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Mol Cell Biol. Jan 2004; 24(1): 352–361.
PMCID: PMC303368

Mutations in the Gal83 Glycogen-Binding Domain Activate the Snf1/Gal83 Kinase Pathway by a Glycogen-Independent Mechanism


The yeast Snf1 kinase and its mammalian ortholog, AMP-activated protein kinase (AMPK), regulate responses to metabolic stress. Previous studies identified a glycogen-binding domain in the AMPK β1 subunit, and the sequence is conserved in the Snf1 kinase β subunits Gal83 and Sip2. Here we use genetic analysis to assess the role of this domain in vivo. Alteration of Gal83 at residues that are important for glycogen binding of AMPK β1 abolished glycogen binding in vitro and caused diverse phenotypes in vivo. Various Snf1/Gal83-dependent processes were upregulated, including glycogen accumulation, expression of RNAs encoding glycogen synthase, haploid invasive growth, the transcriptional activator function of Sip4, and activation of the carbon source-responsive promoter element. Moreover, the glycogen-binding domain mutations conferred transcriptional regulatory phenotypes even in the absence of glycogen, as determined by analysis of a mutant strain lacking glycogen synthase. Thus, mutation of the glycogen-binding domain of Gal83 positively affects Snf1/Gal83 kinase function by a mechanism that is independent of glycogen binding.

The Saccharomyces cerevisiae Snf1 kinase and mammalian AMP-activated protein kinase (AMPK) are highly conserved kinases with roles in metabolic stress responses (for review, see references 14 and 24). Both Snf1 and AMPK monitor nutritional status. AMPK is activated by increases in AMP during metabolic stress, by the hormones leptin and adiponectin (32, 51), and by drugs used in the treatment of type 2 diabetes (52). Snf1 is also activated by stresses, notably glucose limitation (22, 29, 48, 50). Snf1 and AMPK regulate glucose and lipid metabolism both by controlling the activity of metabolic enzymes and by controlling transcription. Snf1 is required for the expression of glucose-repressed genes involved in respiration, gluconeogenesis, peroxisome biogenesis, and metabolism of alternate carbon sources (4, 12).

Snf1 and AMPK are heterotrimeric kinases with multiple subunit isoforms. Snf1 kinase comprises the catalytic subunit Snf1, a β subunit (Gal83, Sip1, or Sip2), and the regulatory subunit Snf4. Snf1 kinase containing the Gal83 β subunit will be referred to here as Snf1/Gal83 kinase. For AMPK, the corresponding subunit isoforms are designated α1, α2, β1, β2, γ1, γ2, and γ3. Individual β subunits of Snf1 and AMPK have distinct subcellular localizations (44, 47) and thereby presumably regulate access of the kinase to substrates. The Gal83 β subunit is known to mediate interaction of the kinase with Sip4, a transcriptional activator of gluconeogenic genes that is dependent on Snf1 kinase for its function (27, 38, 42).

Recent studies identified a glycogen-binding domain in the AMPK β1 subunit that is related to isoamylase domains found in glycogen and starch branching enzymes (21, 34). Mutation of conserved residues abolishes binding to glycogen in vitro (34). In mammalian cells, AMPK β1 colocalizes with glycogen phosphorylase (34) and glycogen synthase (21); with the caveat that AMPK β1 was overexpressed, these results suggest that AMPK binds to glycogen. Addition of glycogen does not affect AMPK catalytic activity in vitro (34). Previous biochemical and genetic evidence has implicated AMPK in regulation of glycogen metabolism, which in mammalian systems is under complex control by hormones and nutritional signals (36). Mutations of the γ subunit affect glycogen storage in pigs (5, 30) and cause a glycogen storage disease associated with cardiac abnormalities in humans (1). However, the physiological roles of the glycogen-binding domain of the β1 subunit in AMPK function and glycogen metabolism remain unclear.

Glycogen is also an important storage carbohydrate in yeast (for review, see reference 11). Levels are low during exponential growth on glucose and rise rapidly before the onset of stationary phase, and the accumulated glycogen is then degraded slowly during prolonged starvation. Snf1 is one of several kinases that regulate glycogen metabolism, and Snf1 is required for glycogen accumulation (3, 19, 20, 41), regulation of the expression and catalytic activity of glycogen synthase (15, 49), and maintenance of glycogen stores (46).

We have taken advantage of yeast genetics to investigate the roles of the glycogen-binding domains of the β subunits in vivo. The sequence of the glycogen-binding domain is conserved in two of the yeast β subunits, Sip2 and Gal83. We found that Gal83 binds glycogen strongly in vitro, whereas Sip2 binds very weakly. We mutated conserved residues that are essential for glycogen binding of AMPK β1 (34) and also analyzed another mutation that maps to a conserved residue (G235) within the glycogen-binding domain of Gal83 (9, 28). Alteration of Gal83 abolished glycogen binding in vitro and caused diverse phenotypes in vivo, positively affecting not only accumulation of glycogen but also transcriptional regulation. Surprisingly, the transcriptional regulatory phenotypes did not depend on the presence of glycogen within the cell, and the differences between the mutant and wild-type Gal83 were manifest even in a mutant strain (gsy1Δ gsy2Δ) lacking glycogen synthase. Thus, mutation of the glycogen-binding domain positively affects Snf1/Gal83 kinase function by a mechanism that is independent of glycogen binding.


Strains and genetic methods.

S. cerevisiae strains used in this study are listed in Table Table1.1. Strain MCY4099 has the W303 genetic background; gal83Δ::TRP1 (42) and sip1Δ::kanMX6 (44) have been described before, and the sip2Δ::kanMX4 allele was amplified by PCR from strain BY24574. MCY4101 was derived from strains BY4741, BY15694, and BY15167 by genetic crossing, and the genotype was verified by using PCR. MCY4622 and MCY4626 are derivatives of Σ1278b; the gal83Δ allele has been described elsewhere (45). Rich medium was yeast extract-peptone-dextrose, and synthetic (SD) and synthetic complete (SC) media contained appropriate supplementation to maintain selection for plasmids (37).

S. cerevisiae strains used in this study


pOV22 and pPL50 (43), pRJ55 (22), and pRJ217 (42) were described previously. Plasmids used to express Sip2 or Gal83 contained the native promoter (~700 bp) and the native terminator (~500 bp). Proteins were tagged at the C terminus with green fluorescent protein (GFP).

To introduce the R214Q mutation into pRT12, which expresses Gal83-GFP (44), PCR was performed using pRT12 as a template. Oligonucleotide Gal831A (5′-CCGATCTCGTAGGATTTGGG-3′) and oligonucleotide Gal831B (5′-CTGAATCTTAACTCATTGTCAACAATAAACTGGAAACGATGAGTACCTGGAGGC-3′) were used to generate a mutated fragment corresponding to nucleotides −119 to +671. Letters in bold indicate altered nucleotides. Oligonucleotides Gal832A (5′-GCCTCCAGGTACTCATCGTTTCCAGTTTATTGTTGACAATGAGTTAAGA-3′) and Gal832B (5′-CAGTATTTGGGTCACGTATTTTG-3′) were used to generate a mutated fragment corresponding to nucleotides 618 to 1233. These two fragments were cotransformed into yeast with pRT12 gapped with XcmI and NarI. The recombinant plasmid pHW33 was recovered in Escherichia coli and sequenced. We next introduced the W184A mutation by PCRs using pHW33 as a template. Oligonucleotide Gal831C (5′-CATAAGTCCAGGCTGTCCAGGGACTGGTACTAACCCGATCATCTTTCTCGCTCCGTAAAAGACCCAGT-3′) and oligonucleotide Gal831A were used to generate a mutated fragment from nucleotides −119 to 600. Oligonucleotides Gal832C (5′-GGGGGGTAATAAAGTGTACGTTACTGGGTCTTTTACGGGAGCGAGAAAGATGATCGGGTTAGTACC-3′) and Gal832B were used to generate a mutated fragment from nucleotides 510 to 1233 of the GAL83-R214Q sequence. These two fragments were used as described above to generate pHW39. pRT13, pHW40, and pHW41 are identical to pRT12, pHW39, and pOV81, respectively, except that the vector is pRS315 (39).

pHW30 is identical to pRT9, which expresses Sip2-GFP (44), except that the vector is pRS316 (39). To introduce the R216Q mutation, a mutagenic PCR was performed using pRT9 as a template. OL65 and oligonucleotide Sip21B (5′-CTCTAAGCTCATTATCCACTATAAACTGGAATCTATGTGTGCCTGGAAGC-3′) were used to generate a mutated fragment corresponding to nucleotides −711 to 673. Oligonucleotide Sip22A (5′-CTAAGGCTGCTTCCAGGCACACATAGATTCCAGTTTATAGTGGATAATGAGCTTAGAG-3′) and oligonucleotide Sip22B (5′-CGAACGATGGAGGCTACACAAAGTG-3′) were used to generate a mutated fragment corresponding to nucleotides 615 to 1193. These two fragments were cotransformed into yeast with pHW30 gapped with MluI and BglII. pHW35 was recovered in E. coli and sequenced. We then introduced the W186A mutation by PCR using pHW35 as a template. OL65 and oligonucleotide Sip21C (5′-GCCATTATTGTCAGAATCAGGTATCAAACCGATCATTTTCCTCGCTTTGGTTGAATGAGCCTGTCAC-3′) were used to generate a mutated fragment spanning nucleotides −711 to 590. Oligonucleotides Sip22C (5′-GGTGGTTCAAAAGTTTACGTGACAGGCTCATTCACCAAAGCGAGGAAAATGATCGGTTTCAT-3′) and Sip22B were used to generate a mutated fragment corresponding to nucleotides 517 to 1193 of the SIP2-R216Q sequence. These two fragments were used as described above to generate pHW38. pOV81 is a derivative of pRT12 containing the GAL83-G235R sequence from pRJ324 (23).

To express the glycogen-binding domains of Gal83 and Sip2 in bacteria, DNA sequences corresponding to codons 152 to 251 and 154 to 253, respectively, were obtained by PCR using pRT12, pHW30, pHW38, pHW39, and pOV81 as templates. The amplified DNAs were cloned into the EcoRI/HindIII sites of the pProEX HT vector (Invitrogen), and clones were confirmed by sequencing.

Bacterial expression of the glycogen-binding domains of Gal83 and Sip2.

Proteins corresponding to the glycogen-binding domains were expressed as His6 fusion proteins in BL21 cells. Cultures were grown to an optical density at 600 nm (OD600) of 0.6, and protein expression was induced with 1 mM isopropyl-β-d-thiogalactopyranoside for 3 h at 37°C. Cells were harvested by centrifugation and lysed in phosphate-buffered saline (PBS) using an Emulsiflex-C5 high-pressure homogenizer (Avestin). Lysates were clarified by centrifugation and chromatographed on Ni-agarose. Nonspecific proteins were removed by washing the Ni-agarose with PBS containing 20 mM imidazole. Gal83 and Sip2 proteins were eluted with PBS containing 300 mM imidazole, concentrated to 0.5 ml, and purified by gel filtration on G-75 Sephadex.

Assay of glycogen binding.

In vitro glycogen-binding assays were performed as described previously (34). Briefly, purified glycogen-binding domains of Gal83 or Sip2 (5 μg each) were incubated with 0.4% (wt/vol) bovine liver glycogen (type IX; Sigma) in a total volume of 0.25 ml in PBS and 0.5% Triton X-100 for 30 min at 4°C with mixing, followed by centrifugation at 200,000 × g for 60 min at 4°C. The supernatant was removed, and the pellet was briefly washed with 500 μl of PBS prior to resuspension to the original volume (0.25 ml). Gal83- or Sip2-containing fractions were identified by electrophoresis of proteins in Tris-tricine gels and immunoblotting with an anti-His6 tag antibody (Rockland Immunochemicals).

Assay of glycogen content.

Glycogen assays were performed as described previously (33) with minor modifications. Mid-log cultures grown in selective SD plus 2% glucose were used to inoculate 25 to 100 ml of selective SD plus 2% glucose to an OD600 of 0.06. During growth of the culture, aliquots of cells (0.5 to 10 ml) were harvested by centrifugation and stored at −70°C. Cells were then resuspended in 25 μl of 0.25 M sodium carbonate, covered with 35 μl of mineral oil, and placed in a heating block at 95°C for 4 h. Fifteen microliters of 1 M acetic acid was added to neutralize the base, and then 60 μl of 0.2 M sodium acetate, pH 5.2, was added. Aspergillus niger amyloglucosidase (catalog number A-7420; Sigma) was added to a final concentration of 6.7 U/ml, and the mixture was incubated at 57°C for 12 h. The amount of glucose released was quantified using the glucose oxidase reaction (13).

Immunoblot analysis.

Protein extracts were prepared for immunoblot analysis as described elsewhere (42). Proteins were separated by sodium dodecyl sulfate (SDS)-polyacrylamide gel electrophoresis in 8% polyacrylamide and detected with anti-GFP antibody (Invitrogen) and ECL Plus reagents (Amersham).


Cells were grown to mid-log phase in SD plus 2% glucose. Nuclei were stained by addition of 4′,6-diamidino-2-phenylindole (DAPI) (0.8 μg/ml) for 5 min. Cells were collected by centrifugation and resuspended in ~20 μl of residual medium. A 1.4-μl aliquot of the cell suspension was placed on a microscope slide. Cells were viewed using a Nikon Eclipse E800 fluorescence microscope. Images were taken with an Orca 100 camera (Hamamatsu) using Open Lab software (Improvision) and were processed in Adobe PhotoShop 5.5.

Kinase assays.

Immunoprecipitation and kinase assays were performed as described previously (26), except that cultures were grown selectively in SD plus 2% glucose. Polyclonal anti-LexA antibody was purchased from Invitrogen. For assays of kinase activity by phosphorylation of the SAMS peptide (HMRSA MSGLHLVKRR) substrate, cells were grown in SC plus 2% glucose to an OD600 of 0.7, collected by filtration, incubated in SC plus 0.5% glucose-2% glycerol-2% ethanol for 30 min, and then collected by filtration. Extracts were prepared and assays were performed as described previously (18). Briefly, Snf1 kinase was partially purified by chromatography on DEAE-Sepharose (Amersham Biosciences), and pooled fractions were assayed in triplicate for phosphorylation of the SAMS peptide in the presence of [γ-32P]ATP (8).

β-Galactosidase assays.

Transformants were grown to mid-log phase in selective SC or SD plus 2% glucose and were shifted to SC or SD plus 2% glycerol plus 2% ethanol for the time indicated. β-Galactosidase activity was assayed in permeabilized cells and is expressed in Miller units (31).

Preparation of RNA and Northern blot analysis.

Cells (50 ml) were collected by filtration, resuspended in 0.7 ml of TES (10 mM Tris-HCl [pH 7.5], 10 mM EDTA, 0.5% SDS), and frozen in liquid nitrogen. An equal volume of acid phenol was added to the sample, and cells were incubated at 65°C for 1.5 h with vortexing for 30 s 10 times at 3-min intervals and then for 30 s every 10 min. Samples were extracted four times with an equal volume of phenol and then twice with chloroform. RNA was precipitated with ethanol and resuspended in water. RNAs (40 μg) were separated by electrophoresis on a 1.2% agarose-morpholinepropanesulfonic acid gel containing formaldehyde and were transferred to a Hybond N+ membrane (Amersham Pharmacia Biotech). Probes were 32P-labeled using Ready-To-Go DNA labeling beads (Amersham Biosciences). The U3 small nucleolar RNA, encoded by SNR17A and SNR17B, was used as a loading control. GSY1 and GSY2 probes encompassed nucleotides 77 to 675 of the coding sequence.

Invasive growth assay.

Transformants were streaked for single colonies on selective SC plus 2% glucose. Single colonies were resuspended in 100 μl of water, and 5 μl was spotted onto a yeast extract-peptone-dextrose plate. After 2 days at 30°C, plates were photographed, then placed under a gentle stream of tap water, rubbed with a gloved finger, and photographed again.


Mutation of conserved residues within the glycogen-binding domain abolishes glycogen binding of Gal83 in vitro.

We constructed double mutations in GAL83 and SIP2 corresponding to those shown to abolish glycogen binding of the β1 subunit of AMPK (Trp100 to Gly and Lys126 to Gln [34]) (Fig. (Fig.1A).1A). These mutations altered Trp184 to Ala and Arg214 to Gln in Gal83, yielding a mutant protein designated Gal83W184A,R214Q, and altered Trp186 to Ala and Arg216 to Gln in Sip2, yielding Sip2W186A,R216Q. GFP-tagged mutant proteins were expressed from the native promoters on centromeric plasmids. The mutations did not inactivate the proteins, as each restored growth of a sip2Δ gal83Δ strain on glycerol (data not shown).

FIG. 1.
Mutation of the glycogen-binding domains of Sip2 and Gal83. (A) Conservation of the glycogen-binding domain among β subunits. Amino acid sequences of rat AMPK β1, Gal83, and Sip2 were aligned with CLUSTAL W (40) and formatted by an Excel ...

To test for the ability to bind glycogen, we expressed the glycogen-binding domains of the wild-type and mutant proteins in bacteria. We also included the Gal83G235R domain, because G235 is a conserved residue (Fig. (Fig.1A).1A). The purified domain of wild-type Gal83 bound glycogen in vitro, whereas those of Gal83W184A,R214Q and Gal83G235R did not bind (Fig. (Fig.1B).1B). Surprisingly, wild-type Sip2 bound weakly to glycogen, while Sip2W186A,R216Q did not bind at all (Fig. (Fig.1B).1B). To understand why Gal83 and Sip2 have different binding affinities for glycogen, despite an alignment suggesting similar glycogen-binding domains (Fig. (Fig.1A),1A), we constructed three-dimensional homology models of the two domains as described for β-GBD, the domain of AMPK β (34). Inspection of the putative glycogen-binding domains revealed that Sip2 contains a less defined and somewhat shallower glycogen-binding site than does Gal83, which may explain why Sip2 failed to bind tightly to glycogen (data not shown). The mutant Sip2 protein was not studied further.

Double sip2Δ gal83Δ deletion reduces glycogen accumulation in vivo.

To assess the roles of the β subunits in glycogen storage, we examined a sip2Δ gal83Δ strain expressing Sip2 and Gal83, separately and together. Cultures of all transformants showed identical growth kinetics in selective SD plus 2% glucose (Fig. (Fig.2A),2A), and accumulation of glycogen was assayed during growth of the cultures (Fig. (Fig.2B).2B). The sip2Δ gal83Δ double mutant had significantly diminished glycogen levels relative to transformants expressing both Sip2 and Gal83. Gal83 alone conferred normal glycogen accumulation, but transformants expressing only Sip2 may have a minor defect (Fig. (Fig.2B).2B). Thus, both the Gal83 and Sip2 β subunits contribute to Snf1 kinase function in promoting glycogen accumulation.

FIG. 2.
Mutation of SIP2 and GAL83 alters glycogen accumulation. Strain MCY4626 (sip2Δ gal83Δ) was transformed with plasmids encoding the indicated GFP-tagged proteins (pRT12, pHW30, pHW39, and pOV81) and with vectors pRS313 or pRS316, as needed ...

Mutation of the glycogen-binding domain of Gal83 enhances glycogen accumulation.

To examine the effects of mutations in the glycogen-binding domain on glycogen accumulation, we compared transformants of a sip2Δ gal83Δ strain expressing wild-type and mutant Gal83 and Sip2. Transformants expressing Gal83W184A,R214Q contained about twofold more glycogen than those expressing Gal83 at all phases of growth, independent of the presence of Sip2 (Fig. 2C and D). Gal83G235R caused similarly elevated glycogen levels (Fig. (Fig.2D).2D). Levels were higher even during early exponential growth (9 h); values for Gal83, Gal83W184A,R214Q, and Gal83G235R were (mean ± standard error) 4.5 ± 0.6, 11.4 ± 2.5, and 12.4 ± 3.0 μg of glycogen/108 cells, respectively (Fig. (Fig.2D).2D). These results indicate that mutation of critical residues in the glycogen-binding domain of Gal83 enhance glycogen accumulation.

Protein levels, nucleocytoplasmic distribution, and Snf1 catalytic activity are normal for Gal83W184A,R214Q.

To assess the possibility that the elevated glycogen levels simply reflect elevated levels of the mutant protein, we carried out immunoblot analysis, but we detected no difference (Fig. (Fig.3A).3A). Another possibility is that these mutations alter the subcellular localization of Gal83, hence affecting the proximity of Snf1 kinase to particular substrates. GFP-tagged Gal83 is localized in the cytoplasm and excluded from the nucleus when cells are growing on glucose, but upon a shift to low glucose or a nonfermentable carbon source, Gal83 rapidly accumulates in the nucleus (44). GFP-tagged Gal83W184A,R214Q exhibited the same nucleocytoplasmic distribution (Fig. (Fig.3B).3B). We cannot exclude differential association with glycogen, as glycogen granules in yeast cells can be resolved only by electron microscopy (6).

FIG. 3.
Protein levels, subcellular localization, and Snf1 catalytic activity for Gal83W184A,R214Q. (A) Protein extracts were prepared from cultures of MCY4626 expressing Gal83, Gal83W184A,R214Q, or no protein (vector) at the 9-h time point (OD600 = 0.6) ...

To determine whether the glycogen-binding domain mutation increases Snf1/Gal83 catalytic activity, we first used an immunoprecipitation kinase assay. Protein extracts were prepared from gal83Δ cells expressing Gal83 or Gal83W184A,R214Q and LexA-tagged Snf1. Snf1 kinase was immunoprecipitated with anti-LexA and incubated with [γ-32P]ATP. Mutation of Gal83 did not increase the phosphorylation of Snf1 or Gal83 (Fig. (Fig.3C).3C). The catalytically inactive LexA-Snf1T210A served as a control to confirm that phosphorylation reflected Snf1 activity.

We also assayed Snf1/Gal83 activity by phosphorylation of the SAMS peptide substrate (8). Snf1 kinase was partially purified from sip1Δ sip2Δ gal83Δ mutant cells (MCY4099) expressing Gal83 or Gal83W184A,R214Q and was incubated with SAMS peptide in the presence of [γ-32P]ATP. Incorporation of radiolabeled phosphate into the peptide was not significantly different (2.6 ± 0.6 and 1.8 ± 0.3 nmol/min/mg for Gal83 and Gal83W184A,R214Q, respectively, and 0.1 for the vector control; values are means of determinations for duplicate extracts). Thus, no alteration of catalytic activity was detected in vitro.

Gal83W184A,R214Q elevates expression of RNAs encoding glycogen synthase.

Previous work implicating Gal83 in transcriptional control (38, 42, 45) raised the possibility that increased transcription of key glycogen biosynthetic genes might be responsible for the increased glycogen accumulation. We examined levels of the GSY1 and GSY2 RNAs, which encode the two isoforms of glycogen synthase (10), in a gal83Δ mutant expressing Gal83W184A,R214Q or Gal83. Northern blot analysis showed that the W184A,R214Q mutation caused a twofold increase in the level of both GSY RNAs relative to a control RNA (Fig. (Fig.4);4); note that levels of the loading control RNA were lower in the mutant sample. Thus, an increase in expression of glycogen synthase likely contributes to the increase in glycogen accumulation.

FIG. 4.
Northern blot analysis of GSY1 and GSY2 RNAs. Transformants of strain MCY4622 (gal83Δ) expressing Gal83 (pRT12) or Gal83W184A,R214Q (pHW39) were grown in SD plus Leu plus Ura to OD600 = 2.0. RNAs were prepared and analyzed by Northern ...

Mutation of the glycogen-binding domain of Gal83 enhances haploid invasive growth.

We explored whether the glycogen-binding domain mutations affect another Gal83-dependent phenotype. A gal83Δ mutant is impaired in haploid invasive growth (45), a cellular response to carbon stress (7), at least in part due to effects on Snf1 kinase-dependent transcription of the FLO11 adhesin gene (25, 45). We expressed Gal83, Gal83W184A,R214Q, or Gal83G235R in a gal83Δ mutant of the invasive strain Σ1278b. The mutant proteins caused increased invasiveness (Fig. (Fig.5).5). This phenotype is dominant, as it was also observed in a GAL83 strain (Fig. (Fig.55).

FIG. 5.
Assay of haploid invasive growth. Strains MCY4702 (wild type) and MCY4622 (gal83Δ), expressing Gal83, Gal83W184A,R214Q, or Gal83G235R were assayed for invasive growth. Photographs were taken after 2 days and are representative of data from three ...

Gal83W184A,R214Q improves transcriptional activation by Sip4.

Previous studies showed that the GAL83-G235R allele confers partially glucose-insensitive GAL gene expression in certain strains (9, 28). This allele also affects the function of Sip4, a Snf1 kinase-dependent transcriptional activator that binds to the carbon source response element (CSRE) of gluconeogenic gene promoters (27, 43). Activation by Sip4 is impaired in a gal83Δ mutant, whereas GAL83-G235R partially relieves glucose inhibition of Sip4 and also improves activation (42).

To address the possibility that Gal83W184A,R214Q similarly affects Sip4, we assayed the ability of LexA-Sip4 to activate transcription of a lacZ reporter with LexA-binding sites (42). LexA-Sip4 was expressed from the ADH1 promoter in a gal83Δ mutant expressing Gal83, Gal83W184A,R214Q, or Gal83G235R. During growth in glucose, cells expressing either mutant protein produced two- to threefold higher β-galactosidase activity than those expressing wild-type Gal83 (Fig. (Fig.6A).6A). Following a shift to inducing conditions (nonfermentable carbon source), β-galactosidase activity increased substantially, and the mutant proteins continued to confer higher activity. The role of Gal83 in upregulating invasive growth is distinct from its role in Sip4 activity, because sip4Δ mutants are hyperinvasive (7).

FIG. 6.
Mutations in the glycogen-binding domain of Gal83 affect Sip4 transcriptional activator function and regulation of the CSRE. (A) Strain MCY4024 (gal83Δ), which carries an integrated lacZ reporter with LexA-binding sites in the promoter, expressed ...

Mutation of the glycogen-binding domain of Gal83 improves activation of the CSRE promoter element.

We next sought to examine the effect of the glycogen-binding mutations on the function of native transcription factors. The CSRE binds both Sip4 and Cat8, a related transcriptional activator that is also glucose inhibited and Snf1 kinase dependent (16, 17, 35, 43). Activation of a CSRE-lacZ reporter was defective in a gal83Δ mutant (Fig. (Fig.6B),6B), suggesting that Cat8, like Sip4 (42), is dependent on the Snf1/Gal83 form of the kinase. We assayed activation of the CSRE-lacZ reporter in a gal83Δ mutant expressing Gal83, Gal83W184A,R214Q, or Gal83G235R. All strains showed normal glucose repression of the reporter, as expected because expression of both activators from their native promoters is glucose repressed. When cells were shifted to glycerol plus ethanol to induce activation of the CSRE, both Gal83W184A,R214Q and Gal83G235R conferred higher activation than wild-type Gal83 (Fig. (Fig.6B).6B). These results show that mutations in the Gal83 glycogen-binding domain upregulate the function of native, Snf1 kinase-dependent transcription factors.

Mutation of the glycogen-binding domain affects the Snf1/Gal83 pathway in gsy1Δ gsy2Δ cells lacking glycogen.

The diverse phenotypes caused by the glycogen-binding domain mutations and, in particular, their effects on Snf1-dependent transcription under different growth conditions, including conditions when cells contain very little glycogen, cannot easily be accommodated by models involving glycogen binding. We therefore considered the possibility that some of these phenotypes are caused not by the failure to bind glycogen but rather by some other property of Gal83 that is altered by these mutations and affects Snf1/Gal83 kinase function. To test this model, we constructed a gsy1Δ gsy2Δ double mutant, which lacks glycogen synthase and does not synthesize glycogen (10). We reasoned that if glycogen binding is the critical factor responsible for the different phenotypes, then the mutant and wild-type Gal83 proteins should behave the same in a strain lacking glycogen altogether. If, on the other hand, the mutant and wild-type Gal83 proteins differ with respect to some other property, then they should still cause different phenotypes even in the absence of glycogen.

We compared isogenic wild-type and gsy1Δ gsy2Δ strains expressing Gal83 or Gal83W184A,R214Q with respect to LexA-Sip4 function and activation of the CSRE. We used reporter assays similar to those described above, but this pair of strains has a different genetic background. Mutation of the glycogen-binding domain caused increased β-galactosidase expression not only in the wild-type control strain but also in the glycogen-deficient strain (Fig. (Fig.7).7). Moreover, the effect is dominant, as both strains carried a wild-type chromosomal GAL83 allele. Thus, mutation of the glycogen-binding domain confers phenotypes even in the absence of glycogen, indicating that the effect on Snf1/Gal83 function occurs by a glycogen-independent mechanism.

FIG. 7.
Gal83W184A,R214Q affects Sip4 activator function and regulation of the CSRE in glycogen-deficient cells. Strains were MCY4101 (gsy1Δ gsy2Δ) and BY4741 (wild type) expressing Gal83 (gray bars) or Gal83W184A,R214Q (white bars). Cultures ...


Previous studies identified a glycogen-binding domain in the AMPK β1 subunit, thereby implicating this subunit in the regulation of glycogen metabolism by AMPK (21, 34). We have used genetic analysis to assess the physiological role of the glycogen-binding domain of the Gal83 β subunit of Snf1 kinase in vivo. We have shown that mutations in conserved residues of the glycogen-binding domain of Gal83 abolish glycogen binding in vitro and confer broad phenotypic effects in vivo. Various Snf1/Gal83-dependent processes were upregulated, including glycogen accumulation, expression of RNAs encoding glycogen synthase, haploid invasive growth, the transcriptional activator function of Sip4, and activation of the CSRE promoter element by native Cat8 and Sip4. In those cases tested, the effect was dominant. Thus, these mutations positively affect Snf1/Gal83 kinase function with respect to multiple targets.

Unexpectedly, the transcriptional regulatory phenotypes tested were not dependent on the presence of glycogen in the cell. First, mutant phenotypes were observed under growth conditions when cells contained very little glycogen. More importantly, mutation of the glycogen-binding domain of Gal83 conferred the same increase in Sip4 activator function and activation of the CSRE in a gsy1Δ gsy2Δ mutant strain lacking glycogen synthase as in a wild-type strain. Thus, mutation of the glycogen-binding domain affects Snf1/Gal83 kinase function by a mechanism that is independent of glycogen binding.

How do mutations in the glycogen-binding domain affect Snf1/Gal83 kinase? There are a number of possible mechanisms that we consider unlikely. First, we detected no alteration in the nucleocytoplasmic distribution of mutant Gal83. Although modest quantitative changes could have escaped notice, it is difficult to imagine such changes accounting for increased transcription during growth in glucose, when most Gal83 is cytoplasmic, and also in nonfermentable carbon sources, when Gal83 is nuclear. Second, in vitro assays revealed no increase in Snf1/Gal83 catalytic activity, suggesting that these mutations do not cause a conformational change in the kinase that favors its activation or augments activity. Third, mutant Gal83 could interact better with the catalytic subunit, thus enhancing Snf1 kinase function in Gal83-dependent processes. However, the glycogen-binding domain appears distinct from the site(s) of interaction with the catalytic subunit. A region that interacts (called the KIS region) was localized by evidence that residues 149 to 350 or 198 to 417 sufficed for interaction, but Sip2 residues 152 to 248, which encompass the glycogen-binding domain, did not interact with Snf1 (23), suggesting that the interacting site lies C-terminal to the domain. Fourth, mutation of the glycogen-binding domain could affect the interaction of Gal83 with particular targets; however, Sip4 interacts with residues 336 to 417 (42). It remains possible that mutation of the glycogen-binding domain indirectly affects the conformation of adjacent regions.

A final, and interesting, possibility is that this domain binds some other molecule that is closely related to glycogen and serves a signaling function. Mutation of the domain would reduce or abolish binding of this unknown molecule. This model is particularly attractive, because the binding of such a molecule could modulate Snf1/Gal83 catalytic activity in vivo, thereby nicely accounting for the manifestation of mutant phenotypes in the glycogen-deficient strain. Such a molecule could dissociate from the kinase during purification and hence not affect catalytic activity in our in vitro assays.

The existence of a glycogen-independent regulatory mechanism does not preclude direct effects of glycogen binding on some functions of Snf1/Gal83 kinase. For example, the increased accumulation of glycogen caused by these mutations may in part reflect the loss of binding to glycogen (an idea that cannot be tested genetically). However, it is also possible that the increased glycogen accumulation largely reflects the increased expression of GSY1 and GSY2 RNAs, which may, like other transcriptional phenotypes examined here, be controlled by glycogen-independent mechanisms. Glycogen is an important reserve carbohydrate that accumulates during conditions of nutrient stress and is then mobilized during periods of starvation, and Snf1 kinase has multiple roles in regulating glycogen metabolism (11, 15, 19, 20, 46, 49); hence, one can easily imagine the utility of a glycogen-sensing mechanism in regulating Snf1 kinase and a physiological role for the glycogen-binding domain in targeting the kinase to glycogen. Nonetheless, our genetic analysis provided no evidence for serious anomalies in glycogen storage or mobilization, but rather revealed unanticipated transcriptional regulatory phenotypes reflecting activation of the Snf1/Gal83 pathway.

These findings support a role for the glycogen-binding domain in modulating Snf1/Gal83 kinase function. This regulatory mechanism unexpectedly proved independent of glycogen binding. The interesting possibility that this domain binds an unidentified signaling molecule, related to glycogen, merits further investigation.


We thank V. Vyas and K. Hedbacker for strains and O. Vincent for plasmids. We thank S.-P. Hong for assistance with kinase assays and V. Vyas for discussion.

This work was supported by National Institutes of Health grant GM34095 to M.C., by the NHMRC (D.S. and B.E.K), and by the National Heart Foundation and Australian Research Council (B.E.K.). D.S. is an NHMRC RD Wright Fellow, and B.E.K. is a senior principal NHMRC Fellow.


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