©1995 by The American Society for Biochemistry and Molecular Biology, Inc.
Analysis of Positive Elements Sensitive to Glucose in the Promoter of the FBP1 Gene from Yeast (*)

(Received for publication, December 8, 1994; and in revised form, March 3, 1995 )

Olivier Vincent (§) Juana M. Gancedo (¶)

From the  Instituto de Investigaciones Biomédicas del Consejo Superior de Investigaciones Cientficas, Arturo Duperier 4, E-28029 Madrid, Spain

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
FOOTNOTES
ACKNOWLEDGEMENTS
REFERENCES

ABSTRACT

We have identified in the promoter of the FBP1 gene from Saccharomyces cerevisiae, which codes for fructose-1,6-bisphosphatase, two elements which can form specific DNAbulletprotein complexes and which confer glucose-repressed expression to an heterologous reporter gene. Complex formation and activation of transcription by either element require a functional CAT1 gene and are not blocked by a hap2-1 mutation, although this mutation interferes with maximal expression of the FBP1 gene. A sequence from one of the elements acts as a weak upstream activating sequence, but its activity can be stimulated up to 10-fold by neighboring sequences. A further element of the promoter has been characterized, which forms a specific DNAbulletprotein complex only when a nuclear extract from derepressed cells is used. This element does not activate transcription in a heterologous promoter. The DNA sequences of the three elements involved in protein binding, defined by DNase I footprinting, have no homology with consensus sequences for known activating factors.


INTRODUCTION

The yeast Saccharomyces cerevisiae is able to utilize a variety of carbon sources, but the preferred one, at least in laboratory conditions, is glucose. Glucose produces a variety of physiological effects in S. cerevisiae(1) , among them a strong repression of many enzymes not required for growth on glucose. The glucose repression system in yeast (for reviews, see Refs. 2-4) is quite complex, and its mechanism is far from understood. It operates mainly at the level of gene transcription, although effects on mRNA stability have been also reported (5, 6, 7, 8) . Studies with different mutants have clearly established that not all the genes affected by catabolite repression are controlled by the same set of regulatory proteins but that there are different circuits of repression. Therefore, to understand the mechanisms of catabolite repression in yeast, a variety of systems ought to be studied.

Up to now, most studies have been directed to the elucidation of the control of the genes necessary for the utilization of sugars like galactose or sucrose. It has been found that expression of the GAL genes is dependent on the transcriptional activator Gal4 (9). Glucose represses the GAL4 gene through the action of Mig1 (10, 11) and also facilitates the binding of Mig1 to upstream repressing sequences in the promoters of the GAL genes (11-13). In the case of the SUC2 gene, Mig1 is also able to bind to the promoter and to inhibit transcription (14) , but a transcriptional activator binding to the upstream activating sequences (UAS)^1(^1) of the promoter has not been yet identified (4) .

While different mutations, including mig1, have been identified which relieve the SUC2 and GAL genes from catabolite repression, none of them allows the expression, in the presence of glucose, of gluconeogenic genes such as FBP1, PCK1, and ICL1, coding for fructose-1,6-bisphosphatase (Fru-1,6-P(2)ase), phosphoenolpyruvate carboxykinase, and isocitrate lyase, respectively. Since this indicates that the mechanisms of regulation of these genes have characteristics different from those of the SUC2 and GAL genes, we have undertaken a study to define the cis- and trans-acting control elements of FBP1 and PCK1. Deletion analysis of the FBP1-lacZ and PCK1-lacZ fusion genes showed a complex picture of both promoters, with regions of apparent functional redundancy (15) . We have also identified in the FBP1 promoter two sites able to bind nuclear proteins (16) . The capacity of the region -431/-420 to bind proteins was subsequently confirmed, using a synthetic oligonucleotide (17) . In this later work, it was also shown that a region further upstream, at -506/-477 was required for maximal activation of transcription.

In the present study, we have undertaken a detailed analysis of different regions of the FBP1 promoter. We have investigated the behavior of these regions both with respect to their ability to act as UAS in the CYC1-lacZ reporter system (18) and to their capacity to form specific complexes with nuclear proteins.

We conclude that the FBP1 promoter contains at least two activating regions regulated by glucose; one of them appears to require the binding of proteins at two different sites to fully activate transcription.


EXPERIMENTAL PROCEDURES

Yeast Strains and Media

Strains of S. cerevisiae used in this work are listed in Table I. Escherichia coli TG1 was used for plasmid manipulations. For the preparation of nuclear extracts, the yeasts were grown on 1% yeast extract, 1% peptone, and 2% glucose. Repressed cells were collected when the culture reached 2-4 mg of wet yeast/ml. Derepressed cells were prepared by overnight incubation of repressed cells in 1% yeast extract, 1% peptone, and 2% ethanol at a cell density of 20 mg/ml. Yeasts carrying plasmids were grown on Difco yeast nitrogen base supplemented with 2% glucose. Repressed cells were obtained by harvesting them in the exponential phase of growth (below 2.5 mg of wet yeast/ml). Derepressed cells were prepared as described above.

Oligonucleotides

Oligonucleotides were purchased from MedProbe A.S. (Oslo, Norway) or synthesized in house. To obtain synthetic double-stranded DNA fragments (shown in Fig. 1), single-stranded oligonucleotides of reverse complementary sequence were mixed in equimolar amounts.


Figure 1: Structure of the FBP1 promoter and of the oligonucleotides OL1 and OL2. The approximated position of the DNAbulletprotein complexes observed in the FBP1 promoter in vitro (A, B, C, D, and E) is shown. Oligonucleotides: the sequence of the FBP1 promoter is written in uppercase; the additional SalI protruding ends are written in lowercase; protected bases in the corresponding footprint of the FBP1 promoter are shown in brackets.



Recombinant DNA Methods

Cloning procedures and additional recombinant DNA techniques were performed according to established protocols (19) .

Plasmid Constructions

Plasmid pJJ11b was obtained from plasmid pJJ11 (16) by substituting the fragment containing the FBP1 promoter until the position -480 by a fragment from pRG6 (20) containing the FBP1 promoter until the position -530. To obtain the plasmid pLG669-ZX, a XhoI-XhoI 0.4-kilobase fragment containing the two UAS of the CYC1 gene promoter was removed from plasmid pLG669-Z (18) . To insert different fragments of the FBP1 promoter into the unique XhoI site of pLG669-ZX, the auxiliary plasmid pOV10, which has a polylinker with a variety of unique sites between a XhoI and a SalI site, was used (21) . Sequences of the FBP1 promoter were taken from plasmid pRG6 and subcloned into pOV10. Restriction ends were blunt-ended in the case where compatible sites were absent from the polylinker. The 5`-phosphorylated double-stranded oligonucleotides had protruding ends which allowed direct insertion in the XhoI site of pLG669-ZX. Orientation and copy number of the oligonucleotide inserts were determined by DNA sequencing.

Transformation

Bacterial competent cells were prepared, stored, and transformed by standard techniques (22) . Yeast transformations were performed according to Ito et al. (23) .

beta-Galactosidase Assay

Yeast extracts were prepared by shaking with glass beads as described (24) omitting the centrifugation step, and the enzyme was assayed as in Ref. 25, samples being centrifuged before reading. Protein was determined by the method of Lowry et al. (26) using the BCA Protein Assay Reagent (Pierce).

Preparation of Yeast Nuclear Extracts

Nuclear extracts were obtained as described in Schneider et al. (27) using repressed or derepressed cells.

Band-shift Assays

Protein-nucleic acid complexes were allowed to form in 10 mM Tris-HCl, pH 7.5, 5 mM MgCl(2), 2 mM EDTA, 10 mM 2-mercaptoethanol, 1 mM phenylmethylsulfonyl fluoride, 50 mM NaCl, and 12.5% glycerol (v/v). In a volume of 20 µl, nuclear extract (4-30 µg of protein) was preincubated with 1 µg of poly(dI-dC) for 15 min in ice. The DNA probe, labeled with the Klenow fragment of DNA polymerase I (19) , was then added (1 ng, approximately 40,000 cpm), and incubation was continued for 30 min at room temperature. Electrophoresis was performed in 4% polyacrylamide gels in Tris borate, pH 7.6 (19) , at 12.5 V/cm for 90 min to 4 h at 4 °C.

Indirect DNase I Footprint Assay

Six parallel samples were prepared, as described above, using 2 ng (80,000 cpm) of DNA probe, 1 µg of poly(dI-dC), and protein at the concentration indicated in the legend for Fig. 4, in a final volume of 40 µl. After the 30-min incubation, 1 µl of 200 mM MgCl(2), 100 mM CaCl(2), and 1 µl of DNase I (10 to 100 ng) were added to each sample, which was incubated for 60 s at room temperature. Digestion was stopped in ice by the addition of 1/10 volume of 0.5 M EDTA, and samples were loaded onto a 4% polyacrylamide gel and electrophoresed as above. The positions of free and protein-bound DNA were identified by autoradiography, the corresponding fragments were excised, the equivalent fragments of the six samples were pooled together, and DNA was isolated by electrophoresis in a gel of agarose and binding to a DEAE-membrane (28) . After elution, the DNA was precipitated with ethanol, denatured, and applied to an 8% polyacrylamide sequencing gel.


Figure 4: DNase I protection analysis of the DNAbulletprotein complexes E (a, fragment -527/-476), B and D (b, fragment -479/-313), and C (c, fragment -316/-176). The T+C and A+G lanes correspond to the Maxam-Gilbert sequencing ladder. Free DNA (a, lanes 1 and 3; b, lanes 2 and 5; c, lane 1) and DNAbulletprotein complexes (a, lanes 2 and 4; b, lanes 1 and 4 (complex D) and lanes 3 and 6 (complex B); c, lane 2) were separated in a nondenaturing 4% polyacrylamide gel after incubation with a nuclear protein extract from wild-type yeast derepressed cells (a, 1 mg/ml; b, 0.4 mg/ml) or from mig1 derepressed cells (c, 0.8 mg/ml) and DNase treatment. Protected bases are marked; hypersensitive bases are shown with a + sign.




RESULTS

Since an activating sequence appeared to be located between positions -527 and -480 of the FBP1 promoter (17) , we examined the effect of its deletion on a fusion gene FBP1-lacZ. To avoid possible artifacts due to variations in the number of copies of the gene, we placed the constructions in a centromeric plasmid. We found that the level of beta-galactosidase was 2.5-fold higher in a yeast carrying the fusion gene pJJ11b, containing the promoter up to position -527, than in a yeast carrying plasmid pJJ11 (16) , which contains a promoter starting at position -480. In both cases, expression occurred only in derepressed cells.

The -527 to -480 sequence could have activating capacity by itself or reinforce the action of a downstream activator. To evaluate the capacity of this and other regions of the FBP1 promoter to activate transcription, fragments of the promoter (Fig. 1) were inserted into a UAS-less reporter plasmid (18) , and the capacity of the plasmids to direct the synthesis of beta-galactosidase was tested (Table II). Both fragments -527 to -475 and -480 to -312 were able to activate transcription and this only in the absence of glucose. The orientation of the inserted DNA had a moderate effect on expression for the smaller fragment, but a very strong one for the larger fragment. This could indicate an influence of the distance between the UAS and the TATA box on the rate of transcription; an alternative possibility would be the presence of a repressing sequence which would be more effective when placed between the UAS and the TATA box.

As Fru-1,6-P(2)ase is not expressed in a cat1/snf1 mutant (29) , and levels of Fru-1,6-P(2)ase are decreased in hap2 and hap4 mutants,^2(^2) the effects of cat1 and hap2 mutations in the expression of the fusion genes were tested. No expression at all was seen in cat1 mutants while in hap2 mutants there was no decrease in beta-galactosidase levels, but rather an increase of about 50% (Table II).

The fragment -317 to -176 of the FBP1 promoter was also inserted in the reporter plasmid and did not show UAS activity. Such an activity could have been masked by the negative regulatory protein Mig1 binding to a site around position -200 (16) . However, we found that even in a mig1 background, the fusion gene was not expressed (data not shown).

We had observed that a fusion gene FBP1-lacZ required part of the coding sequence of FBP1 to be expressed, the first 57 base pairs being sufficient (15) . This suggested the existence of a downstream activating sequence and therefore we tested the fragment -53 to +57 for UAS activity. No expression of beta-galactosidase was observed, showing that this fragment could not activate transcription by itself.

To look for a possible correlation between the effect of different fragments of the FBP1 promoter on transcription and their capacity to bind specific proteins, we performed band-shift assays. The DNA segment from -527 to -476 yielded a single major retarded band (E) as shown in Fig. 2a. This is in contrast with the results of Schöler and Schüller (30) who observed 4 specific DNAbulletprotein complexes in their band-shift experiments. This could be due to the fact that they used whole yeast extracts while we use nuclear proteins. The gel retardation pattern observed by Niederacher et al.(17) using an oligonucleotide which covers the same region comprises also a single band of low mobility in addition to some blurred bands which do not seem specific. The DNAbulletprotein complex E was observed only when nuclear extracts from derepressed cells were used for the band shift, it was also absent when the nuclear proteins used were from a cat1 mutant. In competition experiments, a 50-fold molar excess of the unlabeled DNA fragment prevented the formation of the complex, while the same amount of the fragment -479 to -313 had no effect (results not shown). The DNA fragment from -479 to -313 gave two strong specific retardation bands, B and D, as shown in Fig. 2b. Complex B was observed only when a nuclear extract from derepressed cells was used, while complex D was also formed with the protein extracted from repressed cells. Both complexes were specific, competition was observed with an excess of the corresponding unlabeled DNA, while the DNA from -527 to -476 did not affect the formation of the complexes (not shown). When extracts from a cat1 mutant were used, complex D decreased or almost disappeared while complex B appeared to change its mobility (Fig. 2b). A weaker specific complex migrating between B and D was observed only when extracts from a derepressed, wild-type yeast were used.


Figure 2: Gel mobility shift analysis with fragments -527/-476 (a) and -479/-313 (b) from the FBP1 promotor. The DNA fragment was labeled and incubated as described under ``Experimental Procedures.'' Samples in lanes 1 were incubated without added protein, and samples in lanes 2 and 3 were incubated with the amount of nuclear protein indicated. The protein extract was from derepressed cells (D), from repressed cells (R), or from a derepressed cat1 mutant strain (Dcat1).



In the case of the fragment -316 to -176, two complexes were observed which were in fact two doublets. They did not depend on the state of repression or derepression of the cells (Fig. 3). All of them are designated collectively as complex A and were not detectable when a nuclear extract from repressed cells of a mig1 mutant was used. In the absence of Mig1, a faint band remained which was much stronger when derepressed cells from a mig1 mutant were used. Competition assays indicated that this new complex, C, is specific (results not shown). In a cat1 background, the two doublets corresponding to complex A showed changes in mobility, while in a cat1/mig1 double mutant, complex C was barely visible.


Figure 3: Gel mobility shift analysis with the fragment -316/-176 from the FBP1 promotor. General experimental conditions were as described in Fig. 2. Samples in lanes 1 were incubated without added protein, and samples in lanes 2-4 were incubated with the amount of nuclear protein indicated. The protein extract was from derepressed cells (D), from repressed cells (R), from a repressed mig1 mutant strain (Rmig1), from a derepressed cat1 mutant strain (Dcat1), from a derepressed cat1/mig1 double mutant strain (Dcat1 mig1), or from a derepressed mig1 mutant strain (Dmig1).



When the fragments of the FBP1 promoter -179 to -73 and -76 to +57 were used in electrophoretic mobility shift assays, no clear specific band shifts were observed.

To identify the DNA sequences where nuclear proteins bind specifically, footprinting experiments were carried out. For the -527 to -476 fragment, indirect footprinting was performed in both the coding and noncoding strands with the results shown in Fig. 4a. The different footprints on the two strands of the promoter indicated that the protein(s) in the complex E bound asymmetrically to the DNA.

For the -479 to -313 fragment, indirect footprinting was attempted with both B and D complexes. For B, the results for both strands are shown in Fig. 4b. They show that protection from DNase I cleavage is completely asymmetric, from -431 to -424 in the coding strand and from -422 to -417 in the noncoding strand. In the case of the D complex, no protection could be observed in our experimental conditions. In an attempt to define more precisely the region of the promoter where this complex could be formed, a band shift was performed with a DNA fragment from -480 to -413. As shown in Fig. 5b, this fragment was sufficient for the formation of both the B and D complexes. An oligonucleotide covering the region -432 to -415 acted as a competitor for the formation of the B complex, but did not affect the formation of the D complex, which therefore is likely to be in the region between -480 and -432.


Figure 5: Gel mobility shift analysis with the oligonucleotide OL1 (a), the fragment -480/-413 from the FBP1 promotor (b), and the oligonucleotide OL2 (c). General experimental conditions were as described in Fig. 2. Samples in lanes 1 were incubated without added protein, and samples in lanes 2-6 were incubated with nuclear protein from derepressed cells (a, 32 µg; b, 8 µg; c, 24 µg). Competition experiments were performed with a 50-fold or 100-fold excess of the unlabeled fragment used as probe (*) or of a different unlabeled fragment (OL1 or OL2).



Using extracts from a mig1 mutant, we could localize the DNA region where complex C is formed by indirect footprinting. A clear footprint was observed at positions -239 to -232 on the noncoding strand (Fig. 4c). Although this sequence is very similar to the consensus sequence for the binding of the protein complex Hap2-Hap3-Hap4, ACCAAYNA (31) , we could still observe complex C when extracts from a double mutant mig1/hap2 were used (results not shown).

The results from the footprinting experiments, together with those of sequential deletions (15, 17) , suggested that the regions -507 to -489 and -432 to -415 of the FBP1 promoter contained upstream activating sequences. We therefore tested the synthetic oligonucleotides OL1 and OL2 (Fig. 1) which include these sequences, both for their capacity to activate transcription and to bind proteins in a specific manner.

As shown in Table III, OL1 inserted in a UAS-less fusion gene activated transcription to the same degree as the fragment -527 to -476 (see Table II), and this activation occurred only in the absence of glucose. It appears, therefore, that the sequences within the larger fragment responsible for the regulated activation of transcription are all included in the oligonucleotide. In contrast, the oligonucleotide OL2 activated transcription only weakly, and the effect of glucose on transcription was also weak, only about 2.5-fold repression. This suggests that other regions outside the -432 to -415 sequence are required for proper control of transcription. We therefore tested the fragments -450 to -413 and -480 to -408. The first one was only slightly more active than the oligonucleotide while the larger fragment acted as a strong UAS. In all cases, the activation was observed only in the absence of glucose. We checked also the fragment -480 to -448 and found that, by itself, it did not stimulate transcription at all.

In band-shift assays, OL1 was able to bind nuclear proteins to form complex E. The binding was specific as shown in Fig. 5a, and the complex was also completely dependent on a functional CAT1 gene (Fig. 6a). In the case of OL2, two strong complexes were formed, but only one of them was specific and likely corresponded to complex B (Fig. 5c). The nonspecific complex was independent of CAT1, while the specific one was only formed in a CAT1 background (Fig. 6b).


Figure 6: Gel mobility shift analysis with the oligonucleotides OL1 (a) and OL2 (b). General experimental conditions were as described in Fig. 2. Samples in lanes 1 were incubated without added protein, and samples in lanes 2-4 were incubated with the amount of nuclear protein indicated. The protein extract was from wild-type yeast derepressed cells (WT) or from a derepressed cat1 mutant strain (cat1).




DISCUSSION

The FBP1 promoter appears to be composed of a variety of elements. An element located between positions -507 and -489 can act as UAS, and our footprint experiments situate the sequence involved in the protein-DNA interaction between positions -506 and -492 (Fig. 1). Very similar sequences are found in the promoters for FBP1, ICL1, and PCK1 (Fig. 7); they include the carbon source-responsive element, CSRE, previously described by Schöler and Schüller (30) but also three additional bases, TTC, common to the three promoters. Since it has been reported (30) that the sequence from -506 to -493 works as a strong UAS while the sequence from -503 to -488 is inactive, it can be concluded that the G at position -504 cannot be replaced by a T, while the T at position -492 is dispensable. Therefore, we suggest that the consensus sequence for an activating element in gluconeogenic genes should include two additional bases as shown in Fig. 7. This sequence has no homology with consensus sequences for known activating factors. We have tried to identify a protein able to bind to this sequence by screening a gt11 expression library with a labeled DNA probe (results not shown). Such an experiment has been unsuccessful, as well as our attempt to perform a Southwestern blot (32) .


Figure 7: Comparison of sequences from the promoters of genes encoding gluconeogenic enzymes. Sequences from the promoters of FBP1 (36), ICL1 (30), and PCK1 (15) are shown. Bases which differ from the FBP1 sequence are written in lowercase. The carbon source-responsive element, CSRE (30), is shown in brackets. R stands for A or G, W for A or T, and Y for C or T.



A second activating element is located in the region -480 to -408 of the FBP1 promoter. This element can form two complexes, B and D, and acts as a strong UAS completely repressed by glucose. It contains the sequence -432 to -415, sufficient to form complex B, but which acts only as a weak UAS. This sequence presents some similarities with those recognized by Mig1 (33) , Reb1 (=Grf2) (34) , and Rap1 (=Grf1) (35). However, it has been observed previously that the protein or proteins binding this sequence and called Dap1 could not be Rap1 or Mig1 (17) . As for Reb1, it is a large protein (128 kDa) which would form a DNAbulletprotein complex with a lower mobility than the B complex. As in the case of complex E, neither a Southwestern probe nor a screen of a gt11 expression library with a labeled DNA probe allowed the identification of a protein taking part in complex B (results not shown). When band-shift experiments are performed with the oligonucleotide OL2, two protein-DNA complexes are formed, but only one of them is specific (Fig. 5c). As the nonspecific complex is not detected with the larger fragment -480 to -413, it would seem that sequences adjacent to the region -432 to -415 are not required for the formation of complex B but play a role in avoiding the formation of nonspecific complexes. In addition, the neighboring sequences could allow the formation of complex D which may act as a coactivator. We have examined whether the sequence -450 to -413, which includes a potential site for the activator of transcription Yap1 (15), was a stronger UAS than the shorter -432 to -415 sequence (Table III). The difference is small, thus indicating that nucleotides upstream of -450 are required to activate transcription.

We have found that neither complex E nor complex B is formed when nuclear extracts from repressed cells are used in a band-shift assay, in agreement with observations from other groups (17, 30) . When the extract is from cells with an interrupted CAT1 gene, E is not formed and B is either not formed or has an altered mobility, depending on the DNA probe used ( Fig. 2and Fig. 6). All these results are consistent with the in vivo situation where the sequences containing the -507 to -489 or -432 to -415 elements act as an UAS only in derepressed cells with an intact CAT1 gene. Hap2 which is required for maximal derepression of Fru-1,6-P(2)ase^2 is not necessary for the activity of the two upstream elements of the FBP1 promoter. In fact, their activity was slightly increased in a hap2 background, but the significance of this observation is not clear.

The identification in the region -243 to -228 of a complex C, which is formed only when extracts from derepressed cells are used, suggests the presence of a potential UAS. While the large region -317 to -176 has no UAS activity in a reporter plasmid, when tested as a potential upstream repressing sequence, it represses much less in the absence of glucose than in its presence (21) . A possible interpretation of these results is that complex C facilitates transcription in the absence of glucose through a modification of the chromatin structure.

Up to now, the only protein able to bind to the FBP1 promoter which has been characterized is the repressing protein Mig1. We have found that the mobility of the Mig1bulletDNA complexes changes when extracts from a cat1 mutant are used in the band-shift assay but is the same for extracts from repressed or derepressed cells. This observation could indicate that, in the absence of Cat1, Mig1 exists in a modified form but that in wild-type cells the same form of Mig1 is found, whether glucose is present or not.

Although in the gluconeogenic gene ICL1 all the stimulatory influence of the promoter appears to act via a single upstream element (30), a variety of regulatory elements are present in the FBP1 promoter. These elements, however, seem to play similar roles, since they are affected in the same way by the presence of glucose and by the Cat1 protein. It cannot be yet ascertained whether they are truly redundant or serve for a fine tuning of Fru-1,6-P(2)ase expression.

  
Table: 38

  
Table: Regulation of the expression of fusion genes containing fragments of the FBP1 promoter

The plasmids were constructed as described under ``Experimental Procedures'' and used to transform strains W303-1A (WT), H366 (Deltacat1) and W303 hap2 (hap2-1).


  
Table: Effect of the insertion of different fragments of the FBP1 promoter into a DeltaUAS CYC1-lacZ plasmid

The plasmids were constructed as described under ``Experimental Procedures'' and used to transform strain W303-1A. The sequence of the oligonucleotides OL1 and OL2 is shown in Fig. 1.



FOOTNOTES

*
This work was supported by Grant PB91-0056 from the Dirección General de Investigación Cientfica y Técnica. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked ``advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§
Recipient of a fellowship from the Dirección General de Investigación Cientfica y Técnica (DGICYT), Ministerio de Educación y Ciencia, Spain, Program ``Acciones de Formación de Personal Investigador. Estancias Temporales de Cientficos y Tecnólogos Extranjeros en Espaa.''

To whom correspondence and reprint requests should be addressed. Tel.: 34-1-585-4622; Fax: 34-1-585-4587.

(^1)
The abbreviations used are: UAS, upstream activating sequence(s); Fru-1,6-P(2)ase, fructose-1,6-bisphosphatase.

(^2)
J. M. Gancedo, unpublished observations.


ACKNOWLEDGEMENTS

We thank L. Guarente for plasmid pLG669-Z, H. Ronne for yeast strains H190, H366, and H368, and P. Santisteban and M. J. Mazón for comments on the manuscript.


REFERENCES

  1. Gancedo, C., and Serrano, R.(1989) in The Yeasts (Rose, A. H., and Harrison, J. S., eds) Vol. 3, pp. 205-259, Academic Press, London
  2. Gancedo, J. M.(1992) Eur. J. Biochem. 206, 297-313 [Medline] [Order article via Infotrieve]
  3. Trumbly, R. J.(1992) Mol. Microbiol. 6, 15-21 [Medline] [Order article via Infotrieve]
  4. Johnston, M., and Carlson, M.(1993) in The Molecular and Cellular Biology of the Yeast Saccharomyces: Gene Expression (Jones, E. W., Pringle, J., and Broach J. R., eds) pp. 193-281, Cold Spring Harbor Laboratory, Cold Spring Harbor, NY
  5. Zitomer, R. S., and Nichols, D. L.(1978) J. Bacteriol. 135, 39-44 [Medline] [Order article via Infotrieve]
  6. Federoff, H. J., Eccleshall, T. R., and Marmur, J.(1983) J. Bacteriol. 156, 301-307 [Medline] [Order article via Infotrieve]
  7. Lombardo, A., Cereghino, G. P., and Scheffler, I. E.(1992) Mol. Cell. Biol. 12, 2941-2948 [Abstract]
  8. Mercado, J. J., Smith, R., Sagliocco, F. A., Brown, A. J. P., and Gancedo, J. M.(1994) Eur. J. Biochem. 224, 473-481 [Abstract]
  9. Keegan, L., Gill, G., and Ptashne, M.(1986) Science 231, 699-704 [Medline] [Order article via Infotrieve]
  10. Griggs, D. W., and Johnston, M.(1991) Proc. Natl. Acad. Sci. U. S. A. 88, 8597-8601 [Abstract]
  11. Nehlin, J. O., Carlberg, M., and Ronne, H.(1991) EMBO J. 10, 3373-3377 [Abstract]
  12. Flick, J. S., and Johnston, M.(1992) Genetics 130, 295-304 [Abstract/Free Full Text]
  13. Lamphier, M., and Ptashne, M.(1992) Proc. Natl. Acad. Sci. U. S. A. 89, 5922-5926 [Abstract]
  14. Nehlin, J. O., and Ronne, H.(1990) EMBO J. 9, 2891-2898 [Abstract]
  15. Mercado, J. J., and Gancedo, J. M.(1992) FEBS Lett. 311, 110-114 [CrossRef][Medline] [Order article via Infotrieve]
  16. Mercado, J. J., Vincent, O., and Gancedo, J. M.(1991) FEBS Lett. 291, 97-100 [CrossRef][Medline] [Order article via Infotrieve]
  17. Niederacher, D., Schüller, H.-J., Grzesitza, D., Gütlich, H., Hauser, H. P., Wagner, T., and Entian K.-D.(1992) Curr. Genet. 22, 363-370 [Medline] [Order article via Infotrieve]
  18. Guarente, L., and Ptashne, M.(1981) Proc. Natl. Acad. Sci. U. S. A. 78, 2199-2203 [Abstract]
  19. Sambrook, J., Fritsch, E. F., and Maniatis, T.(1989) Molecular Cloning: A Laboratory Manual , 2nd Ed, Cold Spring Harbor Laboratory, Cold Spring Harbor, NY
  20. De la Guerra, R., Valdés-Hevia, M. D., and Gancedo, J. M.(1988) FEBS Lett. 242, 149-152 [CrossRef][Medline] [Order article via Infotrieve]
  21. Vincent, O., and Gancedo, J. M.(1995) Current Genet. 27, 387-389 [Medline] [Order article via Infotrieve]
  22. Hanahan, D.(1985) in DNA Cloning (Glover, D. M., ed) pp. 109-135, IRL Press, Oxford
  23. Ito, H., Fukuda, Y., Murata, K., and Kimura, A.(1983) J. Bacteriol. 153, 163-168 [Medline] [Order article via Infotrieve]
  24. Blázquez, M. A., Lagunas, R., Gancedo, C., and Gancedo, J. M. (1993) FEBS Lett. 329, 51-54 [CrossRef][Medline] [Order article via Infotrieve]
  25. Miller, J.(1972) Experiments in Molecular Biology , Cold Spring Harbor Laboratory, Cold Spring Harbor, NY
  26. Lowry, O. H., Rosebrough, N. J., Farr, A. L., and Randall, R. J.(1951) J. Biol. Chem. 193, 265-275 [Free Full Text]
  27. Schneider, R., Gander, I., Müller, U., Mertz, R., and Winnacker, E. L.(1986) Nucleic Acids Res. 14, 1303-1317 [Abstract]
  28. Ausubel, F. M., Brent, R., Kingston, R. E., Moore, D. D., Seidman, J. G., Smith, J. A., and Struhl, K.(1988) Current Protocols in Molecular Biology , pp. 12.3.1-12.3.6. Wiley, New York
  29. Entian, K.-D., and Zimmermann, F. K.(1982) J. Bacteriol. 151, 1123-1128 [Medline] [Order article via Infotrieve]
  30. Schöler, A., and Schüller, H.-J.(1994) Mol. Cell. Biol. 14, 3613-3622 [Abstract]
  31. Olesen, J., Hahn, S., and Guarente, L.(1987) Cell 51, 953-961 [Medline] [Order article via Infotrieve]
  32. Francis-Lang, H., Price, M., Polycarpou-Schwarz, M., and Di Lauro, R. (1992) Mol. Cell. Biol. 12, 576-588 [Abstract]
  33. Lundin, M., Nehlin, J. O., and Ronne, H.(1994) Mol. Cell. Biol. 14, 1979-1985 [Abstract]
  34. Wang, H., Nicholson, P. R., and Stillman, D. J. (1990.) Mol. Cell. Biol. 10, 1743-1753
  35. Buchman, A. R., Kimmerly, W. J., Rine, J., and Kornberg, R. D.(1988) Mol. Cell. Biol. 8, 210-225 [Medline] [Order article via Infotrieve]
  36. Rogers, D. T., Hiller, E., Mitsock, L., and Orr, E.(1988) J. Biol. Chem. 263, 6051-6057 [Abstract/Free Full Text]
  37. Thomas, B. J., and Rothstein, R.(1989) Cell 56, 619-630 [Medline] [Order article via Infotrieve]
  38. Guarente, L., Lalonde, B., Gifford, P., and Alani, E.(1984) Cell 36, 503-511 [Medline] [Order article via Infotrieve]

©1995 by The American Society for Biochemistry and Molecular Biology, Inc.