GABA and ?-ketoglutarate (?-KG) into succinate semialdehyde (SSA) and Glu, and succinate semialdehyde dehydrogenase (SSADH, encoded by the UGA2 gene) catalyzing the NAD(P)-dependent conversion of SSA into succinate. These three enzymes act in concert to move carbons from ?-ketoglutamate to succinate bypassing two reactions of the tricarboxylic acid cycle. 85 In yeast Saccharomyces cerevisiae, the UGA1 and UGA2 genes have been shown to play an essential role in the metabolism of GABA (Ramos et al., 1985) since these two enzyme activities are GABA-inducible. Such GABA-induced regulation is at least partially regulated through the GABA-specific transcription factor encoded by the UGA3 gene (Vissers et al., 1989). The import of GABA into cells is regulated by three GABA up-take systems: the general amino acid permease (GAP1p), the proline specific permease (PUT4p), and the GABA permease (UGA4p) (Grenson et al., 1987). The transcriptional activation of UGA1, UGA2, and UGA4 is positively regulated by the GABA-specific transcription factor, UGA3p, and a general nitrogen utilization related protein, UGA35p or DAL81p (Coornaert et al., 1991). The enzyme activity of UGA4 has been shown to be induced by the presence of GABA (Ramos et al., 1985) and it was also demonstrated that the induction of UGA4 transporter activity by GABA correlates with strong accumulation of UGA4 RNA transcript (Andre et al., 1993). In addition, in the absence of GABA, UGA4p activity was also shown to be induced by acidic pH (Moretti et al., 2001). To cope with the damaging effects from the environmental and physiological stresses such as heat stress, osmotic stress, or oxidative stress, yeast cells have developed rapid molecular responses by changing their gene expression patterns (Estruch, F.. 2000). Microarray analysis has shown that the expression of the three GABA shunt genes are up- regulated under mild heat stress at 37?C (Gasch et al., 2000; Sakaki et al., 2003), while the expression of UGA3 is unaffected (Gasch et al., 2000). However, these results have not been directly confirmed, and there is a lack of information on how the GABA shunt 86 genes are regulated under lethal heat stress temperature and the role of the GABA regulatory and transport genes (UGA3 and UGA4) in the heat stress response. Previously, we have shown that the three genes of GABA shunt are involved in thermotolerance under lethal heat stress at 45?C by restricting the accumulation of reactive oxygen species and altering metabolite levels during episodes of lethal heat stress (Chapter II). Here, we examine the role the GABA specific transcription factor and transport genes of UGA3 and UGA4 in lethal heat stress and report their mediation on the expression of the GABA shunt genes under lethal heat stress temperature. Additionally, we examined the role of the UGA3 regulated UGA4 (GABA transporter) in lethal heat stress. Cell survival and RT-PCR analysis were conducted in the wild-type and deletion mutants of uga3 and uga4. Materials and Methods Yeast strains and growth media Yeast strains used in this study were all derived from stain W303-1A (Mat a leu2- 3,112 ura3-1 trp1-1 his3-11,15 ade2-1, lys2, met) and the precise genotypes are listed in Table 1. In various experiments as detailed in text, table and figure legends, the strains were grown on either YPD medium or YNB medium at 30?C and shaken at 250-300 rpm. YPD medium contains: 2% glucose, 1% yeast extract, 2% yeast bactopeptone, and was used for expression and cell survival analyses under heat stress. YNB medium contains: 0.67% [wt/vol] yeast nitrogen base, 2% glucose, supplemented with essential amino acids (Sherman et al., 1979). 87 Gene disruptions The GABA specific regulatory gene UGA3 and GABA permease gene UGA4 were disrupted similarly as GAD1 as described in Chapter II by replacing the open reading frame with TRP1 gene. The TRP1 gene was amplified from plasmid pRS414 template DNA using a forward primer (named UGA3UP45TRP1A for UGA3 and UGA4UP45TRP1A for UGA4, Table 2) that consisted of the 45 nucleotides immediately upstream of the UGA3 or UGA4 gene fastened upstream of the 5?most portion of the TRP1 gene contained in pRS414. The reverse primer (named UGA3DN45TRP1D for UGA3 and UGA4DN45TRP1D for UGA4, Table 2) consisted of the 45 nucleotides immediately downstream of the UGA3 or UGA4 gene ligated to the 22 nucleotides at the 3? end of the TRP1 gene contained in pRS414. The PCR product was then transformed into yeast strain W303-1A by the lithium acetate method as previously described (Geitz and Woods, 2002), and TRP+ colonies were selected for genomic DNA isolation. The correct integration was verified by isolating genomic DNA (Hofman and Winston, 1987) from each selected strain and conducting a series of PCR reactions using primers that land 300-500 nucleotides upstream and downstream of UGA3 or UGA4 open reading frame and within the TRP1 gene (Table 2). Cell growth with different nitrogen sources and pH The standard minimal medium (without nitrogen source) as described by Ramos et al. (1985) was used for wild-type cell growth with ammonium sulfate, GABA, or glutamate as sole nitrogen source at pH 6.0. Minimal medium with ammonium sulfate as 88 sole nitrogen source at pH 4.0 was used to evaluate the acidic pH effect on gene expression. Nitrogen sources were added to a final concentration of 0.1%. Mid-log phase cells grown at 30?C on a rotary shaker (200 rpm) were collected by centrifugation at 3000 x g for 5 min at room temperature, frozen rapidly in liquid nitrogen and stored at -80?C for total RNA isolation. Lethal heat stress at 45?C For cell survival analysis, cells of wild-type and mutants of uga3 and uga4 were grown in YPD, YNB, minimal media with sole nitrogen sources of GABA, Glu and NH4+ (Ramos et al., 1985) respectively until mid-log phase. Cell survival assay was performed and evaluated as described in Chapter II except sample cells were stressed for 30 min. For gene expression analysis, mid-log phase cells of wild-type and mutants of ?uga3 and ?uga4 grown in YPD at 30?C were shifted to 45?C for 0, 5, 15, 30, 45 and 60 min. After heat stress, cells were put on ice for 1 min. Samples were then collected by centrifugation at 3000 ?g for 5 min at room temperature, frozen rapidly in liquid nitrogen and stored at - 80?C for total RNA isolation. Total RNA extraction and reverse transcriptase PCR (RT-PCR) Total yeast RNA was isolated as described by Peper (1997) with minor modification. Harvested yeast cells were washed with DEPC (diethylpyrocarbonate) treated water and then resuspended in 0.6 ml RNA extraction buffer (10 mM EDTA (ethylenediaminetetracetic acid), 50 mM Tris-HCl pH 7.5, 0.1M NaCl, 5% SDS), 0.6 ml of phenol:chloroform:isoamyl alcohol (50:50:1) mixture and 2 g of glass beads (0.45 mm diameter). Cells were broken by vigorous agitation for 5 min on a vortex mix set at 89 maximum speed with 1 min on ice between each 1 min intervals. The organic phase was separated from upper aqueous phase by centrifugation at 3000 x g for 5 min at room temperature. The supernatant was collected, extracted twice with equal volume of phenol: chloroform: isoamyl alcohol (50:50:1) and once with equal volume of chloroform: isoamyl alcohol (24:1). The RNA was precipitated by the addition of 0.1 volume of 3M NaOAc, pH 5.2, plus 2.5 volumes of ice-cold absolute ethanol and resuspended in DEPC treated water. RNA concentration was determined by measuring OD260 spectrophotometrically. First strand cDNA was synthesized from 4 ?g of total RNA using the SuperScript first-strand synthesis kit (Invitrogen, CA, USA) following manufactory's instructions. cDNA was amplified in a volume of 25 ?l containing 1U Taq DNA polymerase, 1? reaction buffer, 20 pmol each gene specific primer set for GAD1, UGA1, UGA2, UGA3, UGA4, SOD1, SOD2 and ACT1 (Table 2), 200 ?M dNTP, 1.5 mM MgCl2, and 2 ?l cDNA RT product. PCR products were separated on 0.8% agarose gels, and photographed under a UV light. ACT1 gene was used for internal control. Results Gene expression responses to different nitrogen sources and pH To examine the expression of the GABA shunt genes GAD1, UGA1 and UGA2, UGA3, and UGA4 by different nitrogen sources and different pH?s, RT-PCR experiments were carried out in strain W303-1A growing on minimal medium with GABA, minimal medium with glutamate, and minimal medium with ammonia as the sole nitrogen source respectively. Fig. 1 shows that the expression of all of these genes was the lowest with 90 NH4+ (pH 6.0) as sole nitrogen source. The expression of all of these genes was strongly induced by the presence of GABA or glutamate when compared to ammonia, but the expression of all of the genes was greatest with GABA as nitrogen source as compared with glutamate. These results demonstrate that the GABA shunt and the transcription factor UGA3 and the GABA permease UGA4 play an important role in utilizing GABA as a nitrogen source in yeast Saccharomyces cerevisiae. When cells were grown in minimal medium with NH4+ at pH 4.0, the expression of UGA1 and UGA4 which encode GABA transaminase and GABA permease respectively was highly induced compared to growth on the same medium at pH 6.0. The expression of UGA2 which encodes succinate semialdehyde dehydrogenase (SSADH) was also induced, but at a lower level than UGA1 and UGA4. However, GAD1, the first enzyme of GABA shunt and UGA3, which encodes a GABA specific transcription factor, were not regulated by the acidic pH. These data suggest that UGA1, UGA2 and UGA4 play more important roles than GAD1 and UGA3 in acid growth condition. Even though the expression of GAD1 and UGA3 are GABA inducible, they are regulated by a different mechanism from the UGA1, UGA2, and UGA4 genes. Cell survival under lethal heat stress of ?uga3 and ?uga4. UGA3 is a transcription factor that is known to positively regulate the transcription of UGA1, UGA2, and UGA4 (Coornaert et al., 1991). UGA4 is a GABA transport protein (Grenson et al., 1987). In Chapter II, we have shown that the GABA shunt mutant strains of ?gad1, ?uga1 and ?uga2 were sensitive to lethal heat stress at 45?C compared to the wild-type. Here, we investigated the role of the genes of UGA3 and UGA4 in response to lethal heat stress. 91 A UGA3 and UGA4 wild type strain (W303-1A), and the isogenic deletion strains ?uga3 and ?uga4 grown in YPD, YNB, and minimal medium with different nitrogen sources (GABA, Glu and NH4+) were subjected to lethal heat stress at 45oC for 30 min. As shown in Fig. 2., for the cells grown in rich YPD medium, after 30 min, the wild type cells showed 78% survival compared to the unstressed wild-type cells. The ?uga3, and ?uga4 deletions showed 90%, and 85% survival respectively, indicating that deletions of UGA3 and UGA4 result in cells that are more heat tolerant than wild-type. Cells grown in other GABA-free media such as YNB, minimal medium with Glutamate as the sole nitrogen source and minimal medium with NH4+ as the sole nitrogen source showed similar heat-tolerance results for ?uga3, and ?uga4 (data not shown). However, when cells were grown in minimal medium with GABA as the sole nitrogen sources, ?uga3 was found to have a defective, slow growth compared to the wild-type (data not shown), while the ?uga4 strain was found to grow normally. When cells grown on GABA as the sole nitrogen source were submitted to lethal heat stress as described above, after 30 min, wild-type and ?uga4 cells did not have significant survival change compared to the cells grown in rich YPD medium, showing 75% and 82% survival respectively. However, ?uga3 had a dramatic decrease in survival rate, showing 60% survival, indicating a heat sensitive phenotype when grown with GABA as the sole nitrogen source. Analysis of gene expression pattern for GABA shunt enzymes and the GABA specific regulatory and transport genes (UGA3 and UGA4) under lethal heat stress To examine the possible effect of lethal heat stress on the expression pattern of GAD1, UGA1, and UGA2 and the GABA specific regulatory (UGA3) and transport 92 (UGA4) genes, RT-PCR was performed to evaluate and compare the expression levels of transcripts during lethal heat stress at 45?C in wild type and the ?uga3 and ?uga4 mutants. Changes were observed at 5, 15, 30, 45, and 60 min following the initiation of heat stress. In the wild type strain, GAD1 was rapidly induced upon heat stress and the expression was gradually increased at each time point (Fig. 3A). The expression level was increased above 6- fold after 60 min of heat stress. UGA1 showed a similar gradual induction during 60 min heat treatment, but exhibited a more rapid induction upon 5 min of heating (3- fold increase) and peaked at 60 min (above 6- fold increase). UGA2 and UGA3 showed a similar induction pattern to GAD1, but the induction rates were slower for the first 15 min (1- to 2- fold increase), and then rapidly induced after 15 min. A 4- fold increase and above a 5- fold increase were observed for UGA2 and UGA3 respectively at 60 min. The basal expression level for UGA4 was low, almost undetectable without heat stress. Upon heat stress, UGA4 was gradually induced at each time point, but exhibited a slower rate of induction than any of the other genes investigated. Only a 2-fold induction was detected after 60 min heat stress. Thus, these results indicate that in the wild-type yeast strain, the expression of the three genes in the GABA shunt pathway and the GABA specific regulatory and transport genes of UGA3 and UGA4 respectively are all induced during heat-stress even though at slightly different levels. To further examine how the UGA3 and UGA4 genes regulate the expression of GABA shunt enzymes under heat stress, the expression patterns of the same genes as above were determined in the ?uga3 and ?uga4 deletion mutant strains under lethal heat stress at 45?C. In ?uga3 (Fig. 3B), the loss of a functional UGA3 coding sequence 93 resulted in the elimination of a detectable UGA3 transcript. GAD1 was rapidly and transiently induced upon heat stress at 45?C for 5 min compared to the unstressed control, and then the expression rapidly decreased to the control level at the15 min time point. After 15 min, the expression again increased through 60 min after heat stress. UGA2 gene expression showed a gradual induction pattern, but the induction was slow, the expression was non-detectable at the 0 and 5 min time points. However, no expression of UGA1 and UGA4 was observed either before or after heat stress in ?uga3. These results support the hypothesis that the transcription factor UGA3 gene is required for the transcriptional induction of the UGA1 and UGA4 genes which is in agreement with Vissers et al.(1989). Additionally, the transcriptional induction of UGA1 and UGA4 but not GAD1 and UGA2, is also mediated by UGA3 during episodes of heat stress. The transcriptional regulation of GAD1 and UGA2 appears to be involved in the mechanism of heat stress defense although this regulation appears to involve other mechanisms of activation. In the deletion mutant of ?uga4 (Fig. 3C), loss of the functional coding sequence resulted in non detectable expression of UGA4 before or after heat stress. The expression levels of GAD1 and UGA2 were relatively low before heat stress compared to those in the wild-type but gradually increased between 15 and 60 min. and between 30 and 60 min. respectively. UGA1 was also rapidly and gradually induced upon heat stress but to a relative higher level than in the wild-type. The expression of UGA3 was rapidly increased after 5 min heat stress, but then decreased to a level slightly higher than the control and maintained that level until 60 min following the initiation of heat stress. These data indicate that the induction of GAD1, UGA1, UGA2 and UGA3 at the transcriptional level 94 is result of heat stress, but the induction is independent of UGA4 although the failure to properly transport GABA may result in quantitative effects on the expression of these genes. Disruption of the regulatory and transport genes (UGA3 and UGA4) and antioxidant gene expression analysis under lethal heat stress. Heat stress has been documented to produce oxidative stress (Davidson et al., 1996; Davidson and R. H. Schiestl, 2001a; 2001b), and to some extent, heat stress is equivalent to oxidative stress (Sugiya et al., 2000). In chapter II, it was shown that cell survival of wild-type and all GABA shunt mutant strains were correlated to the levels of ROS accumulated under lethal heat stress. Here, we analyzed the gene expression of the antioxidant genes SOD1 and SOD2 (coding for cytosolic and mitochondrial localized superoxide dismutases respectively) in the wild-type and mutant cells under lethal heat stress. As shown in Fig. 4, before heat stress, the expression level of mitochondrial SOD (SOD2) was higher than the cytosolic SOD (SOD1) in all wild-type and mutant strains of ?uga3 and ?uga4. However, both SOD1 and SOD2 had much higher levels of expression in the mutant strains of ?uga3 and ?uga4 (Fig. 4B and Fig. 4C) compared to the wild- type (Fig. 4A). Upon lethal heat stress at 45?C, both SOD1 and SOD2 were gradually induced for the time points analyzed (Fig. 4A) in the wild type. However, the expression of SOD1 and SOD2 in the mutant strains ?uga3 and ?uga4 were constitutively and highly expressed before and during heat stress (Fig. 4B and Fig. 4C). These results indicate that an intact GABA-specific transcription factor (UGA3) or GABA permease (UGA4) is 95 required for the regulation of expression of both superoxide dismutases, and that when UGA3 or UGA4 are deleted this negative regulation is removed leading to high level, constitutive expression of the superoxide dismutases subsequently affecting the thermotolerance and ROS production by these mutants. Discussion Previously, GABA was reported to induce the enzyme activities of UGA1 and UGA2 (Ramos et al., 1985). The expression of all three of GABA shunt genes (GAD1, UGA1, and UGA2) and the GABA-specific regulatory and transport genes (UGA3 and UGA4 respectively) was determined on media with and without GABA (glutamate, or NH4+) (Fig. 1). The expression of all 5 of the above genes was greatest with GABA as sole nitrogen source. The UGA1, UGA2, UGA3, and UGA4 genes were also up-regulated on media containing GABA plus glutamate or GABA plus NH4+ but to a slightly lower level than on media with GABA alone. The lowest expression levels of these 4 genes were observed with glutamate or NH4+ alone. GAD1 showed essentially the same pattern of regulation except that the expression levels of GAD1 were greater on glutamate- containing media than on NH4+-containing media. These results confirm the essential roles of all 5 of these genes in GABA utilization as a nitrogen source and demonstrate that there may be at least 2 different mechanisms for regulating the expression of the GABA utilization genes. The induced expression of UGA1 and UGA2 is coordinated with the increased enzyme activities of Uga1p and Uga2p in the presence of GABA (Ramos et al., 1985). The induction of UGA3 and UGA4 96 was coupled with the induction of UGA1and UGA2 further confirming the positive regulatory role of UGA3 on the transcription of GABA-related genes. GABA is incorporated into yeast cells through three permeases (Grenson et al., 1987). Among these, UGA4 is the only GABA?specific transporter. The synthesis of UGA4 has also been shown to be GABA-inducible (Grenson et al., 1987) and was regulated by at least two positive factors, UGA3p and the pleiotropic UGA35p factor (also referred to as Dal81p/DurLp) (Andr? et al., 1995). Without the presence of GABA in the medium, the expression of UGA4 has also been shown to be induced by acidic pH (Moretti et al., 2001). However, the expression of the other GABA-related genes by acidic media has not been investigated. Here we have shown that the expression of UGA1, UGA2, and UGA4 were all increased at acidic pH on media containing NH4+ as the nitrogen source. Acidic pH does not appear to have significant effect on the induction of GAD1 and UGA3 genes. These results indicate that the induction of UGA1, UGA2, and UGA4 by low pH is mediated by a mechanism different from the expression of the other two GABA-related genes (GAD1 and UGA3). Whether this mechanism involves Uga3p cannot be determined from the data presented here. However, since acidic pH does not effect UGA3 expression it is a reasonable hypothesis that acid regulation does not involve Uga3p. Mutation of GABA specific transcription factor (UGA3) and GABA-specific permease (UGA4) resulted in an increase in tolerance of yeast cells to a lethal heat stress at 45?C when cells were grown in rich YPD medium (Fig. 2). Deletion of UGA3 resulted in more increase in stress tolerance than the deletion of UGA4. Essentially similar results were obtained when cells were grown on synthetic complete medium or minimal medium 97 with glutamate or ammonium sulfate as sole nitrogen source (data not shown) indicating that the rich source of amino acids present in YPD medium does not contribute to the heat stress tolerance phenotype of wild type or the mutants. However, the UGA3 deletion generated a decrease in heat stress tolerance when cells were grown on GABA as the sole nitrogen source. This is likely because the expression of UGA1 and UGA4 are eliminated in ?uga3 (Fig 3B), and it has previously been shown that the loss of function of GABA transaminase (UGA1) in a deletion mutant (Chapter II) decreases thermotolerance. Notably in ?uga4, GABA transaminase is not down-regulated as it is in ?uga3. In chapter II and IV, we demonstrated that the three enzymes of the GABA shunt were involved in different levels of lethal heat stress tolerance. Under mild heat stress at 37?C, microarray data show that the three enzymes in the GABA shunt pathway are highly up-regulated (Gasch et al., 2000; Sakaki et al., 2003). These results are further substantiated and expanded here by the observation that the transcripts of the three GABA shunt genes were gradually increased under lethal heat stress at 45?C compared to the unstressed control in the wild-type strain (Fig. 3A). The GABA specific transcription factor gene and the GABA permease gene showed expression patterns under lethal heat stress (Fig. 3A) similar to the genes for the GABA shunt enzymes. Deletion of GABA permease (encoded by UGA4) did not change the induction pattern of the three GABA shunt enzymes and the GABA specific transcription factor (encoded by UGA3) under lethal heat stress (Fig. 3C). However, compared to the wild- type, the transcript level of GABA transaminase UGA1 was higher and the induction of GAD1, UGA2 and UGA3 were slower. This effect may be explained by the existence of 98 three GABA permeases in yeast, simultaneous deletion of the three GABA permeases is needed for further investigation. In unstressed control cells, deletion of UGA3 did not result in a change in GAD1 expression. However, UGA1 and UGA4 were virtually unexpressed during heat stress (Fig. 3B). The expression of UGA2 gene was induced more slowly than in the wild-type strain even though the expression was undetectable before heat stress (Fig. 3B). Thus, it can be concluded that under lethal heat stress, the transcription factor UGA3 gene is required to activate the transcription of UGA1, UGA2, and UGA4, but UGA3 not directly involved in the induction of GAD1. Apparently, factors in addition to UGA3 are involved in regulating the expression of UGA2 which are probably activated by prolonged heat stress. Given the gene expression pattern of the ?uga3 mutant, the heat stress tolerance phenotype is unexpected. Thus, an alternative overriding pleiotrophic factor must be involved in the heat stress phenotype of ?uga3. The heat stress tolerant phenotypes of the mutant strains were further supported by the expression levels of the superoxide dismutase SOD1 and SOD2 antioxidant genes in the ?uga3 and ?uga4 mutant strains (Fig. 4). Cells grown in YPD medium showed higher levels of transcripts for both SOD1 and SOD2 in the mutant strains than in the wild-type before heat stress. The expression of both genes was induced upon heat stress in the wild-type strain but after the initiation of stress. These data indicate a negative regulatory role for the GABA specific transcription factor and/or permease proteins on the expression of superoxide dismutases in yeast. The induced expression of SOD1 and SOD2 may be necessary for detoxifying ROS induced by heat assisting in the reduction of damage caused by the initial production 99 of ROS. The higher transcript level of SOD2 than SOD1 in all the wild-type and mutant strains is consistent with the hypothesis that superoxide dismutase is required for the detoxification of ROS in mitochondria more than in cytosol. References 1. Andr? B., D. Talibi, S. Soussi-Boudekou, C. Hein, S. Vissers and D. Coornaert. 1995. Two mutually exclusive regulatory systems inhibit UASGATA, a cluster of 5?-GAT(A/T)A-3? upstream from UGA4 gene of Saccharomyces cerevisiae. Nucleic Acids Res. 23: 558?564. 2. Andre, B., C. Hein, M. Grenson, and J. C. Jauniaux. 1993. Cloning and expression of the UGA4 gene coding for the inducible GABA-specific transport protein of Saccharomyces cerevisiae. Mol. Gen. Genet. 237:17-25. 3. Bouch? N, A. Fait, D. Bouchez, S. G. M?ller, and H. Fromm. 2003. Mitochondrial succinic-semialdehyde dehydrogenase of the ?-aminobutyrate shunt is required to restrict levels of reactive oxygen intermediates in plants. Proc. Natl. Acad. Sci. U.S.A 100: 6843?6848. 4. Coleman S.T., T. K. Fang , S. A. Rovinsky, F. J. Turano, and W. S. Moye- Rowley. 2001. Expression of a glutamate decarboxylase homologue is required for normal oxidative stress tolerance in Saccharomyces cerevisiae. J Biol Chem. 276: 244-50. 5. Coornaert, D., S. Vissers, and B. Andre. 1991. The pleiotropic UGA35(DURL) regulatory gene of Saccharomyces cerevisiae: cloning, sequence and identity with the DAL81 gene.Gene (Amst.) 97: 163-171. 6. Davidson, J. F., and R. H. Schiestl. 2001a. Cytotoxic and genotoxic consequences of heat stress are dependent on the presence of oxygen in Saccharomyces cerevisiae. J. Bacteriol. 183: 4580-4587. 7. Davidson, J. F., and R. H. Schiestl. 2001b. Mitochondrial respiratory electron carriers are involved in oxidative stress during heat stress in Saccharomyces cerevisiae. Mol. Cell. Biol. 21: 8483-8489. 8. Davidson, J. F., B. Whyte, P. H. Bissinger, and R. H. Schiestl. 1996. Oxidative stress is involved in heat-induced cell death in Saccharomyces cerevisiae. Proc. Natl. Acad. Sci. USA. 93: 5116-5121. 100 9. Estruch, F. 2000. Stress-controlled transcription factors, stress-induced genes and stress tolerance in budding yeast. FEMS Microbiol. Rev. 24: 469-486. 10. Gasch, A. P., P. T. Spellman, C. M. Kao, O. Carmel-Harel, M. B. Eisen, G. Storz, D. Botstein, and P. O. Brown. 2000. Genomic expression programs in the response of yeast cells to environmental changes. Mol Biol Cell. 11(12): 4241-57. 11. Geitz, R. D. and R. A. Woods. 2002. Transformation of yeast lithium acetate/single-stranded carrier DNA / polyethylene glycol method. Methods Enzymol. 350:87-96. 12. Grenson, M., F. Muyldermans, K. Broman, and S. Vissers. 1987. 4- Aminobutyric acid (GABA) uptake in baker?s yeast Saccharomyces cerevisiae is mediated by the general amino acid permease, the proline permease and a specific permease integrated into the GABA-catabolic pathway. Biochemistry Life Sci Adv 6: 35?39. 13. Hoffman, C. S., and R. Winston. 1987. A ten-minute DNA preparation from yeast efficiently releases autonomous plasmids for transformation of Escherichia coli. Gene (Amst.) 57: 267-272. 14. Kumar, S. and N. S. Punekar. 1994. Succinicsemialdehyde dehydrogenase from Aspergillus niger. A kinetic enquiry. Abstract P7-343, XVI IUBMB. International Congress, New Delhi. 15. Moretti, M. B., A. Batlle, and S. C. Garcia. 2001. UGA4 gene encoding the ?- aminobutyric acid permease in Saccharomyces cerevisiae is an acid-expressed gene. The International Journal of Biochemistry & Cell Biology. 33(12): 1202- 1207. 16. Piper W. 1997. Measurement of Transcription. In Molecular Genetics of Yeast-A practical Approach Edited by : Johnston JR. Oxoford University Press, New York; 135-138. 17. Ramos F, M. Guezzar, M. Grenson, and J. M. Wiame. 1985. Mutations affecting the enzymes involved in the utilization of 4-aminobutyric acid as nitrogen source by the yeast Saccharomyces cerevisiae. Eur J Biochem 149: 401?404. 18. Sakaki, K., K. Tashiro, S. Kuhara, and K. Mihara. 2003. Response of genes associated with mitochondrial function to mild heat stress in yeast Saccharomyces cerevisiae. J Biochem (Tokyo). 134(3): 373-84. 19. Sherman, F., G.R. Fink, and C.W. Lawrence. 1979. Methods in Yeast Genetics. Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y. 101 20. Sugiyama, K., S. Izawa, and Y. Inoue. 2000. The Yap1p-dependent induction of glutathione synthesis in the heat-shock response of Saccharomyces cerevisiae. J Biol Chem. 275(20): 15535?15540. 21. Vissers, S., B. Andre, F. Muyldermans, and M. Grenson. 1989. Positive and negative regulatory elements control the expression of the UGA4 gene coding for the inducible 4-aminobutyric acid-specific permease in Saccharomyces cerevisiae. European Journal of Biochemistry. 181: 357?361. 102 Table 1. Yeast strains used in this study Strain Genotype Source and description W303-1A Mat a leu2-3,112 ura3-1 trp1-1 his3-11,15 ade2-1, lys2, met ATCCa, WT ?uga3 Mat a leu2-3112 ura3-1 trp1-1 his3-11,15 ade2-1, lys2, met uga3::TRP1 This study, uga3 disruptant ?uga4 Mat a leu2-3112 ura3-1 trp1-1 his3-11,15 ade2-1, lys2, met uga4::TRP1 This study, uga4 disruptant 103 Table 2. Primers used for UGA3 and UGA4 gene deletions and verifications Primer Name Sequence (5? to 3?) UGA3UP45TRP1A CACGTCGCTCTTAACAATCCAGGCT GAACAAAACAAGGAATAATGGGAAGCATTTAATAGACAGCATCGT UGA3DN45TRP1D TACATACATATAGGGGGCGGTA TATTGGATGACCTTTTCAACTCAAGGCAAGTGCACAAACAATACT UGA3/TRP1KO_A GGTAAGAAATGAAACAATAGGACGA UGA3/TRP1KO_B GAATATTTCAATTTCAGCTTCTCCA UGA3/TRP1KO_C GCATTCAGGTAGAAGATATGGAGAA UGA3/TRP1KO_D TCATGATATAGATATGATTGGCGG UGA4UP45TRP1A TTGTGAAGTGTAACAAGGTCTTATA ATTTATTATTACTAACAATGGGAAGCATTTAATAGACAGCATCGT UGA4DN45TRP1D TTATAAACTCTGAATATAAAAT CTTTATAAAGGTTTGAACATTTAAGGCAAGTGCACAAACAATACT UGA4/TRP1KO_A TTTATCGAATAAGGGGAGAACCTAC UGA4/TRP1KO_B AAATTTCCATCTTTGGTTAATGTGA UGA4/TRP1KO_C ATTATTTTGGTTATGTTCCCCTCTC UGA4/TRP1KO_D TAATGGAATGGAGAGTGATGATTTT TRP1B TCTGCAAGCCGCAAACTTT TRP1C AGTTCCTCGGTTTGCCAGTTATT ACT1F GGTATGTGTAAAGCCGGTTTT ACT1R AGGATGGAACAAAGCTTCTGG 104 UGA1 UGA2 UGA3 UGA4 ACT1 GAD1 pH 4.0 Glu Glu + GABA GABA GABA + NH4+ NH4+ NH4+ N sources, pH 6.0 Fig. 1. Transcript levels of GABA shunt genes (GAD1, UGA1, UGA2), the GABA specific transcription factor (UGA3) and GABA permease (UGA4) in response to minimal media with different nitrogen sources and acidic pH. RT-PCR analysis was used to determine transcript abundance in mid-log phase cells growing at 30?C as described in Materials and Methods. The ACT1 gene was served as internal control. 105 WT ?uga3 ?uga4 Ce ll s ur viv al (% ) 0 20 40 60 80 100 0 min 30 min, YPD 30 min, minimal-GABA Ce ll s ur viv al (% ) Fig. 2. Viability of wild-type, ?uga3, and ?uga4 mutant strains after lethal heat treatment at 45oC. Strains were grown to mid log phase (5 ? 106 cells/ml) in either YPD medium or minimal medium with GABA as sole nitrogen source, washed in 0.87% NaCl and concentrated to 2 ? 108 cells/ml. Aliquots of 100 ?l were heated at 45oC in a thermocycler for 30 min. Viability was determined as described in Materials and Methods. Cell survival percentage was determined prior to heat stress (black bar), 30 min after heat stress in YPD medium (light grey bar), or 30 min after heat stress in minimal medium with GABA as the nitrogen source (dark grey bar). Each point represents the mean ? SD of the results from at least three experiments. 106 GAD1 UGA1 UGA3 UGA4 UGA2 ACT1 0 5 15 30 45 60 min wt + 45?C 0 5 15 30 45 60 min ?uga3 + 45? C ?uga4 + 45? C 0 5 15 30 45 60 min A B C Fig. 3. Transcript levels of GABA shunt genes (GAD1, UGA1 and UGA2) and the GABA specific regulatory and transport genes (UGA3 and UGA4 respectively) in response to lethal heat stress at 45?C. RT-PCR analysis (see Materials and Methods) was used to determine transcript abundance in mid-log phase yeast cells grown in YPD from (A) wild-type; (B) ?uga3 mutant; (C) ?uga4 mutant following exposure to heat stress (45?C) for the time points indicated. 107 ACT1 SOD1 SOD2 0 5 15 30 45 60 min ?uga3 + 45?C B 0 5 15 30 45 60 min wt + 45?C A 0 5 15 30 45 60 min ?uga4 + 45?C C Fig. 4. Transcript levels of the cytosolic and mitochondrial superoxide dismutases (SOD1 and SOD2 respectively) in response to lethal heat stress at 45?C. RT-PCR analysis (see Materials and Methods) was used to determine transcript abundance in mid-log phase yeast cells grown in YPD from (A) wild-type; (B) ?uga3 mutant; (C) ?uga4 mutant following exposure to heat stress (45?C) for the time points indicated. 108 IV. GABA TRANSAMINASE: COMPLEMENTATION ANALYSIS AND INTRACELLULAR LOCALIZATION IN SACCHAROMYCES CEREVISIAE Abstract GABA transaminase is the enzyme which catabolizes GABA conversion to succinate semialdehyde (SSA) in the GABA shunt pathway. GABA transaminase is ?- ketoglutarate dependent (GABA-TKG), localized in the cytosol, and encoded by UGA1 gene in yeast while in plants it is pyruvate dependent (GABA-TP) and localized in mitochondria. The web-based utilities TargetP 1.1 and PSORT predicts that GABA transaminase from Arabidopsis (AtGABA-TP) is localized in the mitochondria with a mitochondrial targeting peptide sequence localized in 54 nucleotides at the 5?end of the coding sequence. Yeast GABA transaminase (Uga1p or ScGABA-TKG) is predicted to be localized in cytosol. To examine the impact of localization differences between ScGABA-TKG and AtGABA-TP on physiological function, expression vectors were constructed that altered organellar targeting information to obtain expression of. Physiological function was evaluated by complementation of yeast GABA transaminase mutant ?uga1 and ?uga2 phenotype of: GABA growth defect, thermosensitivity and heat induced production of reactive oxygen species (ROS). Our studies revealed that AtGABA-TP is functionally interchangeable with ScGABA-TKG for GABA growth, 109 thermotolerance, and limiting production of ROS production whether they are located in mitochondria or cytosol in yeast. Introduction GABA is a ubiquitous non protein amino acid which is widely distributed from prokaryotic to eukaryotic organims (Kumar, 1997). It was first identified in 1950 in plants (Hulme & Arthington, 1950). GABA is produced from glutamate by glutamate decarboxylase (GAD), and further transaminated by GABA tranaminase to succinate semialdehyde (SSA) which is subsequently converted to succinate (SUCC) feeding into the TCA (Krebs) cycle. The role of GABA in animals is well established as a neuronal transmission inhibitor, the deficiency in catabolizing GABA through GABA transaminase (GABA-T) and SSADH is characterized by non-specific neurological disorder including psychomotor retardation, language delay, occasional seizures (Hogema et al., 2001; Gropman, 2003). In plants, GABA is catabolized to SSA by a pyruvate-dependent GABA transaminase (AtGABA-TP) (Shelp et al., 1999). Arabidopsis mutants of SSADH are hypersensitive to environmental stress such as heat and accumulate reactive oxygen species (ROS) during episodes of heat stress (Bouche et al., 2003). In yeast, GABA is catabolized to SSA by ?-ketoglutarate-dependent GABA transaminase (GABA-TKG), the product of the yeast UGA1 gene. Subsequently, SSA is converted to SUCC by SSADH, the gene product of the yeast UGA2 gene. Yeast mutants deficient in UGA1 and UGA2 were shown to be sensitive to the oxidative stress (Coleman et al., 2001). Mutant strains lacking UGA1 or UGA2 were found to be incapable of growth on minimal medium with 110 GABA as the sole nitrogen source. This growth defect was not observed during growth on ammonium sulfate as the sole nitrogen source (Ramos et al., 1985). Previously, we have shown yeast uga1 and uga2 deletion mutants were more sensitive to heat stress than is wild type and the uga1 mutant is more sensitive to heat stress than is the uga2 mutant. This thermosensitivity phenotype was shown to correlate with high levels of intracellular ROS; In that the uga1 deletion mutant produces more ROS than does uga2 during heat stress although both produce more ROS than wild type (Chapter II). Bioinformatic analysis utilizing the BLAST search utility at NCBI suggests that plants do not contain orthologs of yeast ScGABA-TKG and that yeast contains no orthologs of plant GABA-TP. TargetP 1.1. and PSORT predicts Arabidopsis AtGABA- TP is localized in the mitochondria and that it contains an 18 amino acid amino-terminal mitochondrial targeting peptide. These same utilities also predict that ScGABA-TKG and ScSSADH enzymes are localized in cytosol. This leaves 2 prominent hypotheses to explain this comparative difference between the behavior of plant and yeast mutants in GABA shunt enzymes during heat stress. First, mitochondrial localization of the plant GABA transaminase and SSADH may result in this difference, or alternatively the fact that the plant GABA transaminase utilizes pyruvate while the yeast transaminase utilizes ?-ketoglutarate or both may result in the stress sensitivity and ROS production differences between plants and yeast. To test these hypotheses, expression plasmids were constructed to alter the cellular locations of plant GABA-TP and yeast GABA-TKG in yeast. Thus, we overexpressed a set of 4 constructs in yeast uga1, uga2, and uga1uga2 deletion mutants. The 4 constructs 111 consisted of the yeast UGA1 coding region; the yeast UGA1 coding region also containing a 54 nucleotide sequence that would code for the mitochondrial transit peptide from AtGABA-TP; the AtGABA-TP coding region; and the AtGABA-TP coding region lacking the 54 nucleotides coding for the mitochondrial targeting peptide. All of these constructs were placed under the control of the strong constitutive GPD promoter. The mutant strains with GABA growth defect, thermosensitivity phenotype and ROS production during heat stress were assayed to determine the phenotype of each of theses constructs. Physiological analyses were conducted to determine the extent that differently localized enzymes compensate for these phenotypes. Materials and methods Bacterial and yeast strains and growth media. The Escherichia coli strain DH5? (Invitrogen, Carlsbad, CA, USA) was used for molecular cloning. Bacteria were grown on standard Luria Broth medium (LB medium) supplemented with antibiotics as required. All yeast strains used in this study and their genotypes are listed in Table 1. The yeast strain W303-1A was purchased from American Type Culture Collection, and ?uga1, ?uga2, and ?uga1?uga2 were derived from W303-1A as previously described (Chapter II). The strains were grown on either YPD medium or YNB minimal medium. YPD medium contains: 2% glucose, 1% yeast extract, 2% yeast bactopeptone, and was used for growth, cell survival, and ROS detection. YNB medium contains: 0.67% [wt/vol] yeast nitrogen base and 2% glucose supplemented with essential amino acids 112 (Sherman et al., 1979). YNB medium was used for cells grown on GABA as the sole nitrogen source. Gene cloning and plasmid construction. The endogenous S. cerevisiae UGA1 open reading frame (UGA1 ORF) was PCR amplified from the genomic DNA isolated from the wild type strain W303-1A using UGA1FOR and UGA1REV primers (Table 2). Yeast UGA1 with the AtGABA-TP (NC_003074.4) 5?-54 nucleotide sequence coding for the 18 amino acid mitochondrial transit peptide added 5? to the UGA1 ORF sequence was constructed by a two-round PCR procedure. In the first round of PCR W303-1A genomic DNA was used as template and mTP15 UGA1FOR and UGA1REV (Table 2) were used as forward and reverse primers respectively. mTP15 UGA1FOR consisted of 15 nucleotides from 3?-end of the mitochondrial targeting peptide 54 nucleotide sequence in front of the 5?-most 27 nucleotides of UGA1. The PCR product was purified using a GENECLEAN kit (Biogene) and used as template for the second round of PCR. The second round of PCR was conducted using mTP54 UGA1FOR which has 54 nucleotides from the mitochondrial targeting peptide nucleotide sequence as the forward primer, and the same reverse primer UGA1REV (Table 2) as above. To facilitate plasmid construction, two restriction sites for BamHI and HindIII (underlined, Table 2) were introduced at each end of the specific primers utilized. Following PCR amplification, the 1.778-kb (UGA1 orf) or 1.832-kb (mTP+UGA1 orf) BamHI / HindIII fragment containing the entire gene was ligated into the pGEM Teasy vector using DNA ligase (Promega) according to manufacturer's procedure, and the ligated vector was transformed into bacterial strain 113 DH5? competent cells by CaCl2 as described (Sambrook and Rusell, 2001). Plasmid DNA was isolated and purified by QIAGEN Plasmid Mini Kit(QIAGEN, USA), and then digested by BamHI / HindIII (Biolabs, New England) according to manufacturer's procedures and ligated into the unique BamHI / HindIII sites of yeast shuttle vector p425 GPD (American Type Culture Collection) using DNA ligase (Promega) according to manufacturer's procedure to create plasmids p425 GPD-UGA1 and p425 GPD-mTP54 UGA1. The full length AtGABA-TP orf sequence (NC_003074.4) was PCR amplified using p416 GPD-pGAT (Barbosa, 2001) as template DNA which contains the entire cDNA sequence of AtGABA-TP using FL-pGATFOR and pGATREV (Table 2) as forward and reverse primers respectively. The truncated pGAT without mitochondrial targeting peptide sequence was also PCR amplified from plasmid p416 GPD-pGAT by using FL- 51-pGATFOR (Table 2) as forward primer and the same reverse primer as above. To facilitate plasmid construction, two restriction sites for BamHI and HindIII (underlined, Table 2) were introduced at each end of the specific primers. Following PCR amplification, the 1.515-kb or 1.464-kb BamHI / HindIII fragment containing the entire gene was replicated in pGEM Teasy vector, digested, and ligated into the unique BamHI / HindIII site of yeast shuttle vector p425 GPD (American Type Culture Collection) as described above to create plasmid p425 GPD-pGAT and p425 GPD-pGAT-mTP. Gene disruptions UGA1 was disrupted by replacing the coding sequences with HIS3 gene as described (Chapter II). UGA2 was disrupted by replacing the coding sequences with URA3 gene as described (Chapter II). 114 Yeast transformation The plasmid constructs p425 GPD-UGA1, p425 GPD-mTP54UGA1, p425 GPD- pGAT and p425 GPD-pGAT-mTP were used to transform the ?uga1, ?uga2, and ?uga1?uga2 strains using lithium acetate method as described (Geitz et al., 1995) to generate the complementation strains of ?uga1-p425 GPD-UGA1, ?uga1-p425 GPD- mTP54UGA1, ?uga1- p425 GPD-pGAT, ?uga1- p425 GPD-pGAT-mTP; ?uga2-p425 GPD-UGA1, ?uga2-p425 GPD-mTP54UGA1, ?uga2- p425 GPD-pGAT, ?uga2- p425 GPD-pGAT-mTP; and ?uga1?uga2-p425 GPD-UGA1, ?uga1?uga2-p425 GPD- mTP54UGA1, ?uga1?uga2- p425 GPD-pGAT, ?uga1?uga2- p425 GPD-pGAT-mTP. To minimize the nutritional effect of amino acids supplied from the medium as nitrogen source which are not needed for the growth of uga1 and uga2 mutants, the empty shuttle vectors which carry HIS( p423 GPD), URA(p426 GPD), LEU(p425 GPD) and TRP(p424 GPD) were used to transform into wild-type strain, URA(p426 GPD) and TRP(p424 GPD) were used to transform into ?uga1 complementation strains, and HIS( p423 GPD) and TRP(p424 GPD) were used to transform into ?uga2 complementation strains. The transformed strains were then grown on minimal media plus essential amino acids (Ade, Lys, Met) with 10mM GABA as sole nitrogen source to test the complementation of GABA growth phenotype for strains lacking UGA1 or UGA2 (Ramos et al., 1985). Lethal heat stress survival assays Lethal heat stress survival assays were performed as described in Chapter II using the strains in Table 1 and the complementation strains outlined above. 115 Measurement of ROS production. Intracellular ROS generated during heat stress were measured as described in Chapter II using the strains in Table 1 and the complementation strains outlined above. Subcellular fractionation An overnight culture grown at 30?C with vigorous shaking was diluted into 500 ml YPD media for wild type and mutant strains and grown until the OD600 reached 2.0. Cellular fractionation to produce mitochondrial and cytosolic fractions was performed as described (Daum et al., 1982). Briefly, mid-log phase cells were harvested by centrifugation at room temperature for 5 min at 3,000 g, washed once with distilled water, resuspended in DTT buffer (0.1 M Tris.SO4, pH 9.4, 10 mM dithiothreitol) at 2 ml per gram of wet weight cells, and incubated at 30?C for 20 min. Cells were centrifuged for 5 min at 3,000 g, and the pellets were then washed once with Lyticase buffer (1.2 M sorbitol, 20 mM KPO4, pH 7.4) to give 7ml/g wet weight. Cells were resuspended in Lyticase buffer containing 0.5 mg Lyticase/ml (Sigma Aldrich), the suspension was incubated at 30?C with gentle shaking for 60 min until all the cells had been converted to spheroplasts. Spheroplasts were harvested by centrifugation at room temperature for 5 min at 3,000 g, and the pellets were washed twice with Lyticase buffer. Cells were centrifuged at room temperature for 5 min at 3,000 g, and pellets were resuspended in ice cold homogenization buffer (0.6 M sorbitol, 10 mM Tris-C1, pH 7.4, 1 mM ethylenediaminetetraacetic acid (EDTA), 0.1% bovine serum albumin (BSA), 1 mM phenylmethylsulfonyl fluoride (PMSF)) to a concentration of 6.5 ml/ g wet weight. Spheroplasts were homogenized by 10 -15 strokes in a tight-fitting Dounce homogenizer. 116 From this point on, all operation were carried at 0 ? 4?C. The homogenate was diluted with 1 volume of ice cold homogenization buffer and centrifuged for 5 min at 1500 g to pellet the cell debris and nuclei. Supernatants were centrifuged for 5 min at 4000 g, and the pellets were discarded. The resulting supernatant was referred to as the total yeast protein fraction. The mitochondrial fraction was isolated by centrifugation of the supernatant at 12,000 g for 15 min. The resulted supernatant was referred to cytosolic protein fraction. The crude mitochondrial pellet was resuspended in cold SEM buffer (250mM sucrose, 1mM EDTA, 10mM MOPS-KOH, pH 7.2) to a final concentration of 5-10 mg/ml protein. For further purification, the mitochondrial suspension was loaded on top of the sucrose step gradients: 1.5 ml 60%, 4 ml 32%, 1.5 ml 23%, and 1.5 ml 15% (w/v) sucrose in EM buffer (1mM EDTA, 10mM MOPS-KOH, pH 7.2), and centrifuged for 60 min at 2?C at 134,000 g. The purified mitochondria was recovered from 60% / 32% interface, concentrated by centrifugation to a protein concentration of 5-10 mg/ml, quickly frozen in liquid nitrogen, and stored at -80?C for further use. For enzyme assays, mitochondrial pellets were broken by thawing and freezing, subcellular fractionations were assayed for cross contamination of fractions using NAD+ dependent isocitrate dehydrogenase (NAD-IDH) as a mitochondrial marker and hexokinase 1 as a cytosolic marker. Enzyme activity assays Published procedures were used to assay GABA-transaminase (GABA-T), succinate semialdehyde dehydrogenase (SSADH), mitochondrial NAD+ dependent isocitrate dehydrogenase (NAD-IDH) and cytosolic hexokinase 1. Protein concentrations were 117 determined by Bradford assay (Bradford, 1976) with crystalline bovine serum albumin (BSA) as the standard. The activity of GABA-T was assayed as described (Ramos et al., 1985). The standard reaction mixture contains 100 mM potassium phosphate buffer, pH 8.2; 0.2 mM EDTA; 7.5 mM potassium 2-oxoglutarate or pyruvate; 0.2 mM ?-NAD+; 1 unit of yeast SSADH (Chapter V); and an appropriate amount of each cell fraction. The reaction was started by the addition of 7.5 mM GABA after pre-incubation of components at 30oC for 5 min. The activity of SSADH was assayed as described (Ramos et al., 1985). The standard reaction mixture contains 100 mM potassium phosphate buffer, pH 8.2; 0.2 mM EDTA; 0.2 mM NAD+; and an appropriate amount of each cell fraction. The reaction was started by the addition of 0.1 mM SSA after pre-incubation of components at 30oC for 5 min. NAD-IDH, a mitochondrial marker, was assayed as described (Illingworth et al., 1972). The standard reaction mixture contains 40 mM Tris-HCl buffer, pH7.6; 4 mM- MgCL2; 0.25 mM NAD+ and an appropriate amount of each cell fraction. The reaction was started by addition of 2.5 mM trisodium DL-isocitrate after pre-incubation of components at 30oC for 5 min. Hexokinase 1, a cytosolic marker, was assayed as described (Sigma Technical Bulletin SPGLYC100, Bergmeyer, 1983). The standard reaction mixture contains 39 mM triethanolamine, 0.74 mM adenosine 5?-triphosphate (ATP), 7.8 mM magnesium chloride, 1.1 mM ?-nicotinamide adenine dinucleotide phosphate (?-NADP), 2.5 units glucose-6- phosphate dehydrogenase (Sigma Aldrich), and an appropriate amount of each cell 118 fraction. The reaction was started by the addition of 216 mM D-glucose after pre- incubation of components at 30oC for 5 min. The enzymatic activity assays were based on the change in absorbance at 340 nm caused by the reduction of NAD(P)+ to NAD(P)H. All enzymatic reactions were performed at 30oC for 60 min and the change of absorbance at 340 nm was monitored on spectrophotometer (Beckman DU 640, USA). One unit of enzyme for GABA transaminase and succinate semialdehyde dehydrogenase was defined as the amount of enzyme required to produce 1 ?mol of NADH per hour at 30?C. One unit of enzyme for NAD-IDH and hexokinase 1 was defined as the amount of enzyme required to reduce 1 ?mol of NAD+ per min at 30?C. All assays were repeated at least three times and data represented the average of the results from at least three experiments. Results Subcellular localization of GABA shunt enzyme activities in Saccharomyces In order to investigate the interchangeable function of Arabidopsis thaliana AtGABA-TP and yeast ScGABA-TKG, plasmid constructs were made as indicated in Materials and Methods that contained AtGABA-TP with (+P) and without the mitochondrial targeting peptide (+P-mtp) or that contained ScGABA-TKG (+Y) or ScGABA-TKG with the AtGABA-TP mitochondrial targeting peptide (+Y+mtp). The intent was to be able to examine functional complementation of a set of yeast mutants overexpressing these two proteins localized in either the mitochondria or the cytosol. Prior to this investigation the proper localization of the proteins made by these constructs was investigated. 119 The TargetP 1.1 (Nielsen et al., 1997; Emanuelsson et al., 2000) and PSORT (Nakai and Horton, 1999) web based utilities predict AtGABA-TP to be localized in mitochondria with an 18 amino acid (54 nucleotide) mitochondrial trageting peptide in Arabidopsis thaliana. These utilities also indicate that these sequences will localize to mitochondria in yeast. The yeast GABA degrading enzymes ScGABA-TKG and ScSSADH were predicted by Target P 1.1 and PSORT to localize to the cytosol. To further verify the subcellular localization predictions of UGA1p, UGA2p and whether AtGABA-TP is localized in yeast mitochondria when expressed in yeast, we tested isolated subcellular fractions derived from appropriate stains for GABA-TKG, SSADH, and GABA-TP activities as described in Methods. Mitochondria could be easily separated from cytosol by differentiation centrifugation after cell lysis (Daum et al., 1982). Cytosolic hexokinase 1 was employed as a marker enzyme for the cytosol fraction, and mitochondrial NAD+-dependent isocitrate dehydrogenase was employed as a marker enzyme for mitochondria. Transformants and wild type strains were cultivated on YPD medium, and whole yeast cells were fractionated into mitochondria and cytosolic fractions. The specific enzyme activities in each fraction for each strain are listed in Table 3. In all strains which contain intact UGA2 gene, over 99% activity of the SSADH activity was found in cytosol fraction with specific activities of 2.22 to 2.52 U/mg protein. SSADH activity was almost non-detectable in the mitochondrial fraction suggesting that yeast UGA2p is localized in cytosol and not in mitochondria. In wild type yeast cells, significant GABA-TKG activity (>99%, 1.63 U/mg protein) was found in the cytosol. GABA-TKG enzyme activity was almost non detectable in the mitochondrial fraction. In the ?uga1 deletion mutant, no GABA-TKG activity was found 120 in either fraction supporting the complete loss of GABA-TKG activity with the absence of a functional UGA1 gene. In the ?uga1 mutant overexpressing full length ScGABA- TKG (?uga1+Y) and overexpressing the truncated AtGABA-TP (?uga1+P-mTP), GABA-TKG and GABA-TP specific activity was increased 6-fold and 0.5-fold relative to wild type respectively suggesting both constructs play a functional role in yeast and the truncated AtGABA-TP is not as highly expressed as the ScGABA-TKG in wild type. Similar to the wild type, no GABA-T activity was found in the mitochondria for both ?uga1+Y and ?uga1+P-mTP. Over 90% of GABA-TP activity was found in cytosol for ?uga1+P-mTP indicating that functional GABA transaminases without mitochondrial transit peptides are located in the cytosol. We then tested whether the plant mitochondrial transit peptide will cause yeast and plant GABA transaminases to localize into yeast mitochondria. Over 90% of the GABA- TKG and GABA-TP activity was found in mitochondria for both ?uga1+Y+mTP and ?uga1+P transformants respectively. For example, ?uga1+Y+mTP had a specific enzyme activity of 9.4 and 0.15 U/mg protein in mitochondria and cytosol respectively and ?uga1+P had a specific enzyme activity of 0.92 and 0.08 U/mg protein in mitochondria and cytosol respectively. These data suggest that both yeast endogenous GABA-TKG and AtGABA-TP were efficiently imported into yeast mitochondria directed by the presence of the AtGABA-TP mitochondrial targeting peptide sequence. Complementation of the GABA growth phenotype by cytosolic and mitochondrial GABA transaminases Previous studies have demonstrated that Saccharomyces mutants lacking UGA1, 121 UGA2, and UGA3 are incapable of growth on minimal medium with GABA as the sole nitrogen source but grow normally on minimal medium with ammonium sulfate as the sole nitrogen source (Ramos et al., 1985; Coleman et al., 2001). To test the extent of interchangeable function of Arabidopsis thaliana AtGABA-TP and yeast ScGABA-TKG in both the cytosol and mitochondria, mutant strains bearing ?uga1, ?uga2, or the ?uga1?uga2 double mutant were transformed by the set of plasmid constructs containing AtGABA-TP with and without the mitochondrial targeting peptide or containing ScGABA-TKG or ScGABA-TKG with the AtGABA-TP mitochondrial targeting peptide. Cells transformed with ScGABA-TKG (?uga1+Y, ?uga2+Y, ?uga1?uga2+Y) were grown on YNB medium containing 10 mM GABA or NH4+ as the sole nitrogen source (Fig 1B). Growth was fully rescued to at least wild type level for ?uga1+Y while growth was restored to the same extent as ?uga2 for the ?uga1?uga2+Y transformants, and as expected, overexpressing ScGABA-TKG did not recover the growth defect for the ?uga2 mutant. Similar growth recovery effects were also observed for cells transformed with plasmids overexpressing ScGABA-TKG with the mitochondrial targeting peptide from AtGABA-TP (?uga1+Y+mTP, ?uga2+Y+mTP, ?uga1?uga2+Y+mTP), indicating that overexpressing ScGABA-TKG with an Arabidopsis mitochondrial targeting peptide is functionally similar to overexpressing ScGABA-TKG alone in recovering the GABA growth defect in the mutants lacking UGA1. Similarly, cells transformed with full length Arabidopsis AtGABA-TP (?uga1+P, ?uga2+P, ?uga1?uga2+P) and AtGABA-TP with a truncated mitochondrial transit peptide (?uga1+P-mTP, ?uga2+P-mTP, ?uga1?uga2+P-mTP) were examined for GABA growth complementation. Transformation with AtGABA-TP (?uga1+P and 122 ?uga1?uga2+P) which putatively localized in mitochondria partially rescued the growth defect for mutants lacking UGA1 (Fig. 1B). A 3- to 4- fold increase in growth was observed in liquid media (data not shown). However, no significant growth recovery was found for the ?uga2 mutant when transformed with the AtGABA-TP constructs (Fig. 1B). Equivalent growth recovery was observed whether the transformants were overexpressing the full length AtGABA-TP or the AtGABA-TP with the truncated mitochondrial targeting peptide. These results indicate that AtGABA-TP targeted to either the mitochondria or cytosol in yeast has a similar function to yeast endogenous ScGABA-TKG in supporting growth on media containing GABA as the sole nitrogen source although the use of endogenous, ?-ketoglutarate-utilizing ScGABA-TKG has a stronger growth supporting effect than does the pyruvate-utilizing AtGABA-TP. Mitochondrial versus cytosolic localization for both enzymes appears to be less critical than either the use of ?-ketoglutarate versus pyruvate as substrate or the expression level of the specific enzyme in yeast cells. Complementation of the thermosensitivity and ROS production phenotype by cytosolic and mitochondrial forms of GABA transaminases Our previous studies showed that under lethal heat at 45?C, yeast mutants of ?uga1 lacking GABA transaminase and ?uga2 lacking succinate semialdehyde dehydrogenase are more heat sensitive than wild type. and ?uga1 is dramatically more heat sensitive than ?uga2. The ?uga1?uga2 double mutant is more heat sensitive than either of the single mutants. The thermo sensitivities were correlated to the level of ROS production (Chapter II). To test the function and localization of AtGABA-TP and ScGABA-TKG in yeast in thermotolerance and limiting ROS production, the three mutants above which are 123 defective in GABA degradation were transformed with the four plasmid constructs as described and characterized above. Overexpression of ScGABA-TKG in ?uga1 (?uga1+Y) led to heat stress survival comparable to the wild type after 60 min (Fig 2A). Comparably, overexpression of ScGABA-TKG in the double mutant ?uga1?uga2 (?uga1?uga2+Y) led to heat stress survival comparable to the ?uga2 mutant alone after 60 min. However, overexpression of ScGABA-TKG in the ?uga2 mutant (?uga2+Y) did not enhance heat stress cell survival. In all cases, ROS production was comparable to that expected from the survival data (Fig 2A). Overexpression of ScGABA-TKG with the AtGABA-TP mitochondrial transit peptide attached also produced comparable results to the expression of ScGABA-TKG in the cytosol (Fig. 2B compared to Fig 2A and Fig 3B compared to Fig. 3A). To assay the complementation by AtGABA-TP yeast mutants were transformed with plasmid constructs containing either full length AtGABA-TP (?uga1+P, ?uga2+P, and ?uga1?uga2+P) or the truncated GABA-TP lacking a mitochondrial targeting peptide (?uga1+P-mtp, ?uga2+P-mtp, and ?uga1?uga2+P-mtp), these strains were tested for their ability to compensate for the thermosensitivity (Fig. 4) and ROS production (Fig. 5) phenotypes. When ?uga1+P was heat stressed for up to 60 min at 45?C, cell survival for ?uga1+P significantly improved (19% to 50%, survival) but did not achieve wild type levels (71%) of survival (Fig. 4A). Clearly AtGABA-TP did not complement the yeast ?uga1 deletion to the same extent as did ScGABA-TKG or ScGABA-TKG plus the mitochondrial targeting peptide. Similar partial levels of complementation were observed for ?uga1?uga2+P (Fig 4A) and ?uga1?uga2+P-mtp (Fig 4B) since the levels of complementation did not reach the survival level of ?uga2. No significant cell survival 124 recovery was observed for the ?uga2 mutant transformants (?uga2+P or ?uga2+P-mtp). Additionally, the production of ROS (Fig 5) in these mutants and transformants was appropriately proportional to the heat stress survival data as was the case for the ScGABA-TKG transformations above. Taken together these results show that AtGABA-TP with or without a mitochondrial targeting peptide compensated for ScGABA-TKG under lethal heat stress for both survival and ROS accumulation. However, AtGABA-TP, regardless of its intracellular location was less effective than the native ScGABA-TKG form of GABA transaminase (regardless of its location). Discussion In this study, the cellular localization of yeast ScGABA-TKG and ScSSADH in wild type yeast strain were investigated based on the location of enzyme activity in the mitochondrial or cytosol subfractions. For both proteins, over 90% of specific enzyme activities were detected in the cytosol fraction (Table 3), the mitochondrial fractions possessed almost non measurable enzyme activities (<0.01 U/mg protein), suggesting both GABA-TKG and SSADH are localized and functioning in cytosol, not mitochondria. This finding is consistent with the localization predicted by TargetP and PSORT, and the cytosolic location of SSADH previously reported (Huh et al., 2003). Deletion of GABA-TKG in yeast resulted in completely loss of enzyme activities in cytosol. TargetP and PSORT predict that AtGABA-TP is localized in the mitochondria in Arabidopsis and would also be in yeast based on the presence of an 18 amino acid mitochondrial targeting peptide at the amino-terminus of the protein. Using both native 125 and recombinant DNA constructs and transformation of appropriate yeast gene deletions we have shown that the mitochondrial localization of both ScGABA-TGK and AtGABA- TP is mediated by this 18 amino acid mitochondrial targeting peptide. Since all of our plasmid constructs placed the cloned sequences under the control of the strong constitutive GPD promoter it is unclear why AtGABA-TP activity levels were reduced when compared to even wild type expression of ScGABA-TKG. The addition of the mitochondrial targeting peptide did not appear to reduce the stability of ScGABA-TGK, nor did the deletion of the mitochondrial targeting peptide from AtGABA-TP appreciably alter specific activity relative to the full length construct. Thus it seems most likely that the lower expression of AtGABA-TP is the result of either posttranscriptional processing or the posttranslational stability of the Arabidopsis protein in yeast. In complementation assays, transformants targeting ScGABA-TKG to mitochondria were found to be as efficient as the endogenous or ectopically expressed cytosolic form of ScGABA-TKG in supporting growth on GABA medium (Fig. 1A ), thermotolerance (Fig 2), or ROS accumulation (Fig 3). Thus, the production of succinate semialdehye from GABA which is normally a cytosolic process in Saccharomyces appears to be equally functional when it occurs in mitochondria. Thus, it can be concluded that the production of GABA and/or succinate semialdehyde in the mitochondria is no more responsible for heat stress tolerance or ROS accumulation than the production of these compounds in the cytosol. Complementation of the ?uga1 mutant by AtGABA-TP with or without its mitochondrial targeting peptide was found to be partially effective in supporting growth on GABA as the sole nitrogen source (Fig 1), heat stress tolerance (Fig 4), and ROS 126 accumulation (Fig. 5) supporting again the idea that localization in the mitochondria was not critical nor detrimental to the function of this pathway in Saccharomyces. This finding was not consistent with our hypothesis that the conversion of pyruvate to alanine by AtGABA-TP ectopically expressed in mitochondria should limit carbon flux through the TCA cycle, and thus reduce mitochondrial ROS production. Rather this finding is consistent with the hypothesis that carbon flux through the TCA cycle at least from pyruvate to ?-ketoglutarate is not related to the production of ROS during heat stress, and that the accumulation of succinate semialdehyde or some metabolite downstream is likely the major factor playing a role in abating ROS production during heat stress. The fact that AtGABA-TP only partially complements the ?uga1 mutation while ScGABA-TKG fully complements the mutant further supports the above hypothesis. However, it is unclear whether this is from the difference in substrate specificity of the two enzymes or from the difference in enzyme activity content in cells transformed with the two different constructs (Table 3). References 127 1. Barbosa, J. M.. 2002. Physiological biochemical and molecular aspects of ?- aminobutyric acid (GABA): a stress-responsive non-protein amino acid. Dissertation. Auburn university. 2. Bergmeyer, H. U.. M. Grassl, and H. E.Walter. 1983. In Methods of Enzymatic Analysis, 3rd ed., Volume II, pp 222-223, Verlag Chemie, Deerfield Beach, FL. 3. Bouch?, N., A. Fait, S.G. Moller and H. Fromm. 2003. Mitochondrial succinic- semialdehyde dehydrogenase of the ?-aminobutyrate shunt is required to restrict levels of reactive oxygen intermediates in plants, Proc. Natl. Acad. Sci. USA 100: 6843?6848. 4. Bradford, M. M.. 1976. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding, Analytical Biochemistry 72: 248-254. 5. Coleman, S.T., T. K. Fang, S. A. Rovinsky, F. J. Turano, and W. S. Moye- Rowley. 2001. Expression of a glutamate decarboxylase homologue is required for normal oxidative stress tolerance in Saccharomyces cerevisiae. J Biol Chem. 276: 244-50. 6. Daum, G. P., C. Bohni and G. Schatz. 1982. Import of proteins into mitochondria. Cytochrome b2 and cytochrome c peroxidase are located in the intermembrane space of yeast mitochondria. J. Biol. Chem. 257: 13028-13033. 7. Emanuelsson, O., H. Nielsen, S. Brunak, and G. von Heijne. 2000. Predicting subcellular localization of proteins based on their N-terminal amino acid sequence, J.Mol. Biol., 300: 1005-1016. 8. Geitz, R. D., and R. A. Woods. 2002. Transformation of yeast lithium acetate/single-stranded carrier DNA / polyethylene glycol method. Methods Enzymol. 350: 87-96. 9. Gropman, A.. 2003. Vigabatrin and newer interventions in succinic semialdehyde dehydrogenase deficiency, Ann. Neurol. 54: S66?S72. 10. Hogema, B. M., M. Gupta, H. Senephansiri, T.G. Burlingame, M. Taylor, C. Jakobs, R.B. Schutgens, W. Froestl, O.C. Snead, R. Diaz-Arrastia, T. Bottiglieri, M. Grompe and K.M. Gibson. 2001. Pharmacologic rescue of lethal seizures in mice deficient in succinate semialdehyde dehydrogenase, Nat. Genet. 29: 212?216. 11. Huh, W. K.. J. V. Falvo, L. C. Gerke, A. S. Carroll, R. W. Howson, J. S.Weissman, E. K. O'Shea. 2003. Global analysis of protein localization in budding yeast. Nature. 425: 686-691. 128 12. Hulme, A. C., and W. Arthington. 1950. ?-Aminobutyric acid and ?-alanine in plant tissues. Nature 165 : 716?717. 13. Illingworth, J. A.. 1972. Purification of Yeast Isocitrate Dehydrogenase Biochem. J. 129: 119-1124. 14. Kumar, S., and N. S. Punekar. 1997. The metabolism of 4-aminobutyrate (GABA) in fungi. Mycol. Res. 101: 403?409. 15. Nakai, K., and P. Horton. 1999. PSORT: a program for detecting the sorting signals of proteins and predicting their subcellular localization, Trends Biochem. Sci, 24: 34-35. 16. Nielsen, H., J. Engelbrecht, S. Brunak, and G. von Heijne. 1997. Identification of prokaryotic and eukaryotic signal peptides and prediction of their cleavage sites, Prot. Eng.. 10: 1-6. 17. Ramos, F., M. E. Guezzar, M. Grenson, and J. M. Wiame. 1985. Mutations affecting the enzymes involved in the utilization of 4-aminobutyric acid as nitrogen source by the yeast Saccharomyces cerevisiae. Eur. J. Biochem, 149: 401-404. 18. Sambrook, J., and David W. Russel. Molecular cloning. 2001. Cold Spring Harbor, N.Y.: Cold Spring Harbor Laboratory. 19. Shelp, B. J., A. W. Bown and M. D. McLean. 1999. Metabolism and functions of gamma-aminobutyric acid. Trends Plant Sci. 4: 446?452. 20. Sherman, F. G.R. Fink, and C.W. Lawrence. 1979. Methods in Yeast Genetics. Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y. Table 1. Yeast strains used in this study 129 Strain Genotype W303-1A Mat a leu2-3,112 ura3-1 trp1-1 his3-11,15 ade2-1, lys2, met ?uga1 Mat a leu2-3112 ura3-1 trp1-1 his3-11,15 ade2-1, lys2, met uga1::HIS3 ?uga2 Mat a leu2-3112 ura3-1 trp1-1 his3-11,15 ade2-1, lys2, met uga2::URA3 ?uga1?uga2 Mat a leu2-3112 ura3-1 trp1-1 his3-11,15 ade2-1, lys2, met uga1::HIS3 uga2::URA3 Table 2. Primers used for yeast cytolic and mitochondrial forms of GABA-TKG and plant GABA-TP expression Primer Name Sequence (5? to 3?) UGA1FOR GAGAGGATCCATGTCTATTTGTGAACAATACTACCCA mTP15 UGA1FOR ACTCAGGTTCATTTGATGTCTATTTGTGAACAATACTAC CCA mTP54 UGA1FOR GGATCCATGGTCGTTATCAACAGTCTCCGACGCTTGGC GCGTACCACTCAGGTTCATTTG UGA1REV GTGTAAGCTTTCATAATTCATTAACTGATTTGGCTAA FL-pGATFOR GAGAGGATCCATGGTCGTTATCAACAGTCTCCG FL-51-pGATFOR GAGAGGATCCATGCACAGTAGGTATGCCACTT pGATREV GTGTAAGCTTTCACTTCTTGTGCTGAGCCTT Table 3. Enzyme activities in yeast wild type, ?uga1 deletion mutant strain and ?uga1 mutant transformantsa 130 Specific activityb (U/mg protein) Preparation Protein wild type ?uga1 ?uga1+Y ?uga1+Y+MTP ?uga1+P ?uga1+P-MTP NAD-IDH 0.03 0.028 0.032 0.029 0.030 0.033 Hexokinase 3.1 2.5 3.0 3.08 3.1 3.2 GABA-T(P) 0.16 0 0.80 0.88 0.10 0.09 Total cell extract SSADH 0.29 0.27 0.238 0.24 0.26 0.26 Mitochondria NAD-IDH 0.19 0.185 0.20 0.0188 0.0195 0.21 Hexokinase 0 0 0 0 0 0 GABA-T(P) <0.01 0 0 9.4 1.52 0.06 SSADH <0.01 <0.01 <0.01 <0.01 <0.01 <0.01 Cytosol NAD-IDH 0 0 0 0 0 0 Hexokinase 25 23 28 27.5 28 26 GABA-T(P) 1.63 0 9 0.15 0.08 1.48 SSADH 2.52 2.48 2.22 2.35 2.42 2.45 a Yeast strains were grown on rich YPD medium. Cell fractionation was performed as described by Daum et al (Daum, 1982). b The enzymes assayed as described in Materials and Methods, were NAD+ dependent isocitrate dehydrogenase (NAD-IDH) as mitochondrial marker protein, hexokinase as cytosolic marker protein, yeast endogenous GABA-TKG or plant GABA-TP, succinate semialdehyde dehydrogenase (SSADH). The strains used were W303-1A (wild type), yeast deletion mutant ?uga1, and the mutant overexpressing yeast endogenous GABA- TKG (?uga1+Y), GABA-TKG with plant mitochondrial targeting peptide sequences (?uga1+Y+MTP), plant GABA-TP ( ?uga1+P), or GABA-TP without mitochondrial targeting peptide sequences (?uga1+P-MTP). The values shown represent averages of three independent determinations. 131 WT ?uga1 ?uga1+Y ?uga1+Y+mTP ?uga1+P ?uga1+P-mTP ?uga2 ?uga2+Y ?uga2+Y+mTP ?uga1?uga2 ?uga1?uga2+Y ?uga1?uga2+Y +mTP YNB+10mM GABA YNB+10mM NH4+ WT ?uga1 ?uga2 ?uga1 ?uga2 A B Fig. 1. (A) Growth of wild type and mutant strains on YNB medium with NH4+ as the sole nitrogen source; (B) YNB medium with GBAB as the sole nitrogen source. Each strain contains vectors of HIS( p423 GPD), URA(p426 GPD), LEU(p425 GPD), TRP(p424 GPD) and LEU(p425 GPD), LEU(p425 GPD) vector for mutant strains carries the cytolic and mitochondrial forms of yeast endogenous GABA-TKG or cytosolic and mitochondrial forms of plant GABA-TP. All strains are able to grow on YNB media with NH4+ as the sole nitrogen sources, however, ?uga1, ?uga2 and ?uga1uga2 can't grow on GABA as the sole nitrogen source as well as the wild type. Overexpression of plasmids bearing cytosolic and mitochondrial forms of yeast endogenous GABA-TKG fully recovered the GABA growth phenotype (B, left panel); Overexpression of plasmids bearing cytosolic and mitochondrial forms of plant GABA-TP partially recovered the GABA growth phenotype (B, right panel). "+Y" indicates expression of P425 GPD-UGA1, "+Y+mTP" indicates expression of P425 GPD-mTP54UGA1, "+P" indicates expression of P425 GPD-pGAT; "+P-mTP" indicates expression of P425 GPD-pGAT-mTP. 132 Time at 45oC (min) 0 10 20 30 40 50 60 Ce ll s uv iva l (% ) 0 20 40 60 80 100 W303-1A (WT) ?uga1 ?uga1+Y+mTP ?uga2 ?uga2+Y+mTP ?uga1?uga2+Y+mTP?uga1 ?uga2 W303-1A (WT) ?uga1 ?uga1+Y ?uga2 ?uga2+Y ?uga1?uga2+Y?uga1 ?uga2 Time at 45oC (min) 0 10 20 30 40 50 60 Ce ll s ur viv al (% ) 0 20 40 60 80 100 A B Ce ll s uv iva l (% ) Ce ll s ur viv al (% ) Ce ll s uv iva l (% ) Ce ll s ur viv al (% ) Ce ll s ur viv al (% ) Fig. 2. Viability of wild-type, mutant strains and mutant strains transformed with (A) cytosolic and (B) mitochondrial forms of yeast endogenous GABA-TKG after lethal heat treatment at 45oC. Strains were grown to mid log phase (5 ? 106 cells/ml) in YPD, washed in 0.87% NaCl, and concentrated to 2 ? 108 cells/ml in fresh YPD. Aliquots of 100 ?l were heated at 45oC in a thermocycler for 0, 15, 30, 45 and 60 min. Viability was determined as described. Each point represents the mean ? SD of the results from at least three experiments, some error bars are smaller than the symbol sizes. "+Y" indicates expression of P425 GPD-UGA1, "+Y+mTP" indicates expression of P425 GPD- mTP54UGA1. 133 W303-1A (WT) ?uga1 ?uga1+Y+mTP ?uga2 ?uga2+Y+mTP ?uga1?uga2+Y+mTP?uga1 ?uga2 W303-1A (WT) ?uga1 ?uga1+Y ?uga2 ?uga2+Y ?uga1?uga2+Y?uga1 ?uga2 Time at 45oC (min) 0 10 20 30 40 50 60 He at ind uc ed flu or es ce nc e i nc re as e 0 200 400 600 Time at 45oC (min) 0 10 20 30 40 50 60 He at ind uc ed flu or es ce nc e i nc re as e 0 200 400 600 A B He at ind uc ed flu or es ce nc e i nc re as e He at ind uc ed flu or es ce nc e i nc re as e He at ind uc ed flu or es ce nc e i nc re as e He at ind uc ed flu or es ce nc e i nc re as e Fig. 3. Intracellular ROS levels induced by lethal heat stress at 45?C in wild type, mutant strains and mutant strains transformed with cytosolic (A) and mitochondrial (B) forms of yeast endogenous GABA-TKG. Fluorescence was measured at an excitation wavelength of 504nm and an emission wavelength of 524nm in crude cell extracts as described. As a control, fluorescence was recorded during heat stress without cells and cells without DCFH-DA (date not shown). Data shown represent the mean ? SD of the results from at least three experiments, some error bars are smaller than the symbol sizes. "+Y" indicates expression of P425 GPD-UGA1, "+Y+mTP" indicates expression of P425 GPD- mTP54UGA1. 134 Time at 45oC (min) 0 10 20 30 40 50 60 Ce ll s ur viv al (% ) 0 20 40 60 80 100 W303-1A (WT) ?uga1 ?uga1+P-mTP ?uga2 ?uga2+P-mTP ?uga1?uga2+P-mTP?uga1 ?uga2 Time at 45oC (min) 0 10 20 30 40 50 60 Ce ll s ur viv al (% ) 0 20 40 60 80 100 W303-1A (WT) ?uga1 ?uga1+P ?uga2 ?uga2+P ?uga1?uga2+P?uga1 ?uga2 A B Ce ll s ur viv al (% ) Ce ll s ur viv al (% ) Ce ll s ur viv al (% ) Ce ll s ur viv al (% ) Ce ll s ur viv al (% ) Ce ll s ur viv al (% ) Fig. 4. Viability of wild-type, mutant strains and mutant strains transformed with (A) cytosolic and (B) mitochondrial forms of plant GABA-TP after lethal heat treatment at 45oC. Strains were grown to mid log phase (5 ? 106 cells/ml) in YPD, washed in 0.87% NaCl, and concentrated to 2 ? 108 cells/ml in fresh YPD. Aliquots of 100 ?l were heated at 45oC in a thermocycler for 0, 15, 30, 45 and 60 min. Viability was determined as described. Each point represents the mean ? SD of the results from at least three experiments, some error bars are smaller than the symbol sizes. "+P" indicates expression of P425 GPD-pGAT; "+Y-mTP" indicates expression of P425 GPD-pGAT-mTP. 135 W303-1A (WT) ?uga1 ?uga1+P ?uga2 ?uga2+P ?uga1?uga2+P?uga1 ?uga2 Time at 45oC (min) 0 10 20 30 40 50 60 He at ind uc ed flu or es ce nc e i nc re as e 0 200 400 600 W303-1A (WT) ?uga1 ?uga1+P-mTP ?uga2 ?uga2+P-mTP ?uga1?uga2+P-mTP?uga1 ?uga2 Time at 45oC (min) 0 10 20 30 40 50 60 He at ind uc ed flu or es ce nc e i nc re as e 0 200 400 600 A B He at ind uc ed flu or es ce nc e i nc re as e He at ind uc ed flu or es ce nc e i nc re as e He at ind uc ed flu or es ce nc e i nc re as e He at ind uc ed flu or es ce nc e i nc re as e He at ind uc ed flu or es ce nc e i nc re as e He at ind uc ed flu or es ce nc e i nc re as e Fig. 5. Intracellular ROS levels induced by lethal heat stress at 45?C in wild type, mutant strains and mutant strains transformed with (A) cytosolic and (B) mitochondrial forms of plant GABA-TP. Fluorescence was measured at an excitation wavelength of 504nm and an emission wavelength of 524nm in crude cell extracts as described. As a control, fluorescence was recorded during heat stress without cells and cells without DCFH-DA (date not shown). Data shown represent the mean ? SD of the results from at least three experiments, some error bars are smaller than the symbol sizes. "+P" indicates expression of P425 GPD-pGAT; "+P-mTP" indicates expression of P425 GPD-pGAT-mTP. 136 V. MOLECULAR CLONING, EXPRESSION AND CHARACTERIZATION OF RECOMBINANT YEAST SUCCINIC SEMIALDEHYDE DEHYDROGENASE Abstract The yeast succinic semialdehyde dehydrogenase gene (SSADH; EC 1.2.1.16) was cloned and overexpressed in E. coli. Based on SDS-PAGE, the molecular mass of the subunit was around 54 kDa, and the purified recombinant enzyme has a tetrameric molecular mass of approximately 200 kDa. The specific activity of the recombinant enzyme was 1.90 ?mol NADH formed min-1 mg-1, and showed maximal activity at pH 8.4. The recombinant protein was highly specific for succinate semialdehyde (Km = 15.48 ? 0.14 ?M, can use both NAD+ and NADP+ as a cofactor with Km values of 579.06 ? 30.1 ?M and (1.017 ? 0.46) mM respectively. Initial velocity studies show NADH was a competitive inhibitor with respect to NAD+ (Ki = 129.5 ?M ), but non-competitive inhibitor with respect to succinate semialdehyde. Adenine nucleotides of AMP, ADP and ATP inhibited the yeast SSADH activity with Ki of 1.13 ~ 10.2 mM and showed competitive inhibition with respect to NAD+ and mixed-competitive and non-competitive, and non-competitive inhibition respectively with respect to succinate semialdehyde. The kinetic data suggest a ping-pong mechanism. 137 Introduction ?-Aminobutyrate (GABA) is a ubiquitous non protein amino acid widely found in prokaryotic and eukaryotic organisms. It is well known as a major inhibitory neurotransmitter in mammals. GABA plays a role in the nitrogen and carbon metabolism in bacteria. In plants, GABA accumulates rapidly in response to various abiotic and biotic stresses such as hypoxia, cold, and mechanical stimulation. However, in most organisms, its physiological role is uncertain. In yeast, GABA is produced from glutamate in the cytosol, catalyzed by glutamate decarboxylase (GAD; EC 4.1.1.15 ) in the GABA shunt pathway. GABA is then further transaminated to succinate semialdehyde (SSA) by GABA transaminase (GABA-T; EC 2.6.1.19), and finally, SSA is oxidized by succinate semialdehyde dehydrogenase (SSADH; EC 1.2.1.16) to the final product of succinate which then enters the tricarboxylic acid cycle. Clearly, SSADH is involved in the utilization of GABA as a nitrogen source. In human, SSADH deficiency results in 4-hydroxybutyric aciduria, an autosomal recessive disorder due to an accumulation of GABA and 4-hydroxybutyric acid in the central nervous systems (Jakobs et al., 1981). Recently, mutation studies show SSADH in plants (Jakobs et al., 2003) and yeast (Coleman et al., 2001) is critical for normal oxidative stress tolerance. SSADH has been shown to be present and has been purified from mammalian tissues (Blaner and Churchich, 1979; Ryzlak and Pietruszko, 1988; Chambliss and Gibson, 1992; Lee et al., 1995), plants (Busch and Fromm, 1999; Satya and Nair, 1989; Yamaura et al., 1988) and microorganisms (Steinbuchel and Lutke-Eversloh, 1999; Hidalgo et al., 1991; Ramos et al., 1985). The reaction catalyzed by SSADH is 138 essentially irreversible, and the enzyme activity is inhibited by the substrate succinate semialdehyde (SSA) and the product NADH in mammalian brain tissues (Ryzlak and Pietruszko, 1988; Chambliss and Gibson, 1992; Lee et al., 1995). In plants, SSADH activity is also inhibited by adenine nucleotides (Busch and Fromm, 1999). In fungi, the first enzyme of the GABA shunt (GAD) was purified and best studied in N. crassa (Christensen and Schmit, 1980), GABA-T was purified and characterized from Candida species (Der et al., 1986). However, the last enzyme of pathway (SSADH) has received relatively little attention. Its properties could not be determined in crude cell extracts (Baldy, 1977; Vissers et al., 1989), and it has only been partially purified in yeast and not well characterized (Ramos et al., 1985). We reasoned that cloning of the yeast gene encoding SSADH enzyme and detailed characterization of the purified recombinant protein would provide significant information on the physiological and functional properties of SSADH, and its specific regulation in the GABA shunt pathway. Here, we describe the cloning and purification of a yeast SSADH and detailed kinetic studies of the purified recombinant enzyme. Materials and methods Cloning of yeast SSADH gene The Saccharomyces Genome Database (SGD) showed a reference sequence (GenBank Accession No. NC_001134) encoding a full-length ORF sequence for yeast SSADH. This sequence was used to design genomic PCR primers for the amplification of a DNA fragment of the SSADH gene containing this ORF. To facilitate expression vector construction, two restriction sites of XhoI and BamHI were introduced at each end 139 of primers. The sense primer for PCR amplification contained an XhoI site: 5'-GAGAC TCGAGATGACTTTGAGTAAGTATTCTAAACCAACTC-3?, and the antisense primer contained a BamHI site: 5?-GAGAGGATCCTTAAATGCTGTTTGGCAAATTCC-3?. Yeast genomic DNA was prepared as described by Sherman et al.(1989). PCR was carried out using a PCR Thermal Cycler PTC-1000TM for 30 cycles in a 25 ?l reaction mixture containing 10 pmol of primers, 1 unit of High Fidelity Taq DNA polymerase (Roche Applied Sciences), and 100 ng of yeast genomic DNA under the following conditions: denaturation at 95oC for 10 s, annealing at 50oC for 30 s and extension at 72oC for 2 min, each for 30 cycles. The PCR product was purified on 0.8% agarose gel, cloned into the p-GEM?-T Easy vector (Promega, Madison, WI) and sequenced. Expression and purification of His6-tagged yeast SSADH A yeast SSADH expression clone was constructed by inserting the corresponding open reading frame after amplification by PCR into the bacterial expression vector pET- 16b (Novagen, Madison, WI). The amplified PCR product and the pET-16b expression vector were digested with XhoI and BamHI, and the SSADH ORF was ligated into the vector using T4 DNA ligase (Promega) according to manufacturer's procedure. Expression of the recombinant protein was initiated by adding 0.5 mM IPTG to the cultures of transformed BL21 cells in LB medium containing 50 mg/L ampicillin. After the addition of IPTG, the culture was further grown for 6 h to induce the expression of His6-tagged SSADH protein. Cells were harvested and resuspended in Binding Buffer (pH8.0, 50 mM sodium phosphate containing 300 mM NaCl, 10 mM 2-mercaptoethanol, and 10 mM imidazole and 200 ?g/ml lysozyme). Final lysis was achieved by incubation 140 the cell suspension with lysozyme for 15 min at 30oC. The lysate was cleared by centrifugation at 12,000 rpm in a SS34 rotor for 20 min. The supernatant was loaded onto a nickel-nitrilo triacetic acid (Ni-NTA) agarose column (Qiagen). The column was washed three times with Washing Buffer (pH 8.0, 50 mM sodium phosphate containing 10% (v/v) glycerol, 300 mM NaCl, 10 mM 2-mercaptoethanol, and 50 mM imidazole). The enzyme was then eluted with 250 mM imidazole. The purity of the eluted protein was evaluated by 12% SDS-PAGE using Coomassie blue staining to visualize the protein. Enzyme Activity Assay Succinate semialdehyde dehydrogenase activity was assayed in the direction of aldehyde oxidation by measuring the reduction of NAD+ or NADP+ spectrophotometrically at 340 nm (Beckman, model DU 640) in a 600 ?l reaction volume at 25?C. The standard enzyme assay contained 100 ?M potassium pyrophosphate buffer (PH 8.4), 50 ?M SSA, and 714 ?M NAD+. The reaction was started with the addition of succinate semialdehyde dehydrogenase (0.13 ?g). One unit of enzyme activity was defined as the amount of enzyme required to reduce 1 ?mol of NAD(P)+ per minute at 25?C and specific activity is expressed as units / mg protein. The kinetic parameters, Km and Kcat values were calculated from Lineweaver-Burk plots. Protein concentrations were determined by the method of Bradford (Bradford, 1976) using bovine serum albumin as a standard. The dependence of succinate semialdehyde dehydrogenase on pH was determined in 0.1 M potassium phosphate buffer (pH 6 to 9), 0.1 M potassium pyrophosphate buffer (pH 8.5 to 9.5) and 0.1 M glycine-NaOH buffer (pH 9 to 12). Kinetic studies were repeated at least three times with different preparations of the purified recombinant SSADH. 141 Zymogram Staining Analysis Purified recombinant protein was separated on 8% native PAGE at 4?C. The gel was then stained for SSADH activity by incubation at room temperature in a solution containing 100 mM potassium pyrophosphate, pH 8.4, 300 ?M SSA, 0.2 mg/ml nitroblue tetrazolium (NBT), 0.06 mM phenazine methosulfate (PMS), and 1.5 mM NAD(P)+ at room temperature for 15min. The excess stain was removed by washing the gels in ddH2O. SSADH activity appeared as a dark band. FPLC gel filtration analysis FPLC gel filtration analysis was performed on a Bio-Rad HPLC equipped with an OD280 monitor at a flow rate of 0.4 ml / min. A 0.5 x 30 cm column packed with Sephacryl S-300-HR (Sigma Aldrich) was used in the FPLC analysis. The mobile phase was 50 mM sodium phosphate buffer (pH 7.0) containing 150 mM sodium chloride (Jeong et al., 2005). A molecular mass standard curve was constructed by calibrating the column with protein standards from Sigma Aldrich (Blue dextran: 2,000 kDa, Thyroglobulin: 670 kDa, Apoferritin: 443 kDa, ?-Amylase: 200 kDa, Alcohol dehydrogenase: 150 kDa, Albumin: 66 kDa). The molecular mass of the native recombinant SSADH was estimated from the equation for the line of best fit (y=- 0.39x+7.79, where y is log10 molecular mass and x is the retention time [Ve / Vo]). Results Expression and purification of SSADH in E. coli The yeast SSADH gene was cloned from genomic DNA isolated from the W303-1A wild- type strain. The full sequence of the SSADH DNA encodes a 497-amino acid 142 protein with a molecular mass of 54,189 Da, and a computer calculation predicts the isoelectric point is 6.61. Expression of SSADH in bacterial strain BL21/pET-16b-SSADH was induced by 0.5 mM IPTG at 37?C. Most of the expressed protein was found in the soluble extract after cell lysis. A nickle metal-affinity resin column was used for the purification of a His-tagged yeast SSADH. The purity of the protein was estimated by SDS-PAGE (Fig. 1.). The His6-tagged recombinant SSADH yielded one single band with an apparent molecular mass of 54-kDa, and the overall yield of the enzyme in the purification procedure was 76% (Table 1). The purification resulted in approximately 10 mg of pure SSADH from 100 ml of cultured cells. The addition of 5% glycerol was essential, but 5 mM ?-mercaptoethanol was not required for maintaining the stability of the protein for a long time. Properties of yeast SSADH Gel filtration on a Sephacryl S-300-HR column (FPLC) was used to determine the native molecular weight of recombinant yeast SSADH. In three independent runs, the purified enzyme preparation eluted as a single peak with an elution volume of 20 ?l. A line of best fit equation (y=-0.39x+7.79) from the protein standards was used to calculate the native molecular weight. It was found to be 200 kDa (Fig. 2). These data suggest that the recombinant yeast SSADH is a homotetramer with four 54-kDa subunits. The effect of pH on the SSADH activity was determined with the purified enzyme, the optimum pH was 8.4 in 0.1 M potassium pyrophosphate buffer. 143 Lactate, butonal and a number of aldehydes including formaldehyde, glyceraldehyde, propanal, hexanal, valeraldehyde and acetalaldehyde were tested as substrates for the purified recombinant yeast succinate semialdehyde dehydrogenase. The purified enzyme was highly specific for SSA and had no activity for the other tested substrates. The recombinant yeast SSADH could utilize both NAD+ or NADP+ as a coenzyme. Howeverr, the enzyme had a 2.5-fold lower activity observed with NADP+ than with NAD+ at a concentration of 0.2 mM, which was in good agreement with the crude preparations and partially purified yeast SSADH (Ramos et al., 1985). Kinetic experiments Kinetic studies for characterizing the recombinant yeast SSADH were carried out in the direction of NAD(P)+ reduction. Initial velocity experiments were carried out by varying one substrate at several fixed substrate concentrations of the other substrate. Data from experiments with varied moderate concentrations of SSA at different fixed concentrations of NAD(P)+ resulted in intersecting lines as analyzed by double-reciprocal plots (Fig. 3). Further experiments with increasing the varied substrate SSA concentration (above 100 ?M) resulted in reciprocal plots which curve up as they approach the 1/v axis indicating substrate inhibition. In contrast, varied concentrations of NAD(P)+ at different fixed concentrations of SSA revealed a set of parallel lines (Fig. 4 and Fig. 5). No substrate inhibition was observed when NAD+ or NADP+ was used as substrate. Kinetic constants obtained from double-reciprocal plots are presented in table 2. The GABA shunt bypasses 2 reactions in the tricarboxylic acid cycle (?- ketoglutarate dehydrogenase and succinate thiokinase) by removing ?-ketoglutarate from 144 the cycle and feeding succinate and NADH produced by SSADH back into the cycle. An inhibitory effect of NADH on SSADH activity is widely known for several mammalian and plant SSADH enzymes (Blaner and Churchich, 1979; Duncan and Tipton, 1971; Rivett and Tipton, 1981; Busch and Fromm, 1999; Jeong et al., 2005). Similar inhibition of recombinant yeast SSADH with increasing NADH was observed here. SSADH activity was decreased significantly (Fig. 6 and Fig. 7). Further investigation revealed that NADH was a strong competitive inhibitor (Ki=129.5?M) with respect to NAD+, while it was found to be a non-competitive inhibitor (Ki=110.7 ?M) with respect to SSA at fixed concentrations of NAD+ (Table 3). Interestingly, succinate did not cause detectable inhibition at concentrations of up to 10 mM at pH 8.6 with respect to either NAD+ or SSA which was similar to the rat brain SSADH (Rivett and Tipton, 1981). This finding suggests that the concentrations of SSA, NAD+, and NADH in yeast cells might partly regulate the GABA shunt through SSADH activity, but not the product, succinate. The inhibitory effect of adenine nucleotides on the regulation of yeast SSADH was investigated. It has been reported that SSADH activity was inhibited by AMP (Rivett and Tipton, 1981; Busch and Fromm, 1999; Satya and Nair, 1989), ADP and ATP (Busch and Fromm, 1999), but no report for yeast SSADH. Similar to Arabidopsis SSADH, the activity of yeast SSADH was inhibited by all three adenine nucleotides. 5'-AMP was found to be a competitive inhibitor with respect to NAD+ with a Ki value of 4.18x103 ?? (Fig. 8) and a mixed inhibitor with respect to SSA (Fig. 9). Secondary intercept and/ or slope re-plots were linear indicating mixed inhibition. The inhibition by ADP was competitive with respect to NAD+ (Fig. 10) and non-competitive with respective to SSA (Fig. 11) and Ki values of 5.92 x 103 ?? and 7.57 x 103 ?? respectively were 145 determined. Since most adenine-dependent enzymes require Mg2+ bound to the nucleotides, we then tested the inhibition by ADP with the addition of Mg2+. Similar to Arabidopsis SSADH, in the presence or absence of 5 mM Mg2+, no difference was detected for the inhibition by ADP on the yeast SSADH activity suggesting that the binding of NAD+/ SSA was not hindered by ADP and vice versa. Inhibition by ATP was competitive with respect to NAD+ (Fig. 12) and non-competitive with respective to SSA (Fig. 13). The Ki value was 8.89 x 103 ?? for NAD+ and 10.21 x 103 ?? for SSA. In the presence or absence of 5 mM Mg2+, no difference was detected for the inhibition by ATP on the yeast SSADH activity again suggesting that the binding of NAD+ / SSA was not hindered by ATP and vice versa. Discussion Yeast SSADH plays a pivotal role in the metabolism of GABA, providing the mitochondria with succinate. Physiological and biochemical studies on the role of the GABA shunt have been hampered by the lack of detailed characterization of GABA shunt enzymes. In fungi, the first two enzymes, GAD1 and UGA1, have been purified and well characterized (Christensen and Schimt, 1980; Der Garabedian, 1986). However, limited information was reported on the last enzyme SSADH (Ramos et al., 1985). In this study, we report the biochemical characterization of the recombinant yeast SSADH. The enzyme is constituted of four subunits of similar size with an apparent relative molecular mass of 200 kDa (Fig. 2). This is comparable with several other SSADH enzymes from barley, Arabidopsis, and potato (Blaner and Churchich, 1979; Busch and Fromm, 1999; Satya and Nair, 1989) and mammalian species (Ryzlak and 146 Pietruszko, 1988; Chambliss and Gibson, 1992; Jeong et al., 2005). However, different results have been reported for other organisms, in E. coli SSADH was a dimeric protein (Doonelly and Cooper, 1981) while human brain SSADH was a heterotetramer composed of non-identical sized subunits (Ryzlak and Pietruszko, 1988). The results presented here suggest that yeast SSADH is a homotetramer. The optimum pH is similar to that reported from partially purified yeast SSADH (Ramos et al., 1985), human SSADH (Jeong et al., 2005), and Arabidopsis SSADH (Busch and Fromm, 1999). Like the partially purified yeast SSADH preparation (Ramos et al., 1985), the purified recombinant yeast SSADH is highly specific for SSA. None of the other aldehyde analogs (C3 to C6 straight chain aldehyde) gave any detectable activity as a substrate. The Km value obtained for SSA is 15.3 ?M. This is similar to, but slightly higher than the previously reported recombinant enzymes from human, Km = 6.3 ?M (Jeong et al., 2005) and rat, Km = 3.5 ?M (Murphy et al., 2003), and slightly lower than the recombinant Arabidopsis SSADH, K0.5 = 15 ? 5 (Busch and Fromm, 1999). Generally, it seems that eukaryotic SSADHs uses NAD+ as cofactor (Rivett and Tipton, 1981; Busch and Fromm, 1999), rather than NADP+ which is typically required by bacterial SSADHs. The recombinant yeast SSADH can utilize both NAD+ and NADP+ as a cofactor but has a much higher affinity for NAD+, which is in good agreement with the partially purified yeast SSADH (Ramos et al., 1985), and SSADH from rat (Cash et al., 1977) and human brains (Cash et al.,1978). However, the mechanism leading to the variance with these results between these organisms is unknown. In the direction of aldehyde oxidation, substrate inhibition was observed with SSA at concentrations above 100 ?M (Fig. 3). No substrate inhibition with NAD+ or NADP+ 147 was observed. Similarly, substrate inhibition with higher concentrations of SSA was also reported for the crude and partially purified yeast SSADH, the SSADHs from mammalian species, the SSADHs from plants, and bacterial SSADHs. SSADH product inhibition by NADH was reported from mammalian species (Blaner and Churchich, 1979; 1980; Rivett and Tipton, 1981; Jeong et al., 2005) and plants (Busch and Fromm, 1999). Here, similar result for a yeast SSADH is reported that it has a comparatively high affinity for NADH. NADH is a strong competitive inhibitor with respect to NAD+ (Ki = 129.5 ?M), but is a non-competitive inhibitor with respect to SSA (Ki = 110.7 ?M). However, similar to rat brain SSADH (Rivett and Tipton, 1981), no product inhibition by succinate could be detected for yeast SSADH, which makes it difficult to accurately determine the inhibition by NADH at the SSA substrate concentrations at Km levels for NAD+. The high substrate inhibition is more likely caused by a compulsory-order mechanism in which NADH is the last product to be released from the enzyme and the inhibition is resulted from the formation of an abortive ternary complex between enzyme, NADH and SSA (Rivett and Tipton, 1981). Adenine nucleotide inhibition by AMP was reported for rat-brain SSADH (Rivett and Tipton, 1981) and potato SSADH (Satya and Nair, 1989). Arabidopsis SSADH was inhibited by all three adenine nucleotides (Busch and Fromm, 1999), and it was also reported that succinate dehydrogenase in mitochondria is regulated by ATP (Singer et al., 1973). Our kinetic studies show that yeast SSADH was inhibited by AMP, ADP and ATP similar to the plant enzyme. The inhibition pattern given by AMP, ADP and ATP is competitive with respect to NAD+, mixed-competitive and non-competitive, and non- competitive respectively with respect to SSA. We may reason that AMP inhibition is 148 regulated by an ordered or random mechanism through which AMP could bind to the free enzyme or bind to the enzyme-SSA complex or both in the latter case. For the type of inhibition by ADP and ATP, we assume that the inhibitor could only bind to the enzyme- NAD+ binary complex in a compulsory-order mechanism rather than in a random-order mechanism. The inhibition pattern given by AMP is comparable with rat-brain SSADH (Rivett and Tipton, 1981). However, the inhibition by ADP and ATP differs from that previously reported for Arabidopsis SSADH (Busch and Fromm, 1999) with respect to NAD+ which was mixed-competitive and non-competitive respectively. However, like Arabidopsis SSADH (Busch and Fromm, 1999), it was found that yeast SSADH was not dependent on the complex of the nucleotide with Mg2+. The inhibition constants of yeast SSADH for all the three adenine nucleotides were in the milimolar range. It was found in plants that ATP regulates some enzymes in the TCA cycle in the millimolar concentration range (Raymond et al., 1987). In addition, AMP and ATP inhibited Arabidopsis SSADH in the milimolar range (Busch and Fromm, 1999). In animals, the concentrations of total nucleotides in mitochondria are in the milimolar range (Pradet and Raymond, 1983; Hutson et al., 1989). In yeast, the levels of ATP determined by Theobald et al (1996) during steady state growth in both cytosol and mitochondrial were in milimolar range (2.1 ? 0.1 mM and 7.8 ? 0.3 mM respectively), and the levels of AMP and ADP were 0.11 ? 0.03 mM and 0.47 ? 0.05 mM respectively in cytosol and for ADP the level in mitochondria was 0.8 ? 0.1 mM. The levels of NAD+ in both cytosol and mitochondria were 1.07 mM and 1.73 mM, while the concentration of NADH was 0.04 mM in the cytosol and 6.28 mM in mitochondria. These values indicate 149 the possibility that adenine nucleotides and pyridine nucleotides inside yeast cells and other organisms are in the range to affect the SSADH activity. Kinetic experiments with recombinant yeast SSADH showed an interesting substrate saturation pattern in double reciprocal plots versus initial velocity. These experiments revealed intersecting lines with SSA as the varied substrate, while parallel lines were obtained with various concentrations of NAD(P)+. Two ping pong mechanisms previously reported (Brigitte and Gerhard, 1993; Cleland, 1979; Fromm, 1967) are in accord with our kinetics, in which, the binding of the substrate which gives parallel lines in double reciprocal plots is followed by the release of a product. The substrate may bind before the last product is released or bind followed by the first product released (Brigitte and Gerhard, 1993). In our kinetic data for yeast SSADH, varied NAD(P)+ gave parallel lines, we predict that binding of the NAD(P)+ is followed by the release of NAD(P)H at the beginning or in the end of the reaction. However, more investigation is needed to provide evidence for this proposed mechanism. 150 References 1. Baldy, P. 1977. Metabolisme du c-aminobutyrate chez Agaricus bisporus. III. La succinate-semialdehyde: NAD(P)+ oxydoreductase PHysiologia Plantarum. 40:91?97. 2. Blaner, W. S., and J. E. Churchich. 1979. Succinic semialdehyde dehydrogenase, reactivity of lysyl residues. J. Biol. Chem. 254:1794?1798. 3. Bradford, M. M. 1976. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Analytical Biochemistry 72:248-254. 4. Brigitte, S., and G. Gerhard. 1993. Purification and characterization of a coenzyme-A-dependent succinate-semialdehyde dehydrogenase from Clostridium kluyver. Eur. J. Biochem. 212:121-127. 5. Busch, K. B., and H. Fromm. 1999. Plant succinic semialdehyde dehydrogenase. Cloning, purification, localization in mitochondria, and regulation by adenine nucleotides. Plant PHysiol. 121(2):589-97. 6. Cash, C., L. Ciesielski, M. Maitre, and P. Mandel. 1977. Purification and properties of rat brain succinic semialdehyde dehydrogenase. Biochimie. 59(3): 257-68. 7. Cash, C, M. Maitre, and P. Mandel. 1978. Purification of 2 succinic semi- aldehyde reductases from the human brain. C R Acad Sci Hebd Seances Acad Sci D. 286 (24):1829-32. 8. Chambliss, K. L., and K. M. Gibson. 1992. Succinic semialdehyde dehydrogenase from mammalian brain: subunit analysis using polyclonal antiserum. Int. J. Biochem. 24:1493?1499. 9. Christensen, R. L., and J. C. Schmit. 1980. Regulation of glutamic acid decarboxylase during Neurospora crassa conidial germination. Journal of Bacteriology 144:983?990. 10. Cleland, W. W. 1979. Substrate inhibition. Methods Enzymol. 63:500-513. 11. Coleman, S. T., T. K. Fang, S. A. Rovinsky, F. J. Turano, and R. W. S. Moye. 2001. Expression of a glutamate decarboxylase homologue is required for normal oxidative stress tolerance in Saccharomyces cerevisiae. J Biol Chem 276(1):244- 50. 151 12. Der Garabedian, P. A. 1986. Candida d-aminovalerate: a-ketoglutarate aminotransferase: purification and enzymologic properties. Biochemistry. 25:5507?5512. 13. Doonelly, M. I., and R. A. Cooper. 1981. Succinic semialdehyde dehydrogenases of Escherichia coli: their role in the degradation of p- hydroxypHenylacetate and gamma-aminobutyrate. Eur. J. Biochem. 113:555?561 14. Duncan, R. J. S., and K. F. Tipton. 1971. The kinetics of pig brain aldehyde dehydrogenase. Eur. J. Biochem. 22:538?543. 15. Fromm, H. J. 1967. The use of competitive inhibitors in studying the mechanism of action of some enzyme systems utilizing three substrates. Biochim. BiopHys. Acta. 139:221 -230. 16. Hidalgo, E., J. Aguilar, Y.M. Chen, and E.C. Lin. 1991. Molecular cloning and DNA sequencing of the Escherichia coli K-12 ald gene encoding aldehyde dehydrogenase. J. Bacteriol. 173:6118-6123. 17. Hutson, S. M., D. Berkich, G. D. Williams, K. F. LaNoue, and R. W. Briggs. 1989. 31P-NMR visibility and characterization of rat liver mitochondrial matrix adenine nucleotides. Biochemistry. 28:4325?4332. 18. Jakobs, C., A. Fait, D. Bouchez, S. G., Moller, and H. Fromm. 2003. Mitochondrial succinic-semialdehyde dehydrogenase of the gamma- aminobutyrate shunt is required to restrict levels of reactive oxygen intermediates in plants. Proc Natl Acad Sci USA. 100 (11):6843-8. 19. Jakobs, C., M. Bojasch, E. Monch, D. Rating, H. Siemes, and F. Hanefeld. 1981. Urinary excretion of gamma-hydroxybutyric acid in a patient with neurological abnormalities.The probability of a new inborn error of metabolism. Clin. Chim. Acta 111:169?178. 20. Jeong, H. K., B. P. Yong, L. H. Tae, H. L. Won, S. C. Myung, and S. K. Oh. 2005. High-level expression and characterization of the recombinant enzyme, and tissue distribution of human succinic semialdehyde dehydrogenase. Protein Expression and Purification. 44(1):16-22. 21. Lee, B. R, J. W. Hong, B. K. Yoo, S. J. Lee, S. W. Cho, and S.Y. Choi. 1995. Bovine brain succinic semialdehyde dehydrogenase: purification, kinetics and reactivity of lysyl residues connected with catalytic activity. Mol. Cells 5:611? 617. 22. MurpHy, T. C., V. Amarnath, K. M. Gibsion, and M. J. Picklo. 2003. Oxidation of 4-hydroxy-2-nonenal by succinic semialdehyde dehydrogenase (ALDH5A). J. Neurochem. 86:298?305. 152 23. Pradet, A., and P. Raymond. 1983. Adenine nucleotide ratios and adenylate energy charge in energy metabolism. Annu Rev Plant PHysiol. 34:199?224. 24. Ramos, F., M. E. Guezzar, M. Grenson, and J. M. Wiame. 1985. Mutations affecting the enzymes involved in the utilization of 4-aminobutyric acid as nitrogen source by the yeast Saccharomyces cerevisiae. Eur. J. Biochem. 149:401- 404. 25. Raymond, P., X. Gidrol, C. Salon, and A. Pradet. 1987. Control involving adenine nucleotides and pyridine nucleotides. In: Stump PK, Conn EE., editors. The Biochemistry of Plants. New York: Academic Press. 11:129?176. 26. Rivett, A. J., and K. F. Tipton. 1981. Kinetics studies with rat-brain succinic semialdehyde dehydrogenase. Eur. J. Biochem. 117:187?193. 27. Ryzlak, M. T., and R. Pietruszko. 1988. Human brain ?high Km? aldehyde dehydorogenase: purification, characterization, and identification as NAD+- dependent succinic semialdehyde dehydrogenase. Arch. Biochem. BiopHys. 266:386?396. 28. Satya, N. V., and P. M. Nair. 1989. Potato tuber succinate dehydrogenase: purification and characterization. Arch Biochem BiopHys 275:469-477. 29. Sherman, F., G.R. Fink, and C.W. Lawrence. 1979. Methods in Yeast Genetics. Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y. 30. Singer, T. P., G. Oestreicher, P. Hogue, J. Contreiras, and I. Brandao. 1973. Regulation of succinate dehydrogenase in higher plants. Plant PHysiol. 52:616? 621. 31. Steinbuchel, A., and T. Lutke-Eversloh. 1999. Biochemical and molecular characterization of a succinate semialdehyde dehydrogenase involved in the catabolism of 4-hydroxybutyric acid in Ralstonia eutropHa. FEMS Microbiol. Lett. 181:63-71. 32. Theobald, U., W. Mailinger, M. Baltes, M. Rizzi, and M. Reuss. 1996. In Vivo Analysis of Metabolic Dynamics in Saccharomyces cerevisiae: I. Experimental Observations. Biotechnology and Bioengineering. 55(2):305-316. 33. Vissers, S., B. Andre, F. Muyldermans, and M. Grenson. 1989. Positive and negative regulatory elements control the expression of the UGA4 gene coding for the inducible 4-aminobutyric acid-speci?c permease in Saccharomyces cerevisiae. European Journal of Biochemistry 181:357?361. 34. Yamaura, I., T. Matsumoto, M. Funatsu, and T. Shinohara. 1988. Purification and some properties of succinic semialdehyde dehydrogenase from barley seeds. Agric Biol Chem 52:2929-2930. 153 Table 1. Single step purification of recombinant yeast SSADH Table 2. Kinetic parameters of recombinant yeast SSADH Purification step Total protein Total activity Specific activity Purification Yield mg units Units mg-1 fold % Crude extract Ni-NTA agarose 10 2 5 3.80 0.5 1.90 1 3.8 100 76 substrate Km Vmax kcat/Km ?M ?mol.min-1.mg-1 M-1s-1 SSA 15.48 ? 0.14 15.93 ? 0.55 7.28 x 105 ? 210 NAD+ 579.06 ? 30.1 2.38 x 104 ? 108 NADP+ (1.017 ? 0.46) x 103 1.15 x 104 ? 152 154 Table 3. Inhibition of recombinant yeast SSADH by the products and nucleotides aKi, competitive inhibition constant; bKui, non-competitive inhibition constant; cNo inhibition, no inhibition at succinate concentration up to 10 mM. Inhibitor Varied substrates Pattern Kia or Kuib (?M) NADH SSA Non-competitive 110.7 NADH NAD+ competitive 129.5 SUCC SSA No inhibitionc SUCC NAD+ No inhibitionc AMP SSA Mixed-competitive Ki = 1.13 x 103, Kui = 2.37 x 103 AMP NAD+ competitive 4.18 x 103 ADP SSA Non-competitive 7.57 x 103 ADP NAD+ Competitive 5.92 x 103 ATP SSA Non-competitive 10.21 x 103 ATP NAD+ Competitive 8.89 x 103 155 mature SSADH ~(54 kDa)66 45 36 29 24 20 kDa 1 2 3 4 5 Fig. 1. Expression and purification of recombinant yeast SSADH. 12% of SDS-PAGE analysis of crude cell extracts of BL21 transformed with expression vector pET16b containing the SSADH coding sequence. Lane 1, marker protein; Lanes 2, 3 and 4, cell free extracts (50 ?g) before induction, without and with the presence of 0.5 mM IPTG; Lane 5, purified His6-tagged recombinant enzyme (5 ?g). 156 Ve/Vo 0.9 1.0 1.1 1.2 1.3 1.4 1.5 1.6 1.7 log M .W . 4 5 6 Thyroglobulin Appoferrintin ?-Amylase Alcohol dehydrogenase BSA yeast SSADH Yeast SSADH activity log M .W . log M .W . log M .W . Fig. 2. Molecular Mass determination of native SSADH by FPLC gel filtration on Sephacryl S-300-HR. Sephacryl S-300-HR was used to determine the molecular mass of purified SSADH as described under "Materials and Methods". The molecular mass of the proteins used to calibrate the column are: Thyroglobulin (670 kDa), Appoferrintin (443 kDa), ?-Amylase (200 kDa), Alcohol dehydrogenase (150 kDa), BSA (66 kDa). The void volume (Vo) was determined with blue dextran (2000 kDa). The arrow points to the position of SSADH on the standard curve. Inset shows the zymogram staining of yeast SSADH activity as described under "Materials and Methods". 157 [1/SSA] (mM-1) -50 0 50 100 150 5 10 15 20 25 30 1/V (? mo l x m in- 1 ) 1/V (? mo l x m in- 1 ) Fig. 3. Double reciprocal plot from initial velocity experiments of yeast recombinant succinate semialdehyde dehydrogenase with varied concentrations of SSA. Enzyme activities were assayed in the direction of SSA oxidation as described in Materials and Methods. Substrate concentrations for SSA were varied over 0.0067-0.317 mM as indicated in the figure. Concentrations of NAD+ were 0.225 mM (?), 0.416 mM (?) and 0.712 mM (?). Note the decrease in velocity at the highest substrate levels indicating substrate inhibition. 158 1/[NAD+] (mM-1) -1 0 1 2 3 4 5 2 4 6 8 10 12 14 16 18 20 1/V (? mo l x m in- 1 ) 1/V (? mo l x m in- 1 ) Fig. 4. Double reciprocal plot from initial velocity experiments of yeast recombinant succinate semialdehyde dehydrogenase with varied concentrations of NAD+. Enzyme activities were assayed in the direction of SSA oxidation as described in Materials and Methods. Substrate concentrations for NAD+ were varied over 0.227-2.5 mM as indicated in the figure. Concentrations of SSA were 0.090 mM (?), 0.045 mM (?) and 0.030 mM (?). 159 [NADP+] (mM-1) -1 0 1 2 3 5 10 15 20 25 30 1/V (? mo l x m in- 1 ) 1/V (? mo l x m in- 1 ) Fig. 5. Double reciprocal plot from initial velocity experiments of yeast recombinant succinate semialdehyde dehydrogenase with varied concentrations of NADP+. Enzyme activities were assayed in the direction of SSA oxidation as described in Materials and Methods. Substrate concentrations for NADP+ were varied over 0.417-2.5 mM as indicated in the figure. Concentrations of SSA were 0.085 mM (?), 0.027 mM (?) and 0.015 mM (?). 160 1/ [NAD+] (mM-1) 0 1 2 3 4 5 0 5 10 15 20 25 1/V (? mo l x m in- 1/ mg pr ot ein ) [NADH] (mM) -0.1 0.0 0.1 0.2 0.3 0.4 0.5 Sl op e 1 2 3 4 5 1/V (? mo l x m in- 1/ mg pr ot ein ) 1/V (? mo l x m in- 1/ mg pr ot ein ) Sl op e Fig. 6. Inhibition of yeast recombinant SSADH by NADH. Double-reciprocal plots of the rate of NADH formation vs NAD+ concentration at different NADH concentrations: 0 ?M NADH (?), 0.1 mM NADH (?), 0.2 mM NADH (?), 0.35mM NADH ( ? ), 0.45 mM NADH (?). Assays were carried out as described in Fig. 2 in the presence of 85 ?M of SSA. The inset shows the secondary plot of the line slope vs NADH concentration to determine the inhibition constant. 161 1/[SSA] (mM-1) 0 20 40 60 80 0 10 20 30 40 50 60 70 [NADH] (mM) -0.1 0.0 0.1 0.2 0.3 0.4 0.5 Int er ce pt 2 4 6 8 10 12 1/V ( ? mo l x m in- 1/ mg pr ot ein ) Int er ce pt 1/V ( ? mo l x m in- 1/ mg pr ot ein ) Fig. 7. Inhibition of yeast recombinant SSADH by NADH. Double-reciprocal plots of the rate of NADH formation vs SSA concentration at different NADH concentrations: 0 ?M NADH (?), 100 ?M NADH (?), 200 ?M NADH (?), 350 ?M NADH (? ), 450 ?M NADH (?). Assays were carried out as described in Fig. 2 in the presence of 712 ?M of NAD. The inset shows the secondary plot of the line intercept vs NADH concentration to determine the inhibition constant. 162 1/[NAD+] (mM-1) -1 0 1 2 3 4 5 0 5 10 15 20 25 [AMP] (mM) -4 -2 0 2 4 6 8 10 Sl op e 1 2 3 4 5 1/V ( ? mo l x m in- 1 / m g p ro tei n) S lop e Sl op e 1/V ( ? mo l x m in- 1 / m g p ro tei n) Fig. 8. Inhibition of yeast recombinant SSADH by AMP. Double-reciprocal plots of the rate of NADH formation vs NAD+ concentration at different AMP concentrations: 0 mM AMP (?), 2mM AMP (?), 4 mM AMP (?), 6 mM AMP (? ), 7.5 mM AMP (?). Assays were carried out as described in Fig. 2 in the presence of 85 ?M of SSA. The inset shows the secondary plot of the line slopes vs AMP concentration to determine inhibition constant. 163 [AMP] (mM) -2 -1 0 1 2 3 4 5 Int er ce pt 1 2 3 4 5 6A [AMP] (mM-1) -2 -1 0 1 2 3 4 5 Sl op e 0.1 0.2 0.3 0.4 0.5 0.6B 1/[SSA] (mM-1) 0 20 40 60 80 0 10 20 30 40 50 1/V ( ? mo l x m in- 1/ mg pr ot ein ) Inte rc ep t Sl op e Int er ce pt Int er ce pt Sl op e Sl op e 1/V ( ? mo l x m in- 1/ mg pr ot ein ) 1/V ( ? mo l x m in- 1/ mg pr ot ein ) Fig. 9. Inhibition of yeast recombinant SSADH by AMP. Double-reciprocal plots of the rate of NADH formation vs SSA concentration at different AMP concentrations: 0 mM AMP (?), 1mM AMP (?), 2 mM AMP (?), 3 mM AMP ( ? ), 4 mM AMP (?). Assays were carried out as described in Fig. 2 in the presence of 712 ?M of NAD. The insets show the secondary plot of the line intercepts (1/Vmax) (A) or slope (B) vs AMP concentration to determine the inhibition constants. 164 1/[NAD+] (mM-1) 0 1 2 3 4 0 5 10 15 20 25 [ADP] (mM) -5 0 5 10 15 20 Sl ope 1 2 3 4 5 6 1/V ( ? mo l x m in- 1 / m g p ro tei n) Sl ope 1/V ( ? mo l x m in- 1 / m g p ro tei n) Fig. 10. Inhibition of yeast recombinant SSADH by ADP. Double-reciprocal plots of the rate of NADH formation vs NAD concentration at different ADP concentrations: 0 mM ADP (?), 4mM ADP (?), 8 mM ADP (?), 12 mM ADP (? ), 16 mM ADP (?). Assays were carried out as described in Fig. 2 in the presence of 85 ?M of SSA. The inset shows the secondary plot of the line slopes vs ADP concentration to determine the inhibition constant. 165 1/[SSA] (mM-1) 0 20 40 60 80 0 5 10 15 20 25 30 1/V ( ? mo l x m in- 1 / m g p ro tei n) [ADP] (mM) -5 0 5 10 15 20 Int er ce pt 1 2 3 4 5 6 7 1/V ( ? mo l x m in- 1 / m g p ro tei n) Int er ce pt Fig. 11. Inhibition of yeast recombinant SSADH by ADP. Double-reciprocal plots of the rate of NADH formation vs SSA concentration at different ADP concentrations: 0 mM ADP (?), 4mM ADP (?), 8 mM ADP (?), 12 mM ADP (? ), 16 mM ADP (?). Assays were carried out as described in Fig. 2 in the presence of 712 ?M of NAD. The inset shows the secondary plot of the line intercept (1/Vm) vs ADP concentration to determine the inhibition constant. 166 1/[NAD+] (mM-1) 0 1 2 3 4 5 0 5 10 15 20 25 1/V (? mo l x m in- 1 / m g p ro tei n) [ATP] (mM) -5 0 5 10 15 20 Sl op e 1 2 3 4 5 1/V (? mo l x m in- 1 / m g p ro tei n) Sl op e Fig. 12. Inhibition of yeast recombinant SSADH by ATP. Double-reciprocal plots of the rate of NADH formation vs NAD concentration at different ATP concentrations: 0 mM ATP (?), 4mM ATP (?), 8 mM ATP (?), 12 mM ATP (? ), 16 mM ATP (?). Assays were carried out as described in Fig. 2 in the presence of 85 ?M of SSA. The inset shows the secondary plot of the line slopes vs ATP concentration to determine the inhibition constant. 167 1/[SSA] mM-1 0 20 40 60 80 0 10 20 30 40 50 60 [ATP] (mM) -10 -5 0 5 10 15 20 Int er ce pt 2 4 6 8 10 12 1/V (? mo l x m in- 1/ mg pr ot ein ) Int er ce pt 1/V (? mo l x m in- 1/ mg pr ot ein ) Fig. 13. Inhibition of yeast recombinant SSADH by ATP. Double-reciprocal plots of the rate of NADH formation vs SSA concentration at different ATP concentrations: 0 mM ATP (?), 4mM ATP (?), 8 mM ATP (?), 12 mM ATP (? ), 16 mM ATP (?). Assays were carried out as described in Fig. 2 in the presence of 712 ?M of NAD. The inset shows the secondary plot of the line intercept (1/Vmax) vs ATP concentration to determine the inhibition constant. 168 VI. IN GEL STAINNING METHOD FOR DETECTING GABA TRANSAMINASE ACTIVITY Abstract A method was developed to detect GABA transaminase activity in native polyacrylamide gels. This method was then tested using homogenates from Arabidopsis thaliana for the identification of plant GABA-TP substrates. GABA transaminases in cell-free crude extract samples were resolved by native polyacrylamide gel and then the protein gels were incubated with the succinate semialdehyde dehydrogenase coupling substrate solution containing GABA as the amino donor and ?-ketoglutarate or pyruvate as the amino acceptor, the resulting product succinate semialdehyde was reduced by succinate semialdehyde dehydrogenase causing the oxidation of nitroblue tetrazolium. The resulting enzyme activity appeared as a dark band against an opaque background. The whole procedure can be completed within 15 to 20 min after protein gel electrophoresis, and is a rapid, semiquantitative method to examine the activity of GABA aminotransferase activity in crude extracts. 169 Introduction GABA is a ubiquitous, four carbon non protein amino acid which is widely found from all prokaryotic and eukaryotic organisms (Satya and Nair 1990; Bown and Shelp 1997; Shelp et al., 1999). It is well known as a neurotransmission inhibitor in mammals (Varju et al., 2001), but its role in most organisms remains unknown. For decades, GABA has been found to accumulate rapidly in plants in response to various biotic and abiotic stresses, such as hypoxia, cold, heat, and mechanical stimulation (Kinnersley and Turano, 2000). GABA is produced from glutamate by glutamate decarboxylase (GAD), and then transaminated by GABA aminotransferase (GABA-TP) to succinate semialdehyde (SSA) which is finally converted to succinate by succinate semialdehyde dehydrogenase (SSADH) (Bown and Shelp, 1997). These reactions constitute the GABA shunt pathway, which moves carbon from ?-ketoglutarate through glutamate to succinate in the TCA (Krebs) cycle. In Saccharomyces, GABA transaminase was found to play an important role in stress tolerance (Coleman et al., 2003; Chapter 1) and is highly specific for ?- ketoglutarate as a substrate (Ramos et al., 1985). In plants, the only well characterized GABA-T is a pyruvate-dependent form (Van Cauwenberghe et al., 1999). However, an ?-ketoglutarate-dependent GABA-T activity has been observed in tobacco crude preparations (Van Cauwenberghe et al., 1999). Pyruvate-dependent activity (GABA-TP) may have been partially separated from an ?-ketoglutarate-dependent GABA-T activity by FPLC anion exchange chromatography (Van Cauwenberghe et al., 1999). However, 170 the ?-ketoglutarate dependent GABA-T activity was so unstable that such conclusions are uncertain and have not been verified. To date no plant (in the Viridiplantae) cDNA or genomic sequences with homology to bacterial, fungal, or animal ?-ketoglutarate- dependent GABA-T are found in the nonredundant database at the National Center for Biotechnology information or have been found including plants with complete genome sequences. Thus, it is unclear whether an ?-ketoglutarate-dependent GABA-T exists in any higher plant. A number of assay methods for GABA-T activity have been recorded. The transamination activity was detected by measuring product formation such as alanine, glutamate, or SSA which can be determined either by HPLC (Van Cauwenberghe et al., 1995) or a second enzyme coupled assay based on the formation of NAD(P)H which absorbs at 340 nm spectrophotometrically (Ansari et al., 2005; Ramos et al., 1985). An activity staining of a filter paper print of electrophoresis resolved proteins has been used to identify GABA-T from plant tissues based on the fluorescence of NADPH under short wave ultraviolet light (Van Cauwenberghe et al., 2002), but none of the above techniques has met wide acceptance. In-gel activity staining or zymography is a useful technique for the detection of enzyme activities in non-denaturing polyacrylamide gels. It involves protein separation by electrophoresis followed by in gel assay of enzyme activities (Gabriel and Gersten, 1992; Gabriel and Gersten, 1992). It has proved invaluable in assessing the enzyme activity in non-fractionated cell extracts and for estimating the molecular weight and isoelectric point of the corresponding polypeptides and their isoforms (Kaberdin and McDowall, 2003). An in gel activity method for the detection of succinate semialdehyde 171 dehydrogenase from rat brain has been reported (Kammeraat and Veldstra, 1986). In this paper, we have extended the in-gel activity staining for succinate semialdehyde dehydrogenase activity to the detection of GABA transaminase activity by using our previously characterized yeast GABA-T knockout and overexpression of yeast strains and Arabidopsis plants as the enzyme sources to demonstrate the ability of the in gel staining assay system to detect GABA-T activity from crude protein mixtures. Materials and methods Plant material, growth condition, and Media Seeds of Arabidopsis thaliana ecotype ?Columbia? were obtained from Lehli Seeds.. Standard procedures were employed for plant germination and growth. The plants were grown on basal salts medium of Murashige and Skoog (Murashige, 1962) at approximately 22?C under continuous light provided by cool-white fluorescent lamps for two weeks. The collected plant seedlings were rapidly frozen in liquid N2 and stored at - 80?C before extraction. Yeast strains, growth conditions and media GABA transaminase deletion mutant ?uga1 (Mat a leu2-3112 ura3-1 trp1-1 his3- 11,15 ade2-1, lys2, met uga1::HIS3) derived from wild type yeast strain W303-1A (Mat a leu2-3,112 ura3-1 trp1-1 his3-11,15 ade2-1, lys2, met) was generated as described (Chapter I). A plasmid P425 GPD-UGA1 bearing yeast GABA transaminase gene UGA1 was transformed into ?uga1 and wild type W303-1A as described (Chapter II) to generate the overexpression of UGA1p. The wild type, mutant and overexpression transformant strains were grown on YPD medium containing 2% glucose, 1% yeast extract, 2% yeast 172 bactopeptone, or YNB medium containing 0.67% [wt/vol] yeast nitrogen base and 2% glucose supplemented with essential amino acids (Sherman et al., 1979). Enzyme and protein crude extract preparations The enzyme succinate semialdehyde dehydrogenase (SSADH) cloned and purified as described (Chapter V) was used for the assay of GABA-transaminase activity. Two weeks old Arabidopsis seedlings were homogenized as described (Wattebled et al., 2005) in cold extraction buffer (100 mM MOPS, pH 7.2; 1 mM EDTA; 1 mM dithiothreitol; and 10% glycerol). The homogenate was centrifuged twice for 20 min at 10,000g at 4?C. The resulting supernatant was immediately used to perform zymograms (after protein assay). Mid-log phase yeast cells were harvested and washed once in ddH2O. The washed cells were suspended in in lysis buffer containing 53.35 mM sodium phosphate, pH 8.0; 47.8 mM sodium chloride; 5 mM potassium chloride; 61 mM glucose, 0.1% Triton X-100 and 0.1 mM EDTA and broken using glass beads as described (Kharade et al., 2005). The homogenate was centrifuged at 4?C for 10 min at 10,000g. The protein content of the resulting supernatant was determined, and it was immediately used to perform zymograms as described below. Protein concentration was determined spectrophotometrically at 595 nm by Bradford method (Bradford, 1976) using the Bio-Rad Protein Assay Dye Reagent concentrate (Bio-Rad, Hercules, CA). 173 Zymograms of GABA transaminase activities Protein gels were prepared according to the standard protocol (Sambrook et al., 1989). Protein mixtures were separated under non-denaturing conditions using 8% polyacrylamide gels with 5% stacking gel. For detection of GABA transaminase activity, gels was soaked at room temperature with gentle shaking in a 10 ml solution containing 100 mM potassium pyrophosphate buffer, pH 8.4, 0.1mM pyridoxal 5-phosphate (PLP), 16mM GABA and 4 mM pyruvate or ?-ketoglutarate for 5 min for the first step GABA transaminase reaction, the resulting product succinate semialdehyde (SSA) was determined based on the oxidation of nitroblue tetrazolium (NBT) with the addition of 0.2 mg/ml NBT, 0.06 mM phenazine methosulfate (PMS), 1.5 mM NAD+ and 1.5 unit of SSADH. The gel was incubated with this solution at room temperature for another 10 to 15 min until dark bands appeared. The excess stain was removed by washing the gels in ddH2O. Results and Discussion Published methods for assaying the enzyme activity of GABA transaminse are mostly based on the production of NADH or NADPH (Ramos et al., 1985; Ansari et al., 2005) or measuring the formation of product of alanine, glutamate or SSA. Most of these assays do not work well on crude cell extracts because the assays are not very specific requiring separation of the protein mixture to accurately assay the activity of interest, and such assays can also be so time consuming. Zymogram staining is a common technique for the detection of many enzyme activities in acrylamide gels under non-denaturing conditions based on the production of colored products immediately following protein 174 separation by gel electrophroesis. We have developed a novel method using zymogram in gel staining to overcome many of these drawbacks and avoid possible artifacts from spectrophotometric measurements. The detection GABA-T activity requires relatively short incubation time (15 ~ 20 minutes), and many samples can be assayed at once. The detection of GABA-transaminase from various yeast sources indicated that in gel staining can be effectively employed. Yeast GABA-transaminase is ?-ketoglutarate specific and encoded by the UGA1 gene. The zymogram shown in Fig. 1 demonstrates only a single band of UGA1p activity in the cell free crude extracts from yeast cells. One single band was detected in the wild type cells, and no band was observed in the ?uga1 deletion mutant due to the deletion of the coding sequence, confirming the loss of function of UGA1p in the ?uga1 deletion mutant. When the plasmid bearing UGA1 gene under a strong constitutive promoter (P425 GPD-UGA1) was transformed into the wild type and mutant strains, the wild type cells showed a much stronger band with UGA1p activity. Similarly, more UGA1p activity was recovered in the mutant transformant cells than detected wild type level. However, it should be noted that some similar enzyme isoforms may migrate together with the detected protein because the separation of the proteins by native polyacrylamide gel is based on the electric charge density. The specificity of the detection of UGA1p activity was confirmed by the reaction when no GABA was added, no band was detected (data not shown). It has been widely accepted that plant GABA-transaminase is pyruvate specific and can only use pyruvate as a substrate. Interestingly, ?-ketoglutarate dependent plant GABA-T activity was shown to be present in the crude extracts from tobacco leaf, but not detected in the partially purified preparation (Van Cauwenberghe et al., 1999). This 175 suggests the existence of a second ?-ketoglutarate-specific GABA-T in plants, or alternatively that the ?-ketoglutarate-dependent plant GABA-T activity was an artifact due to some combination of aminotransferase activities in crude preparations. To further identify the substrates of plant GABA-T and identify the number of GABA-T activities in plants crude cell-free extracts from 2 weeks old Arabidopsis seedlings were separated on 8% native polyarcylamide gels, and zymogram staining was conducted for GABA-T activity. In Fig. 2, when pyruvate was present as substrate in the staining solution, a single dark band was detected showing the pyruvate-dependent GABA-T activity (lane 1). When ?-ketoglutarate instead of pyruvate was present, even darker bands were detected in a different location (lane 2) showing the ?-ketoglutarate-dependent enzyme activities. To further investigate whether the detected bands were related to GABA-T activity, SSADH (lane 3, Fig. 2) and GABA (lane 4, Fig. 2) were separately removed from the staining solution. Unexpectedly, the enzyme activity-related bands were not observed at the same location as in lane 1, suggesting that ?-ketoglutarate-dependent GABA-T activity was not observed. However, it was interesting to find bands were disappeared without the presence of co-factor NAD+ (lane 6, Fig. 2), ?-ketoglutarate (lane 7) or both (data not shown). Possibly an ?-ketoglutarate dependent dehydrogenase activity was detected instead. These results from in gel staining indicated to us that the plant GABA-T from Arabidopsis was highly pyruvate-dependent, and that there was no detectable GABA-T activity for the substrate ?-ketoglutarate, which was contradictory to Van Cauwenberghe et al.'s finding (1999). 176 Conclusion We describe a simple and rapid zymogram technique for detection of GABA- transaminase activity after electrophoresis under native conditions in polyacrylamide gels. The technique involves incubation of the native gel with a SSADH staining solution containing GABA-T and SSADH substrates. The assays were based on the production of SSA from the first reaction by GABA transaminase which was subsequently detected by excess SSADH leading to the production of NADH and the subsequent reduction of NBT. The reduced product in the gel appeared as a dark blue band. The method was shown to be useful for monitoring of GABA-T activities from cell crude extracts including plants and yeast and possibly other organims as well as for characterization of GABA-T and checking the biochemical purity and substrate specificity of GABA-T preparations. 177 References 1. Ansari, M. I., R. H. Lee, and S. C. G. Chen. 2005. A novel senescence- associated gene encoding ?-aminobutyric acid (GABA): pyruvate transaminase is upregulated during rice leaf senescence. Physiologia Plantarum. 123(1): 1-8. 2. Bown, A. W., and B. J. Shelp. 1997. The metabolism and function of ?- aminobutyric acid. Plant Physiol. 115: 1?5. 3. Bradford, M. M. 1976. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Analytical Biochemistry. 72: 248-254. 4. Coleman, S. T., T. K. Fang, S. A. Rovinsky, F. J. Turano, and R. W. S. Moye. 2001. Expression of a glutamate decarboxylase homologue is required for normal oxidative stress tolerance in Saccharomyces cerevisiae. J Biol Chem. 276(1):244- 50. 5. Gabriel, O., and D. M. Gersten. 1992. Staining for enzymatic activity after gel electrophoresis, I. Anal. Biochem. 203: 1-21. 6. Gersten, D. M., and O. Gabriel. 1992. Staining for enzymatic activity after gel electrophoresisII. Enzymes modifying nucleic acids. Anal. Biochem. 203: 181- 186. 7. Kaberdin, V. R., and K. J. McDowall. 2003. Expanding the Use of Zymography by the Chemical Linkage of Small, Defined Substrates to the Gel Matrix. Genome Res. 13: 1961-1965. 8. Kammeraat, C., and H. Veldstra. 1986. Characterization of succinate semialdehyde dehydrogenase from rat brain.Biochim Biophys Acta. 151(1): 1-10. 9. Kharade, V. N. Mittal, S. P. Das, P. Sinha, and N. Roy. 2005. Mrg19 depletion increases S. cerevisiae lifespan by augmenting ROS defence. FEBS Letters, 579 (30): 6809-6813. 10. Kinnersley, A. M., and F. J. Turano. 2000. ?-Aminobutyric acid (GABA) and plant responses to stress. Crit Rev Plant Sci . 19: 479?509. 11. Murashige, T., and F. Skoog. 1962. A revised medium for rapid growth and bioassays with tobacco tissue cultures. Physiol. Plant. 15: 472-497. 12. Ramos, F., M. E. Guezzar, M. Grenson, and J. M. Wiame. 1985. Mutations affecting the enzymes involved in the utilization of 4-aminobutyric acid as 178 nitrogen source by the yeast Saccharomyces cerevisiae. Eur. J. Biochem. 149: 401-404. 13. Sambrook, J., E. F. Fritsch, and T. Maniatis. 1989. Molecular cloning: A laboratory manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. 14. Satya, N. V., and P. M. Nair. 1990. Metabolism, enzymology and possible roles of ?-aminobutyrate in higher plants. Phytochemistry. 29: 369?375. 15. Shelp, B. J., A. W. Bown, and M. D. MacLean. 1999. Metabolism and function of ?-aminobutyric acid. Trends Plant Sci. 4: 446?452. 16. Sherman, F., G. R. Fink, and C.W. Lawrence. 1979. Methods in Yeast Genetics. Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y. 17. Van Cauwenberghe, O. R., A. Makhmoudova, M. D. McLean, S. M. Clark, and B. J. Shelp. 2002. Plant pyruvate-dependent gamma-aminobutyrate transaminase: identification of an Arabidopsis cDNA and its expression in Escherichia coli. Can. J. Bot. 80(9): 933?94. 18. Van Cauwenberghe, O. R., and B. J. Shelp. 1999. Biochemical characterization of partially purified gaba:pyruvate transaminase from Nicotiana tabacum. Phytochemistry. 52(4): 575-581. 19. Varju, P., Z. Katarova, E. Madarasz, and G. Szabo. 2001. GABA signaling during development: new data and old questions. Cell Tissue Res. 305: 239?246. 20. Wattebled, F., Y. Dong, S. Dumez, D. Delvalle, V. Planchot, P. Berbezy, D. Vyas, P. Colonna, M. Chatterjee, S. Ball, and C. D?Hulst. 2005. Mutants of Arabidopsis Lacking a Chloroplastic Isoamylase Accumulate Phytoglycogen and an Abnormal Form of Amylopectin1. Plant Physiology 138: 184-195. 179 1 2 3 4 Fig. 1. Zymogram staining analysis of GABA transaminase activities from various yeast sources. Cell-free extracts prepared from WT yeast cells (lane 1, 100 ?g), WT yeast overexpressing yeast GABA transaminase cells (lane 2, 50 ?g), yeast GABA transaminase deletion mutant cells (lane 3, 100 ?g), and yeast GABA transaminase deletion mutant cells overexpressing yeast GABA transaminase (lane 4, 50 ?g). The cells were all grown on YPD medium. After electrophoresis, the gel was soaked in staining solution containing 100 mM potassium pyrophosphate buffer, pH 8.4, 0.1mM pyridoxal 5-phosphate (PLP), 16mM GABA and 4 mM a-ketoglutarate for 5 min at room temperature. Then 0.2 mg/ml nitroblue tetrazolium (NBT), 0.06 mM phenazine methosulfate (PMS), 1.5 mM NAD+ and 1.5 unit of SSADH were added into the solution, followed by incubation at room temperature for another 15 min. 180 1 2 3 4 5 6 7 Fig. 2. Zymogram staining analysis for the substrate identification of Arabidopsis GABA transaminase. Approximately 100 ?g of crude cell extract from 2 week-old Arabidopsis seedlings were loaded onto an 8% native polyacrylamide gel. After electrophoresis under native conditions, gels were soaked as described in Fig. 1. The staining solution components vary from each lane: lane 1, pyruvate as substrate instead of ?-ketoglutarate; lane 2, ?-ketoglutarate as substrate instead of pyruvate; lane 3, ?-ketoglutarate as substrate but without SSADH; lane 4, ?-ketoglutarate as substrate but without GABA; lane 5, ?-ketoglutarate as substrate but without GABA and SSADH; lane 6: ?- ketoglutarate as substrate but without NAD+; lane 7, no ?-ketoglutarate as substrate. 181 APPENDIX I. GABA METABOLISM DURING STRESS IN YEAST Mitochondria Cytosol FADH2 FAD Fumarate H2O Oxaloacetate NAD+ NADH+H+ Malate Pyruvate Pyruvate CoA CO2 NADH+H+ NAD+ Succinate Succinyl-CoA CoA CO2 NADH+H+ NAD+ Isocitrate Acetyl-CoA Citrate 2-ketoglutarate NAD+ NADH+H+CO2 IDH1 CoA ADP+Pi ATP LSC1 2-ketoglutarate CO2GABA H+ glutamate GOAT UGA1 GAD YMR250W Succinyl- semialdehyde Succinyl- semi- aldehyde NADP+ NADPH +H+ SSADH UGA5 GABA Metabolism During Stress in Yeast 182 APPENDIX II. GABA METABOLISM DURING STRESS IN PLANTS NADPH + H+ CoA CO2 NADH+H+ NAD+ 2-ketoglutarate Mitochondria Cytosol Succinate FADH2 FAD CoA 2-ketoglutarate CO2 NADH+H+ NAD+ Acetyl-CoA Citrate Isocitrate NAD+ NADH+H+CO2 Pyruvate Pyruvate Glutamine NAPD+ or Fdox NADPH + H+ or Fdred NH4+ ATP ADP + Pi GABA H+CO2 Glutamate Isocitrate CO2 NADP+ NADPH + H+ GAD Alanine AAT CoA ADP+Pi Succinyl-CoA ATP Oxaloacetate NAD+ NADH+H+ Fumarate Malate H2O GABA GlycineSerine GlyoxylateHydroxypyruvate AGT Alanine Succinyl- semialdehyde NADP+ GPAT SSADH GABA Metabolism During Stress in Plants