Vol. 11, No. 10

MOLECULAR AND CELLULAR BIOLOGY, OCt. 1991, p. 5251-5258

0270-7306/91/105251-08$02.00/0 Copyright C 1991, American Society for Microbiology

Quantitation of ot-Factor Internalization and Response during the Saccharomyces cerevisiae Cell Cycle BETTINA ZANOLARI AND HOWARD RIEZMAN* Biocenter, University of Basel, Klingelbergstrasse 70, CH4056 Basel, Switzerland Received 15 April 1991/Accepted 22 July 1991

The a-factor pheromone binds to specific cell surface receptors on Saccharomyces cerevisiae a cells. The pheromone is then internalized, and cell surface receptors are down-regulated. At the same time, a signal is transmitted that causes changes in gene expression and cell cycle arrest. We show that the ability of cells to internalize ot-factor is constant throughout the cell cycle. a cells are also able to respond to pheromone throughout the cycle even though there is cell cycle modulation of the expression of two pheromone-inducible genes, FUSI and STE2. Both of these genes are expressed less efficiently near or just after the START point of the cell cycle in response to ot-factor. For STE2, the basal level of expression is modulated in the same manner.

Saccharomyces cerevisiae exists in two haploid mating a, that cycle through a vegetative life cycle. Haploid cells are also capable of switching their developmental program to enter the conjugation pathway. The two developmental pathways are mutually exclusive because entry into the conjugation pathway implies an arrest of the cell cycle, and cells arrested at other points of the cycle cannot conjugate. The mating program is initiated by the action of secreted peptide pheromones that bind to cell surface receptors and cause changes in the pattern of gene expression, an arrest at START in the G1 phase of the cell cycle, and a morphological change called the shmoo. At the tips of these shmoos, a single a and a cell fuse to form the zygote. After nuclear fusion, the diploid cell can also enter into a vegetative cycle very similar to the haploid cell cycle (reviewed in reference 11). The pheromone receptors are polytopic membrane proteins (8, 19, 38) that transduce the extracellular signal by interacting with a G protein (12, 36, 53). The enzyme activity regulated by the G protein has not yet been identified. Subsequent steps in the signal transduction pathway involve the products of several genes including STE5, -7, -11, and -12 (5, 22, 33, 37) and FUS3 (16). The point of the cell cycle at which pheromone arrests cells is late in G1 and has been defined as START (7, 23, 54). The morphological definition of START is unbudded cells having a spindle pole body satellite that has not yet been duplicated (41). This point has also been found to be the arrest point of cell division cycle mutants cdc28, cdc36, cdc37, and cdc39 (42). Progression through START depends on the product of the CDC28 gene, a serine/threonine protein kinase with homology to cdc2+ in Schizosaccharomyces pombe and MPF in starfish and Xenopus laevis (1, 18, 31, 45). This p34cdc28 associates with smaller subunits called cyclins to form an active complex (6, 29, 55). Addition of a-factor to a cells prevents the expression of two cyclin homologs encoded by the CLNJ and CLN2 genes (55). The resulting inactivation of the kinase could be the cause of the pheromone-induced arrest at

is subsequently degraded by a mechanism that depends on the presence of active vacuolar hydrolases (13, 46). These data suggest that a-factor is internalized by receptor-mediated endocytosis. The detection of a vesicular intermediate involved in the transport of a-factor from the plasma membrane to the vacuole (46) lends strong support to this hypothesis. In mammalian cells, endocytic uptake and subsequent membrane traffic are blocked during mitosis (3, 4, 44, 52). Studies performed in vitro implicate the vertebrate homolog of the p34Cdc28 protein kinase in the inhibition of endocytic traffic during mitosis (50). As the pathways of endocytosis and progression through the cell cycle are seemingly coupled in mammalian cells and a receptor that is taken up by endocytosis in yeast cells is involved in the pheromoneinduced arrest of the cell cycle, we decided to examine the ability of cells to internalize and respond to pheromone during the cell cycle. We show that receptors are present at the cell surface throughout the cell cycle and are able to internalize ac-factor. Even though cells can respond to a-factor throughout the cycle, the level of the induced expression is lowest near or just after START and highest near the G2 phase of the cycle.

types, a and

MATERIALS AND METHODS

Strains, media, and reagents. The following yeast strains used: RH448 (MATa ura3 his4 leu2 lys2 bar]-1), RH210-3C (MATa cdc15-2 ade2 ura3 leu2 his4 trpl bar]-i), RH209-4A (MATa cdc13-1 ade2 leu2 his4 bar]-1), RH178-2B (MATa cdc35-20 leu2 his4 ura3 barl-i), RH211-1C (MATa cdc284 ade2 his4 leu2 ura3 barl-1), and RH294-7A (MATa cdc20-1 lys2 ural ura3 tyri barl-i). Strains containing the cdc13, -15, and -28 alleles were obtained from Kim Nasmyth (Vienna, Austria), and a strain containing the cdc20 allele was from Peter Philippsen (Basel, Switzerland). These strains were crossed once or twice with our strain background (RH448 and isogenic strains) to introduce the barl-i mutation. The cdc35 allele (encoding adenylate cyclase [28]) was isolated in our laboratory, and its identity was confirmed by complementation analysis and demonstration of centromere linkage. Measurements of intracellular cyclic AMP in a radioimmunoassay (New England Nuclear) indicated that the defect in adenylate cyclase affected the enzyme as soon as the cells were shifted to 37°C (36a). Plasmid pSB231 were

START. After binding to its receptor on a cells, the a-factor pheromone is internalized (10, 26) concomitant with a downregulation of active cell surface receptors (26). The a-factor *

Corresponding author. 5251

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(URA3 CEN4-ARS FUSI-lacZ) (49) was introduced into yeast strains as described previously (25). When plasmidcarrying strains were used, precultures were grown in selective medium (15). Cells were then grown overnight to log phase in YPUAD (1% yeast extract, 2% peptone, 2% glucose, 30 mg each of adenine and uracil per ml). a-Factor was chemically synthesized at the Institut de Biochimie, Universite de Lausanne, Lausanne, Switzerland, and 35S-a-factor was purified from culture supernatants of 35S-labelled yeast cells as described previously (15). 4-Methylumbelliferyl-PD-galactoside, 4-methylumbelliferylphosphate, and monoclonal mouse antibodies against ,-galactosidase were obtained from Sigma Chemical Co. (St. Louis, Mo.). Mouse monoclonal antibodies against ,-tubulin and affinity-purified sheep immunoglobulin G to mouse immunoglobulin labelled with rhodamine were obtained from Boehringer Mannheim. Assays. a-Factor internalization was assayed as described previously (15), using 35S-a-factor in a pulse-chase protocol. Briefly, cells were incubated with 35S-a-factor on ice for 40 min in 50 mM potassium phosphate (pH 6.0-1% bovine serum albumin, washed, and resuspended in 50 mM potassium phosphate (pH 6.0)-2% glucose at 24°C. At 2, 10, 15, and 20 min, cells were diluted into ice-cold pH 6 or pH 1 buffer and subsequently washed with the same to determine total cell-associated or internalized ac-factor, respectively. The percentage of bound ax-factor internalized was calculated as (cpm at pH 1)/(cpm at pH 6) x 100. All determinations were performed in duplicate. FUS1-p-galactosidase induction was assayed as described previously (14) during the cell cycle. Cells were adjusted to approximately 3 x 106/ml (10 ml) and incubated for 22.5 min at 24°C with 10-6 M a-factor. In our genetic background (bar]-1), this is a saturating amount of at-factor. The cells were harvested and lysed in ice-cold lysis buffer (0.1 M sodium phosphate [pH 7.0], 1 mM MgCl2, 10 mM P-mercaptoethanol) by agitation with glass beads in the presence of 2 mM phenylmethylsulfonyl fluoride, using a Vortex mixer. Triton X-100 was then added to a final concentration of 0.1% (vol/vol). The lysate was assayed in lysis buffer for f-galactosidase activity, using 2 x 10' M 4-methylumbelliferyl-pD-galactoside as a substrate (30). To correct for cell lysis and cell number, we measured alkaline phosphatase activity by using 2 x 10-' M 4-methylumbelliferylphosphate in 0.1 M Tris HCl (pH 9.0)-i mM MgCl2. The assay was performed in a total volume of 200 [LI and was stopped after 20 min of incubation at room temperature by adding 50 Fl of 0.5 N NaOH. The methylumbelliferone fluorescence was read in a Microfluor reader (Dynatech). The P-galactosidase activity is expressed as a relative value. The fluorescence units obtained from the P-galactosidase assay were divided by those obtained from the alkaline phosphatase assay. FUS1,-galactosidase induction in cdc28 and cdc35 cells was performed as described above 15 and 150 min after the shift to 37°C except that the incubation with ax-factor was at 37°C. For Northern (RNA) analysis, synchronized cells were incubated with 10-6 M (for FUSI) or 108 M (for STE2) a-factor for 22.5 min. RNA was extracted as described previously (27). RNA was electrophoresed, transferred to nitrocellulose, and detected with 32P-labelled probes for FUS1, STE2, BIKJ, and URA3 as described previously (14). For STE2 and URA3, 20 p.g of total RNA was used per lane. Staining of the filters with methylene blue indicated that equal amounts of rRNA were present in all lanes. For FUSJ and BIKI, 5 ,ug of poly(A) RNA was used per lane. X-ray films were quantified in the linear exposure range, using a Molecular Dynamics computing densitometer (model 300A).

MOL. CELL. BIOL.

All mRNAs detected were expressed from their native chromosomal loci, not from plasmid-borne genes. Cell synchronization and immunofluorescence. RH210-3C cells harboring plasmid pSB231 were inoculated from saturated precultures in selective medium and grown to exponential phase in YPUAD medium at 24°C. They were then shifted to 37°C, incubated for 150 min, harvested, and resuspended in fresh YPUAD medium at 24°C. At 15-min intervals, cells were removed to assay for a-factor internalization or FUS1-,-galactosidase induction or were fixed in formaldehyde and prepared for immunofluorescence (21). Briefly, the cells were fixed at room temperature for 2 h with 3.7% formaldehyde, washed, and attached to poly-L-lysinetreated slides. They were then incubated with the primary antibody for 30 min, and after washing were incubated with the rhodamine-labelled secondary antibody for 30 min. DNA was localized by incubation with 4',6-diamidino-2-phenylindole (DAPI) at 10 ,ug/ml in phosphate-buffered saline for 5 min in the dark. After washing, the cells were covered with Mowiol mounting medium (Hoechst) and a coverslip and viewed under a Zeiss Axiophot Microscope with appropriate Zeiss filters for rhodamine and DAPI. The primary antibodies were monoclonal mouse antibodies against P-tubulin or ,B-galactosidase. In all cases, cells were also viewed by using phase-contrast or Nomarski interference optics. To examine FUS1-,-galactosidase induction in asynchronous cells, RH448 cells carrying plasmid pSB231 were inoculated from a saturated preculture in selective medium (15) into YPUAD, grown overnight, harvested from a logphase culture, washed, and resuspended to 106 cells per ml in fresh YPUAD. At this point, less than 10% of the cells had lost the plasmid (data not shown). After 30 min of incubation at 24°C with or without 10-6 M a-factor, the cells were fixed and FUS1-p-galactosidase was revealed using indirect immunofluorescence as described above. For quantitation of the experiment, photographs were taken over random areas and the cells were classified into four groups: unbudded cells, budded cells with the DNA staining still entirely in the mother cell, budded cells with the DNA migrating into the daughter cell, and large budded cells in which the DNA was clearly in both mother and daughter cell. Immunofluorescence of FUS1-,-galactosidase was then scored as positive or negative by comparison with the control cells that were not treated with a-factor. A total of 1,250 cells from the a-factor-treated culture were scored, using 883, 203, 39, and 125 cells from the four groups, respectively. RESULTS To measure a-factor internalization during mitosis, we synchronized cdc13, cdcl5, and cdc2O cells by a 150-min incubation at 37°C. The cdcl3 and cdc2O (data not shown) mutants arrest at an early stage of mitosis, probably corresponding to metaphase with a short spindle as detected by indirect immunofluorescence using antitubulin antibodies (Fig. 1). The cdclS mutant arrests at a later stage of mitosis, probably corresponding to anaphase with a fully extended spindle (Fig. 1) (41). The synchronized cdc mutants and wild-type cells similarly treated at 37°C were then incubated with 35S-a-factor at 0°C, washed, and allowed to internalize the at-factor at 24°C. External a-factor was removed by an acid treatment, and the internalized ao-factor was then determined by scintillation counting. Figure 2 shows that both wild-type and mutant cells were capable of rapid internalization of a-factor. The rate of a-factor internalization is a little slower in the cdc2O strain. This effect is not due to the arrest

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FIG. 1. Immunofluorescence detection of tubulin as a diagnostic marker of cell cycle position and synchrony. cdcl5 cells were grown in YPUAD medium to log phase at 24°C, shifted to 37°C for 150 min, harvested, and resuspended in fresh medium at 24°C to release the mitotic block synchronously. At 15-min intervals, cells were fixed and tubulin was localized by indirect immunofluorescence. The time (in minutes) after the shift to 24°C is indicated on the appropriate panels. The cdc13 and cdc28 mutants were treated like the cdc15 mutant but were fixed just after the 150-min incubation at 37°C. The bar represents 10 ,um.

in mitosis because cdc2O cells incubated for only 30 min at 37°C, and thus not synchronized in mitosis, show the same a-factor internalization curve (data not shown). It is clear from Fig. 2 that the cdcl5 mutant cells did not internalize all of the bound a-factor. This is a strain-dependent effect and is not due to a block at this particular position in the cell cycle

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FIG. 2. Internalization of a-factor during M phase. Wild-type (RH448; 0), cdc13 (RH209-4A; +), cdclS (RH210-3C; *) and cdc2O (RH294-7A; *) cells were grown to log phase in YPUAD medium at 24°C, shifted to 37°C for 150 min to block the cdc mutants in mitosis, harvested, and washed, and a-factor internalization was measured at 24°C as described in Materials and Methods. Each curve represents the average results of two separate experiments performed in duplicate. The curve presented for cdc2O is very similar to that found if the mutant cells are incubated for only 30 min at 37°C, which is not enough time to achieve a synchrony in mitosis.

(see Table 1). The clear ability of the mutant strains to internalize a-factor suggests that yeast cells do not stop receptor-mediated endocytosis during mitosis. To measure whether a-factor internalization is modulated at some other point of the cell cycle, we used the temperature-sensitive cdcl5 mutation to make synchronous cell cultures. cdcl5 cells were synchronized by incubation at 37°C for 150 min, harvested by centrifugation, and resuspended in fresh medium at 24°C (permissive temperature). At various time points after the shift to 24°C, the cells were fixed and tubulin was visualized by immunofluorescence (Fig. 1). After a 30-min incubation at 24°C, the mitotic spindles were beginning to break down; by 45 min, this breakdown was complete. The cells were in the G1 phase of the cell cycle because they appeared similar to cells arrested at START by the cdc28 mutation (Fig. 1). At 60 min, the tubulin staining was still homogeneous and the cells had tiny buds. By 90 min, the cells had large buds and the tubulin staining was mainly localized near the neck of the bud, indicating that the nucleus was starting to migrate into the daughter cell. After 120 min, the cells had returned to a mitotic state similar to the cdcl5-induced block, showing a strong staining of a fully extended spindle. At 150 min, the cells still showed a reasonable synchrony and had returned to the typical asterlike tubulin pattern of G1 near START. The time for a complete round of the cell cycle was approximately 105 min.

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TABLE 1. a-factor internalization during the cell cyclea Time (min) at 24oCb

Initial uptake ratec

% Uptake at 20 min

0 15 30 45 60 75 90 105 120 135 150

7.2 7.0 7.5 6.9 6.9 7.0 7.4 6.5 6.6 6.5 6.5

85 84 87 91 80 88 87 82 92 90 89

a RH210-3C (cdcl5) cells were accumulated at mitosis by incubation at 37°C and then released synchronously from the block. At 15-min intervals, aliquots of cells were harvested and a-factor internalization was measured as described in Materials and Methods. b Time after shift to 24°C when the aliquot of cells was harvested. c Percentage of the bound a-factor internalized per minute. The percentage of the input a-factor that binds to the cells does not change to a large extent during the cycle.

We next tested the synchronized cells for a-factor uptake. Every 15 min after the shift to 24°C, cells were harvested and washed, and 35S-a-factor uptake was measured over a 20min period. The results (Table 1) show that the rate of a-factor uptake at all points measured during the cell cycle was constant. We have not detected any period of the cycle when cells could not internalize a-factor, nor have we been able to detect any large fluctuation (greater than fourfold) in a-factor binding during the cell cycle (data not shown), indicating that receptors were most likely present in roughly similar amounts throughout the cell cycle. As receptors are constantly present during the cell cycle and capable of internalizing a-factor, we were interested in determining whether the signal transduction pathway was also intact. To perform such experiments, it was necessary to quantify the response after a relatively short incubation with pheromone. Therefore, we monitored the a-factordependent induction of the FUSI gene (49). FUSI induction can be measured 15 to 20 min after a-factor addition and is a direct measure of signal transduction since no new protein synthesis is necessary for its induction (35). As a preliminary experiment, we transformed wild-type cells with pSB231, a plasmid carrying a FUSI-lacZ chimeric gene, that has been used before to measure FUSI induction (14, 49). Unsynchronized wild-type cells were incubated with or without a-factor for 30 min and then fixed, and ,B-galactosidase was detected by indirect immunofluorescence (Fig. 3). One can clearly see that the detection of ,-galactosidase was specific, since cells that had not been incubated with a-factor showed only background staining. Upon close examination, one notices that few of the cells with small buds show specific labelling for 3-galactosidase. To quantify this difference, we stained the cells with DAPI (data not shown), classified the cells into four groups, and scored them for positive immunofluorescence. The four groups were unbudded cells, budded cells with the nucleus in the mother cell, budded cells with the nucleus starting to migrate into the daughter cell, and cells that had begun mitosis but had not divided. The percentages of cells in these groups that showed positive immunofluorescence for ,B-galactosidase were 55, 37, 79, and 78, respectively. These differences were probably not due to differential permeabilization of the cells at the different stages

FIG. 3. FUS1-p-galactosidase induction in asynchronous cells. RH448 (wild-type) cells carrying plasmid pSB231 were grown in YPUAD medium at 24°C to log phase, treated with (+) or without (-) 10-6 M a-factor for 30 min at 24°C, and then fixed, and P-galactosidase was detected by indirect immunofluorescence. The exposure time was 100 s. The bar represents 10 L.m. Plasmid loss was less than 10% under these conditions.

because our immunofluorescence experiments using tubulin antibody never showed this property. These results suggest that there is a point in the cell cycle near the end of G1 or beginning of S phase (unbudded cells and cells with small buds at the end of the incubation) at which yeast cells do not respond efficiently to a-factor. To examine this matter in more detail, we assayed 3-galactosidase after synchronization of pSB231-transformed cdcl5 cells. At 15-min intervals after release from the cdcl5-induced block, cells were incubated for 22.5 min with a-factor, lysed, and assayed for 3-galactosidase activity. The results of this experiment (Fig. 4) show that induction of the FUSI gene varies considerably during the cell cycle. At the time of release from the cdcl5-imposed block, the

PHEROMONE UPTAKE AND RESPONSE DURING THE CELL CYCLE

VOL. 11, 1991

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FIG. 4. FUS1-0-galactosidase induction in synchronous cells. RH210-3C (cdcl5) cells carrying plasmid pSB231 were grown to log phase in YPUAD medium at 24°C, incubated for 150 min at 37°C, harvested, and resuspended in YPUAD medium at 24°C to release the mitotic block synchronously. At 15-min intervals, 106 M a-factor was added and the cells were further incubated for 22.5 min. The cells were harvested and lysed with glass beads, and ,3-galactosidase and alkaline phosphatase activities were measured. The abscissa represents the time at which a-factor was added. Values graphed on the ordinate are the fluorescence units obtained from the assay of p-galactosidase divided by those obtained from the assay of alkaline phosphatase (with [*] and without [0] a-factor). Data are those from one experiment; data from other experiments confirm these results, and assays of several other points without a-factor show that these values are the same throughout the cell cycle.

FUSJ-lacZ induction was very low. This effect was due to the cdc15-imposed block and not to the position of the cells in the cell cycle because cells in a similar position after one full round of the cell cycle (120 min) responded well. Maximal induction was seen when the cells had large buds (90 and 195 min), whereas minimal induction occurred when the cells were unbudded (150 min). The cells that responded the least (150 min) showed a tubulin staining typical of START when a-factor was added. At 15 min before or after this time point, the tubulin immunofluorescence looked different (data not shown; compare the tubulin immunofluorescence patterns at 30 and 60 min with the pattern at 45 min). This result agrees well with the immunofluorescence data shown above (Fig. 3). At the end of the incubation with a-factor, those cells that responded the least had small buds. Our data suggest that cells in all stages of the cycle can respond to a-factor, but that cells located at or just after the START point in the cell cycle are less efficient in induction of the FUSI gene. We cannot rule out that pheromone induction of FUSJ is blocked for a very short period of the cycle. To check specifically the START point of the cycle, we analyzed FUSI induction in cdc35 (START II [11]) and cdc28 (START I [11]) arrested cells by Northern blotting and enzyme assay. FUSI mRNA was induced by a-factor at both of these arrest points (data not shown). Our results with FUSI induction could be explained by a modulation of the signal transduction pathway during the cell cycle or an overlayed cell cycle control of FUSI expression. Therefore, we decided to examine the induction of another pheromone-responsive gene during the cell cycle. mRNA levels for the STE2 gene, encoding the a-factor receptor, also increase upon short exposure of a cells to a-factor. To test this induction during the cell cycle, cdcl5 cells were synchronized and exposed to a-factor for 22.5 min at 24°C starting at 45, 90, 150, and 195 min after release of the cdcl5 block. These times correspond to the troughs and peaks in FUS1 induction. After incubation with a-factor,

a "

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135 90 1S0 MINUTES AT 240C

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FIG. 5. STE2 and FUSI induction during the cell cycle. Synchronous cells (see Fig. 4) without plasmid were treated with a-factor at 24°C for 22.5 min beginning at the times plotted on the graph. RNA was analyzed by Northern blotting as described in Materials and Methods, using radioactive probes for STE2, URA3, FUSI, and BIKI. (a) Results for the different mRNAs without (-) and with (+) a-factor. For STE2 induction, a-factor was added to 10-8 M; for FUSI induction, it was added to 106 M. These quantities give a maximal response in asynchronous cells (14). (b) Quantification of the results from less exposed films by scanning densitometry, plotted by using arbitrary units. The values plotted for STE2 are the actual values measured, whereas the values for FUSI are divided by those for BIKI. This correction was necessary for FUS1 only because poly(A) RNA was probed to detect FUSI.

RNA was extracted and analyzed by Northern blotting to quantify the levels of STE2 mRNA. We also analyzed the expression of FUS1, URA3, and BIKI mRNAs. As can be seen in Fig. 5, basal mRNA levels for STE2 varied during the cell cycle, being lower at 45 and 150 min than at 90 and 195 min. However, the mRNA levels were increased at all times after exposure to a-factor. The levels of induction by a-factor of STE2 mRNA levels were eight-, four-, eight-, and fivefold at 45, 90, 150, and 195 min, respectively. This result shows that the signal transduction pathway to the STE2 gene was intact even when FUSI induction was the lowest. FUSI mRNA was induced to levels that are fully compatible with the results that we found measuring the induction of the FUSIJ--galactosidase fusion protein. Quantitation of the results shows that the amounts of FUSI, basal STE2, and induced STE2 mRNAs varied with approximately the same pattern at these points of the cell cycle. We were unable to detect any basal expression of the FUSI transcript even with very long exposures to X-ray film. BIKI and URA3 did not vary in this manner, even though we consistently found some modulation of URA3 mRNA. This mRNA was always slightly more abundant near START than near G2. DISCUSSION

Three main conclusions can be drawn from this work. First, a-factor binding to its receptor on a cells and subsequent endocytosis are not regulated to a large extent during the S. cerevisiae cell cycle. Second, the ability to transduce the pheromone signal does not seem to be strongly regulated

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during the cell cycle. Third, even though pheromone signalling can occur throughout the cell cycle, there is a cell cycle control on the expression of at least two pheromone-inducible genes. The finding that receptor-mediated endocytosis can occur in yeast cells at all stages of the cell cycle, in particular during mitosis, is in sharp contrast to the results found in mammalian cells (3, 4), in which endocytosis is blocked during mitosis. However, the continuation of endocytosis throughout mitosis in yeast cells is consistent with the finding that secretion by yeast cells also continues normally during mitosis (34, 39). This difference in membrane traffic between yeast and animal cells probably reflects a difference in the strategy for organelle division during mitosis. In mammalian cells, the nucleus undergoes an open mitosis and organelle components become distributed throughout the cytoplasm (32, 51). S. cerevisiae cells undergo a closed mitosis with the nucleus being divided between the mother and daughter cell (9). The dispersal of the compartments during mitosis in mammalian cells is most likely necessary to ensure that both cells receive a full complement of organelles. Evidence is accumulating from in vitro and in vivo studies in mammalian cells that the p34Cdc28 protein kinase is involved in the down-regulation of endocytic traffic (50) and breakdown of the nuclear membrane (24, 40). Even though this kinase is expressed and plays a role in mitosis in S. cerevisiae (43, 47), it does not seem to have the same effect on membrane traffic. The basis for this difference is unknown. In our analysis of pheromone response during the cell cycle, we did not expect to find a modulation of the pheromone-stimulated expression of the FUSI transcript. Induction was greatest near G2 and lowest near or just after START. This was shown in three ways: by immunofluorescence analysis of FUS1 induction in asynchronous wild-type cells and by enzyme assay and Northern blotting in synchronous cdcl5 cells. We found a similar modulation in the expression of the STE2 mRNA, both in the absence and in the presence of pheromone. These levels parallel the FUS1 levels quite well, which would be consistent with a common control mechanism. The variations that we measured are not likely to be due to differential rates of macromolecular synthetic capacity during the cell cycle because the measured rates do not vary (17). This modulation of expression of FUSI and STE2 during the cell cycle is probably not dependent on a modulation of pheromone signal transduction because the fold induction of the STE2 mRNA near START was even higher than near G2. These results suggest that some type of general cell cycle control on pheromoneinducible genes may be operative. Upon pheromone induction, this cell cycle control would remain. If this is the case, one would also expect to see a cell cycle modulation of the basal FUS1 mRNA levels. Unfortunately, we were unable to detect any FUS1 mRNA in the absence of pheromone. Since we cannot detect the basal FUSI mRNA, we cannot rule out the possibility that the signal transduction pathway to FUSI and STE2 is branched and that one arm of this pathway, leading to FUSI, is under a separate type of cell cycle control. Nor can we rule out the possibility that the STE2 gene is regulated during the cell cycle and that this is responsible for the modulation of the FUSI response. However, this latter possibility seems unlikely because transformation of a cells with an additional copy of the STE2 gene, leading to an enhanced number of cell surface receptors, does not trigger a greater response (14). In this study, we have measured the levels of two phero-

MOL. CELL. BIOL.

mone-inducible genes, but our findings are likely to be expandable to other pheromone-inducible genes because the genes that we studied are needed for very different functions and at different times in the conjugation pathway. The a-factor receptor (STE2 gene product) is needed early in the pheromone response and has to be present at least near the point of cell cycle arrest since a-factor must be present for 90 min in order to induce the full response (2). The FUSI gene product acts late in the conjugation pathway, playing a role in cell-cell fusion (49). If the control mechanism that we have uncovered is truly expandable to all conjugation-related genes, then the candidate controlling proteins, those that are modulated during the cell cycle, would have to recognize, directly or indirectly, all of the appropriate genes or mRNAs. Therefore, some of the candidates would be proteins of the signal transduction pathway since some of the corresponding genes exert a control on the basal expression of haploidspecific gene products. It is perhaps too early to speculate which of the many known genes that affect expression of conjugation-related gene products may be involved. A few studies have examined pheromone-induced gene expression during the cell cycle. Hagen and Sprague (20) used cells blocked in the cycle to examine a-factor induction of the STE3 (a-factor receptor) gene. Their results are consistent with ours. They showed that the STE3 gene could be induced at several different arrest points in the cell cycle. Terrance and Lipke (48) showed that agglutinins could be induced equally in G1 and G2 phases; however, these studies used cells obtained by centrifugal elutriation, and the G, cells could be from a wide portion of this phase. In conclusion, our findings demonstrate that there is a cell cycle control on the expression of some pheromone-inducible genes, with their expression being lowest near the Gl-to-S transition. This is the point in the cell cycle at which the decision is made whether to embark on a new round of the cell cycle or to enter into the conjugation pathway. One possibility would be that the same process that promotes the decision to embark upon a new round of the cell cycle also inhibits the expression of pheromone-induced genes. In this manner, progression through START and into S phase would suppress the expression of conjugation-related genes at a time when the cells have just committed themselves to an alternate developmental pathway. This could be advantageous in two ways. First, it would result in an economy due to the low expression of conjugation-related genes that are not immediately needed. Second, it would still allow the cells to sense and respond to pheromone. This response would increase during S and G2 phases as the cells progress toward the next decision point for entry into the conjugation pathway. ACKNOWLEDGMENTS We thank Fabienne Crausaz for technical assistance, Grace Parraga, Mark Egerton, and Birgit Singer for critical reading of the manuscript, and Karin Vogt for her secretarial skills. This work was supported by a grant from the Swiss National Science Foundation and the Canton of Basel. REFERENCES 1. Arion, D., L. Meier, L. Brizuela, and D. Beach. 1988. cdc2 is a component of the M phase-specific histone Hi kinase: evidence for identity with MPF. Cell 55:371-378. 2. Baffi, R. A., P. Shenbagamurthi, K. Terrance, J. M. Becker, F. Naider, and P. N. Lipke. 1984. Different structure-function relationships for a-factor-induced morphogenesis and agglutination in Saccharomyces cerevisiae. J. Bacteriol. 158:1152-1156. 3. Berlin, R. D., and M. J. Oliver. 1980. Surface functions during

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