From the Department of Physiology and Biophysics,
University of Iowa, Iowa City, Iowa 52242 and the ¶ United States
Department of Agriculture, Agricultural Research Service, Climate
Stress Laboratory, Beltsville, Maryland 20705
Received for publication, August 6, 2000, and in revised form, October 5, 2000
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ABSTRACT |
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The action of Regulation of GABA levels is a consequence of the balance between GABA
synthesis, catalyzed by GAD, and GABA degradation, catalyzed by the
concerted action of the enzymes GABA transaminase and succinate
semialdehyde dehydrogenase (SSADH) (reviewed in Ref. 5). Human genetic
diseases have been linked to loss of each enzyme in this pathway,
underscoring its importance in normal physiology (6). The specific
mechanisms that regulate the activity of these GABA metabolic enzymes
are not well described except for plant GAD (7). This GAD enzyme has
been found to bind the Ca2+ regulatory protein calmodulin
(CaM), a binding step that acts to elevate the specific activity of the
plant GAD (8), perhaps by inducing dimerization (9). This tight
regulation is important in plants, as loss of CaM-regulated GAD
activity elicits developmental abnormalities in plants (10).
Although the role of GAD-produced GABA as an intercellular signal is
well established, relatively little is known of the intracellular function of this glutamate metabolite. In this work, we present initial
characterization of the GAD/GABA metabolic pathway in the yeast
Saccharomyces cerevisiae. The use of GABA as a nitrogen source in S. cerevisiae has been well studied, especially
with regards to the transcriptional control of the GABA transaminase and GABA transporter loci (11-13). UGA1 encodes the GABA
transaminase (14), whereas UGA4 produces the GABA
transporter (15). Genetic evidence argues that UGA2 may
encode the enzyme SSADH (16), but definitive data are lacking to
substantiate this suggestion.
Here we provide evidence that the YMR250w locus is the gene
that encodes the S. cerevisiae glutamate decarboxylase. Data
are also presented that YBR006w encodes the SSADH enzyme in
this organism. Importantly, we find that the S. cerevisiae
L-glutamate metabolism pathway is involved in oxidant
tolerance. These experiments provide important evidence that, in
addition to the roles of GABA as a neurotransmitter and endocrine
regulator, glutamate/GABA metabolism is a key contributor to the
ability of cells to tolerate oxidative insult.
Yeast Strains and Media--
The yeast strains used in this
study were SEY6210 (MAT Isolation of the GAD1 Gene--
A YEp24-based genomic library
(21) was introduced into SEY6210 by a high efficiency transformation
method (22). Ura+ transformants were selected on minimal
medium and tested for oxidant hyper-resistance by replica plating onto
YPD medium containing diamide. Ura+ and diamide-resistant
colonies were then streaked onto SC media containing a range of diamide
concentrations. A plasmid conferring increased levels of resistance to
diamide was recovered and designated HRD12. This plasmid was
re-introduced into SEY6210, resulting in increased diamide and
H2O2 tolerance.
Plasmids--
HRD12 is 9.8 kb of yeast genomic DNA cloned as a
Sau3AI fragment into the BamHI site of YEp24. The
YEp352-GAD1 plasmid was constructed by inserting a 3.6-kb
Asp718 fragment containing the GAD1 gene into
Asp718-digested YEp352. A 3.0-kb
SmaI-SalI fragment was inserted into YEp351 to
form YEp351-GAD1. The Asp718 fragment from YEp352-GAD1 was
moved into pBluescript KSII+ to form pBS-GAD1. The coding
sequence of GAD1 was PCR-amplified (primers: CCG AGA TCT ATG
TTA CAC AGG CAC GGT TC and CCG GTC GAC TCA ACA TGT TCC TCT ATA GT) and
cloned into pTrcHis2-TOPO (Invitrogen Corp., Carlsbad, CA) generating
the plasmid pBG1. The GAD1 coding sequence was then moved
into BamHI/SalI-digested pGEX-KG (23)
as a BglII/SalI fragment (pBG2) to form the
GST-GAD1 fusion. The plasmid pUGA4-lacZ was
constructed by PCR. A 579-base pair fragment of the UGA4
promoter and translation start signal was produced with an upstream
primer (GCG AAT TCT TTG GGA TCT ATT TTT CTC TTT AG) and a downstream
primer (CGG GAT CCT TAT TCT CGT TTT TGC TTG A) corresponding to
positions Gene Disruption--
A SmaI-SalI fragment
from pUC19-HIS3 was inserted into
SmaI-SalI-digested pBS-GAD1 to produce a gene
disruption of GAD1. The resulting plasmid was designated
p Characterization of Yeast GAD-Glutathione S-Transferase (GST)
Fusion Protein--
E. coli transformants expressing the
GST-Gad1p fusion protein were grown overnight, ~16 h, at 37 °C in
LB with 100 µg/ml ampicillin. Overnight cultures were diluted 1:100
in fresh LB with 100 µg/ml ampicillin and incubated at 25 °C with
shaking at 200 rpm. After 4 h,
isopropyl-1-thio- Cloning of a Diamide and H2O2
Resistance-conferring Locus--
As part of our ongoing studies on
oxidative stress in S. cerevisiae, we carried out a screen
of a high copy number plasmid library to identify genes that were
capable of increasing resistance to the oxidants diamide and
H2O2. Diamide causes oxidative stress by
shifting the majority of intracellular glutathione from the reduced
form to the oxidized form (28), whereas H2O2
elicits oxidative damage to proteins, lipids and nucleic acids
(29).
The wild-type yeast strain SEY6210 was transformed with a S. cerevisiae YEp24 genomic DNA library. Approximately 20,000 Ura+ transformants were recovered and tested for diamide
tolerance by replica plating to YPD plates containing various
concentrations of diamide. Colonies that survived increased
concentrations of diamide were purified and plasmids recovered. The
recovered plasmids were then reintroduced into SEY6210 to further
assess their diamide tolerance. Plasmids that were able to reproducibly
confer resistance to diamide were also tested for resistance to
H2O2. We focused our studies on a plasmid that
conferred resistance to both diamide and H2O2
that was designated HRD12, for hyper-resistance
to diamide. The genomic DNA insert carried by the HRD12
plasmid was ~9.8 kb (data not shown).
To localize the gene responsible for resistance to diamide and
H2O2 from HRD12, we first sequenced both ends
of the genomic DNA insert. We then performed a BLAST search of the
S. cerevisiae genomic sequence (30) using these DNA
sequences to determine the bounds of the segment of cloned genomic DNA
and found that this segment of genomic DNA originated from chromosome
XIII. Further computer analysis detected the presence of several open
reading frames (ORFs) present in this segment of genomic DNA. One of
these ORFs (YMR250w) was subcloned as a 3.6-kb
Asp718 fragment into YEp352 (31), and the resulting
construct was designated YEp352-GAD1. This construct was then
transformed into wild-type cells and found to confer the same level of
tolerance to diamide and H2O2 as HRD12 (data
not shown). A BLAST search of the GenBankTM data base with the
predicted amino acid sequence of YMR250w indicated the
presence of sequence similarity with glutamate decarboxylase proteins. The two most closely related sequences were the Petunia
hybrida GAD with 39% and A. thaliana GAD with 38%
identity, respectively (Fig. 1). Key
lysine and histidine residues were conserved in the plant and yeast
sequences, again supporting the identification of YMR250w as
encoding the S. cerevisiae glutamate decarboxylase enzyme,
and we have designated this locus as GAD1.
Biochemical Characterization of Gad1p--
To examine the
biochemical properties of S. cerevisiae Gad1p, the open
reading frame from YMR250w was expressed in E. coli as a fusion protein. The recombinant protein was purified by
affinity chromatography (Fig. 2) using a
CaM-Sepharose column. CaM-binding proteins were eluted with 2 mM EGTA. Purified CaM-binding proteins were analyzed by
silver staining SDS-polyacrylamide gels and by immunoblot analysis. A
91-kDa protein was found to be retained by the CaM-Sepharose column
from extracts expressing the GST-Gad1p fusion protein. This result
agreed well with the combined molecular masses of GST (26.5 kDa) and
the estimated molecular mass of the deduced amino acid sequence from
the yeast Gad1p (66 kDa). To demonstrate that GST did not bind to CaM,
protein extracts from E. coli overexpressing GST were
subjected to CaM affinity purification and no proteins were found to be
bound by this resin. To confirm the identity of the CaM-binding
peptide, affinity-purified peptides were subjected to immunoblot
analysis with antiserum to Petunia GAD (7). The 91-kDa
fusion peptide cross-reacted with the Petunia GAD antiserum.
Western blot analysis of crude extracts expressing GST or the GST-Gad1p
fusion also demonstrated that the Petunia antiserum would
specifically detect the yeast protein (data not shown). Collectively,
these results confirm that the recombinant fusion protein was expressed
in E. coli and that the yeast Gad1p was responsible for CaM
binding of the fusion peptide. The purified recombinant fusion protein
was assayed for GAD activity, and no activity was observed. Several
attempts, including excision of the Gad1p from GST fusion protein or
expression of the yeast Gad1p as an unfused protein, were unsuccessful.
Possible reasons for the failure to detect enzymatic activity of this
recombinant protein include unique properties of the yeast enzyme and
possible differences in cofactor requirements (see below).
Normal H2O2 Resistance Requires the
Presence of the GAD1 Locus--
To examine the physiological role of
Gad1p, a strain lacking the GAD1 gene was constructed. A
fragment from
Growth of gad1 Normal Oxidant Tolerance Requires a Functional Glutamate Metabolism
Pathway in Yeast--
Our finding that oxidant resistance of S. cerevisiae is proportional to the gene dosage of GAD1
could be explained by the action of this enzyme to form GABA, which in
turn acts to signal a stress response. Alternatively, increased GAD
production could simply elevate the catabolism of glutamate. Increased
glutamate catabolism is believed to allow bacterial cells to tolerate
low pH conditions in the medium through the removal of acid by
elimination of glutamate (32). Importantly, elevated flux through the
glutamate catabolic pathway defined by GAD, GABA transaminase, and
SSADH could also elevate production of NADPH, an important antioxidant (33-35). NADPH can be produced by the action of SSADH (16). If this
last model is correct, then the entire glutamate catabolic pathway
would be predicted to be required for the observed increase in
oxidative stress tolerance provided by enhanced production of Gad1p.
To test this idea, we wanted to construct strains that lacked the
S. cerevisiae GABA transaminase and SSADH loci.
UGA1 encodes the GABA transaminase (14), but the
SSADH-encoding gene in this organism has not yet been identified. A
mutation called uga2 was isolated that eliminated SSADH
activity, but the gene corresponding to this mutation was never
isolated (16). A BLAST analysis of the protein data base from S. cerevisiae using the E. coli SSADH protein indicated
that the YBR006w ORF shared 52% identity with the bacterial
protein (Fig. 4). The
YBR006w-encoded protein also exhibited 47% identity with
human SSDH. Based on this high degree of sequence identity and other
properties (see below) of YBR006w, we designated this locus
UGA5 and believe it encodes the S. cerevisiae SSADH.
Deletion mutant strains were constructed that lacked either
UGA1 or UGA5. Primers were generated to
amplify a PCR product that replaced nucleotides +53 to +1370 of the
UGA1 coding region with the HIS3 gene employing
the same strategy used above to disrupt GAD1. This same
method was used to substitute nucleotides +53 to +1259 of
UGA5 with the HIS3 gene. The
uga1-
The ability to utilize GABA as the sole
nitrogen source in the medium defines the UGA series of
genes in S. cerevisiae (16). To provide support for our
placement of UGA5 in the glutamate catabolic pathway, we
tested the ability of the uga1
We next compared the oxidative stress resistance phenotypes of cells
lacking GAD1, UGA1, or UGA5. These
strains were also transformed with a high copy number plasmid that
contains the wild-type GAD1 gene. In a strain containing
UGA1 and UGA5, elevation of GAD1 gene
dosage increases oxidative stress tolerance (Fig. 3). To determine
whether this increased resistance required the presence of genes
downstream in the glutamate catabolic pathway, we assayed the ability
of these strains to tolerate H2O2 (Fig. 3).
Strains were grown to mid-log phase and assayed for tolerance to
varying concentrations of H2O2.
Loss of either UGA1 or UGA5 reduced
H2O2 resistance of cells with single or
multiple copies of the GAD1 locus. Thus, the oxidative stress phenotypes of uga1 UGA5 Expression Is Induced by Oxidants and GABA--
The
UGA1 gene and the GABA transporter-encoding UGA4
locus are expressed at a 30-100-fold higher levels when GABA is used as the nitrogen source rather than ammonium sulfate (11-13). This induction involves binding of the transcriptional regulatory protein Gln3p to the promoter elements of these GABA-regulated loci (36). Gln3p
recognizes elements centered around a GATAA sequence, a binding site
found upstream of genes involved in metabolism of poor nitrogen sources
like GABA (12). There are two close relatives of the GATAA sequence in
the UGA5 5'-flanking region, suggesting that this gene, like
UGA1 and UGA4, might be transcriptionally regulated by the nitrogen source in the medium. To examine this possibility, we obtained a lacZ fusion constructed by
transposon insertion into UGA5 from the Yale Genome Analysis
Center (37). This transposon insertion produced an in-frame fusion
between lacZ and UGA5 at codon 67 and was carried
in a diploid strain to allow the presence of a functional copy of the
gene as well as the UGA5-lacZ fusion gene. This strain was
grown in medium using either ammonium sulfate or GABA as the nitrogen
source and
UGA5-lacZ expression increased by ~240-fold when GABA
replaced ammonium sulfate as the nitrogen source (Fig.
6). This large induction of gene
expression is similar to those reported previously for
UGA1 and UGA4, other GABA metabolic proteins
(11-13). Since UGA5 is required for normal
H2O2 tolerance (Fig. 3), expression of the
UGA5-lacZ fusion was also examined in the presence of this oxidant. A nearly 3-fold higher level of
UGA5-dependent Genetic Evidence Supporting the Role of Gad1p as the S. cerevisiae
Glutamate Decarboxylase Protein--
Since we believe that
GAD1 encodes the S. cerevisiae glutamate
decarboxylase protein, elevation in the copy number of this gene would
be expected to increase the ability of a cell to produce GABA. To test
the ability of changes in GAD1 gene dosage to influence intracellular
GABA level, we utilized an indirect assay for GABA production.
UGA4 encodes the GABA transporter protein and is highly induced upon an increase in the level of GABA in the medium (15). We
constructed an UGA4-lacZ gene fusion to facilitate
measurement of UGA4 expression and introduced this
fusion gene into several different strains. These strains were
generated to provide varying levels of intracellular GABA by either
allowing (UGA1) or blocking (uga1
When GABA breakdown was blocked by the presence of a uga1 The participation of glutamate decarboxylase proteins in
intercellular signaling pathways has received a great deal of attention due to the important role of GABA in neurotransmission (1) and in plant
development (10). Our work provides insight into the intracellular
involvement of GAD in oxidative stress tolerance. Increasing the gene
dosage of the S. cerevisiae GAD1 locus produced an increased
tolerance to two different oxidative agents, diamide and
H2O2. This increased tolerance was
strictly dependent on the presence of the intact glutamate catabolic
pathway leading to the production of succinate from glutamate. Genetic
elimination of either enzymatic reaction downstream from glutamate
decarboxylase rendered cells hypersensitive to oxidants. Genetic
diseases have been described in humans that are associated with loss of
either GABA transaminase or SSADH enzymes, further underscoring the
importance of maintenance of the intact glutamate catabolic pathway
(6).
The identification of GAD1 as encoding S. cerevisiae glutamate decarboxylase was made on three independent
criteria. First, the highest degree of sequence similarity was found
between Gad1p and GAD proteins. Second, an antiserum directed against a
plant GAD enzyme cross-reacted with bacterially expressed Gad1p.
Finally, an increase in the gene dosage of GAD1 elicited an
increase of the expression of a gene known to be responsive to GABA
levels (UGA4). Another interesting biochemical similarity
between the plant and yeast enzymes was the observation that Gad1p was
able to bind to calmodulin like several plant GAD enzymes (7, 8, 27,
38-40). This finding suggests that perhaps S. cerevisiae GAD activity will be controlled by calcium/calmodulin as are these plant enzymes and may provide an explanation for the failure to detect
enzyme activity of the yeast enzyme in vitro. These
experiments suggest that, although Gad1p can bind to the mammalian
calmodulin protein, perhaps proper regulation of the yeast enzyme
requires the authentic S. cerevisiae calmodulin, a
possibility currently under investigation.
Evidence is also provided that the S. cerevisiae succinate
semialdehyde dehydrogenase is encoded by the YBR006w locus
that we propose be designated UGA5. As mentioned above,
earlier work defined a mutation called uga2 that eliminated
SSDH activity, but the locus defined by this mutation was not described
(16). Genetic analysis will be required to determine whether
YBR006w and uga2 are allelic. Several independent
pieces of evidence support identification of UGA5 as the
SSDH locus in S. cerevisiae. UGA5 shares the highest degree
of sequence similarity with SSDH enzymes from other organisms of any
ORF in the genome, it is required for utilization of GABA as a nitrogen
source and it is highly inducible when GABA is present in the medium.
SSADH is also a key participant in the ability of elevated Gad1p levels
to influence oxidative stress tolerance. Since loss of either
UGA1 or UGA5 is epistatic to amplification of the
GAD1 locus, we argue that the entire glutamate catabolic
pathway is required for the observed increase in oxidant resistance
conferred by high copy number GAD1 plasmids. SSADH enzymes
produce either NADH or NADPH during their oxidation of succinate
semialdehyde to succinate (41). This production of reduced nicotinamide
adenine dinucleotide can then be coupled to buffering redox changes
that would otherwise occur in the presence of oxidants like
H2O2 or diamide.
The precise nature of the reduced nicotinamide adenine dinucleotide is
not known in the case of S. cerevisiae SSADH. Measurements of the enzymatic properties of the yeast SSADH found that this protein
had a 2.5-fold higher activity when assayed in the presence of
NAD+ rather than NADP+ (16). Since the
[NADPH+NADP+]/[NADH+NAD+] ratio is 35 in
yeast (42), it seems more likely that NADP+ will serve as
the cofactor for the SSADH reaction in this organism. NADPH is critical
for maintenance of normal redox balance and is required for
regeneration of reduced glutathione and thioredoxin by glutathione
reductase and thioredoxin reductase, respectively (43). Previous work
has shown that activity of the pentose phosphate pathway, which also
leads to production of NADPH, is critical for normal oxidative stress
tolerance (33, 34). Ensuring proper levels of key antioxidants like
glutathione and NADPH is crucial for maintenance of normal oxidative
stress resistance (33, 34, 44). The data described here indicate
that activity of the glutamate catabolic pathway is also an essential
contributor to the intracellular pool of antioxidants.
The plant and human SSADH enzymes carry out their catalysis in the
mitochondrion (41, 45). We believe this is unlikely to be the case for
the S. cerevisiae protein. As expected for a protein
targeted to the mitochondrion, both the plant and human SSADH proteins
possess N-terminal targeting sequences (41, 45). However, the alignment
of yeast, bacterial, human, and plant enzymes indicates that the yeast
protein does not exhibit an N-terminal targeting extension and instead
shows strong sequence similarity with the bacterial enzymes. These data
suggest that the S. cerevisiae protein will be found in an
extra-mitochondrial location. Localization of the yeast proteins is the
focus of future work.
-aminobutyrate (GABA) as an
intercellular signaling molecule has been intensively studied, but the
role of this amino acid metabolite in intracellular metabolism is
poorly understood. In this work, we identify a Saccharomyces
cerevisiae homologue of the GABA-producing enzyme glutamate
decarboxylase (GAD) that is required for normal oxidative stress
tolerance. A high copy number plasmid bearing the glutamate
decarboxylase gene (GAD1) increases resistance to two
different oxidants, H2O2 and diamide, in cells
that contain an intact glutamate catabolic pathway. Structural
similarity of the S. cerevisiae GAD to previously studied
plant enzymes was demonstrated by the cross-reaction of the yeast
enzyme to a antiserum directed against the plant GAD. The yeast GAD
also bound to calmodulin as did the plant enzyme, suggesting a
conservation of calcium regulation of this protein. Loss of either gene
encoding the downstream steps in the conversion of glutamate to
succinate reduced oxidative stress tolerance in normal cells and was
epistatic to high copy number GAD1. The gene encoding
succinate semialdehyde dehydrogenase (UGA5) was identified and found to be induced by H2O2 exposure.
Together, these data strongly suggest that increases in activity of the
glutamate catabolic pathway can act to buffer redox changes in the cell.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
-Aminobutyric acid
(GABA)1 is a metabolite of
glutamate that is a major inhibitory neurotransmitter in animals (see
Ref. 1 for a review). GABA is generated by the decarboxylation of
L-glutamate by the enzyme glutamate decarboxylase (GAD) and
has been implicated in hormone release from endocrine cells of several
target tissues in mammals (2). Biosynthesis and secretion of GABA is a
critical step in assuring normal neural function, and defects in
metabolism of this glutamate derivative have been linked with clinical
manifestations including epilepsy and Parkinson's disease (3, 4).
MATERIALS AND METHODS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
leu2-3,-112 ura3-52
his3-
200 trp1-
901 lys2-801
suc2-
9, Mel
), YSC11 (MAT
leu2-3,-112 ura3-52 his3-
200
trp1-
901 lys2-801 suc2-
9 Mel
gad1-
1::HIS3),
YSC29 (MAT
leu2-3,-112 ura3-52
his3-
200 trp1-
901 lys2-801
suc2-
9 Mel
uga1-
1::HIS3),
YBB1 (MAT
leu2-3,-112 ura3-52
his3-
200 trp1-
901 lys2-801
suc2-
9 Mel
uga5-
1::HIS3), and BYv22B1
(MATa/MAT
leu2-98/leu2-98
cry1R/CRY1 ade2-101/ade2-101 HIS3/his3-200 ura3-52/ura3-52
lys2-801/lys2-801 can1R/CAN1
trp1-1/TRP1 CYH2/cyh2R
(CEN-LEU2-pGAL-cre) uga5-lacZ-URA3). Yeast cells
were grown in rich, nonselective medium (yeast extract-peptone-dextrose (YPD)), minimal medium synthetic dextrose (SD) with required
supplements, or synthetic complete (SC) medium (17). Transformation was
performed by the lithium acetate technique of Ito et al.
(18). Diamide and H2O2 tolerance assays
were carried out by spot tests (19). Assays for
-galactosidase
assays were carried out on permeabilized cells as described previously
(20). The growth rate experiments were performed over a 12-h period
with time points taken every hour in minimal medium with either
ammonium sulfate or 0.1% GABA as the nitrogen source.
550 to +29. The resulting product was cloned into the
lacZ fusion plasmid pSEYC102 (24) as an
EcoRI-BamHI fragment. All PCR products were
sequenced to ensure that no errors occurred during amplification.
GAD1 and replaced the entire GAD1 coding sequence with
the HIS3 gene. The p
GAD1 plasmid was digested with
AgeI and XbaI prior to transformation into
SEY6210. His+ transformants were selected and purified.
Genomic DNA was prepared (25), and the correct integration event was
verified by PCR and Southern blotting. A representative
gad1
mutant was used in these studies and designated
YSC11. Disruption of the UGA1 gene was produced by a
PCR-based method using the primers TCT ATT TGT GAA CAA TAC TAC CCA GAA
GAG CCA ACC AAA CCA ACT GTT AAT TGT
ACT GAG AGT GCA
CC and TCT ATT TGT GAA CAA TAC TAC CCA GAA GAG CCA ACC AAA
CCA ACT GTT AAT TGT ACT
GAG AGT GCA CCA
T. These primers were used to amplify via the underlined
sequences the HIS3 gene from plasmid pRS303 (26). The
resulting PCR product replaces the UGA1 coding sequence from
+53 to +1370 with the HIS3 gene. His+
transformants were selected and purified. As described above, the
correct integration event was verified by PCR using a primer upstream
of the UGA1 deletion (TTC GCG CTA TCT CGA TTT CTA CCT A) and
in the middle of HIS3 (TTC TTC GAA GAA ATC ACAT TAC TTT ATA
TA). The UGA5-lacZ fusion gene was obtained from the Yale Genome Analysis Center in plasmid form and used to transform the uga1
strain to analyze expression of UGA5 in response to
elevation of GAD1 copy number. Disruption of the
UGA5 gene was accomplished as above using the primers: ACT
TTG AGT AAG TAT TCT AAA CCA ACT CTA AAC GAC CCT AAT TTA TTC
AGT TGT ACT GAG
AGT GCA CCA T and TTA
AGT GTT GAC ATT TTT AGA AAA GAC ATA TGC TGC TAA ACC AAA CTC AGG GTA TTT CAC
ACC GCA TA. The resulting PCR
product replaces the UGA1 coding sequence from +53 to +1259
with HIS3. His+ transformants were selected and
purified. The correct integration event was verified from genomic DNA
by PCR using a primer upstream of the UGA5 deletion (ACC CAC
CGG AGA GGG CAA AGG TAA A) and in the middle of HIS3 (TTC
TTC GAA GAA ATC ACA TTA CTT TAT ATA).
-D-galactopyranoside was added to a
final concentration of 100 µM, to induce expression of
the recombinant protein, incubation continued 16 h. Bacterial
cells were collected by centrifugation at 6000 × g for
10 min. Pellets were immediately frozen in liquid N2 and
stored at
80 °C. As a positive control, rGAD1 from
Arabidopsis thaliana was prepared as described previously by
Turano and Fang (27), and, as negative control, a control GST was
prepared. Pellets containing cells were resuspended in 5-7 ml of
extraction buffer (50 mM Tris-HCl, pH 7.5, 0.2 mM EDTA, 2.5% (v/v) glycerol, 2 mM DTT, 0.05 mM pyridoxal 5'-phosphate, 0.05% (v/v) Triton, 10 µM leupeptin, 20 µg/ml lysozyme, and 1 mM
PMSF. The cells were lysed by sonication with 12 2-s bursts at 150 watts. Cellular debris was removed by centrifugation at 12,000 rpm for
20 min. The supernatant was filtered through a 0.45-µm filter.
CaCl2, DTT, and PMSF were added to final concentrations of
1, 2, and 1 mM, respectively. The supernatant was loaded on to a CaM-Sepharose column (Amersham Pharmacia Biotech, ~1-ml bed volume), pre-equilibrated in binding buffer (25 mM
Tris-HCl, pH 7.5, 150 mM NaCl, 2.5% (v/v) glycerol, 1 mM CaCl2, 2 mM DTT, 10 µM leupeptin, and 1 mM PMSF). The column was
washed with 20 bed volumes of wash buffer minus calcium (25 mM Tris-HCl, pH 7.5, 150 mM NaCl, 2.5% (v/v)
glycerol, 2 mM DTT, 10 µM leupeptin, and 1 mM PMSF). CaM-binding proteins were eluted with (50 mM Tris-HCl, pH 7.5, 2 mM EDTA, 150 mM NaCl, 2.5% (v/v) glycerol, and 1 mM PMSF).
Samples were immediately assayed as described above and analyzed by
SDS-PAGE and immunoblot analysis. Protein concentrations, estimates of
GAD activity, SDS-PAGE, silver staining, and immunoblot analysis were
conducted as described by Turano and Fang (27), except that
immunoreactive peptides were detected using a chemiluminescent detection system (SuperSignal West Dura, Pierce).
RESULTS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
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Fig. 1.
Computer alignment of glutamate decarboxylase
sequences from S. cerevisiae and plants. The
primary amino acid sequences of the glutamate decarboxylase enzymes
from Petunia and Arabidopsis were aligned with
the predicted gene product of the YMR250w (GAD1)
locus from S. cerevisiae. The alignment was generated by the
Megalign (Lasergene, DNA Star) program using the Clustal algorithm.
Amino acids are listed using the one-letter code,
and residues conserved in all three proteins are boxed. A
histidine residue important in enzyme function and the adjacent lysine
required for the binding of the pyridoxal-phosphate cofactor (46) are
indicated by asterisks.
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Fig. 2.
Calmodulin binding and immunological
cross-reactivity are shared properties of a plant GAD enzyme and
S. cerevisiae Gad1p. Protein extracts from
bacterial cells expressing either a fusion protein between GST and
Gad1p or GST alone were subjected to chromatography on
calmodulin-Sepharose beads. Specifically bound material was eluted with
EGTA and analyzed by SDS-PAGE, followed by visualization of total
eluted proteins using silver stain or Western blotting using an
anti-Petunia GAD antiserum. Bacterially expressed A. thaliana GAD (rGad1p) was used as a positive control for
immunological cross-reaction. Molecular mass standards are indicated on
the right.
530 to +2482 base pairs, relative to the
GAD1 translation start, was replaced with the
HIS3 gene. The resulting plasmid, p
GAD1, was digested with AgeI and XbaI before being transformed into
the wild-type strain SEY6210. His+ transformants were
recovered and confirmed to contain the desired gene disruption allele.
The resulting S. cerevisiae strain was designated YSC11.
cells on rich medium was indistinguishable
from wild-type cells. Differences in growth do arise, however, when
growth is compared on medium containing oxidants. We assayed the growth
of wild-type and gad1
strains on YPD plates containing either diamide or H2O2. Unlike wild-type cells,
the gad1
strain was unable to grow on plates containing 3 mM H2O2 (Fig.
3) or on increased diamide concentrations
(data not shown). These findings indicate that GAD1 is
required for normal resistance to oxidants.
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Fig. 3.
The glutamate catabolic pathway is required
for normal oxidative stress tolerance. A, the enzymatic
steps for conversion of glutamate to succinate in S. cerevisiae is shown with the corresponding structural genes listed
below. B, strains with either high (2-µm GAD1)
or low (GAD1) dosages of the GAD1 gene along with
the indicated UGA1 and UGA5 alleles were analyzed for their
ability to tolerate H2O2 by spot test assay.
Equal numbers of cells of each genotype were placed on rich medium
(YPD) or the same medium containing 3 mM
H2O2 and allowed to grow at 30 °C.
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Fig. 4.
Uga5p shares strong sequence similarity with
plant and bacterial SSADH enzymes. A computer alignment performed
as described above was carried out to analyze the extent of sequence
conservation between S. cerevisiae Uga5p (YBR006w
gene product), A. thaliana SSADH, and E. coli GabD. Identical residues are indicated as
boxes. Note the N-terminal extension on the plant protein
that is believed to serve a as mitochondrial targeting signal.
1::HIS3-containing strain and
the uga5-
1::HIS3-containing
strain were designated YSC29 and YBB1, respectively.
and uga5
strains to grow with either ammonium sulfate or GABA as the nitrogen source in the medium. These strains, along with the isogenic wild-type, were pre-grown in synthetic complete medium and then diluted into minimal medium with either 0.1% ammonium sulfate or GABA as the nitrogen source. Growth was monitored over 12 h to assess the ability of each strain to grow. Strains grown in medium with ammonium sulfate as the nitrogen source were able to grow normally (Fig. 5). However, in GABA media, only
wild-type cells were able to grow. Wild-type cells reached an optical
density of nearly 2, whereas uga1
and uga5
cells only grew to an absorbance value of 0.16. This result is
consistent with our identification of the UGA5 locus as
encoding S. cerevisiae SSADH since uga1
and uga5
mutant strains have similar defects during growth on
GABA-containing medium.
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Fig. 5.
Both UGA5 and
UGA1 are required for use of GABA as a nitrogen
source. An isogenic series of strains with the indicated
UGA1 or UGA5 alleles was tested for the ability
to grow in minimal medium using either ammonium sulfate or GABA as the
sole nitrogen source. Strains were grown to saturation in ammonium
sulfate medium and then diluted into fresh minimal medium containing
the nitrogen source indicated on the right. Cultures were
incubated at 30 °C with shaking and the optical density at 600 nm
(A600) was determined each hour.
and uga5
mutant
strains were epistatic to that conferred by high copy number
GAD1, again supporting both our placement of UGA5
downstream from GAD1 in the glutamate catabolic pathway and
the requirement for this pathway in the effect of Gad1p on oxidative
stress tolerance.
-galactosidase activity determined.
-galactosidase activity was
found when this strain was exposed to 1 mM
H2O2, supporting the idea that UGA5
expression may be increased to elevate flux through the glutamate
catabolic pathway and increase NADPH pools in the cell.
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Fig. 6.
UGA5 expression is regulated by
nitrogen source and oxidative stress. A, a diagram of
the UGA5 locus on chromosome II is shown. The potential
GATAA binding sites required for Gln3p-dependent gene
regulation are indicated upstream of the ATG for the UGA5
open reading frame. The approximate position for insertion of the
lacZ-containing transposon is shown. B, a diploid
cell carrying a single copy of the UGA5-lacZ gene fusion was
grown in minimal medium with either ammonium sulfate or GABA as the
sole nitrogen source. The ammonium sulfate-grown culture was split in
two aliquots, and one was treated with the addition of 1 mM
H2O2 to induce oxidative stress.
UGA5-dependent -galactosidase activities were
then determined for all strains. The numbers
above the bars represent the average of two
assays for two different transformants.
) consumption
of GABA. We anticipated that GABA levels should increase as the
UGA1 gene is removed and GAD1 gene dosage level increases as GABA is expected to be produced at a higher rate but can
no longer be broken down. Strains carrying the UGA4-lacZ fusion gene and varying in their copy number of GAD1 and
UGA1 were grown in medium containing ammonium sulfate or
glutamate as the nitrogen source and then assayed for expression of
UGA4 (Table I).
Elevation in GAD1 copy number induces UGA gene expression
) copy of
UGA1 were grown in media using the indicated nitrogen source
to mid-log phase. Each strain was also transformed with a
UGA4-lacZ gene fusion carried on a low copy number plasmid.
UGA4-dependent
-galactosidase activity was
determined in each case.
allele, UGA4-lacZ expression was found to increase by nearly
5-fold when the GAD1 copy number was elevated in the
presence of glutamate as a nitrogen source. If ammonium sulfate
replaced glutamate as the nitrogen source,
UGA4-dependent
-galactosidase levels were much lower and only increased from 0.4 units/OD to 1.3 units/OD when
GAD1 was present on a high-copy-number plasmid. Although UGA4 expression was induced when glutamate was used as the
nitrogen source rather than ammonium sulfate in a UGA1
genetic background, the extent of this induction was not significantly
affected by elevation of GAD1 copy number, consistent with
the intact glutamate catabolic pathway limiting the accumulation of
GABA. We also assayed the response of the UGA5-lacZ fusion
gene described above carried in uga1
cells to varying the
copy number of GAD1. A uga1
,
UGA5-lacZ strain produced 196 ± 42 units/OD
-galactosidase in the presence of high copy GAD1, which
was reduced to 12 ± 2.5 units when GAD1 was maintained
at chromosomal levels. These data support identification of
GAD1 as encoding the S. cerevisiae glutamate
decarboxylase enzyme and indicate that coordinate induction of
UGA1 and UGA5 is likely to occur when Gad1p
activity increases with accompanying GABA accumulation.
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
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ACKNOWLEDGEMENTS |
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We thank Terry Cooper for important discussions, the Yale Genome Analysis Center for rapidly providing reagents, and Belinda Baxa for technical assistance.
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FOOTNOTES |
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* This work was supported in part by National Institutes of Health Grant GM49825 (to W. S. M.) and was performed while W. S. M. was an Established Investigator of the American Heart Association.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
§ Present address: Dept. of Biology, University of the Ozarks, Clarksville, AR 72830.
Present address: Dept. of Biological Sciences, George
Washington University, Washington, DC 20052.
** To whom correspondence should be addressed: Dept. of Physiology and Biophysics, 5-612 Bowen Science Bldg., University of Iowa, Iowa City, IA 52242. Tel.: 319-335-7874; Fax: 319-335-7330; E-mail: moyerowl@blue.weeg.uiowa.edu.
Published, JBC Papers in Press, October 12, 2000, DOI 10.1074/jbc.M007103200
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ABBREVIATIONS |
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The abbreviations used are:
GABA, -aminobutyric acid;
GAD, glutamate decarboxylase;
YPD, yeast
extract-peptone-dextrose;
SD, minimal medium synthetic dextrose;
SC, synthetic complete;
PAGE, polyacrylamide gel electrophoresis;
CaM, calmodulin;
SSADH, succinate semialdehyde dehydrogenase;
DTT, dithiothreitol;
PMSF, phenylmethylsulfonyl fluoride.
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REFERENCES |
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---|
1. | Mody, I., De Koninck, Y., Otis, T. S., and Soltesz, I. (1994) Trends Neurosci. 17, 517-525[CrossRef][Medline] [Order article via Infotrieve] |
2. | Satin, L. S., and Kinard, T. A. (1998) Endocrine 8, 213-223[Medline] [Order article via Infotrieve] |
3. | Sherwin, A. L. (1999) Neurochem. Res. 24, 1387-1395[Medline] [Order article via Infotrieve] |
4. | Nisbet, A. P., Eve, D. J., Kingsbury, A. E., Daniel, S. E., Marsden, C. D., Lees, A. J., and Foster, O. J. (1996) Neuroscience 75, 389-406[CrossRef][Medline] [Order article via Infotrieve] |
5. | Shelp, B. J., Bown, A. w., and McLean, M. D. (1999) Trends Plant Sci. 4, 446-452[CrossRef][Medline] [Order article via Infotrieve] |
6. | Jakobs, C., Jaeken, J., and Gibson, K. M. (1993) J. Inherit. Metab. Dis. 16, 704-715[Medline] [Order article via Infotrieve] |
7. |
Baum, G.,
Chen, Y.,
Arazi, T.,
Takatsuji, H.,
and Fromm, H.
(1993)
J. Biol. Chem.
268,
19610-19617 |
8. |
Snedden, W. A.,
Koutsia, N.,
Baum, G.,
and Fromm, H.
(1996)
J. Biol. Chem.
271,
4148-4158 |
9. |
Yuan, T.,
and Vogel, H. J.
(1998)
J. Biol. Chem.
273,
30328-30335 |
10. | Baum, G., Lev-Yadun, S., Fridmann, Y., Arazi, T., Katsnelson, H., Zik, M., and Fromm, H. (1996) EMBO J. 15, 2988-2996[Abstract] |
11. | Vissers, S., Andre, B., Muyldermans, F., and Grenson, M. (1990) Eur. J. Biochem. 187, 611-616[Abstract] |
12. | Cunningham, T. S., Dorrington, R. A., and Cooper, T. G. (1994) J. Bacteriol. 176, 4718-4725[Abstract] |
13. | Talibi, D., Grenson, M., and Andre, B. (1995) Nucleic Acids Res. 23, 550-557[Abstract] |
14. | Andre, B., and Jauniaux, J. C. (1990) Nucleic Acids Res. 18, 3049[Medline] [Order article via Infotrieve] |
15. | Andre, B., Hein, C., Grenson, M., and Jauniaux, J. C. (1993) Mol. Gen. Genet. 237, 17-25[Medline] [Order article via Infotrieve] |
16. | Ramos, G., el Guezzar, M., Grenson, M., and Wiame, J. M. (1985) Eur. J. Biochem. 149, 401-404[Abstract] |
17. | Sherman, F., Fink, G., and Hicks, J. (1979) Methods in Yeast Genetics , Cold Spring Harbor Laboratory, Cold Spring Harbor, NY |
18. | Ito, H., Fukuda, Y., Murata, K., and Kimura, A. (1983) J. Bacteriol. 153, 163-168[Medline] [Order article via Infotrieve] |
19. |
Wu, A.,
Wemmie, J. A.,
Edgington, N. P.,
Goebl, M.,
Guevara, J. L.,
and Moye-Rowley, W. S.
(1993)
J. Biol. Chem.
268,
18850-18858 |
20. | Guarente, L. (1983) Methods Enzymol. 101, 181-191[Medline] [Order article via Infotrieve] |
21. | Carlson, M., and Botstein, D. (1982) Cell 28, 145-154[Medline] [Order article via Infotrieve] |
22. | Gietz, D., St. Jean, A., Woods, R. A., and Schiestl, R. H. (1992) Nucleic Acids Res. 20, 1425[Medline] [Order article via Infotrieve] |
23. | Guan, K.-L., and Dixon, J. E. (1991) Anal. Biochem. 192, 262-267[Medline] [Order article via Infotrieve] |
24. | Emr, S. D., Vassarotti, A., Garret, J., Geller, B. C., Takeda, M., and Douglas, M. G. (1986) J. Cell Biol. 102, 523-533[Abstract] |
25. | Hoffman, C. S., and Winston, R. (1987) Gene (Amst.) 57, 267-272[CrossRef][Medline] [Order article via Infotrieve] |
26. |
Sikorski, R. S.,
and Hieter, P.
(1989)
Genetics
122,
19-27 |
27. |
Turano, F. J.,
and Fang, T. K.
(1998)
Plant Physiol.
117,
1411-1421 |
28. | Kosower, N. S., and Kosower, E. M. (1987) Methods Enzymol. 143, 264-270[Medline] [Order article via Infotrieve] |
29. | Cadenas, E. (1989) Annu. Rev. Biochem. 58, 79-110[CrossRef][Medline] [Order article via Infotrieve] |
30. | Goffeau, A., Barrell, B. G., Bussey, H., Davis, R. W., Dujon, B., Feldmann, H., Galibert, F., Hoheisel, J. D., Jacq, C., Johnston, M., Louis, E. J., Mewes, H. W., Murakami, Y., Phillipsen, P., Tettelin, H., and Oliver, S. G. (1996) Science 274, 563-567 |
31. | Hill, J. E., Myers, A. M., Koerner, T. J., and Tzagaloff, A. (1986) Yeast 2, 163-167[Medline] [Order article via Infotrieve] |
32. |
Castanie-Cornet, M. P.,
Penfound, T. A.,
Smith, D.,
Elliott, J. F.,
and Foster, J. W.
(1999)
J. Bacteriol.
181,
3525-3535 |
33. | Nogae, I., and Johnston, M. (1990) Gene (Amst.) 96, 161-169[CrossRef][Medline] [Order article via Infotrieve] |
34. | Juhnke, H., Krems, B., Kotter, P., and Entian, K.-D. (1996) Mol. Gen. Genet. 252, 456-464[CrossRef][Medline] [Order article via Infotrieve] |
35. |
Salvemini, F.,
Franze, A.,
Iervolino, A.,
Filosa, S.,
Salzano, S.,
and Ursini, M. V.
(1999)
J. Biol. Chem.
274,
2750-2757 |
36. | Minehart, P. L., and Magasanik, B. (1991) Mol. Cell. Biol. 11, 6216-6228[Medline] [Order article via Infotrieve] |
37. | Burns, N., Grimwade, B., Ross-Macdonald, P. B., Choi, E.-Y., Finberg, K., Roeder, G. S., and Snyder, M. (1994) Genes Dev. 8, 1087-1105[Abstract] |
38. |
Ling, V.,
Snedden, W. A.,
Shelp, B. J.,
and Assmann, S. M.
(1994)
Plant Cell
6,
1135-1143 |
39. | Gallego, P. P., Whotton, L., Picton, S., Grierson, D., and Gray, J. E. (1995) Plant Mol. Biol. 27, 1143-1151[Medline] [Order article via Infotrieve]. |
40. | Zik, M., Arazi, T., Snedden, W. A., and Fromm, H. (1998) Plant Mol. Biol. 37, 967-975[CrossRef][Medline] [Order article via Infotrieve] |
41. |
Busch, K. B.,
and Hromm, H.
(1999)
Plant Physiol.
121,
589-597 |
42. |
Anderlund, M.,
Nissen, T. L.,
Nielsen, J.,
Villadsen, J.,
Rydstrom, J.,
Hahn-Hagerdal, B.,
and Kielland-Brandt, M. C.
(1999)
Appl. Environ. Microbiol.
65,
2333-2340 |
43. | Jamieson, D. J. (1998) Yeast 14, 1511-1527[CrossRef][Medline] [Order article via Infotrieve] |
44. | Grant, C. M., MacIver, F. H., and Dawes, I. W. (1996) Curr. Genet. 29, 511-515[CrossRef][Medline] [Order article via Infotrieve] |
45. |
Hearl, W. G.,
and Churchich, J. E.
(1984)
J. Biol. Chem.
259,
11459-11463 |
46. |
Tramonti, A.,
De Biase, D.,
Giartosio, A.,
Bossa, F.,
and John, R. A.
(1998)
J. Biol. Chem.
273,
1939-1945 |