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Molecular and Cellular Biology, November 1998, p. 6273-6280, Vol. 18, No. 11
0270-7306/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
Snf1 Protein Kinase Regulates Phosphorylation of
the Mig1 Repressor in Saccharomyces cerevisiae
Michelle A.
Treitel,1
Sergei
Kuchin,1 and
Marian
Carlson1,2,*
Departments of Genetics and
Development1 and
Microbiology,2 Columbia University, New
York, New York 10032
Received 13 February 1998/Returned for modification 26 March
1998/Accepted 28 July 1998
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ABSTRACT |
In glucose-grown cells, the Mig1 DNA-binding protein recruits the
Ssn6-Tup1 corepressor to glucose-repressed promoters in the yeast
Saccharomyces cerevisiae. Previous work showed that Mig1 is
differentially phosphorylated in response to glucose. Here we examine
the role of Mig1 in regulating repression and the role of the Snf1
protein kinase in regulating Mig1 function. Immunoblot analysis of Mig1
protein from a snf1 mutant showed that Snf1 is required for
the phosphorylation of Mig1; moreover, hxk2 and
reg1 mutations, which relieve glucose inhibition of Snf1, correspondingly affect phosphorylation of Mig1. We show that Snf1 and
Mig1 interact in the two-hybrid system and also coimmunoprecipitate from cell extracts, indicating that the two proteins interact in vivo.
In immune complex assays of Snf1, coprecipitating Mig1 is
phosphorylated in a Snf1-dependent reaction. Mutation of four putative
Snf1 recognition sites in Mig1 eliminated most of the differential
phosphorylation of Mig1 in response to glucose in vivo and improved the
two-hybrid interaction with Snf1. These studies, together with previous
genetic findings, indicate that the Snf1 protein kinase regulates
phosphorylation of Mig1 in response to glucose.
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INTRODUCTION |
In Saccharomyces
cerevisiae the Ssn6 (Cyc8)-Tup1 complex represses transcription of
genes regulated by glucose, cell type, oxygen, DNA damage, and other
signals (27, 34, 45, 46, 48, 52, 54, 56, 57, 61). Ssn6-Tup1
is recruited to these promoters by specific DNA-binding proteins,
including
2-Mcm1, a1-
2, Mig1, Mig2, Rox1, and Rgt1
(1, 27, 31, 39, 47, 51), and mediates repression by
interacting with chromatin (43) and/or the general
transcriptional machinery (20, 21, 41). In this work we have
focused on the role of Mig1 in regulating repression by Ssn6-Tup1 in
response to the glucose signal.
Mig1 is a Cys2-His2 zinc finger protein
(36) that binds to the promoters of SUC,
GAL, MAL, and other glucose-repressible genes;
mutation of Mig1 or its binding sites partially relieves glucose
repression (15, 17, 22, 25, 35, 36, 44, 53, 55). A LexA-Mig1
fusion protein represses transcription of a CYC1-lacZ
reporter containing lexA operators. Such repression requires
Ssn6-Tup1 and occurs only in glucose-grown cells (47, 51).
Mig1 is differentially phosphorylated in response to glucose availability (11, 47), and the localization of Mig1 to the nucleus requires glucose (11). In contrast, no difference in modification was detected for Ssn6 or Tup1 (46, 57), and
Ssn6 resides in the nucleus regardless of glucose availability
(46). These findings strongly suggest that the recruitment
of Ssn6-Tup1 to a promoter by Mig1 is regulated by glucose. However, it
remains possible that other mechanisms also contribute to regulation of repression by the Mig1-Ssn6-Tup1 complex. Here we present evidence that
LexA-Mig1 confers glucose-regulated repression to a promoter that is
not otherwise glucose repressed, thereby excluding any requirement for
other promoter-bound glucose-regulated factors. We also show that
repression by LexA-Ssn6 is not glucose regulated, indicating that the
repressor function of the Ssn6-Tup1 complex is not directly regulated
by the glucose signal.
The differential phosphorylation of Mig1 in response to glucose
suggests that phosphorylation controls its activity in repression, and
genetic evidence implicates the Snf1 (Cat1) protein kinase in
regulating Mig1. The Snf1 kinase is activated by glucose starvation and
is required for expression of glucose-repressed genes (9, 23, 58,
59). Mig1 is thought to function downstream from Snf1 in the
pathway, because a mig1 mutation suppresses the
snf1 mutant defects in SUC2 and GAL1
expression (25, 53). Thus, Snf1 appears to inhibit
repression by Mig1. Snf1 also inhibits the function of a hybrid
Mig1-VP16 activator in the absence of glucose (37). Deletion
analysis of Mig1 defined regions that both inhibit repression by Mig1
in the absence of glucose and confer inhibition of Mig1-VP16 by Snf1
(37). Finally, mutation of SNF1 causes
constitutive nuclear localization of Mig1 (11).
In this study, we have examined the role of the Snf1 protein kinase in
regulating Mig1 function. We show that Snf1 is required for the
phosphorylation of Mig1 in vivo and that the two proteins interact in
the two-hybrid system and coimmunoprecipitate. We present evidence that
Mig1 is phosphorylated in vitro in a Snf1-dependent reaction. Finally,
we show that mutation of four putative Snf1 recognition sites in Mig1
eliminates most of the differential phosphorylation of Mig1 in response
to glucose and improves the two-hybrid interaction with Snf1.
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MATERIALS AND METHODS |
Strains and genetic methods.
The S. cerevisiae
strains used are listed in Table 1. The
Escherichia coli strains used for propagation of plasmid DNA
were XL1-Blue and DH5
. Standard genetic methods were used, and yeast cultures were grown in synthetic complete (SC) medium lacking appropriate supplements to maintain selection for plasmids
(42).
Oligonucleotides.
Oligonucleotides used for PCR are as
follows, with serine-to-alanine conversions underlined: OL-H1,
5'-ACTACCATAGCCATGGGCGGCCGCCAAAGCCCATATCCAATG-3'; OL-L1,
5'-TCGAGTGCTGTATATAAAACCAGTGGTTATATGTACAGTACC-3'; OL-L2, 5'-TCGAGGTACTGTACATATAACCACTGGTTTTATATACAGCAC-3'; OL-S1,
5'-AATAGCCATAGTGGCGCTAGACTGAAACTGAAC-3'; OL-S2,
5'-ATATTACCAGGTCCGCGAGCTTTAACGGATTTTCAA-3';
OL-S3,
5'-CAGTTGAAGAGACCAGCTGCTGTTTTAAGTTTGAAC-3'; and
OL-S4,
5'-ATGCTAAGTAGAGCTGCTGCTGGTACGAATTTGCAC-3'.
Plasmids.
To construct pHA-Mig1, the
SmaI-KpnI fragment from pMIG1 (36) was
cloned into the cognate sites of pKB174, a derivative of pRS426 lacking
the NotI site (2). The resulting plasmid was
subjected to site-directed mutagenesis with OL-H1 to introduce a
NotI site 3' to the initiating ATG of Mig1. The resulting
DNA was digested with NotI and ligated to a NotI
fragment from pGTEP encoding a triple-hemagglutinin (HA) epitope tag
(50). pHA-Mig1 partially complements a mig1
mutation.
To mutate sites in Mig1, the
BamHI-
SalI fragment
from pLexA-Mig1, a derivative of pSH2-1 (
47), was cloned
into pKB174. Site-directed
mutagenesis was carried out by using
oligonucleotides OL-S1, -S2,
-S3, and -S4. In multiply mutated
constructs, alterations were
added sequentially and confirmed by
restriction digestion or sequence
analysis. To make LexA fusions, the
BamHI-
SalI fragment was then
recloned into pSH2-1
(
19) or pJH106 (pSH2-1 with
URA3 replacing
HIS3). pLexA-Mig1

Z is pLexA-Mig1 with a deletion between
the
EcoRI site in the polylinker and the
XhoI
site at codon 96 (Fig.
1A). pGAD-Mig1
contains the
BamHI-
SalI fragment of pLexA-Mig1
cloned into the same sites of pGAD-Not (
29). An
EcoRI-
SalI fragment
from pGAD-Mig1 was cloned
into pACTII (
28) (
EcoRI at codon 88),
to create
pGAD-Mig1

Z. To construct pGAD-Mig1

ZS222*S278*S311*
and
pGAD-Mig1

ZS278*S311*S381*, an
EcoRI-
SalI
fragment from the
corresponding mutant derivative of pLexA-Mig1 was
cloned into
pACTII. HA-Snf1 and HA-Snf1K84R were expressed from pSK119
and
pSK120, which contain the wild-type and K84R mutant
SNF1
BamHI
fragments from pRJ55 and pRJ215, respectively, cloned into
pWS93,
which expresses a triple-HA epitope from the
ADH1
promoter (a
gift of W. Song, Columbia University). pSK117 is derived
from
pSK37, which is pACTII with the Gal4 activation domain (GAD)
deleted,
and expresses untagged Snf1. Other proteins were expressed
from
the following plasmids: LexA
87, pSH2-1
(
19); LexA-Mig1, pLexA-Mig1
(
47); GAD, pACTII
(
28); GAD-Ssn6, pGAD-Ssn6 (
47); LexA-Ssn6,
CK23
(
27); LexA-Snf1, pRJ55 (
23); LexA-Snf1K84R,
pRJ215 (a
gift of R. Jiang, Columbia University); and HA
3,
pWS93.

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FIG. 1.
(A) Structures of Mig1 and Mig1 Z. Stippled boxes
represent the two zinc fingers. The two regions that are involved in
the inhibition of Mig1 function are shown as hatched boxes, and the
solid box represents the C-terminal 24 amino acids required for
repression (37). Serine residues that are potentially
phosphorylated by Snf1 are marked. (B) Potential Snf1 recognition
sites. The consensus Snf1 recognition sequence (10) is
shown. , hydrophobic residue. Preferred residues: position 5,
L > F = I = M > V; position +4, L > I > F > M > V. Potential Snf1 recognition sites in Mig1 are
shown and are numbered according to the position of the
phosphorylatable serine. The alanine substitutions resulting from
site-directed mutagenesis are indicated, and the mutated sites are
designated by an asterisk.
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pMT27 contains one LexA operator 5' to the
HIS3 upstream
activation sequence (UAS) in pBM2762 (
40).
SalI-digested pBM2762
was ligated to a fragment composed of
two complementary oligonucleotides
(OL-L1 and OL-L2) which recreate a
high-affinity ColE1 LexA binding
site (
12,
26) flanked by
mutant
SalI sites.
All LexA fusions contain the LexA DNA-binding domain,
LexA
87, except LexA-Snf1 fusions, which contain the entire
LexA sequence.
Invertase and
-galactosidase assays.
Invertase activity
was assayed as previously described (7, 16) and expressed as
micromoles of glucose released per minute per 100 mg of cells (dry
weight).
-Galactosidase activity was assayed in permeabilized cells
(42) and expressed in Miller units (32) or was
assayed in protein extracts (8) and expressed as units per
milligram of protein (3).
Immunoblot analysis.
Cells were grown to mid-log phase in
selective SC medium containing 5% glucose (repressed) and derepressed
by a shift to 0.05% glucose for 1 h. Cells were collected by
centrifugation for 2 min and frozen immediately at
70°C without
washing. For Fig. 2C, cells were collected by rapid filtration onto a
0.8-µm-pore-size filter (Micron Separations), and the cell cake was
scraped off into methanol at
80°C. Protein extracts were prepared
as described previously (8). Proteins were separated by
sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE)
and analyzed by immunoblotting. Primary antibodies were polyclonal LexA
antibody (a gift of J. Kamens and R. Brent, Massachusetts General
Hospital, Boston) or monoclonal HA antibody (Boehringer Mannheim
Biochemical). Antibodies were detected by enhanced chemiluminescence
with ECL or ECL Plus reagents (Amersham).
Coimmunoprecipitation assays.
Preparation of protein
extracts and immunoprecipitation procedures were essentially as
described previously (8). The extraction buffer was 50 mM
HEPES (pH 7.5)-150 mM NaCl-0.1% Triton X-100-1 mM
dithiothreitol-10% glycerol, containing 1 or 2 mM
phenylmethylsulfonyl fluoride and complete protease inhibitor cocktail
(Boehringer Mannheim). rProtein A immobilized on Sepharose beads
(RepliGen) was added to protein lysates, which were rotated for 20 min
and then cleared by centrifugation at 12,000 rpm for 10 min. Anti-HA antibody was added, and samples were mixed for 30 min and cleared by
centrifugation for 5 min at 10,000 rpm. The supernatant was mixed with
immobilized rProtein A for 1.5 h. The beads were collected by
brief centrifugation and washed four times with 1 ml of extraction buffer without protease inhibitor cocktail. The entire procedure was
done at 4°C or on ice.
Immune complex kinase assays.
Preparation of protein
extracts and immunoprecipitation were as described above. Beads were
then washed in kinase buffer (50 mM Tris-HCl [pH 7.5], 10 mM
MgCl2, 1 mM dithiothreitol, 0.1% Triton X-100) and
resuspended in 20 µl of kinase buffer. The kinase reaction was
initiated by the addition of 20 µCi of [
-32P]ATP
(3,000 Ci/mmol; NEN). Reaction mixtures were incubated at room
temperature for 30 min, and reactions were terminated by the addition
of 30 µl of 2× sample buffer. Proteins were separated by SDS-PAGE.
After electrophoresis, the gel was stained, washed extensively in
destaining solution containing 10 mM sodium pyrophosphate, dried, and
exposed to film at
70°C with an intensifying screen.
 |
RESULTS |
Glucose-regulated repression by LexA-Mig1.
Previous work
showed that LexA-Mig1 represses transcription of a
lexAop-CYC1-lacZ reporter only in glucose-grown cells and that repression depends on Ssn6-Tup1 (47, 51) (Table
2). These findings suggested that
recruitment of Ssn6-Tup1 by Mig1 is regulated by the glucose signal.
However, the CYC1 promoter responds to glucose, and it
remained possible that other factors bound to this reporter contribute
to the regulation of repression. To address this issue, we tested the
ability of LexA-Mig1 to repress transcription of a reporter driven by
the LEU2 UAS and HIS3 promoter, with no or one
lexA operator 5' to the UAS. In glucose-grown cells, LexA-Mig1 repressed LEU2-HIS3-lacZ expression 11-fold,
whereas in raffinose-grown cells, LexA-Mig1 did not repress
transcription better than LexA87 alone (Table 2); levels of
the LexA-Mig1 protein were comparable (data not shown). Thus,
glucose-regulated repression by LexA-Mig1 is not promoter specific or
dependent on other glucose-regulated factors bound at the reporter.
Mig1 is a likely candidate to mediate the glucose signals controlling
repression by Ssn6-Tup1, because Mig1 functions specifically
at
glucose-regulated promoters, whereas Ssn6-Tup1 serves as a
global
repressor of differently regulated genes. However, no data
exclude the
possibility that the repressor function of Ssn6-Tup1
is also regulated
by glucose. To test this possibility, we compared
the abilities of
LexA-Ssn6 (
27) to repress transcription in
cells grown in
glucose or raffinose (Table
2). LexA-Ssn6 repressed
lexAop-CYC1-lacZ expression 30-fold in glucose-grown cells,
consistent
with previous results (
27). In contrast to
LexA-Mig1, LexA-Ssn6
repressed transcription with comparable efficiency
(27-fold) in
raffinose-grown cells. These findings suggest that
regulation
of repression is achieved solely via regulated recruitment
of
Ssn6-Tup1.
Consistent with these findings, we detected a two-hybrid interaction
(
14) between LexA-Ssn6 and GAD-Mig1

Z, a derivative
lacking the zinc finger DNA-binding domain (Fig.
1A), in glucose-grown
cells but not in raffinose-grown cells (data not shown). The same
results were found with LexA-Mig1 or LexA-Mig1

Z paired with
GAD-Ssn6.
Effect of snf1 on LexA-Mig1
Z function.
Genetic
evidence suggests that the Snf1 protein kinase inhibits repression by
Mig1 during glucose limitation (25, 37, 53). To further
examine the role of Snf1 in regulating Mig1 function, we used
LexA-Mig1
Z, which was stably expressed in a snf1 mutant;
for unknown reasons, LexA-Mig1 was not detectable. In wild-type cells,
LexA-Mig1
Z conferred glucose-dependent, Ssn6-dependent repression of
a reporter (Table 2 and data not shown); thus, the zinc finger domain
is dispensable for regulated repression.
LexA-Mig1

Z repressed transcription of the
CYC1-lacZ
reporter in glucose-grown
snf1 mutant cells, indicating that
the Mig1
repressor function does not require Snf1 (Table
2). To achieve
glucose-limiting conditions, we shifted cells from high to low
(0.05%)
glucose, because a
snf1 mutant does not grow in
nonrepressing
carbon sources. A shift to low glucose did not relieve
repression
of
CYC1-lacZ, consistent with a role for Snf1 in
inhibiting repression
by Mig1 (Table
2).
Requirement for the Snf1 protein kinase in phosphorylation of Mig1
in vivo.
Previously, we showed that LexA-Mig1 is differentially
phosphorylated in response to glucose availability (47)
(Fig. 2A). To test whether the Snf1
protein kinase is required for this phosphorylation, we used immunoblot
analysis to examine tagged Mig1 proteins in a snf1 mutant.
In wild-type cells, LexA-Mig1
Z was also differentially modified,
similarly to LexA-Mig1 (Fig. 2A). In snf1 mutant cells, however, the major band migrated at the position predicted for the
unmodified protein in both glucose-repressed and derepressed cells
(Fig. 2A). We next examined the HA-Mig1 protein expressed from its own
promoter. HA-Mig1 was differentially modified in wild-type cells but
not in snf1 mutants (Fig. 2B). In control experiments, a
snf1 mutation did not alter the phosphorylation of an
unrelated protein encoded by SFH1 (4) (data not
shown). Thus, the Snf1 protein kinase is required for the
phosphorylation of Mig1.

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FIG. 2.
Immunoblot analysis of Mig1 fusion proteins. Cultures
were grown in selective SC medium plus 5% glucose (repressed) (lanes
R). Mid-log-phase cultures were derepressed by a shift to SC medium
plus 0.05% glucose for 1 h (lanes D). Extracts were prepared, and
proteins were separated by SDS-PAGE in 7.5% polyacrylamide and
subjected to immunoblot analysis with anti-LexA (A, C, and D) or
anti-HA (B). (A and B) Protein extracts (25 and 50 µg for wild-type
[WT] and snf1-K84R strains, respectively) were prepared
from strains MCY829 and MCY2692 transformed by pLexA-Mig1 or
pLexA-Mig1 Z (A) and pHA-Mig1 (B). (C) Protein extracts (25 µg)
were prepared from strain YM4738 expressing LexA-Mig1 from vector
pJH106. Cells were collected by rapid membrane filtration (Filt.) or by
centrifugation for 2 min (Cent.) (see Materials and Methods). (D)
Protein extracts (25 µg for the wild type and 50 µg for
reg1 and hxk2 strains) were prepared from strains
FY250, MCY829, MCY3278, and MCY3541 transformed with pLexA-Mig1. The
position of the 66-kDa size marker is indicated.
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The increased phosphorylation of Mig1 that occurs upon glucose
deprivation is compatible with evidence that the Snf1 kinase
is more
active in glucose-deprived cells (
58,
59). However,
the
phosphorylation observed in glucose-grown cells could result
from
partial derepression during sample preparation. Because Wilson
et al.
(
58) reported that harvesting of glucose-grown cells
by
rapid membrane filtration, followed by freezing, minimizes
activation
of the Snf1 kinase, we also examined LexA-Mig1 from
glucose-grown cells
prepared in this manner. The pattern was the
same as that obtained with
our usual procedure (centrifugation
for 2 min without washing, followed
by freezing) (Fig.
2C). These
findings suggest that the Snf1-dependent
phosphorylation observed
in glucose-grown cells is unlikely to result
from partial derepression,
but they cannot exclude this possibility.
In addition, these results strongly suggest that phosphorylation of
Mig1 does not occur simply as a consequence of transcriptional
activation of an adjacent promoter. LexA-Mig1

Z, which lacks the
native DNA-binding domain, is still modified in cells with no
LexA
binding sites.
Phosphorylation of Mig1 in reg1 and hxk2
mutants.
We next examined mutants in which Snf1 is active in
glucose-grown cells. If differential phosphorylation of Mig1 reflects the functional status of Snf1, then mutations that affect the regulation of Snf1 should also influence Mig1 phosphorylation. The
REG1 gene encodes a targeting subunit that directs the
function of protein phosphatase 1 in the glucose response
(49). Mutation of REG1 relieves glucose
repression of Snf1-dependent genes (24) and causes the Snf1
protein kinase complex to assume an active conformation even in
glucose-grown cells (23, 30). Mutation of HXK2,
encoding hexokinase PII, causes similar phenotypes (23, 24).
Immunoblot analysis showed that the LexA-Mig1 species present in
glucose-grown
reg1 and
hxk2 mutants are similar
to those
found in derepressed cells (Fig.
2D). These results indicate
that
Reg1 and hexokinase PII affect phosphorylation of Mig1 in a manner
consistent with their roles in modulating Snf1 kinase activity.
Two-hybrid interaction between Mig1 and both wild-type and
kinase-dead Snf1 proteins.
The preceding data show that Snf1 is
required for phosphorylation of Mig1 but do not address whether Snf1
phosphorylates Mig1 directly or controls the phosphorylation of Mig1 by
another kinase. To assess the interaction of Snf1 with Mig1 in vivo, we
used the two-hybrid system (14). In glucose-grown cells,
LexA-Snf1 did not interact significantly with GAD-Mig1
Z, but
LexA-Snf1K84R interacted strongly (Table
3). The mutant Snf1K84R protein contains a substitution of Arg for the conserved Lys84 in the ATP binding site
and exhibits no catalytic activity (8). After cells were shifted to 0.05% glucose for 3 h,
-galactosidase activity
could be detected for both combinations. When cells were grown in
raffinose, no significant interaction was detected (data not shown),
consistent with evidence that Mig1 is cytoplasmic under derepressing
conditions (11). These data support the view that Mig1 is a
substrate of Snf1 in vivo and suggest that for the wild-type Snf1,
glucose deprivation transiently enhances interaction with Mig1.
Coimmunoprecipitation of Snf1 and Mig1.
To obtain biochemical
evidence for the interaction of Snf1 and Mig1 in vivo, we tested
LexA-Snf1 for coimmunoprecipitation with HA-Mig1, expressed from the
MIG1 promoter (Fig. 3).
Whole-cell extracts were prepared from cells expressing both proteins,
and HA-Mig1 was immunoprecipitated with monoclonal anti-HA antibody. Immunoblot analysis of the precipitate showed that LexA-Snf1
coprecipitated; it was expected that only a small fraction of the
LexA-Snf1 would be associated with Mig1. In control experiments,
LexA-Snf1 was not detected when the extract contained HA instead of
HA-Mig1. Similar results were obtained with LexA-Snf1K84R (Fig. 3); the mutant protein did not coprecipitate better than the wild-type LexA-Snf1, probably because the two-hybrid system and
coimmunoprecipitation are not comparable assays.

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FIG. 3.
Coimmunoprecipitation of LexA-Snf1 and LexA-Snf1K84R
with HA-Mig1. Strains MCY3573 and MCY3640 were transformed with
plasmids expressing LexA-Snf1 and LexA-Snf1K84R, respectively, and
either HA-Mig1 (from the MIG1 promoter) or HA (from the
ADH1 promoter). Protein extracts were prepared from cells
grown in glucose, and proteins (200 µg) were immunoprecipitated (IP)
with anti ( )-HA antibody, separated by SDS-PAGE in 7.5%
polyacrylamide, and immunoblotted with -LexA (upper panel). The
input protein (25 µg) is also shown (middle panel); an arrow
indicates the position of the Snf1 fusion. The same immunoblot was
reprobed with -HA to confirm the precipitation of HA-Mig1 (lower
panel). No band was detected at the position of LexA-Snf1 when HA-Mig1
was immunoprecipitated from extracts lacking LexA-Snf1. WT, wild
type.
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Snf1-dependent phosphorylation of Mig1 in vitro.
We next
addressed the ability of Snf1 to phosphorylate Mig1 in vitro. Extracts
were prepared from cells expressing HA-Snf1 and LexA-Mig1 from the
ADH1 promoter, and HA-Snf1 was immunoprecipitated with
anti-HA. The immune complexes were resuspended in kinase assay buffer
and incubated with [
-32P]ATP. The proteins were
separated by gel electrophoresis, and the phosphorylated products were
visualized by autoradiography (Fig. 4A).
In addition to the products usually detected in such assays, including
Snf1, Sip1, and Gal83 (60), a phosphorylated protein
corresponding to LexA-Mig1 was detected (Fig. 4A, lane 1). This product
was absent in assays of extracts containing only the LexA moiety
(expressed from the parental vector) (Fig. 4A, lane 3). Control
experiments with the kinase-dead HA-Snf1K84R mutant protein confirmed
that the kinase activity detected in this assay was dependent on Snf1
(Fig. 4A, lane 2), and no phosphorylated LexA-Mig1 was detected even
upon overexposure (Fig. 4A, lanes 4 and 5). In an independent
experiment, we similarly detected phosphorylation of LexA-Mig1 in
immune complex assays of the wild-type HA-Snf1, but not the mutant
kinase, and also showed that no phosphorylation was detected in
controls with untagged Snf1 expressed at the same level (data not
shown). Thus, LexA-Mig1 is phosphorylated in vitro in a Snf1-dependent
reaction.

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FIG. 4.
Snf1-dependent phosphorylation of Mig1 in vitro. Strain
FY250 (wild type [WT]) was transformed with plasmids expressing
HA-Snf1 or HA-Snf1K84R and LexA-Mig1 or the LexA moiety (from the
parental vector). Protein extracts were prepared from cells grown in
selective SC medium plus 2% glucose; previous studies have shown that
the Snf1 kinase is activated during preparation of the extracts
(13, 58). (A and B) Proteins (200 µg) were
immunoprecipitated with anti-HA antibody. (A) Half of the
immunoprecipitate was resuspended in kinase assay buffer and incubated
with [ -32P]ATP. Proteins were then separated by
SDS-PAGE in 8% polyacrylamide, and the gel was subjected to
autoradiography to detect phosphorylated products. The left panel
(lanes 1 to 3) shows a 3.5-h exposure, and the right panel (lanes 4 and
5) shows a 15-h exposure of lanes 1 and 2. (B) The remaining half of
the sample was subjected to SDS-PAGE and immunoblot analysis with
anti-HA to confirm the immunoprecipitation of HA-Snf1 and HA-Snf1K84R.
We could not reproducibly detect LexA-Mig1 by immunoblot analysis of
the immunoprecipitates. (C) Input proteins (10 µg) were analyzed by
immunoblot analysis to verify the expression of LexA-Mig1. The
phosphorylated LexA-Mig1 product detected in panel A corresponds to the
lower band in this panel. Numbers on the right of each panel are
molecular masses in kilodaltons.
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Mutation of putative Snf1 phosphorylation sites in Mig1.
We
identified potential Snf1 phosphorylation sites in Mig1 based on their
similarity to the consensus substrate recognition sequence
(10), which contains an arginine at position
3 and hydrophobic residues at positions
5 and +4 relative to the
phosphorylated serine (Fig. 1B). This consensus sequence was determined
by assaying the ability of purified Snf1 kinase to phosphorylate
variants of a synthetic peptide that is recognized by the mammalian
Snf1 homolog, AMP-activated protein kinase (5, 33).
Three sites in Mig1, containing serine residues S278, S311, and S381,
match the consensus site, and the site at S222 resembles
the consensus
(Fig.
1B). S278 and S311 are located in a regulatory
region that
appears to mediate the inhibition of Mig1 function
by Snf1
(
37), and residues 261 to 400 are sufficient to confer
glucose-regulated nuclear localization (
11). The sites at
S222,
S278, and S311 are conserved in Mig1 homologs from the yeasts
Kluyveromyces marxianus and
Kluyveromyces lactis
(
6).
We converted the phosphorylatable serine residues to alanine by
site-directed mutagenesis and also altered any adjacent serines
or
threonines (Fig.
1B). The mutant sites are designated with
an asterisk.
We constructed several combinations of mutant sites
including S278* and
S311*, because these two sites match the consensus
sequence, reside
within a regulatory region, and are conserved.
The mutant Mig1 proteins
were expressed as LexA fusions to facilitate
detection of the proteins
and assays of repressor function.
Immunoblot analysis indicated that mutation of all four sites
eliminated most of the differential phosphorylation of Mig1
in response
to glucose (Fig.
5A). Although the
triple-mutant proteins
still displayed a glucose-dependent shift in
mobility, the shift
was less pronounced, and the same was true for the
S278*S311*
double mutant (data not shown). Thus, these sites appear to
be
phosphorylated in vivo. However, mutation of these sites did not
reduce phosphorylation of Mig1 as substantially as did mutation
of
SNF1, suggesting that Mig1 also contains additional sites
for
Snf1-dependent phosphorylation. We were unable to assess the effect
of these mutations on phosphorylation by Snf1 in immune complex
assays
due to the low levels of the mutant proteins.

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|
FIG. 5.
Analysis of mutant LexA-Mig1 fusion proteins. (A) Strain
MCY3640 was transformed with plasmids derived from the vector pJH106.
Cultures were grown selectively in SC medium plus 5% glucose
(repressed) (lanes R) and were derepressed by a shift to SC medium plus
0.05% glucose for 1 h (lanes D), and protein extracts were
prepared. Proteins (25 µg for cells expressing wild-type LexA-Mig1
[WT] and 50 µg for cells expressing mutant LexA-Mig1 proteins) were
separated by SDS-PAGE in 7.5% polyacrylamide and subjected to
immunoblot analysis with anti-LexA antibody. The position of the 66-kDa
size marker is indicated. (B) Transformants of strains MCY3640
(mig1 ) (shaded bars) and YM4738 (mig1
mig2 ) (open bars) expressing the indicated LexA-Mig1 protein
were grown selectively in SC medium plus 2% raffinose and 0.05%
glucose. Derepression of SUC2 was monitored by assaying
invertase activity for 4 to 11 transformants. Standard errors were
<13%. Immunoblot analysis confirmed that mutant proteins were present
at lower levels than wild-type LexA-Mig1 in strain YM4738, consistent
with panel A. Similar assays of transformants shifted from 5 to 0.05%
glucose for 1 h did not reveal any delay in derepression for
strains expressing mutant proteins.
|
|
None of the mutated sites is essential for Mig1 repressor function.
LexA-Mig1 proteins containing these mutations restored
glucose
repression of
SUC2 in a
mig1
mig2
mutant
(Table
4);
Mig2 is a related zinc finger
repressor protein that assists Mig1
(
31). The mutant
LexA-Mig1 proteins also repressed transcription
of the
lexAop-CYC1-lacZ reporter in glucose-grown cells (Table
4).
We then assayed for release of repression by the mutant LexA-Mig1
proteins in response to glucose limitation. Derepressed
invertase
activities of
mig1
and
mig1
mig2
cells
expressing
the mutant proteins were 1.6- to 2.8-fold and 1.4- to
1.7-fold
lower, respectively, than that of cells expressing the
wild-type
LexA-Mig1 (Fig.
5B). The levels of the mutant LexA-Mig1
proteins
were reproducibly lower than that of the wild-type protein
(twice
as much protein was loaded for the mutant extracts in Fig.
5A;
data not shown for
mig1
mig2
transformants). In view
of the
decreased abundance of the mutant proteins, the effects of the
mutations on release of repression of
SUC2 may be
substantial.
Protein levels are likely to be important, because
overexpression
of wild-type LexA-Mig1 reduced derepression relative to
the case
for the control with LexA.
Finally, we determined the effects of these mutations on the two-hybrid
interaction of Mig1 with Snf1. We reasoned that if
a mutation
abolishing the Snf1 catalytic activity (K84R) improves
detection of
this interaction, then mutations that prevent phosphorylation
of Snf1
recognition sites might also affect interaction. In glucose-grown
cells, mutant derivatives of GAD-Mig1

Z showed weak interaction
with
LexA-Snf1; however, substantial

-galactosidase activity
was produced
after a shift to low glucose, which activates the
Snf1 kinase. The two
mutant proteins interacted strongly with
Snf1, producing 1,110 and 810 U of activity, compared to 72 U
for the wild-type GAD-Mig1

Z (Table
3). Thus, Ser-to-Ala substitutions
in these putative Snf1 recognition
sites increased the two-hybrid
interaction between Mig1 and Snf1 11- to
15-fold.
 |
DISCUSSION |
Previous evidence suggested that Mig1 recruits the Ssn6-Tup1
corepressor to glucose-repressed promoters in response to the glucose
signal. Here we have further examined the role of Mig1 in regulating
repression. First, we show that LexA-Mig1 confers glucose-regulated
repression to a LEU2-HIS3-lacZ reporter, thereby excluding
any requirement for other regulatory factors specific to
glucose-regulated reporters. Similar studies of LexA-Mig1
Z further
indicate that the zinc finger region is not required for regulated
repression. Second, we show that repression by LexA-Ssn6 is not
regulated by glucose. These experiments substantiate the model that
regulation is achieved by the regulated recruitment of Ssn6-Tup1 by
Mig1.
The differential phosphorylation of Mig1 in response to glucose
suggested that phosphorylation regulates its repressor function (11, 47), and genetic evidence indicated that the Snf1
protein kinase inhibits Mig1 function during glucose starvation
(25, 37, 53). Here we present evidence that Snf1 regulates
the phosphorylation of Mig1. We show that modification of Mig1 is dramatically reduced in a snf1 mutant, indicating that Snf1
is required for the phosphorylation of Mig1. Consistent with these observations, Snf1 kinase activity increases in glucose-limited cells
(58, 59). Conversely, in glucose-grown hxk2 and
reg1 mutants, which are defective in glucose inhibition of
the Snf1 kinase activity (23) and glucose repression of
Snf1-dependent genes (24), the migration patterns of
LexA-Mig1 resemble that of the derepressed wild type. These data
strongly suggest that the regulation of Snf1 kinase activity is coupled
to the regulation of Mig1 modification, with the caveat that
hxk2 and reg1 may also affect Mig1 by other
mechanisms. During the preparation of this paper, the Snf1-dependent
phosphorylation of a Mig1-VP16 protein containing the N-terminal
two-thirds of Mig1 (residues 1 to 351) was reported; however, this
truncated Mig1 fusion differs from the full-length Mig1 proteins
examined here in that it is not phosphorylated in glucose-grown cells
(38).
Several lines of genetic and biochemical evidence support the view that
Snf1 phosphorylates Mig1 in vivo. First, Snf1 and Mig1 interact in the
two-hybrid system. Moreover, the kinase-dead mutant Snf1K84R gives a
stronger signal than wild-type Snf1, and mutant Mig1 proteins with
Ser-to-Ala substitutions in consensus Snf1 recognition sites give a
stronger signal than wild-type Mig1. A shift to low glucose causes an
increase in interaction between wild-type Snf1 and Mig1, presumably
transient because no interaction was detected in cells grown in
raffinose. Second, Snf1 coimmunoprecipitates with Mig1 from cell
extracts. Third, mutation of all four putative Snf1 recognition sites
eliminates most of the differential phosphorylation of Mig1 in response
to glucose. Finally, functional assays of the mutant LexA-Mig1 proteins
revealed defects of up to 2.8-fold in release of repression of
SUC2, and the magnitude is most likely underestimated due to
the reduced levels of the mutant proteins. Studies of Mig1-VP16
similarly showed that mutation of serines 278, 310, and 311 affects its
phosphorylation and reduces the Snf1 dependence of its activation
function 3.8-fold, although protein levels were not reported
(38).
These studies of the relationship of Snf1 and Mig1 in vivo are further
supported by in vitro evidence that immunoprecipitated Snf1 kinase
phosphorylates coprecipitated Lex-Mig1. This reaction was dependent on
Snf1 activity, and no phosphorylation was detected in immune complex
assays of Snf1K84R. The simple interpretation is that Mig1 is
phosphorylated by Snf1, but a more complicated scenario, in which Snf1
is still intimately involved in the phosphorylation of Mig1, cannot be
excluded. It is possible that Snf1 phosphorylates and activates an
associated Snf1-dependent kinase, which then phosphorylates Mig1;
however, this model is difficult to reconcile with the effects of
mutations in Mig1 on its phosphorylation and two-hybrid interaction
with Snf1.
Although most of the phosphorylation of Mig1 in vivo depends on the
Snf1 kinase, Snf1 may not be directly responsible for all of the
phosphorylation events. Mutation of all four Snf1 consensus recognition
sites did not reduce phosphorylation of Mig1 nearly as substantially as
the snf1 mutation. Mig1 may contain other Snf1 recognition
sites, unrelated to the defined consensus, and/or Snf1 may regulate the
phosphorylation of Mig1 by another protein kinase. Consistent with this
view, analysis of Mig1-VP16 identified a Snf1-dependent phosphorylation
site at serine 108, which does not resemble a Snf1 site
(38).
Phosphorylation of Mig1 could regulate the recruitment of Ssn6-Tup1 to
a promoter by affecting any of several steps: binding of Mig1 to the
promoter, interaction of Mig1 with the Ssn6-Tup1 corepressor, or
localization of Mig1 to the nucleus. It is unlikely that DNA binding is
regulated, because regulated repression was achieved by LexA-Mig1 bound
to lexA operators. The possibility that phosphorylation
disrupts the interaction of Mig1 with Ssn6-Tup1 has not been addressed,
but this cannot be the only mechanism, because Snf1 affects activation
by Mig1-VP16 (37) and affects localization of Mig1 in an
ssn6 mutant (11). Evidence that the differential
localization of Mig1 is Snf1 dependent and correlates with its
differential phosphorylation (11) strongly suggests that
phosphorylation functions as a regulatory signal for localization.
 |
ACKNOWLEDGMENTS |
We thank Aaron Mitchell, Rod Rothstein, and Brehon Laurent for
fruitful discussion and members of the Carlson lab, especially Rong
Jiang, for critical input. We thank Lillian Ho for technical assistance. We are grateful to David Carling and Grahame Hardie for
communication of unpublished results on phosphorylation of Mig1 in
vitro.
This work was supported by grant GM34095 from the National Institutes
of Health to M.C.
 |
FOOTNOTES |
*
Corresponding author. Mailing address: HHSC 922, Box
136, 701 W. 168th St., New York, NY 10032. Phone: (212) 305-6314. Fax: (212) 305-1741. E-mail: mbcl{at}columbia.edu.
 |
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