Skip to main content
NCBI home page
Molecular Biology of the Cell logoLink to Molecular Biology of the Cell
. 2024 Feb 26;35(4):ar46. doi: 10.1091/mbc.E23-05-0174

Evidence for novel mechanisms that control cell-cycle entry and cell size

Amanda Brambila a, Beth E Prichard a, Jerry T DeWitt a, Douglas R Kellogg a,*
Editor: Sophie Martinb
PMCID: PMC11064657  PMID: 38231863

Abstract

Entry into the cell cycle in late G1 phase occurs only when sufficient growth has occurred. In budding yeast, a cyclin called Cln3 is thought to link cell-cycle entry to cell growth. Cln3 accumulates during growth in early G1 phase and eventually helps trigger expression of late G1 phase cyclins that drive cell-cycle entry. All current models for cell-cycle entry assume that expression of late G1 phase cyclins is initiated at the transcriptional level. Current models also assume that the sole function of Cln3 in cell-cycle entry is to promote transcription of late G1 phase cyclins, and that Cln3 works solely in G1 phase. Here, we show that cell cycle−dependent expression of the late G1 phase cyclin Cln2 does not require any functions of the CLN2 promoter. Moreover, Cln3 can influence accumulation of Cln2 protein via posttranscriptional mechanisms. Finally, we show that Cln3 has functions in mitosis that strongly influence cell size. Together, these discoveries reveal the existence of surprising new mechanisms that challenge current models for control of cell-cycle entry and cell size.

  • Current models for cell-cycle entry assume that transcription of late G1 phase cyclins is the critical event that initiates cell-cycle entry. In budding yeast, a cyclin called Cln3 is thought to function solely to initiate transcription of late G1 phase cyclins.

  • The authors show that posttranscriptional mechanisms play a major role in controlling expression of late G1 phase cyclins, and that Cln3 has functions in mitosis that strongly influence cell size.

  • These discoveries reveal the existence of surprising new mechanisms for control of cell-cycle entry and cell size.

INTRODUCTION

The decision to enter a new round of cell division is among the most consequential decisions in the life of a cell. Entry into the cell cycle occurs only when sufficient growth has occurred and only when there are sufficient nutrients for further growth. In animal cells, cell-cycle entry is also controlled by growth factors, which ensure that cell division occurs at an appropriate time and place. Defects in signals that control cell-cycle entry are a primary cause of cancer.

The mechanisms that initiate cell-cycle entry are poorly understood (Rubin et al., 2020). In budding yeast, decades of work led to a canonical model in which a cyclin called Cln3 appears in early G1 phase and activates Cdk1 (Carter and Sudbery, 1980; Cross, 1988, 1989; Nash et al., 1988; Hadwiger et al., 1989; Richardson et al., 1989; Tyers et al., 1992, 1993). The Cln3/Cdk1 complex then directly phosphorylates and inactivates Whi5, a transcriptional repressor that binds and inhibits two transcription factors, referred to as SBF and MBF, that drive transcription of late G1 phase cyclins, as well as hundreds of additional genes (Nasmyth and Dirick, 1991; Tyers et al., 1993; Koch et al., 1996; Spellman et al., 1998; Iyer et al., 2001; Jorgensen et al., 2002; Wijnen et al., 2002; de Bruin et al., 2004; Costanzo et al., 2004; Bean et al., 2005; Ferrezuelo et al., 2010). Thus, Cln3-dependent inactivation of Whi5 has been thought to initiate transcription of late G1 cyclins, which is the critical molecular event that marks cell-cycle entry. Genetic analysis suggests that Cln3 and Whi5 link cell-cycle entry to cell growth. For example, loss of WHI5 causes premature cell-cycle entry before sufficient growth has occurred, leading to a reduced cell size (Jorgensen et al., 2002; Costanzo et al., 2004; de Bruin et al., 2004). Similarly, loss of Cln3 causes delayed cell-cycle entry and increased cell size, while overexpression of Cln3 leads to premature cell-cycle entry and reduced cell size (Sudbery et al., 1980; Cross, 1988, 1989; Nash et al., 1988; Tyers et al., 1993; Jorgensen and Tyers, 2004).

The canonical model has provided an important framework for analysis of cell-cycle entry and has strongly influenced models for how cell growth influences cell-cycle entry. However, a number of observations cannot be explained by the model. For example, overexpression of Cln3 causes a large reduction in the size of whi5∆ cells, which indicates that Cln3 has critical targets other than Whi5 (Costanzo et al., 2004; Wang et al., 2009). Furthermore, recent work has shown that Cln3 is not required for phosphorylation of Whi5 in vivo (Bhaduri et al., 2015; Kõivomägi et al., 2021). Rather, a recent study suggested a revised model in which Cln3/Cdk1 binds to SBF promoters and directly phosphorylates RNA polymerase to stimulate transcription (Kõivomägi et al., 2021). This revised model, as well as the canonical model, presume that Cln3 functions entirely at the transcriptional level to promote synthesis of late G1 phase cyclin proteins. However, this has never been directly tested.

A further concern regarding current models is that they assume that Cln3 exerts all its key functions in G1 phase, especially with respect to control of cell size. However, there are two peaks of Cln3 protein during the cell cycle – one in G1 phase and a second in mitosis (Landry et al., 2012; Zapata et al., 2014; Litsios et al., 2019). The functions of Cln3 during mitosis are unknown. Since most growth of a yeast cell occurs during bud growth in mitosis (Leitao and Kellogg, 2017), it is possible that mitotic functions of Cln3 play a major role in Cln3’s ability to influence cell size. No previous experiments have tested for mitotic functions of Cln3.

Here, we tested current models for cell-cycle entry and whether Cln3 has functions in mitosis. Together, the data indicate that posttranscriptional events play an important role in the mechanisms that control expression of late G1 phase cyclins. The data further show that Cln3 can strongly influence accumulation of late G1 phase cyclins via posttranscriptional mechanisms, and that Cln3 executes functions during mitosis that influence cell size. Together, these observations significantly expand the range of possible models for the mechanisms that control cell-cycle entry and cell size.

RESULTS

Cln3 can influence production of Cln2 protein via Whi5-independent mechanisms

Previous studies examined how Cln3 and Whi5 influence Cln2 mRNA levels. Here, we examine how Cln3 and Whi5 influence production of Cln2 protein, which is the critical output of the mechanisms that drive cell-cycle entry.

We first set out to test how Cln3 and Whi5 contribute to regulation of Cln2 protein expression. Recent work suggests that Cln3/Cdk1 does not phosphorylate Whi5 (Bhaduri et al., 2015; Kõivomägi et al., 2021); however, it remains possible that Cln3 drives dissociation of Whi5 from SBF via other mechanisms that could make a substantial contribution to expression of Cln2 protein. Therefore, we tested whether Cln3 can influence production of Cln2 protein via mechanisms that are independent of Whi5. To do this, we analyzed the effects of loss- or gain-of-function of Cln3 on production of Cln2 protein in whi5∆ cells. If Cln3 influences expression of Cln2 protein primarily via Whi5-dependent mechanisms, then loss- or gain-of-function of Cln3 should have little effect on Cln2 protein expression in whi5∆ cells. In these experiments, and following experiments, we detected Cln2 using several different N-terminal and C-terminal tags to ensure that results are not influenced by the location or composition of the tag. All tagged versions of Cln2 gave similar results, although detection of phosphorylated forms of Cln2 that have reduced electrophoretic mobility differed between the tagged versions.

We used cln3∆ to test the effects of loss of Cln3 function. To achieve gain of Cln3 function, we utilized the cln3-∆177 allele, which lacks C-terminal PEST sequences that target it for rapid turnover via ubiquitin-dependent proteolysis (Nash et al., 1988). Previous work found that cln3-∆177 results in a 10-fold increase in Cln3 protein levels, as well as a large decrease in cell size (Sudbery et al., 1980; Tyers et al., 1992). We synchronized wild-type, whi5∆, whi5∆ cln3-∆177, and whi5∆ cln3∆ cells in G1 phase using mating pheromone and assayed production of Cln2-3XHA during the cell cycle at 10-min intervals (Figure 1, A and B). Controls showed that cln3-∆177 alone accelerated expression of Cln2 protein, whereas cln3∆ alone delayed and reduced expression of Cln2 protein, as expected (Supplemental Figure S1). Loss of WHI5 accelerated production of Cln2 protein by approximately 10 min relative to wild type. Overexpression of Cln3 accelerated production of Cln2 protein in whi5∆ cells, while cln3∆ delayed production of Cln2 in the whi5∆ cells. Thus, Cln3 can strongly influence production of Cln2 protein via mechanisms that are independent of Whi5. Furthermore, Cln2 protein expression in whi5∆ cln3∆ cells was nearly identical to wild-type cells, which indicates that Cln3 and Whi5 are not required for cell cycle−dependent expression of Cln2.

FIGURE 1:

FIGURE 1:

Open in a new tab

Cln3 can influence production of Cln2 protein via Whi5-independent mechanisms. (A) Cells were grown in YPD medium overnight and synchronized in G1 phase using alpha factor. Cells were released from G1 arrest and samples were collected every 10 min to assay for Cln2-3XHA protein levels by western blotting using 12CA5 mouse monoclonal anti-HA. An anti-Nap1 antibody was used as the loading control. (B) Quantification of data in panel (A). Cln2 protein levels were normalized against loading control. Error bars represent the standard error of the mean for three biological replicates. (C) Cells were grown in YPGE medium overnight and synchronized in G1 phase using mating pheromone. 2% galactose was added to cultures at 40 min prior to release from G1 arrest. Cells were released from G1 arrest into YP medium containing 2% galactose and samples were collected every 10 min to assay for Cln2 protein levels by western blotting using 12CA5 mouse monoclonal anti-HA. (D) Quantification of data in panel (C). Error bars represent standard error of the mean for three biological replicates.

A concern with the use of cln3∆ and cln3-∆177 is that they cause substantial cell size defects that could indirectly influence the timing of cell-cycle entry. Therefore, as a further means of testing the effects of Cln3 overexpression, we utilized cells that express an extra copy of wild-type CLN3 from the inducible GAL1 promoter, which allowed us to test the immediate effects of increased Cln3 levels in otherwise normal cells. Wild-type and GAL1-CLN3 cells were arrested in G1 phase in media containing 2% glycerol and 2% ethanol to repress transcription of GAL1-CLN3. Expression of GAL1-CLN3 was induced with 2% galactose 40 min before release from the G1 phase arrest. We again found that overexpression of Cln3 accelerated production of Cln2 protein in whi5∆ cells (Figure 1, C and D). The effects of full-length CLN3 expressed from the GAL1 promoter were substantially stronger than the effects of cln3-∆177, as GAL1-CLN3 caused a large increase in the amount of Cln2 protein (compare Figure 1B and D). This may be due to the fact that cln3-∆177 lacks a nuclear localization sequence (Edgington and Futcher, 2001; Miller and Cross, 2001).

A previous study concluded that Cln3/Cdk1 promotes transcription of CLN2 via direct phosphorylation of RNA polymerase (Kõivomägi et al., 2021). The most straightforward interpretation of this model would predict that loss of Cln3 in whi5∆ cells should cause reduced transcription of CLN2 and a corresponding reduction in Cln2 protein levels. However, we found that cln3∆ did not cause a reduction in peak Cln2 protein levels in whi5∆ cells (Figure 1B, compare whi5∆ and whi5∆ cln3∆). Furthermore, cln3∆ whi5∆ cells express slightly more Cln2 protein than wild-type cells and the timing of expression of Cln2 protein in whi5∆ cln3∆ cells is indistinguishable from wild-type cells (Figure 1B).

Together, these data indicate that Cln3 and Whi5 are not required for robust cell cycle−dependent expression of Cln2. Thus, there are mechanisms for repression of Cln2 expression in early G1 phase and initiation of Cln2 expression at cell-cycle entry that work completely independently of Whi5 and Cln3. The data further indicate that Cln3 can strongly influence Cln2 protein expression via Whi5-independent mechanisms. Finally, the results suggest that Cln3-dependent phosphorylation of RNA polymerase (Kõivomägi et al., 2021) is unlikely to make a major contribution to the mechanisms responsible for cell cycle−dependent expression of Cln2 protein.

Cln3 can influence levels of Cln2 protein via posttranscriptional mechanisms

Cln3 could control transcription of CLN2 via mechanisms that are independent of Whi5, but dependent upon other features of the CLN2 promoter. Alternatively, Cln3 could influence production of Cln2 protein via posttranscriptional mechanisms, which has never been tested. To distinguish these possibilities, we created strains in which transcription of CLN2 is controlled by the MET25 promoter, thereby eliminating normal control of CLN2 transcription. Previous studies have shown that SBF and MBF cannot be detected at the MET25 promoter and that overexpression of Cln3 does not influence transcription of MET25 (Iyer et al., 2001; Ferrezuelo et al., 2010). The MET25 promoter drives a basal level of transcription when methionine is present in the growth medium, which we used to express CLN2.

We found that cln3-∆177 advanced expression of Cln2 protein in MET25pr-3XHA-CLN2 cells, whereas cln3∆ caused delayed and reduced expression of Cln2 protein (Figure 2, A and B). We carried out a further test using a different heterologous promoter. In this case, we used the YPK1 promoter, which does not undergo cell cycle−dependent transcription and does not bind SBF or MBF. In addition, the Ypk1 protein is expressed at constant levels throughout the cell cycle and across different carbon sources (Alcaide-Gavilán et al., 2018; Lucena et al., 2018). We found that GAL1-CLN3 accelerated production of Cln2 protein expressed from the YPK1 promoter (Figure 2, C and D). As in Figure 1, the effects of GAL1-CLN3 appeared to be stronger than the effects of cln3-∆177. Northern blot analysis confirmed that CLN2 mRNA expressed from the YPK1 promoter shows no cell cycle−dependent periodicity and that GAL1-CLN3 does not influence expression of CLN2 mRNA from the YPK1 promoter (Figure 2E).

FIGURE 2:

FIGURE 2:

Open in a new tab

Cln3 can influence levels of Cln2 protein via posttranscriptional mechanisms. (A) Cells were grown in YPD medium overnight and synchronized in G1 phase using alpha factor. Cells were released from G1 arrest and samples were collected every 10 min to assay for 3XHA-Cln2 protein levels by western blotting using 12CA5 mouse monoclonal anti-HA. An anti-Nap1 antibody was used as the loading control. (B) Quantification of the data in panel (A). Plots were generated by normalizing 3XHA-CLN2 to loading control. Error bars represent the standard error of the mean for four biological replicates. (C) Cells were grown in YPGE medium overnight and synchronized in G1 phase using alpha factor. 2% galactose was added to cultures 45 min prior to release from G1 arrest. Cells were then released from G1 arrest into YP medium containing 2% galactose. Western blotting was used to assay protein levels for Cln2 using anti-Myc and anti-Nap1 for loading control. (D) Quantification of the data in panel (D). Plots were generated by normalizing Cln2-9Myc to loading control. Error bars represent the standard error of the mean for three biological replicates. (E) Cells were grown in YPGE medium overnight and then synchronized in G1 phase using alpha factor. 2% galactose was added to cultures at 45 min prior to release from G1 arrest. Cells were released from G1 arrest into YP medium containing 2% galactose and samples were collected every 10 min to assay for CLN2 mRNA by Northern blot.

These results show that Cln3 can influence expression of Cln2 protein via mechanisms that are completely independent of the CLN2 promoter. Furthermore, Cln2 protein expressed from heterologous promoters showed strong cell-cycle periodicity, which provides further evidence that posttranscriptional mechanisms play a substantial role in the mechanisms that repress expression in early G1 phase and initiate cell-cycle entry in late G1 phase. Previous studies have shown that constitutive expression of CLN1 from the very strong GAL1 promoter can rescue the unviability of cln1∆ cln2∆ cln3∆ cells (Richardson et al., 1989). Although GAL1-CLN1 cln2∆ cln3∆ cells have an abnormal morphology, this observation shows that the essential functions of late G1 phase cyclins are not dependent upon regulation of their transcription, which further suggests that posttranscriptional mechanisms play an important role in regulating expression of late G1 phase cyclins and cell-cycle entry.

Cln3 cannot influence Cln2 protein levels in swi6∆ cells

Cln3 is thought to influence transcription of late G1 phase cyclins via two transcription factors, known as SBF and MBF. Both are dimers that include a DNA binding subunit and a shared subunit called Swi6 (Andrews and Herskowitz, 1989; Andrews and Moore, 1992; Siegmund and Nasmyth, 1996). SBF and MBF bind the promoters of late G1 phase cyclins, as well as hundreds of additional genes. Loss of Swi6 causes a large reduction in transcription of CLN2, as well as delayed cell-cycle entry and a large increase in cell size (Nasmyth and Dirick, 1991; Wijnen et al., 2002). Key phenotypic effects of CLN3 overexpression are not seen in swi6∆ cells. For example, overexpression of CLN3 does not cause reduced cell size in swi6∆ cells and fails to drive premature cell-cycle entry or premature transcription of CLN1 (Nasmyth and Dirick, 1991; Wijnen et al., 2002). However, we found that swi6∆ cells are barely viable and rapidly accumulate suppressor mutations. Furthermore, since Swi6 is a component of both SBF and MBF it controls the transcription of hundreds of genes (Iyer et al., 2001; Horak et al., 2002). Among these genes are 16 transcription factors, including several that strongly influence cell cycle−dependent transcription later in the cell cycle (Horak et al., 2002). Transcription factors downstream of SBF and MBF also control the transcription of numerous genes involved in ubiquitin-dependent proteolysis, which plays essential roles in controlling the levels of cyclins and other proteins (Horak et al., 2002). Thus, loss of Swi6 causes pervasive and complex cascading effects upon the expression of thousands of genes, including many that play central roles in cell-cycle control and protein turnover (Horak et al., 2002). Together, these considerations make it difficult to draw clear and rigorous conclusions based on the effects of swi6∆.

No previous studies investigated the effects of Cln3 on production of late G1 cyclin proteins in swi6∆ cells. Here, we tested whether loss of SWI6 influences accumulation of Cln2 in MET25pr-3XHA-CLN2 cells. In the presence of SWI6, GAL1-CLN3 advanced expression of Cln2 protein, as expected (Figure 3). Loss of Swi6 caused a loss of periodic expression of Cln2 protein in MET25pr-3XHA-CLN2 cells, and overexpression of CLN3 had no effect on expression of Cln2 protein in MET25pr-3XHA-CLN2 swi6∆ cells. Together, these data show that Swi6 is required for periodic expression of Cln2 protein. The data further suggest that Cln3 modulates levels of Cln2 protein at least partly via mechanisms that are dependent upon the many genes whose transcription is controlled, directly or indirectly, by SBF. However, the complex and pleiotropic effects of swi6∆ on expression of thousands of genes make it difficult to draw clear conclusions from the data.

FIGURE 3:

FIGURE 3:

Open in a new tab

Cln3 requires Swi6 to influence Cln2 protein levels. (A and B) Cells were grown in YPGE medium overnight and synchronized in G1 phase using alpha factor. 2% galactose was added to the cultures 40 min prior to release from G1 arrest. Cells were released from G1 arrest into YP medium containing 2% galactose and samples were collected every 10 min. Western blotting was used to assay protein levels for 3XHA-Cln2 using mouse monoclonal anti-HA (A) and Clb2 using an anti-Clb2 antibody (B).

Expression of Cln2 protein appeared to be repressed in early G1 phase in all four MET25pr-3XHA-CLN2 strains because very little Cln2 protein is detected at the earliest time point (Figure 3), which suggests that posttranscriptional Swi6-independent mechanisms repress accumulation of Cln2 protein in G1 phase.

Cln3 can influence cell size via mechanisms that are independent of transcriptional control of Cln1 and Cln2

Cln3 has been thought to influence cell size by regulating transcription of the late G1 cyclins (Cross, 1990; Tyers et al., 1993; Wijnen et al., 2002; Ferrezuelo et al., 2010). To further investigate, we tested whether overexpression of CLN3 influences cell size in a strain in which normal transcriptional control of both CLN1 and CLN2 has been eliminated. To do this, we created a strain in which the CLN1 gene was deleted and CLN2 was controlled by the MET25 promoter (MET25pr-3XHA-CLN2 cln1∆). We then integrated a copy of the wild-type CLN3 gene under the control of the GAL1 promoter. When grown in dextrose to repress expression of GAL1-CLN3, the MET25pr-3XHA-CLN2 cln1∆ GAL1-CLN3 cells were larger than wild type (Figure 4A). When grown in 2% galactose, overexpression of CLN3 drove a large decrease in the size of MET25pr-3XHA-CLN2 cln1∆ cells (Figure 4B; Supplemental Figure S2). The fact that Cln3 can drive a decrease in cell size in MET25pr-3XHA-CLN2 cln1∆ cells shows that Cln3 can influence cell size via mechanisms that are independent of transcriptional control of CLN1 and CLN2.

FIGURE 4:

FIGURE 4:

Open in a new tab

Cln3 can influence cell size via mechanisms that are independent of transcriptional control of Cln1 and Cln2. (A and B) Cells were grown in YPD medium or YP medium containing 2% galactose overnight to an OD600 between 0.4 and 0.6 and cell size was measured using a Beckman Coulter Counter Z2. Data shown represents the average of three biological replicates, where each biological replicate is the average of three technical replicates.

Cln3 influences protein levels of targets of the SCFGrr1 ubiquitin ligase complex

The preceding results indicate that Cln3 can influence expression of Cln2 protein via posttranscriptional mechanisms. The only posttranscriptional mechanism currently known to play a major role in controlling Cln2 protein levels is ubiquitin-dependent proteolysis. Cln2 is targeted for ubiquitylation and proteolytic turnover by the SCFGrr1 ubiquitin ligase complex. Loss of SCFGrr1 activity causes a failure in Cln2 protein turnover, leading to accumulation of abnormally high levels of Cln2 (Willems et al., 1996; Skowyra et al., 1997). Thus, one potential explanation for the effects of Cln3 is that it influences activity of the SCFGrr1 complex. Consistent with this, previous studies have found that Cln3 can bind the SCFGrr1 complex. Moreover, cln3-∆177 binds less effectively to components of the SCFGrr1 complex compared with full-length Cln3 (Willems et al., 1996; Landry et al., 2012), and we found that cln3-∆177 also appeared to be less effective than full-length CLN3 at causing increased levels of Cln2 protein, which suggests a model in which Cln3 binds and inhibits SCFGrr1 (compare Figure 1B and D). To begin to test this model, we determined whether levels of other known SCFGrr1 targets are influenced by loss- or gain-of-function of CLN3. Two of the best characterized targets of SCFGrr1 are Hof1 and Ndd1. Hof1 controls cytokinesis, whereas Ndd1 is an essential transcription factor that controls expression of a cluster of mitotic genes that includes mitotic cyclins (Blondel et al., 2005; Li et al., 2006; Edenberg et al., 2015). Both show strong cell cycle−dependent changes in protein levels.

We first analyzed Hof1-3XHA levels in asynchronous rapidly growing cells. Levels of Hof1 were decreased in cln3∆ cells and increased in cln3-∆177 cells (Figure 5A). In synchronized cells, cln3-∆177 led to an increase in Hof1 levels during mitosis, whereas cln3∆ led to a decrease in Hof1 levels (Figure 5B). We next compared the timing of expression of Hof1-3XHA and Cln3-6XHA in synchronized cells. Because the two proteins migrate at different locations in SDS–PAGE we were able to analyze levels of Hof1-3XHA and Cln3-6XHA in the same western blot. We also analyzed levels of the mitotic cyclin Clb2 as a marker for mitotic progression. Hof1-3XHA began to accumulate in early mitosis and reached peak levels late in mitosis, as levels of Clb2 began to decline (Figure 5C). Hof1 protein levels were strongly correlated with Cln3 protein levels, consistent with the possibility that Cln3 modulates levels of Hof1. The 6XHA tag used to detect Cln3 shows much higher sensitivity than the 3XHA tag used to detect Hof1, so relative levels of the two proteins cannot be compared.

FIGURE 5:

FIGURE 5:

Open in a new tab

Cln3 influences levels of Hof1 protein. (A) Western blots showing levels of Hof1-3XHA protein in asynchronous wild type, cln3∆, and cln3-∆177 cells. Bar graphs show quantification of three biological replicates. Error bars show standard error of the mean. (B) Cells were grown overnight in YPD medium and synchronized in G1 phase using alpha factor. Cells were released from G1 arrest and samples were collected every 10 min. The behavior of Hof1-3XHA was assayed by western blot using 12CA5 mouse monoclonal anti-HA. An anti-Nap1 antibody served as a loading control. The graph shows quantification of data from three biological replicates normalized to loading control. Error bars represent the standard error of the mean. (C) Wild-type cells containing Cln3-6XHA and Hof1-3XHA were grown overnight in YPD medium and sychronized in G1 phase with alpha factor. After release from the G1 arrest, the behavior of Cln3-6XHA and Hof1-3XHA was assayed by western blot using mouse monoclonal anti-HA.

We carried out a similar analysis for Ndd1. As with Hof1, cln3∆ caused a decrease in Ndd1 protein levels in both asynchronous and synchronous cells. cln3-∆177 caused an increase in Ndd1 levels, although the increase in this case was not statistically significant (Figure 6, A and B). In contrast to Hof1, Ndd1 accumulates slightly before mitosis, consistent with its essential role in induction of transcription of mitotic cyclins and other mitotic genes. Therefore, Ndd1 levels are not closely correlated with Cln3 levels, and the effects of Cln3 on Ndd1 protein levels could be an indirect consequence of increased levels of late G1 cyclins, which are known to initiate mitotic transcription programs.

FIGURE 6:

FIGURE 6:

Open in a new tab

Cln3 influences levels of Ndd1 protein. (A) Western blots showing levels of Ndd1-3XHA protein in asynchronous wild type, cln3∆, and cln3-∆177 cells using mouse monoclonal anti-HA. Bar graphs show quantification of three biological replicates normalized to loading control. Error bars show standard error of the mean. (B) Cells were grown overnight in YPD medium and synchronized in G1 phase using alpha factor. Cells were released from G1 arrest and samples were collected every 10 min. The behavior of Ndd1-3XHA was assayed by western blot. An anti-Nap1 antibody served as a loading control. The graph shows quantification of data from three biological replicates. Error bars represent the standard error of the mean.

At the least, these results demonstrate that loss- or gain-of-function of Cln3 causes complex effects on the proteome that make it difficult to draw simple conclusions about the effects of Cln3 mutants. Moreover, these results are consistent with the idea that Cln3 influences the activity of SCFGrr1, but do not rule out alternative models. Extensive additional work will be needed to further test whether Cln3 can influence the activity of the SCFGrr1 complex.

Cln3 carries out functions in mitosis that influence cell size

Cln3 has been thought to exert all its effects in G1 phase. However, there is a second peak of Cln3 protein in mitosis (Landry et al., 2012; Zapata et al., 2014; Litsios et al., 2019), and we found that Cln3 can influence the expression of the Hof1 and Ndd1 proteins, which function after G1 phase. Together, these observations suggest that Cln3 could have functions when it appears later in the cell cycle during mitosis. Because most growth of budding yeast cells occurs during mitosis (Leitao and Kellogg, 2017), a mitotic function of Cln3 could make a major contribution to the effects of Cln3 on cell size. To investigate further, we used microscopy to analyze how loss- or gain-of-function of Cln3 influences the duration and extent of bud growth in mitosis. Because effects of Cln3 on cell size in G1 phase could influence growth in mitosis, we used conditional alleles to inactivate or overexpress Cln3 after cells passed through G1 phase. Conditional overexpression of Cln3 was achieved by expression of CLN3 from the inducible GAL1 promoter. Conditional inactivation of Cln3 was achieved with an auxin inducible degron version of CLN3 (cln3-AID) (Nishimura et al., 2009). The cln3-AID allele caused a modest increase in cell size in the absence of auxin, which indicated that the AID tag caused decreased function of Cln3. Prolonged growth of cln3-AID cells in the presence of auxin caused a larger increase in cell size, although the increase was not as large as the increase caused by cln3∆, which indicated that the cln3-AID allele caused a partial loss of function of Cln3 (Supplemental Figure S3).

GAL1-CLN3 cells were arrested in G1 phase in media containing 2% glycerol and 2% ethanol to repress the GAL1 promoter. Cells were released from the arrest and galactose was added to induce expression of CLN3 when 15% of the cells had undergone bud emergence. To analyze loss of function, cln3-AID cells were released from a G1 arrest and auxin was added when 40−50% of the cells had undergone bud emergence. We induced gain- or loss-of-function of CLN3 at different times relative to bud emergence because auxin-induced destruction of AID-tagged proteins occurs relatively rapidly (10−15 min), whereas high-level expression of proteins from the GAL1 promoter occurs more slowly (20−30 min).

The durations of metaphase and anaphase were analyzed in single cells using a fluorescently tagged spindle pole protein, as previously described (Leitao and Kellogg, 2017; Jasani et al., 2020). The spindle poles in wild-type control cells were tagged with mCherry, while the spindle poles in cln3-AID and GAL1-CLN3 were tagged with GFP, which allowed analysis of control and experimental cells simultaneously under identical conditions. Cell growth was analyzed by measuring the size of the daughter bud as a function of time.

Overexpression of Cln3 caused a decrease in the durations of both metaphase and anaphase, as well as a decrease in the size at which daughter buds complete metaphase and anaphase (Figure 7, A and B). Conversely, loss of function of Cln3 caused an increase in the duration of metaphase and an increase in daughter bud size at the end of metaphase (Figure 7, C and D). Loss of function of Cln3 did not cause a statistically significant change in the duration of anaphase.

FIGURE 7:

FIGURE 7:

Open in a new tab

Cln3 carries out functions in mitosis that influence cell size. Mitotic spindle poles in wild type control cells were labeled with Spc42-mRUBY2, whereas the mitotic spindle poles of GAL1-CLN3 or cln3-AID cells were labeled with Spc42-eGFP. For experiments comparing wild-type to GAL1-CLN3, cells were grown in complete synthetic media (CSM) containing 2% glycerol and 2% ethanol overnight. The cells were then arrested in G1 phase using alpha factor and wild-type and mutant cells were mixed before releasing from the arrest. Cells were released in CSM containing 2% glycerol/ethanol and 2% galactose was added 60 min after release. For experiments comparing wild-type cells to cln3-AID, cells were grown overnight in CSM containing 2% dextrose and were arrested in G1 phase using alpha factor. The wild type and mutant cells were mixed before releasing from the G1 phase arrest. 0.5 mM auxin was added at 50 min after release. Cells were imaged by confocal microscopy at 3-min intervals at a constant temperature of 27°C. (A) Graphs comparing the duration of each stage of mitosis in wild-type and GAL1-CLN3 cells. (B) Graphs comparing bud volume at the end of metaphase and the end of mitosis in wild-type and GAL1-CLN3 cells. (C) Graphs comparing the duration of each stage of mitosis in wild-type and cln3-AID cells. (D) Graphs comparing bud volume at the end of metaphase and the end of mitosis in wild-type and cln3-AID cells. Bars indicate the mean and the standard error of the mean for n = 29 (wild type, GAL1CLN3) in panels A and B, and n = 49 (wild type), and n = 45 (cln3-AID) in panels (C) and (D).

Together, these data show that Cln3 can influence the duration and extent of growth in mitosis, which suggests Cln3 has functions outside of G1 phase that strongly influence cell size. The data are consistent with our finding that Cln3 can influence expression of proteins outside of G1 phase (Figures 5 and 6). Previous studies have shown that loss of Hof1 causes a large increase in cell size, which suggests that decreased expression of Hof1 could contribute to the large size of cln3∆ cells. However, the mechanisms by which Cln3 influences cell growth and size in mitosis remain largely unknown.

DISCUSSION

Expression of late G1 phase cyclin proteins is strongly influenced by posttranscriptional mechanisms

Current models suggest that expression of late G1 phase cyclins during cell-cycle entry is controlled primarily at the transcriptional level. Thus, it has been thought that initiation of Cln2 transcription is the critical molecular event that drives cell-cycle entry, and that Cln3 influences cell-cycle entry solely via transcriptional mechanisms that initiate expression of Cln2 and other late G1 phase cyclins (Tyers et al., 1993; Dirick et al., 1995; Costanzo et al., 2004; de Bruin et al., 2004; Kõivomägi et al., 2021). Here, we carried out new tests of these models. Importantly, we analyzed production of Cln2 protein as the critical output of mechanisms that drive cell-cycle entry. We found that Cln3 and Whi5, which are thought to be critical regulators of CLN2 transcription and cell-cycle entry, are not required for robust cell cycle−dependent expression of Cln2 protein. Thus, in cln3∆ whi5∆ cells Cln2 protein expression is fully repressed before cell-cycle entry and accumulation of Cln2 protein is initiated with normal timing relative to wild-type cells. We further discovered that Cln2 protein expression shows strong cell cycle-dependent expression even when it is expressed from heterologous promoters that are not regulated by Whi5 or SBF. Finally, we found that Cln3 can influence production of Cln2 protein when Cln2 is expressed from heterologous promoters, which indicates that Cln3 can influence Cln2 protein levels via posttranscriptional mechanisms.

These surprising discoveries show that there are major mechanisms for repression and initiation of Cln2 protein expression in G1 phase that operate independently of Cln3 and Whi5. Furthermore, it has been thought that growth influences cell-cycle entry via Cln3 and Whi5, yet it appears that there are no substantial defects in cell-cycle entry in cells that lack both proteins, which suggests that it is possible that growth-dependent signals can initiate expression of Cln2 and cell-cycle entry via mechanisms that are independent of Cln3 and Whi5. The data further show that posttranscriptional mechanisms play a major role in regulating expression of Cln2 during cell-cycle entry. A previous study that used orthogonal methods independently concluded that growth-dependent expression of Cln2 protein in late G1 phase does not require the CLN2 promoter and that posttranscriptional mechanisms play an important role in regulation of Cln2 protein expression before cell-cycle entry (Schneider et al., 2004). Overall, these observations show that the mechanisms that control cell-cycle entry remain deeply mysterious.

Two independent studies detected Cln3 at SBF promoters via chromatin immunoprecipitation assays, which suggests that Cln3 carries out functions at SBF promoters (Wang et al., 2009; Kõivomägi et al., 2021). A recent study concluded that a Cln3/Cdk1 complex functions at SBF promoters to directly phosphorylate and activate RNA polymerase (Kõivomägi et al., 2021). This model was supported by previous studies in which it was found that loss of CLN3 in an otherwise wildtype background causes reduced transcription of CLN2 (Cross and Tinkelenberg, 1991; Tyers et al., 1993; Di Como et al., 1995; Dirick et al., 1995). Here, we found that loss of CLN3 does not cause a reduction in peak Cln2 protein levels in whi5∆ cells, which is inconsistent with the idea that Cln3 acts independently of Whi5 to promote activation of RNA polymerase at the CLN2 promoter. We also found that cells that lack both Whi5 and Cln3 express more Cln2 protein than wild-type cells, and the timing of Cln2 expression is indistinguishable from wild-type cells (Figure 1B). Together, the data are more consistent with previous genetic data that led to a model in which the primary function of Cln3 at the CLN2 promoter is to drive release of Whi5 from SBF (Costanzo et al., 2004; de Bruin et al., 2004). The available data do not rule out the possibility that Cln3 drives release of Whi5 via mechanisms that work at the posttranscriptional level to promote production of more Cln2/Cdk1, which is thought to directly phosphorylate proteins at the Cln2 promoter to drive release of Whi5. Loss of Cln3 caused a slight delay in Cln2 protein expression in whi5∆ cells compared with whi5∆ alone, which could be due entirely to posttranscriptional mechanisms (Figure 1B). Overall, however, the mechanisms by which Cln3 drives release of Whi5 remain poorly understood.

The key experiments that led to a model in which Cln3/Cdk1 phosphorylates RNA polymerase were based on experiments that utilized engineered Cln3-Cdk1 fusion proteins (Kõivomägi et al., 2021). The fusion protein approach was used because it has not been possible to purify a Cln3/Cdk1 complex, which is surprising because other cyclins bind Cdk1 to form a tight stoichiometric complex that can only be dissociated under denaturing conditions. The fact that it has not been possible to purify a Cln3/Cdk1 fusion protein indicates that Cln3 binds Cdk1 with dramatically lower affinity. Consistent with this, one well-controlled study did not detect any Cdk1 activity associated with Cln3 (Schneider et al., 2004), whereas several others detected only very weak activity (Tyers et al., 1992, 1993; Levine et al., 1996; Miller et al., 2005). The fact that Cln3 binds Cdk1 with dramatically lower affinity raises the possibility that the Cdk1 in a Cln3-Cdk1 fusion protein could form complexes with other cyclins, which bind Cdk1 with a much higher affinity. Thus, Cln3-Cdk1 fusion proteins could form highly active Cln3-Cdk1/cyclin complexes and recruit them to the CLN2 promoter via the known interaction of Cln3 with SBF promoters. In this case, the recruitment of ectopic highly active Cdk1/cyclin complexes to the CLN2 promoter would be expected to robustly phosphorylate and activate RNA polymerase because it is well known that there are Cdk/cyclin complexes that specifically control transcription via phosphorylation of the RNA polymerase (Lim and Kaldis, 2013). Similarly, the activity of Cln3-Cdk1 fusion proteins in vitro could represent the sum activity of multiple diverse Cln3-Cdk1/cyclin complexes. Additional controls will be necessary to rule out these kinds of scenarios and to more clearly establish the role of Cln3 at SBF promoters.

The discovery that Cln3 can influence production of Cln2 protein via posttranscriptional mechanisms has important implications, as it expands the range of possible models for control of cell-cycle entry and cell size. For example, previous studies have shown that Cln2 promotes its own transcription via a positive feedback loop in which Cln2/Cdk1 is thought to directly phosphorylate and regulate proteins at the CLN2 promoter (Nasmyth and Dirick, 1991; Costanzo et al., 2004; de Bruin et al., 2004; Skotheim et al., 2008; Wagner et al., 2009). Thus, an ability of Cln3 to directly influence Cln2 protein levels could determine when the positive feedback loop is engaged to drive full entry into the cell cycle. Because Cln3 accumulates gradually during G1 phase in a manner that is dependent upon and proportional to cell growth and nutrient availability (Litsios et al., 2019; Sommer et al., 2021), Cln3 could influence the amount of growth required to trigger cell cycle entry, which would provide a mechanism for linking cell cycle entry to cell growth and nutrient availability.

Several models could explain how posttranscriptional mechanisms help enforce periodic expression of late G1 phase cyclins. The rise in Cln2 levels in G1 phase could be explained by mechanisms that inhibit SCFGrr1. For example, signals related to cell growth could inhibit SCFGrr1 to drive an increase in Cln2 protein levels. If Cln3 were to influence the activity of SCFGrr1, it could influence the amount of growth needed to initiate expression of Cln2 protein. The decline in Cln2 levels at the end of G1 phase could be explained by a model in which cyclin/Cdk complexes expressed after cell-cycle entry relay signals that inhibit the SCFGrr1 complex.

Numerous models are possible at this point and considerable additional work will be required to gain a better understanding of the mechanisms that drive cell cycle−dependent expression of Cln2 protein and cell-cycle entry. The mechanisms that drive cell-cycle entry in mammalian cells also remain poorly understood (Rubin et al., 2020). Thus, a better understanding of cell-cycle entry in yeast could lead to the discovery of conserved mechanisms that are relevant to mammalian cells.

Cln3 is likely to have pervasive effects on the proteome

The only major posttranscriptional mechanism that is known to regulate Cln2 protein levels is ubiquitin-dependent proteolysis, which is mediated by the SCFGrr1 ubiquitin ligase complex. Therefore, the most straightforward hypothesis for how Cln3 influences Cln2 levels is via regulation of SCFGrr1. To begin to test this idea, we determined whether Cln3 can influence the levels of other known protein targets of SCFGrr1. We found that protein levels of two of the best characterized targets of SCFGrr1 (Hof1 and Ndd1) are strongly influenced by Cln3 in a manner similar to Cln2. These observations are consistent with the idea that Cln3 influences Cln2 protein levels via SCFGrr1, but do not rule out alternative models. At the least, the data show that Cln3 is likely to have pervasive effects on the proteome, which suggests that the effects of Cln3 on cell size could be due to complex effects on the expression of multiple proteins. The fact that Cln3 modulates levels of multiple SCFGrr1 targets could explain the genetic data that indicate that Cln3 can influence cell size via targets other than Whi5 (Costanzo et al., 2004; Wang et al., 2009).

To test whether Cln3 influences the activity of SCFGrr1, we tried to determine whether loss- or gain-of-function of CLN3 influences the half-life of Cln2 or Hof1. One way to measure protein half-life is to shutdown total cellular translation with cycloheximide and then measure the rate of disappearance of a protein. However, a concern with this approach is that cycloheximide will cause highly pleiotropic effects that make it difficult to draw clear conclusions. For example, inhibiting all translation could cause rapid loss of additional proteins that control turnover of Cln2 protein. In some experiments we saw effects of loss- or gain-of-function of CLN3 on the protein half-life of Cln2 in cycloheximide-treated cells, but the results were not consistent or statistically significant. We also considered expressing CLN2 from the GAL1 promoter, which can be shut off with dextrose to measure protein turnover, but the GAL1 promoter drives massive overexpression of CLN2 that could overwhelm the mechanisms that control turnover of Cln2 protein.

Evidence for mitotic functions of Cln3 that influence cell size

Influential studies carried out over 50 years ago reached the conclusion that cell size is regulated primarily in G1 phase in budding yeast (Hartwell and Unger, 1977). However, more recent work has shown that little growth occurs in G1 phase (Leitao and Kellogg, 2017). For example, cell volume increases by only 10−20% during G1 phase when cells are growing in rich nutrients. Rather, most growth takes place during growth of the daughter bud, which occurs almost entirely during mitosis (Leitao and Kellogg, 2017). Moreover, the extent of bud growth during mitosis is modulated by nutrient availability, which has large impacts on daughter cell size. There is evidence for nutrient-modulated mechanisms that measure the extent of bud growth to ensure that sufficient growth has occurred before cytokinesis (Leitao and Kellogg, 2017; Leitao et al., 2019; Jasani et al., 2020).

The idea that cell size is regulated primarily in G1 phase has strongly influenced current models for the functions of Cln3, which assume that Cln3 exerts its effects on cell size solely in G1 phase. However, the idea that Cln3 functions solely in G1 phase is challenged by several observations. First, there is a second peak of Cln3 protein in mitosis, which suggests that Cln3 has functions in mitosis (Landry et al., 2012; Zapata et al., 2014; Litsios et al., 2019). Second, analysis of Coulter Counter data shows that overexpression of Cln3 causes daughter cells to be born at a dramatically reduced size, which can only occur if daughter buds undergo less growth before mitosis (Costanzo et al., 2004; Zapata et al., 2014). These observations led us to hypothesize that Cln3 could influence cell size via effects on the duration and extent of bud growth in mitosis. To test this, we developed methods to induce conditional loss- or gain-of-function of Cln3 after G1 phase and before mitosis, which showed that loss of Cln3 causes daughter cells to be born at a larger size, while overexpression of Cln3 causes them to be born at a smaller size. These data indicate that Cln3 can influence the duration and extent of bud growth in mitosis, and that mitotic functions of Cln3 cannot be ignored when considering models for how Cln3 influences cell size.

The targets of Cln3 in mitosis that influence cell size are unknown. We found that Cln3 influences levels of Hof1, a mitotic regulator of cytokinesis, and that Hof1 protein levels in mitosis are correlated with Cln3 protein levels, consistent with the idea that Cln3 has functions in mitosis. We and others have found that hof1∆ cells are larger than wild-type cells (Li et al., 2006) and we report here that loss of Cln3 leads to a reduction in Hof1 protein levels. Together, these observations suggest that part of the effects of Cln3 on cell size could be due to effects on Hof1 protein levels; however, Hof1 has not previously been implicated in cell size control and we found that overexpression of HOF1 does not reduce cell size. Additional work will be needed to gain a better understanding of how Cln3 influences cell size in mitosis.

MATERIALS AND METHODS

Request a protocol through Bio-protocol.

Yeast strains, plasmids, media, and cell-cycle time courses.

All strains are in the W303 background and carry a deletion of the BAR1 gene to facilitate arrest with alpha factor (MATa leu2-3,112 ura3-1 can1-100 ade2-1 his3-11,15 trp1-1 GAL+, ssd1-d2 bar1–). The additional genetic features of the strains are listed in Table 1. Genetic alterations, such as epitope tagging, promoter swaps, and gene deletions were carried out using homologous recombination at the endogenous locus (Longtine et al., 1998; Janke et al., 2004).

TABLE 1:

Strains used in this study.

Strain Genotype Figure
DK1295 whi5∆::KanMx4 Figure 1
DK4074 CLN2-3XHA::hphNT1 Figure 1
DK4102 CLN2-3XHA::hphNT1 whi5∆::KanMX4 Figure 1
DK4108 CLN2-3XHA::hphNT1 whi5∆::KanMX4 cln3-∆177-6XHA::HIS3MX6 Figure 1
DK4114 CLN2-3XHA::hphNT1 whi5∆::KanMX4, cln3∆::His3MX6 Figure 1
DK4125 CLN2-3XHA::hphNT1 whi5∆::KanMX4, GAL1-CLN3::URA3 (pDK93) Figure 1
DK4032 MET25pr-3XHA-CLN2::NatNT2 cln3∆::HIS3 Figure 2
DK4034 MET25pr-3XHA-CLN2::NatNT2 cln3-∆177-6XHA::HIS3MX6 Figure 2
DK4026 MET25pr-3XHA-CLN2::NatNT2 Figure 2, 3
DK4714 His3-YPK1pr-CLN2-9XMyc::hphNT1 Figure 2
DK4718 His3-YPK1pr-CLN2-9xMyc::hphNT1 GAL1-CLN3::URA3 (pDK93) Figure 2
DK4527 MET25pr-3XHA-CLN2::NatNT2 GAL1-CLN3::URA3 (pDK93) Figure 3
DK4545 MET25pr-3XHA-CLN2::NatNT2 swi6∆::HIS3 Figure 3
DK4548 MET25pr-3XHA-CLN2::NatNT2 swi6∆::HIS3 GAL1-CLN3::URA3 (pDK93) Figure 3
DK4144 MET25pr-3XHA-CLN2::NatNT2 cln1∆::hphNT1 Figure 4
DK4230 MET25pr-3XHA-CLN2::NatNT2 cln1∆::hphNT1 GAL1-CLN3::URA3 (pDK93) Figure 4
KA65 cln1∆::TRP Figure 4
DK186 Figure 4, Figure 7- supplement
DK3922 HOF1-3XHA::klTRP1 Figure 5
DK3945 HOF1-3XHA:: klTRP1 cln3-∆177-6XHA::HIS3MX6 Figure 5
DK3960 HOF1-3XHA:: klTRP1 cln3∆::His3MX6 Figure 5
DK3991 HOF1-3XHA:: klTRP1 Cln3-6XHA::His3MX6 Figure 5
DK4218 NDD1-6XHA::hphNT1 Figure 6
DK4233 NDD1-6XHA::hphNT1 cln3∆:::His3MX6 Figure 6
DK4235 NDD1-6XHA::hphNT1 cln3-∆177-6XHA::His3MX6 Figure 6
DK3348 MYO1-GFP::TRP SPC42-GFP::His3MX6 GAL1-CLN3-3XHA::URA3 (pAB1) Figure 7
DK3510 his3::HIS3+TIR1 leu2::LEU2+TIR1 SPC42-mRuby2::KanMX4 Figure 7
DK3580 SPC42-yomRuby2::KanMX4 GAL1::URA3 Figure 7
DK3524 his3::HIS3+TIR1 leu2::LEU2+TIR1 cln3-AID::KanMX4 SPC42-GFP::hphNT1 Figure 7, Supplemental Figure 7
SH184 cln3∆::His3MX6 Supplemental Figure 7
Open in a new tab

Plasmid pAB1 was used to integrate GAL1-CLN3-3XHA at the URA3 locus. To create pAB1, the GAL1 promoter was amplified and cloned into the KpnI and Xho1 sites of pRS306 (primers: CGCGGTACCTTATATTGAATTTTCAAAAATTCT and GCGCCTCGAGTATAG­TTTTT-TTCTCCTTGACG) to make pDK20. A 3XHA tag sequence was then cloned into the EagI and SacII sites of pDK20 to create pSH32. The CLN3 coding sequence was amplified and cloned into the XhoI and EagI sites of pSH32 (primers: GCGCTCGAGATGGCCATATTGAAGGATAC-CATAATTAGATACGC and CGCCGGCC-GG­CGAGTTTTCTTGAGGTTGCTACTATC). pAB1 is cut with Stu1 to target integration at the URA3 locus.

To create a plasmid that allows PCR-based replacement of promoters with the YPK1 promoter, the YPK1 promoter was amplified by PCR with BglII and PacI sites and was used to replace the GAL1 promoter in plasmid pFA6a-His3MX6-pGAL with the YPK1 promoter (pDK132, primers: GCGAGATCTGATGTTTTAACTGATCTTAATTTATATGTAGAGGA, GCGTTAATTAA­TTTCAGGAACTGTATTAATGTTTGTTGATAT).

For cell-cycle time courses, cells were arrested with 0.5 µg/ml alpha factor for 3 and 3.5 h at room temperature and released from the arrest by washing cells three times with 25 ml of growth medium. For cell-cycle time courses where GAL1-CLN3 was induced, cells were grown in YP medium containing 2% glycerol and 2% ethanol (YPGE) and were arrested with alpha factor for 3.5−4 h at room temperature. In Figure 7, GAL1-CLN3 was induced by addition of 2% galactose at 60 min after release from alpha factor arrest. In Figures 1C, 2C, and Figure 3, 2% galactose was added to the cultures 40 min before releasing from the arrest and cells were then released from the arrest into YP medium containing 2% galactose. All time courses were done at 30°C.

Cell size analysis

Cell size was measured with a Beckman Coulter Counter Z2. Briefly, cells were grown in 10-ml culture medium to an OD600 between 0.4 and 0.6. Cells were then fixed by addition of 1/10 volume of formaldehyde for 30−60 min. Cells were pelleted and resuspended in 500 µl of PBS, 0.02% sodium azide, and 0.1% Tween-20 and briefly sonicated. In each figure, cell size data represent the average of three biological replicates, in which each biological replicate is the average of three technical replicates.

Single-cell analysis of cell growth and size during the cell cycle

Analysis of cell growth and size during the cell cycle was carried out as previously described (Leitao and Kellogg, 2017; Jasani et al., 2020). Briefly, cells were grown in complete synthetic media (CSM) containing 2% dextrose (CSM-Dex) or 2% glycerol and 2% ethanol (CSM-G/E). For conditional expression of GAL1-CLN3, 2% galactose was added to the media when cells reached approximately 15% bud emergence, indicating that they were 20−30 min away from entering mitosis, which allowed sufficient time for expression CLN3 from the GAL1 promoter before mitosis. For conditional degradation of Cln3 in mitosis, auxin was added to the medium once cells were approximately 40−50% budded to allow for degradation of Cln3 before the beginning of mitosis. Sample preparation, data acquisition, and processing was performed as previously described by (Leitao and Kellogg, 2017; Jasani et al., 2020).

Western blotting

Western blotting was done as previously described (Sommer et al., 2021). All SDS–PAGE gels were 10% polyacrylamide and 0.13% bis-acrylamide. When comparing protein behavior between strains, all gels were transferred together to the same nitrocellulose membrane using Bio-Rad Trans-Blot Turbo Transfer system. Blots were probed overnight at 4°C in 3% dry milk in western wash buffer (PBS + 250 mM NaCl +0.1% Tween-20) using 12CA5 mouse monoclonal anti-HA, anti-Nap1, anti-Clb2, or anti-Myc. Primary antibodies were detected with an HRP-conjugated donkey anti-rabbit secondary antibody or HRP-conjugated donkey anti-mouse secondary antibody. Western blots of samples from different strain within an experiment were imaged together with identical exposures using ECL reagents (K-12045-D50; Advansta) and a Bio-Rad Chemi-Doc Imaging system.

Western blotting quantification

All quantifications were done using Image Lab (Bio-Rad) as previously described (Jasani et al., 2020; Sommer et al., 2021). For alpha factor block and release experiments, the signal at each time point was calculated as a ratio over the signal at the 10-min time point for the control strain. The signal for each time point was then normalized to the loading control. For log phase samples, signals were calculated as a ratio over the WT control and then normalized to loading control.

Northern blotting

Gel-purified PCR products were used to generate 32P-labeled probes to detect CLN2 and ACT1 mRNAs by northern blotting (CLN2 oligonucleotides: TATTACTTGGGTATTGCCCATACCAAAAAGA, TGAA­CCAATGATCAATGATTACGT; ACT1 oligonucleotides: TCATACCTT­CTACAACGAATTGAGA and ACACTTCATGATGGAGTTGTAAGT). Northern blotting was carried out as previously described (Cross and Tinkelenberg, 1991; Kellogg and Murray, 1995). CLN2 blots were reprobed for ACT1 to control for loading.

Experimental replicates

All experiments were repeated for a minimum of three biological replicates. Biological replicates are defined as experiments carried out on different days with cultures started with newly thawed cells.

Material availability

All yeast strains, plasmids, and antibodies described in this study are available upon request.

Data availability

All source data are included within the manuscript files.

Supplementary Material

mbc-35-ar46-s001.pdf (640.7KB, pdf)

Acknowledgments

This work was supported by NIH grant GM131826. A.B. was supported by NIH grants GM058903 and GM133391, and by the ARCS Foundation, Marilyn D. Davis Memorial Scholarship, Denice Denton Award for Promoting Diversity, Café Bustelo El Café Del Futuro Scholarship, and UCSC Slug Support Office. We thank Benjamin Abrams (University of California, Santa Cruz) for providing technical support for the microscopy experiments, and David Toczyski (University of California, San Francisco) for providing 12CA5 mouse monoclonal anti-HA antibody.

Abbreviations used:

Cdk1

cyclin-dependent kinase 1

YP

Yeast extract, Peptone

YPD

Yeast extract, Peptone, 2% Dextrose

YPGE

Yeast extract, Peptone, 2% glycerol, 2% ethanol

Footnotes

This article was published online ahead of print in MBoC in Press (http://www.molbiolcell.org/cgi/doi/10.1091/mbc.E23-05-0174) on January 17, 2024.

REFERENCES

  1. Alcaide-Gavilán M, Lucena R, Schubert KA, Artiles KL, Zapata J, Kellogg DR (2018). Modulation of TORC2 signaling by a conserved Lkb1 signaling axis in budding yeast. Genetics 210, 155–170. [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Andrews BJ, Herskowitz I (1989). The yeast SWI4 protein contains a motif present in developmental regulators and is part of a complex involved in cell-cycle-dependent transcription. Nature 342, 830–833. [DOI] [PubMed] [Google Scholar]
  3. Andrews BJ, Moore LA (1992). Interaction of the yeast Swi4 and Swi6 cell cycle regulatory proteins in vitro. Proc Natl Acad Sci 89, 11852–11856. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Bean JM, Siggia ED, Cross FR (2005). High Functional overlap between MluI cell-cycle box binding factor and Swi4/6 cell-cycle box binding factor in the G1/S transcriptional program in Saccharomyces cerevisiae. Genetics 171, 49–61. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Bhaduri S, Valk E, Winters MJ, Gruessner B, Loog M, Pryciak PM (2015). A docking interface in the cyclin Cln2 promotes multi-site phosphorylation of substrates and timely cell-cycle entry. Curr Biol 25, 316–325. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Blondel M, Bach S, Bamps S, Dobbelaere J, Wiget P, Longaretti C, Barral Y, Meijer L, Peter M (2005). Degradation of Hof1 by SCFGrr1 is important for actomyosin contraction during cytokinesis in yeast. EMBO J 24, 1440–1452. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Carter BLA, Sudbery PE (1980). Small-sized mutants of Saccharomyces Cerevisiae. Genetics 96, 561–566. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Costanzo M, Nishikawa JL, Tang X, Millman JS, Schub O, Breitkreuz K, Dewar D, Rupes I, Andrews B, Tyers M (2004). CDK activity antagonizes Whi5, an inhibitor of G1/S transcription in yeast. Cell 117, 899–913. [DOI] [PubMed] [Google Scholar]
  9. Cross FR (1988). DAF1, a mutant gene affecting size control, pheromone arrest, and cell cycle kinetics of Saccharomyces cerevisiae. Mol Cell Biol 8, 4675–4684. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Cross FR (1989). Further characterization of a size control gene in Saccharomyces cerevisiae. J Cell Sci 1989, 117–127. [DOI] [PubMed] [Google Scholar]
  11. Cross FR (1990). Cell cycle arrest caused by CLN gene deficiency in Saccharomyces cerevisiae resembles START-I arrest and is independent of the mating-pheromone signalling pathway. Mol Cell Biol 10, 6482–6490. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Cross FR, Tinkelenberg AH (1991). A potential positive feedback loop controlling CLN1 and CLN2 gene expression at the start of the yeast cell cycle. Cell 65, 875–883. [DOI] [PubMed] [Google Scholar]
  13. de Bruin RAM, McDonald WH, Kalashnikova TI, Yates J, Wittenberg C (2004). Cln3 activates G1-specific transcription via phosphorylation of the SBF bound repressor Whi5. Cell 117, 887–898. [DOI] [PubMed] [Google Scholar]
  14. Di Como CJ, Chang H, Arndt KT (1995). Activation of CLN1and CLN2 G1 Cyclin Gene Expression by BCK2. Mol Cell Biol 15, 1835–1846. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Dirick L, Böhm T, Nasmyth K (1995). Roles and regulation of Cln-Cdc28 kinases at the start of the cell cycle of Saccharomyces cerevisiae. EMBO J 14, 4803–4813. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Edenberg ER, Mark KG, Toczyski DP (2015). Ndd1 turnover by SCFGrr1 is inhibited by the DNA damage checkpoint in Saccharomyces cerevisiae. PLOS Genet 11, e1005162. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Edgington NP, Futcher B (2001). Relationship between the function and the location of G1 cyclins in S. cerevisiae. J Cell Sci 114, 4599–4611. [DOI] [PubMed] [Google Scholar]
  18. Ferrezuelo F, Colomina N, Futcher B, Aldea M (2010). The transcriptional network activated by Cln3 cyclin at the G1-to-S transition of the yeast cell cycle. Genome Biol 11, R67. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Hadwiger JA, Wittenberg C, Richardson HE, de Barros Lopes M, Reed SI (1989). A family of cyclin homologs that control the G1 phase in yeast. Proc Natl Acad Sci USA 86, 6255–6259. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Hartwell LH, Unger MW (1977). Unequal division in Saccharomyces cerevisiae and its implications for the control of cell division. J Cell Biol 75, 422–435. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Horak CE, Luscombe NM, Qian J, Bertone P, Piccirrillo S, Gerstein M, Snyder M (2002). Complex transcriptional circuitry at the G1/S transition in Saccharomyces cerevisiae. Genes Dev 16, 3017–3033. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Iyer VR, Horak CE, Scafe CS, Botstein D, Snyder M, Brown PO (2001). Genomic binding sites of the yeast cell-cycle transcription factors SBF and MBF. Nature 409, 533–538. [DOI] [PubMed] [Google Scholar]
  23. Janke C, Magiera MM, Rathfelder N, Taxis C, Reber S, Maekawa H, Moreno-Borchart A, Doenges G, Schwob E, Schiebel E, Knop M (2004). A versatile toolbox for PCR-based tagging of yeast genes: new fluorescent proteins, more markers and promoter substitution cassettes. Yeast 21, 947–962. [DOI] [PubMed] [Google Scholar]
  24. Jasani A, Huynh T, Kellogg DR (2020). Growth-dependent activation of protein kinases suggests a mechanism for measuring cell growth. Genetics 215, 729–746. [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Jorgensen P, Tyers M (2004). How cells coordinate growth and division. Curr Biol 14, R1014–R1027. [DOI] [PubMed] [Google Scholar]
  26. Jorgensen P, Nishikawa JL, Breitkreutz B-J, Tyers M (2002). Systematic identification of pathways that couple cell growth and division in yeast. Science 297, 395–400. [DOI] [PubMed] [Google Scholar]
  27. Kellogg DR, Murray AW (1995). NAP1 acts with Clb1 to perform mitotic functions and to suppress polar bud growth in budding yeast. J Cell Biol 130, 675–685. [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Koch C, Schleiffer A, Ammerer G, Nasmyth K (1996). Switching transcription on and off during the yeast cell cycle: Cln/Cdc28 kinases activate bound transcription factor SBF (Swi4/Swi6) at start, whereas Clb/Cdc28 kinases displace it from the promoter in G2. Genes Dev 10, 129–141. [DOI] [PubMed] [Google Scholar]
  29. Kõivomägi M, Swaffer MP, Turner JJ, Marinov G, Skotheim JM (2021). Localized phosphorylation of RNA Polymerase II by G1 cyclin-Cdk promotes cell cycle entry. bioRxiv, 10.1101/2021.03.25.436872. [DOI] [PMC free article] [PubMed] [Google Scholar]
  30. Landry BD, Doyle JP, Toczyski DP, Benanti JA (2012). F-Box protein specificity for G1 cyclins is dictated by subcellular localization. PLoS Genet 8, e1002851. [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Leitao RM, Kellogg DR (2017). The duration of mitosis and daughter cell size are modulated by nutrients in budding yeast. J Cell Biol 216, 3463–3470. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Leitao RM, Jasani A, Talavera RA, Pham A, Okobi QJ, Kellogg DR (2019). A conserved PP2A regulatory subunit enforces proportional relationships between cell size and growth rate. Genetics 213, 517–528. [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Levine K, Huang K, Cross FR (1996). Saccharomyces cerevisiae G1 cyclins differ in their intrinsic functional specificities. Mol Cell Biol 16, 6794–6803. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Li WJ, Wang YM, Zheng XD, Shi QM, Zhang TT, Bai C, Li D, Sang JL, Wang Y (2006). The F-box protein Grr1 regulates the stability of Ccn1, Cln3 and Hof1 and cell morphogenesis in Candida albicans. Mol Microbiol 62, 212–226. [DOI] [PubMed] [Google Scholar]
  35. Lim S, Kaldis P (2013). Cdks, cyclins and CKIs: roles beyond cell cycle regulation. Development 140, 3079–3093. [DOI] [PubMed] [Google Scholar]
  36. Litsios A, Huberts DHEW, Terpstra HM, Guerra P, Schmidt A, Buczak K, Papagiannakis A, Rovetta M, Hekelaar J, Hubmann G, et al. (2019). Differential scaling between G1 protein production and cell size dynamics promotes commitment to the cell division cycle in budding yeast. Nat Cell Biol 21, 1382–1392. [DOI] [PubMed] [Google Scholar]
  37. Longtine MS, Iii AM, Demarini DJ, Shah NG, Wach A, Brachat A, Philippsen P, Pringle JR (1998). Additional modules for versatile and economical PCR-based gene deletion and modification in Saccharomyces cerevisiae. Yeast 14, 953–961. [DOI] [PubMed] [Google Scholar]
  38. Lucena R, Alcaide-Gavilán M, Schubert K, He M, Domnauer MG, Marquer C, Klose C, Surma MA, Kellogg DR (2018). Cell size and growth rate are modulated by TORC2-dependent signals. Curr Biol 28, 196–210.e4. [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Miller ME, Cross FR (2001). Mechanisms controlling subcellular localization of the G 1 cyclins Cln2p and Cln3p in budding yeast. Mol Cell Biol 21, 6292–6311. [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Miller ME, Cross FR, Groeger AL, Jameson KL (2005). Identification of novel and conserved functional and structural elements of the G1 cyclin Cln3 important for interactions with the CDK Cdc28 in Saccharomyces cerevisiae. Yeast 22, 1021–1036. [DOI] [PubMed] [Google Scholar]
  41. Nash R, Tokiwa G, Anand S, Erickson K, Futcher AB (1988). The WHI1+ gene of Saccharomyces cerevisiae tethers cell division to cell size and is a cyclin homolog. EMBO J 7, 4335–4346. [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Nasmyth K, Dirick L (1991). The role of SW4 and SW16 in the activity of Gl cyclins in yeast. Cell 66, 995–1013. [DOI] [PubMed] [Google Scholar]
  43. Nishimura K, Fukagawa T, Takisawa H, Kakimoto T, Kanemaki M (2009). An auxin-based degron system for the rapid depletion of proteins in nonplant cells. Nat Methods 6, 917–922. [DOI] [PubMed] [Google Scholar]
  44. Richardson HE, Wittenberg C, Cross F, Reed SI. (1989). An essential G1 function for cyclin-like proteins in yeast. Cell 59, 1127–1133. [DOI] [PubMed] [Google Scholar]
  45. Rubin SM, Sage J, Skotheim JM (2020). Integrating old and new paradigms of G1/S control. Mol Cell 80, 183–192. [DOI] [PMC free article] [PubMed] [Google Scholar]
  46. Schneider BL, Zhang J, Markwardt J, Tokiwa G, Volpe T, Honey S, Futcher B (2004). Growth rate and cell size modulate the synthesis of, and requirement for, G1-phase cyclins at start. Mol Cell Biol 24, 10802–10813. [DOI] [PMC free article] [PubMed] [Google Scholar]
  47. Siegmund RF, Nasmyth KA (1996). The Saccharomyces cerevisiae Start-specific transcription factor Swi4 interacts through the ankyrin repeats with the mitotic Clb2/Cdc28 kinase and through its conserved carboxy terminus with Swi6. Mol Cell Biol 16, 2647–2655. [DOI] [PMC free article] [PubMed] [Google Scholar]
  48. Skotheim JM, Di Talia S, Siggia ED, Cross FR (2008). Positive feedback of G1 cyclins ensures coherent cell cycle entry. Nature 454, 291–296. [DOI] [PMC free article] [PubMed] [Google Scholar]
  49. Skowyra D, Craig KL, Tyers M, Elledge SJ, Harper JW (1997). F-Box proteins are receptors that recruit phosphorylated substrates to the SCF ubiquitin-ligase complex. Cell 91, 209–219. [DOI] [PubMed] [Google Scholar]
  50. Sommer RA, DeWitt JT, Tan R, Kellogg DR (2021). Growth-dependent signals drive an increase in early G1 cyclin concentration to link cell cycle entry with cell growth. eLife 10, e64364. [DOI] [PMC free article] [PubMed] [Google Scholar]
  51. Spellman PT, Sherlock G, Zhang MQ, Iyer VR, Anders K, Eisen MB, Brown PO, Botstein D, Futcher B (1998). Comprehensive identification of cell cycle–regulated genes of the yeast Saccharomyces cerevisiae by microarray hybridization. Mol Biol Cell 9, 3273–3297. [DOI] [PMC free article] [PubMed] [Google Scholar]
  52. Sudbery PE, Goodey AR, Carter BLA (1980). Genes which control cell proliferation in the yeast Saccharomyces cerevisiae. Nature 288, 401–404. [DOI] [PubMed] [Google Scholar]
  53. Tyers M, Tokiwa G, Futcher B (1993). Comparison of the Saccharomyces cerevisiae G1 cyclins: Cln3 may be an upstream activator of Cln1, Cln2 and other cyclins. EMBO J 12, 1955–1968. [DOI] [PMC free article] [PubMed] [Google Scholar]
  54. Tyers M, Tokiwa G, Nash R, Futcher B (1992). The Cln3-Cdc28 kinase complex of S. cerevisiae is regulated by proteolysis and phosphorylation. EMBO J 11, 1773–1784. [DOI] [PMC free article] [PubMed] [Google Scholar]
  55. Wagner MV, Smolka MB, de Bruin RAM, Zhou H, Wittenberg C, Dowdy SF (2009). Whi5 regulation by site specific CDK-phosphorylation in Saccharomyces cerevisiae. PLoS One 4, e4300. [DOI] [PMC free article] [PubMed] [Google Scholar]
  56. Wang H, Carey LB, Cai Y, Wijnen H, Futcher B (2009). Recruitment of Cln3 cyclin to promoters controls cell cycle entry via histone deacetylase and other targets. PLoS Biol 7, e1000189. [DOI] [PMC free article] [PubMed] [Google Scholar]
  57. Wijnen H, Landman A, Futcher B (2002). The G1 cyclin Cln3 promotes cell cycle entry via the transcription factor Swi6. Mol Cell Biol 22, 4402–4418. [DOI] [PMC free article] [PubMed] [Google Scholar]
  58. Willems AR, Lanker S, Patton EE, Craig KL, Nason TF, Mathias N, Kobayashi R, Wittenberg C, Tyers M (1996). Cdc53 targets phosphorylated G1 cyclins for degradation by the ubiquitin proteolytic pathway. Cell 86, 453–463. [DOI] [PubMed] [Google Scholar]
  59. Zapata J, Dephoure N, MacDonough T, Yu Y, Parnell EJ, Mooring M, Gygi SP, Stillman DJ, Kellogg DR (2014). PP2ARts1 is a master regulator of pathways that control cell size. J Cell Biol 204, 359–376. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

    This section collects any data citations, data availability statements, or supplementary materials included in this article.

    Supplementary Materials

    mbc-35-ar46-s001.pdf (640.7KB, pdf)

    Data Availability Statement

    All source data are included within the manuscript files.

    Articles from Molecular Biology of the Cell are provided here courtesy of American Society for Cell Biology

    RESOURCES