CDK-Independent Initiation of the S.

cerevisiae Cell Cycle –Analysis of BCK2

by

Nazareth Bastajian

A thesis submitted in conformity with the requirements

for the degree of Doctor of Philosophy

Graduate Department of Molecular Genetics

University of Toronto

© Copyright by Nazareth Bastajian, 2012

ii

CDK-Independent Initiation of the S. cerevisiae Cell Cycle –Analysis of BCK2

Nazareth Bastajian

Doctor of Philosophy 2012

Department of Molecular Genetics

University of Toronto

Abstract

Much of the work on how the cell cycle is regulated has focused on Cyclin-Dependent

Kinase (CDK)-mediated regulation of factors that control the coordinate expression of

genes required for entry into the cell cycle. In Saccharomyces cerevisiae, SBF and MBF

are related transcription factors that co-ordinately activate a large group of genes at the

G1/S transition, and their activation depends on the Cln3-Cdk1 form of the cyclin-

dependent kinase. However, cells are viable in the absence of Cln3, or SBF and MBF,

indicating that other regulatory pathways must exist that activate the budding yeast cell

cycle. The known CDK-independent pathways are made up of various phosphatases and

plasma membrane transporters that control ion homeostasis in early G1 phase, a time

when cells assess environmental growth conditions in order to commit to cell cycle entry.

The enigmatic Bck2 protein is thought to act within these CDK-independent pathways,

but the means by which it activates G1/S-regulated genes is not known. Bck2 contains

little sequence homology to any known protein. In order to understand how CDK-

independent pathways operate, I have studied the Bck2 protein using multiple

approaches. In one approach, I have screened for novel SBF/MBF-binding proteins in

order to determine if other non-CDK proteins, such as Bck2, might activate SBF and

MBF. I have also investigated which region of Bck2 is required for its activity in order to

determine if Bck2’s transcriptional activation region is essential. Using one of the

iii

truncation derivatives from this analysis, I have screened for proteins that interact with

Bck2. One of these novel proteins is Mcm1, a global transcriptional activator of genes

involved in cell cycle progression, mating gene transcription and metabolism. My studies

suggest that Bck2 regulates the activity of Mcm1 in early G1 phase to activate the

expression of SWI4, CLN3, and others. My evidence suggests that Bck2 competes for

binding to a specific pocket on Mcm1 that is also bound by an Mcm1 repressor called

Yox1. My findings suggest that CDK-independent pathways function through Bck2, in

order to induce the initial suite of genes required for entry into the cell cycle.

iv

Acknowledgements

I would like to thank my supervisor Brenda Andrews for her advice and support

throughout the duration of this thesis project. I would like to thank the members of my

supervisory committee Brigitte Lavoie and Jack Greenblatt for their helpful guidance. I

would like to thank Oliver Schub for setting up the baculovirus-mediated insect cell

expression system that I used to purify SBF/MBF for biochemical assays. I would also

like to thank Yuen Ho for introducing me to the mysterious genetics of BCK2. In the

laboratory of Brigitte Lavoie, I would like to thank Charly Chahwan for stimulating

discussions on Bck2 function. In the laboratory of Charlie Boone, I would like to thank

Xiaofeng Xin for technical guidance on how to perform the yeast-two-hybrid screen. I

am thankful for the general guidance of Helena Friesen who has been intimately involved

with this thesis project from its infancy through to its end. I would like to thank all the

past and present members of the Andrews laboratory for their help and advice. Finally, I

would like to thank Andrew Becker for giving me the chance to enter the undergraduate

specialist program in the department of molecular genetics.

v

Table of Contents

Abstract ii

Acknowledgements iv

Table of Contents v

List of Tables viii

List of Figures ix

List of Abbreviations x

Chapter 1: Introduction 1

1.1 The cell cycle of S. cerevisiae 2

1.1.1 Cell cycle phases and transition points 4

1.1.2 Cyclin dependent kinases and cyclins 8

1.1.3 Cell cycle – Entry from G0 to G1 10

1.2 Transcriptional circuitry of the cell cycle 11

1.2.1 M/G1 transition 15

1.2.2 G1/S transition 16

1.2.3 S phase 17

1.2.4 S/G2 transition 19

1.2.5 G2/M transition 20

1.2.6 Complexity of cell cycle promoters 20

1.3 CDK-dependent regulation of the G1/S phase transition 22

1.3.1 Cln3-dependent activation of SBF/MBF 24

1.3.2 A positive feedback loop -- G1/S 25

1.3.3 Cln3 abundance as a threshold for cell cycle commitment 26

1.4 A CDK-dependent oscillator 29

1.5 CDK-independent regulation of G1/S and cell cycle progression 32

1.5.1 Sit4 & Ppz1 pathways 35

1.5.2 Ion homeostasis in G1 phase 36

1.5.3 The BCK2 gene 39

1.5.4 Bck2 – the problem of SBF/MBF-dependent transcription 40

1.6 Thesis rationale 42

Chapter 2: A biochemical approach identifies Bck2 as a binding 45

partner of the G1 transcription factors SBF and MBF

2.1 Abstract 46

2.2 Introduction 47

2.3 Materials and methods 49

2.3.1 Yeast strains and plasmids 49

2.3.2 Recombinant protein expression and purification 50

2.3.3 EMSA analyses 51

2.3.4 SBF/MBF affinity resins 52

2.3.5 Affinity capture binding assays 53

2.3.6 ChIP analysis 54

2.3.7 Bck2-DB constructs, complementation and 54

auto-activation assays

vi

2.4 Results 55

2.4.1 Purification of recombinant SBF and MBF from insect cells 55

2.4.2 Purified SBF and MBF are functional in vitro 58

2.4.3 Novel binding partners of SBF and MBF – Elm1, 60

Rtf1, Ykr077w, Bck2

2.4.4 Localization of Bck2 to promoters of genes 63

transcribed at the G1/S transition

2.4.5 Bck2 DB fusion proteins can activate 66

transcription when tethered to DNA

2.5 Discussion 68

2.5.1 SBF and MBF affinity resins for high-throughput 68

screening of G1-specific regulators

2.5.2 Identification of known SBF and MBF transcriptional 69

regulators

2.5.3 Identification of novel SBF and MBF transcriptional 70

regulators

Chapter 3: Dissection of Bck2 protein domains and identification of 73

Bck2 binding partners using a yeast two-hybrid screen

3.1 Abstract 74

3.2 Introduction 74

3.3 Materials and Methods 77

3.3.1 Yeast strains and plasmids 77

3.3.2 Cloning and construction of BCK2 truncations 78

3.3.3 β-Galactosidase assays 79

3.3.4 Complementation analysis 80

3.3.5 Genome-wide Y2H screen 80

3.3.6 Direct pair-wise Y2H assays 81

3.4 Results 83

3.4.1 Truncation analysis of the BCK2 gene 83

3.4.2 A yeast-two-hybrid screen using Gal4 DB-Bck2 as bait 89

identifies six novel interacting proteins

3.4.3 Bck2 and Mot3 activate CYC1 and RPL39 transcription 92

3.5 Discussion 95

3.5.1 Bck2 activity under hyper-osmotic conditions 95

3.5.2 Bck2 activity under hypo-osmotic conditions 97

3.5.3 Tpd3 and Bck2 in nutrient sensing 99

Chapter 4: Bck2 interacts with the MADS box protein Mcm1 on 101

cell cycle-regulated promoters to activate early G1 phase

transcription in budding yeast

4.1 Abstract 102

4.2 Introduction 103

4.3 Materials and Methods 105

4.3.1 Yeast strains and plasmids 105

4.3.2 β-galactosidase assays 107

vii

4.3.3 Cell synchronization 107

4.3.4 mRNA purification and RT-qPCR 107

4.3.5 Yeast two-hybrid assays 108

4.3.6 Mutagenesis 109

4.3.7 ChIP assays 109

4.4 Results 110

4.4.1 Bck2 activates early G1 phase-expressed genes that 110

contain Mcm1-binding sites in their promoters

4.4.2 Bck2 requires ECB elements for transcriptional activation 114

4.4.3 Bck2 localizes to early and late G1 phase promoters 119

4.4.4 Bck2 competes with Yox1 for access to Mcm1 on 121

ECB elements

4.5 Discussion 125

4.5.1 SBF/MBF-independent activation by Bck2 125

4.5.2 Bck2 as an activator of G2/M genes 128

4.5.3 How Bck2 activates the G1/S transition 129

Chapter 5 - Summary and future directions 131

5.1 Thesis summary 132

5.2 Future directions 135

5.2.1 Does Bck2 activate MAT cluster genes at M/G1? 137

5.2.2 Bck2 functional analogs in humans 142

References 150

viii

List of Tables

Table 2-1 S. cerevisiae strains used in Chapter 2 49

Table 2-2 Plasmids used in Chapter 2 50

Table 3-1 S. cerevisiae strains used in Chapter 3 77

Table 3-2 Plasmids used in Chapter 3 77

Table 3-3 Bck2 truncation oligonucleotides 79

Table 4-1 S. cerevisiae strains used in Chapter 4 106

Table 4-2 Plasmids used in Chapter 4 106

Table 4-3 Oligonucleotides used for gene expression 108

Table 4-4 Oligonucleotides used for ChIP assays 109

ix

List of Figures

Figure 1-1 The cell cycle of S. cerevisiae 5

Figure 1-2A Periodic gene expression 12

Figure 1-2B Transcriptional circuitry of the cell cycle 14

Figure 1-3 CDK-dependent regulation of the G1/S phase 23

transition

Figure 1-4 A CDK-dependent oscillator 30

Figure 1-5 CDK-independent regulation of the G1/S phase 33

transition

Figure 2-1 Purification of recombinant SBF and MBF from 57

insect cells

Figure 2-2 DNA-binding by purified SBF and MBF 59

Figure 2-3 Affinity capture analysis using SBF/MBF affinity 61

resins

Figure 2-4 Localization of Bck2 to the promoters of G1/S 64

transcribed genes

Figure 2-5 Bck2 auto-activates transcription when fused to 67

a DNA-binding domain

Figure 3-1 Truncation analysis of the BCK2 gene 84

Figure 3-2 Transcriptional activation by BCK2 truncations 85

in lacZ reporter assays

Figure 3-3 Transcriptional activation by BCK2 truncations 86

in ADE2 reporter assays

Figure 3-4 Complementation of a cln3Δbck2ΔpGAL-CLN3 strain 87

growth defect by BCK2 truncation derivatives

Figure 3-5 Bck2-interacting proteins identified in a 90

genome-wide yeast two-hybrid screen

Figure 3-6 Bck2 and Mot3 activate CYC1 and RPL39 93

transcription

Figure 4-1 Bck2 activates Mcm1-driven lacZ reporter constructs 111

Figure 4-2 Effect of BCK2 deletion on CLN2, ALG9, CLN3, 113

SWI4, BCK2 and CLB2 mRNA accumulation

during the cell cycle

Figure 4-3 Bck2 requires intact ECB elements in the CLN3 115

promoter for transcriptional activation

Figure 4-4 Bck2 requires intact ECB elements in the CLN3 117

and SWI4 promoters for transcriptional activation

of CLN3 and SWI4.

Figure 4-5 Bck2 localizes to the promoters of early and late G1 120

phase-expressed genes.

Figure 4-6 Bck2 competes with Yox1 for interaction with Mcm1 122

Figure 5-1 Early G1 phase signalling in human cells and 143

budding yeast

Figure 5-2 Localization of Bck2 to early G1 phase promoters in 147

the presence of rapamycin

x

List of Abbreviations

Δ deletion (of a gene or region of a gene)

:: gene disruption

°C degrees Celsius

AD activation domain

Ade adenine

ADH alcohol dehydrogenase

α-factor alpha factor mating pheromone

ATP adenosine 5’-triphosphate

β-gal β-galactosidase

bp base pair(s)

BCK bypass of C kinase

BCK2 bypass of C kinase 2

BLAST basic local alignment search tool

CDC cell division cycle

CDK cyclin dependent kinase

cDNA complementary DNA

CE crude extract

ChIP chromatin immunoprecipitation

ChIP-chip ChIP followed by microarray hybridization

CLB B-type cyclin

CLN G1 cyclin

C-terminus carboxyl-terminus

d 2’-deoxy-

DB DNA-binding domain

DBD DNA-binding domain

ddH2O double distilled water

DEPC diethyl pyrocarbonate

DNA deoxyribonucleic acid

dsDNA double-stranded DNA

DTT dithiothreitol

ECB early cell cycle box

E. coli Escherichia coli

EDTA ethylene diamine tetra acetic acid

EMSA elecrophoretic mobility shift assays

FLAG N-DYKDDDDK-C

FOA 5-fluoroorotic acid

G0 gap phase 0

G1 gap phase 1

G2 gap phase 2

Gal galactose

GFP green fluorescent protein

GRE glucose response element

H. sapiens Homo sapiens

HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid

xi

HU hydroxyl urea

IEG immediate early gene

kDa kilodalton(s)

lacZ β-galactosidase gene

Leu leucine

LexA bacterial Op-binding repressor

M mitosis phase

MADS Mcm1 Agamous Deficiens SRF

MAPK mitogen activated protein kinase

MATa mating type “a”

MATα mating type “α”

MBF MCB binding factor

MCB MluI cell cycle box

MKL megakaryoblastic leukemia

ml millilitres

mM millimolar

MPF maturation promoting factor

mRNA messenger ribonucleic acid

MUT mutant

N-terminus amino-terminus

OD optical density

Op operator

ORF open reading frame

PAGE polyacrylamide gel electrophoresis

PBS phosphate buffered saline

PCR polymerase chain reaction

PKC protein kinase C

PMSF phenylmethylsulfonyl fluoride

PP2A protein phosphatase 2A

PPM pre-stained protein marker

pr promoter

PRC pre-replication complex

PRE pheromone response element

Q-PCR quantitative PCR

RiBi ribosome biogenesis

RNA ribonucleic acid

RNAPII RNA polymerase II

S replication phase

SBF SCB binding factor

SC synthetic complete

SCB Swi4/6 cell cycle box

S. cerevisiae Saccharomyces cerevisiae

SD synthetic dropout media dextrose

SDS sodium dodecyl sulphate

SG synthetic dropout media galactose

S. pombe Schizosaccharomyces pombe

xii

SRE serum response element

SRF serum response factor

SUP supernatant

SWI switching gene

TAP tandem affinity purification

TBE tris borate EDTA

TBS tris buffered saline

TOR target of rapamycin

Trp tryptophan

UAS upstream activating sequence

V volts

WT wild-type

X-Gal 5-bromo-4-chloro-3-indolyl-β-D-galactoside

Y2H yeast two-hybrid

YPD yeast peptone dextrose

YPGal yeast peptone galactose

YPRaff yeast peptone raffinose

1

Chapter 1

Introduction

2

1.1 The cell cycle of S. cerevisiae

A cell divides from a single cell into two cells in a manner that preserves the

hereditary material in both the parent and daughter cell. The decision to begin cell

division is of great importance for cell viability, and it is crucial that the division process

is initiated only in the appropriate environmental and genetic conditions. In single-celled

organisms, premature entry into cell division leaves cells lacking in the raw materials and

structures necessary to complete the cell cycle. In metazoan organisms, mis-regulated

cell proliferation can lead to cancer and other diseases. Broadly speaking, the molecular

architecture of cell division is well conserved between unicellular and multicellular

eukaryotes. Therefore, experimentally accessible unicellular organisms have been

productively used to understand the basic mechanisms of cell division. For example, the

unicellular budding yeast, Saccharomyces cerevisiae, has provided researchers with great

insight into the regulation of cell division, including the identification of many genes

deregulated in cancers of H. sapiens (Semple and Duncker 2004; Diaz-Ruiz et al. 2009;

Vavouri et al. 2009; Diaz-Ruiz, Rigoulet, and Devin 2011).

Budding yeast is particularly useful for studying cell division since visible

morphological changes correlate with different stages of the cell division cycle. For

example, at the beginning of DNA replication, a bud emerges which is destined to

become the daughter cell. The correlation between cell cycle position and cell

morphology enabled the identification of a class of mutants dysfunctional in cell division

called the CDC (“Cell Division Cycle”) mutants (Hartwell, Culotti, and Reid 1970;

Hartwell 1971b; Hartwell et al. 1974). These pioneering studies involved mutagenizing

budding yeast cells, and then screening for temperature-sensitive mutants that exhibited a

3

uniform morphological arrest phenotype (e.g. all unbudded cells). The idea behind this

screen was that the WT versions of the mutated genes may encode key regulators of cell

division. Indeed, the CDC genes, in both S. cerevisiae and S. pombe, have subsequently

been shown to encode conserved regulators of cell division, including factors controlling

cell cycle commitment, DNA replication, nuclear division, and cytokinesis. Early genetic

analysis of CDC mutants also revealed that the cell division cycle comprises a series of

events that occur in a specific sequence and are dependent on the proper execution of

prior events. For example, DNA replication must occur once and only once during each

cell cycle in order to maintain genome stability (Dalton 1998). Many labs have studied

CDC genes in a variety of model systems over the past several decades and it now seems

clear that cell division is controlled by a complex network of regulators that must be

integrated with both the intra- and extracellular environments.

In this chapter, I discuss broadly how the cell division cycle is controlled in the

budding yeast S. cerevisiae. I start with an outline of the key molecular events that occur

during the cell cycle, with an emphasis on Cyclin-Dependent Kinases (CDK), whose

periodic activity in cycling cells drives cell division. I then describe the transcriptional

circuit which underlies the cell cycle. I then focus on CDK-dependent regulation of the

G1/S transition. I conclude by discussing the interplay between the extracellular

environment and the decision to enter the mitotic cell cycle. In particular, I discuss

studies related to the BCK2 gene, which encodes a genetically potent regulator of cell

cycle commitment, but whose mechanism of action is poorly understood.

4

1.1.1 Cell cycle phases and transition points

The budding yeast cell cycle can be described as four distinct phases that proceed

in the following order: G1 (gap 1), S (synthesis), G2 (gap 2), and M (mitosis) (Figure 1-

1). During S phase, DNA replication occurs, and mitosis takes place during M phase.

These two phases are separated by gap phases called G1 and G2, which are important

control points for the phases that follow. During the cell cycle, growth occurs co-

ordinately with division (Elliott and McLaughlin 1978; Alberghina and Porro 1993). For

example, G1 phase is the point of cell cycle entry and exit and is characterized by a period

of growth during which the cell increases in size and continually assesses whether

appropriate environmental conditions exist to support a new round of cell division. If

environmental conditions are appropriate, the cell transits the G1/S boundary, called

START in budding yeast (Hereford and Hartwell 1974b; Sudbery, Goodey, and Carter

1980). During G2 phase, the daughter cell continues to grow and the completion of DNA

replication is monitored. After G2, interphase is complete and the cell begins mitosis

(M), during which chromosomes are appropriately segregated to mother and daughter

cells followed by cytokinesis.

Budding yeast can also exit the cell cycle during G1 phase if mating pheromone is

present. S. cerevisiae can exist in one of two mating types, “a” or “α”. The mating of

yeast of two different mating types produces a diploid “a/α” cell, which can reproduce as

a diploid or give rise to spores that germinate into “a” or “α” haploid progeny. Before

conjugation, yeast cells form a shmoo, which is an elongated structure that allows the cell

to extend in the direction of the highest pheromone concentration. Haploid cells of the

“a” mating-type can sense the “α” pheromone secreted from haploid cells of the “α”

5

Figure 1-1. The cell cycle of S. cerevisiae. A schematic of the budding yeast cell cycle is shown. Cell

cycle phases are not proportional to time spent in each cell cycle phase. Black circles represent spindle

pole bodies that orients the mitotic spindle, and dark red circles represent nuclei. The budding yeast cell

cycle is characterized by visible morphological changes that are associated with different phases of the cell

division cycle. For example, the beginning of DNA replication (S phase) is marked by the emergence of a

small bud, destined to become the new daughter cell. The cell cycle can be described as four phases that

proceed in the order G1 (gap 1), S (synthesis), G2 (gap 2), and M (mitosis). G1 phase is the point of cell

cycle entry from G0 (stationary) phase. In G1 phase the cell assesses whether appropriate environmental

conditions exist to support a new round of cell division. Under appropriate conditions, the cell irreversibly

transits the G1/S phase transition, called START. In S phase, chromosomal DNA is replicated in order for

it to be distributed between the mother and daughter cells in the future M phase. The molecular events

underlying cell cycle phase transitions are controlled by cyclin-dependent kinase (CDK)-cyclin complexes.

The principle CDK involved in cell cycle control is Cdk1, which is activated by nine different cyclins. In

G1 phase, Cln3-Cdk1 activates the expression of a cluster of genes including the cyclins CLN1 and CLN2.

Cln1-Cdk1 and Cln2-Cdk1 activity is important for transit through the G1/S transition. Clb3-Cdk1 and

Clb4-Cdk1 promote later processes in S and G2 phases, whereas Clb1-Cdk1 and Clb2-Cdk1 promote

processes at the G2/M transition. After M phase, the cell cycle is reset at G1 when the cell again assesses

whether the proper environmental conditions exist for a new round of division.

Cdk1

Cdk1

G1 G2

M

S

G0

START

Cdk1

Cln3

Clb3

Clb4

Cdk1

Clb2

Cdk1

Clb1

Cdk1

Cln2

Cdk1

Cln1

Cdk1

Cdk1

Clb5,6

Clb6

6

mating-type. The presence of pheromone arrests the cell cycle in G1 phase, and begins

molecular processes prerequisite for mating and formation of a diploid cell (Bardwell

2005).

G1 phase is distinct from the other phases of the cell cycle in that it is temporally

plastic (Rupes 2002). This plasticity occurs because the length of G1 is a function of

environmental growth conditions (Hartwell and Unger 1977; Johnston et al. 1979). If

nutrients are limiting, cells arrest early in G1 phase (Mendenhall and Hodge 1998).

However, after G1 phase is transited, the remaining cell cycle is of fairly constant length

even if nutrients become limiting (Johnston, Pringle, and Hartwell 1977; Jagadish and

Carter 1977). Transiting G1 phase is equivalent to passing START, originally defined as

the point at which a yeast cell acquires resistance to mating pheromone (Hereford and

Hartwell 1974a). The concept of START was developed by studying the pioneering

CDC screens mentioned earlier (Hartwell et al. 1974) and is analogous to the restriction

point in mammalian cells (Zetterberg, Larsson, and Wiman 1995). Prior to the execution

of START, either nutrient limitation or the presence of yeast mating pheromone can

arrest the cells in G1 (Gray et al. 2004). The G1 arrest of cells exposed to pheromone

occurs very late in G1 compared to cells arrested in cells exposed to nutrient deprivation,

where cells arrest much earlier (Barbet et al. 1996). Many mutations resulting in human

cancer are in genes encoding proteins involved in processes that occur in G1 phase

(Sidorova and Breeden 2003a).

During a normal S phase, DNA replication occurs and is completed before mitosis

is initiated (Hartwell and Weinert 1989). Interestingly, recent work from the L. Aragon

laboratory has suggested that the cell may not monitor the completion of replication, and

7

mitosis can initiate with unfinished replication when checkpoint adaptation occurs

(Torres-Rosell et al. 2007). DNA replication initiates from multiple sites on

chromosomes called origins of replication (Remus and Diffley 2009). Within S phase,

replication is generally monitored to ensure faithful chromosome segregation later on in

M phase. Processes important for chromosome segregation occur during S phase, such as

bud emergence, kinetochore assembly, formation of new spindle pole bodies, and sister

chromatid cohesion (Tanaka et al. 2005; Sherwood, Takahashi, and Jallepalli 2010).

Centromeres remain attached to the microtubules during most of the cell cycle except

during early S phase when kinetochores are disassembled to facilitate centromere

replication (Winey and O'Toole 2001; Pearson et al. 2004), after which kinetochores are

reassembled and attached to microtubules (Tanaka et al. 2005).

During M phase, mitosis occurs, which is the intricate process by which the cell

segregates the chromosomes in its nucleus into two identical sets. Sister chromatid

cohesion must be established and maintained to allow bipolar orientation of sister

chromatids, and chromosome condensation must also occur (Hirano 2005; Sakuno, Tada,

and Watanabe 2009). For faithful segregation of chromosomes, the mitotic spindle has to

be oriented perpendicular to the mother-bud neck (Yeh et al. 2000). Sister kinetochores

bi-orient, a bipolar spindle is formed, and the sister chromatids are pulled toward the

opposite spindle poles and into sister cells (Janke et al. 2002; Dewar et al. 2004; Caydasi,

Ibrahim, and Pereira 2010). Later on, a new nuclear envelope forms around each set of

separated sister chromosomes, a cleavage furrow forms, and cells undergo cytokinesis

(Jensen and Johnston 2002; Roncero and Sanchez 2010).

8

1.1.2 Cyclin dependent kinases and cyclins

Cyclin-dependent kinase (CDK)-cyclin complexes promote phase transitions in

the eukaryotic cell cycle (Figure 1-1). CDKs are activated by association with cyclin

subunits, which induce a change in protein conformation (Jeffrey et al. 1995). In S.

cerevisiae, the principle CDK involved in cell cycle regulation is Cdc28 (Cdk1), which is

activated by nine different cyclins. Three G1 cyclins, encoded by CLN1, CLN2, and

CLN3, activate Cdc28 during G1 phase to promote cell cycle commitment, or START,

and events such as bud formation, and any one of the three CLN genes is sufficient for

START (Cross 1990; Nasmyth 1993). Strikingly, various cyclin genes from other

organisms can rescue CLN-deficient S. cerevisiae when expressed from strong promoters

(Xiong et al. 1991; Leopold and O'Farrell 1991; Lahue, Smith, and Orr-Weaver 1991;

Lew, Dulic, and Reed 1991; Koff et al. 1991), indicating significant functional

conservation. Cln-Cdc28 complexes also activate the expression of the B-type (CLB)

cyclins, CLB1, CLB2, CLB3, CLB4, CLB5, and CLB6 (Nasmyth 1993; Cross 1995;

Nasmyth 1996). The S-phase specific CLB5 and CLB6 primarily drive events relating to

DNA replication, such as the initiation of DNA replication and the inhibition of

replication origin reloading (Richardson et al. 1992; Nasmyth 1996), but in certain

genetic backgrounds also function at START or at G2/M (Epstein and Cross 1992;

Schwob and Nasmyth 1993). Despite showing considerable genetic redundancy, the G1

cyclins, CLN1, CLN2, and CLN3 (Dirick, Bohm, and Nasmyth 1995; Stuart and

Wittenberg 1995; Levine, Huang, and Cross 1996; Skotheim et al. 2008), and the B-type

cyclins, CLB1, CLB2, CLB3, CLB4, CLB5, and CLB6 (Segal, Clarke, and Reed 1998;

Stuart and Wittenberg 1998; Cross et al. 1999), are functionally specialized and activate

9

cell cycle processes by distinct mechanisms (Enserink and Kolodner 2010) as

summarized below.

In G1 phase, Cln3-Cdc28 activates expression of the G1 cyclins CLN1 and CLN2

and the B-type cyclins CLB5 and CLB6 (Tyers, Tokiwa, and Futcher 1993; Dirick,

Bohm, and Nasmyth 1995; Levine, Huang, and Cross 1996). The emergence of Cln1-

Cdc28 and Cln2-Cdc28 activity in late G1 phase (Wittenberg, Sugimoto, and Reed 1990;

Tyers, Tokiwa, and Futcher 1993) targets the repressor of Clb-Cdc28 activity, Sic1, for

degradation (Schwob et al. 1994; Schneider, Yang, and Futcher 1996; Verma et al. 1997;

Nash et al. 2001). Apart from their role in the degradation of Sic1, the Cln1-Cdc28 and

Cln2-Cdc28 complexes also control various processes related to bud emergence and cell

polarity (Lew and Reed 1993; Cvrckova and Nasmyth 1993; Benton et al. 1993; Nasmyth

1996). Once Sic1 is degraded, the Clb5-Cdc28 and Clb6-Cdc28 complexes promote

DNA replication early in S phase (Schneider, Yang, and Futcher 1996; Tyers 1996; Nash

et al. 2001) through activation of pre-replication complexes (PRC) on DNA (Tye 1999).

The late S phase accumulation of CLB3 and CLB4 is important for events related to the

initiation of mitosis (Fitch et al. 1992; Richardson et al. 1992). Clb3-Cdc28 and Clb4-

Cdc28 kinases have a minor early role in spindle morphogenesis and a major role in

promoting spindle assembly (Segal et al. 2000). At the G2/M transition, Cdc28 associates

with either one of the B-type cyclins Clb1 or Clb2 to promote mitosis (Fitch et al. 1992;

Grandin and Reed 1993). Proteolytic degradation of key cell cycle regulators such as

cyclins, and dephosphorylation of cyclin targets are other important contributors to cell

cycle phase transitions (Reed 2003; Potapova et al. 2006).

10

1.1.3 Cell cycle – Entry from G0 to G1

As noted earlier, budding yeast cells in early to mid G1 stop active cell division

when external conditions are unfavourable for continued growth (Johnston, Pringle, and

Hartwell 1977; Lillie and Pringle 1980). If nutrients are absent, cells exit the cell cycle

and enter into a quiescent state characterized by very low metabolic activity and very

little protein synthesis (Choder 1991; Jona, Choder, and Gileadi 2000). The genes

encoding many important activators of the G1/S transition, such as CLN3, SWI4 and

CDC28, are down-regulated under conditions of glucose, nitrogen or phosphate

deprivation (Klosinska et al. 2011). In the presence of ample nutrients, the TOR pathway

functions to repress a quiescence program, but if nutrients are absent the TOR pathway is

inactivated and cells enter quiescence (Schmelzle and Hall 2000; Rohde and Cardenas

2004; Powers 2004; Lorberg and Hall 2004; Duvel and Broach 2004; Loewith 2011).

When nutrients are plentiful, TOR proteins promote association of PP2A catalytic

subunits, such as Sit4, Pph21, and Pph22, with the protein Tap42 (Di Como and Arndt

1996; Jiang and Broach 1999). The presence of rapamycin or entry into quiescence

causes dissociation of Tap42 from its phosphatase partners and results in downregulation

of TOR signaling (Di Como and Arndt 1996; Jacinto et al. 2001; Duvel and Broach

2004). Under rich nutrients, the TOR signalling pathway activates a number of genes

within the RiBi regulon (Klein and Struhl 1994; Neuman-Silberberg, Bhattacharya, and

Broach 1995; Cardenas et al. 1999; Hardwick et al. 1999; Powers and Walter 1999;

Martin, Soulard, and Hall 2004). The RiBi regulon consists of genes involved in

ribosome biogenesis that are expressed in early G1 phase by addition of nutrients (Gasch

11

et al. 2000; Hughes et al. 2000; Wade et al. 2001; Jorgensen et al. 2002; Miyoshi, Shirai,

and Mizuta 2003).

1.2 Transcriptional circuitry of the cell cycle

In S. cerevisiae, there are six main clusters of genes that are synchronously

expressed at different points in the cell cycle (Figure 1-2A), and I provide an overview of

the regulation of each of these groups in this section. Early studies of cell cycle-

dependent gene expression in S. cerevisiae focused on the histone genes (Borun et al.

1975; Hereford et al. 1981; Hereford, Bromley, and Osley 1982). Histone gene

expression is induced during S phase to ensure production of sufficient histones to

package newly replicated DNA into chromatin. The induction of gene expression only

when the gene products are needed avoids unnecessary protein synthesis and is a

common strategy in biology. For example, the subunits of the E. coli flagellum are

synthesized in the order they are required for assembly (Kalir et al. 2001), and the precise

temporal control of transcription during E. coli amino acid synthesis ensures that

enzymes are made in the order they are needed (Zaslaver et al. 2004). Since the

discovery of cell cycle-dependent transcription of histone genes, many other genes have

been shown to exhibit clear oscillations in transcript levels at different points in the cell

cycle. Microarray expression profiling and other experiments have allowed grouping of

cell cycle regulated genes into “clusters” (Spellman et al. 1998; Tavazoie et al. 1999),

such that approximately 20% of budding yeast genes is estimated to be transcribed

periodically, with at least a two-fold fluctuation, in the cell cycle (Cho et al. 1998;

Spellman et al. 1998).

12

Figure 1-2A. Periodic gene expression. In S. cerevisiae, six main clusters of genes are synchronously

expressed at distinct points in the cell cycle. Different cyclin-CDK complexes drive the synchronous

expression of these gene clusters. The periodic activity of CDKs is a consequence of the cell cycle-

dependent expression of different cyclin subunits (shown within curves). Distinct transcription factor

binding sites in the promoters of periodically expressed genes confine transcription to a specific cell cycle

phase. Representative members of a gene cluster are shown within the curves. The Y-axis represents the #

of induced genes.

CLB1 CLB2 SWI5 ACE2

CLN1 CLN2 PCL1 PCL2 CLB5 CLB6 NDD1

HTA2 HTB2 CLN3

SWI4

CLB3 CLB4

SIC1 ASH1 RME1

G1 S G2 M

Cdk1 Cdk1 Cdk1 Cdk1 Cdk1

Cln3 Clb3,4 Clb1,2 Clb5,6 Cln1,2

13

Conservation of specific DNA elements in the promoters of these periodically

expressed genes has been successfully used to identify regulatory proteins responsible for

coordinate expression of periodically expressed genes in S. cerevisiae (Kellis et al. 2003;

Harbison et al. 2004). Amazingly, approximately 75% of the regulatory proteins that

control these periodically expressed genes are themselves periodically expressed such

that they peak just before they are required (Simon, Barnett et al. 2001). This serial

(Simon, Barnett et al. 2001) or regulator chain model (Lee et al. 2002) for cell cycle

transcription proposes that regulators at an earlier phase of the cell cycle drive expression

of regulators required at a later phase of the cell cycle. Consequently, aberrant

expression of genes earlier in the cell cycle can cause aberrant expression of genes

expressed later on in the cell cycle. Thus, cell cycle transcriptional control occurs

through a highly connected network of transcriptional regulators (Figure 1-2B), which

function at specific sequences in the promoters of periodically expressed genes (Lee and

Young 2000; Garvie and Wolberger 2001; Orphanides and Reinberg 2002).

14

Figure 1-2B. Transcriptional circuitry of the cell cycle. Cell cycle transcription is coordinated by a

complex network of regulatory proteins, a subset of which bind to specific DNA sequences in the

promoters of periodically-expressed genes. Cell cycle regulatory proteins that control the expression of

distinct gene clusters begin to accumulate at a point in the cell cycle earlier than the time at which they are

required. For example, at the M/G1 transition, Swi5 and Ace2 activate transcription of the SIC1 gene

cluster, which leads to increased production of Sic1, an inhibitor of Clb-Cdk1 complexes. Also at M/G1,

transcription of the MCM1 gene cluster is activated by the DNA-bound complex of Mcm1-Yox1/Yhp1.

Two genes of the MCM1 cluster, SWI4 and CLN3, encode proteins involved in SBF (Swi4, Swi6) and MBF

(Mbp1, Swi6) activity. SBF/MBF activate CLN2 gene cluster gene expression at G1/S. The G1/S-regulated

expression of the CLN2 cluster genes CLN1 and CLN2 causes the initiation of a positive feedback loop in

addition to induction of the CLN2 cluster gene, HCM1, an activator of S phase cluster genes such as WHI5,

YOX1, YHP1, FKH2 and NDD1. Other constituents of the main cell cycle regulatory network are shown.

SWI5

HCM1

WHI5

SWI4

CLN3

SIC1

Cln3

CLN2

ACE2

NDD1

FKH2

CLB2

YHP1

YOX1

G1 G2

M

S

Whi5

Mbp1

Swi6

Swi6

Swi4

Cln2

Sic1

Hcm1

Mcm1

Swi5 Yox1

Yhp1

Ace2

Fkh2

Mcm1

Ndd1

Clb2

15

1.2.1 M/G1 transition

Two waves of gene expression occur during the M/G1 transition, which are

controlled by different regulators (Spellman et al. 1998). The MCM1 cluster contains

approximately 30-40 genes (Spellman et al. 1998) and is induced by Mcm1, a MADS-

box transcription factor (Shore and Sharrocks 1995; Nurrish and Treisman 1995). The

MCM1 cluster contains genes required for expression of genes encoding pre-replicative

complex components, which are necessary for DNA replication (Mcm2-7, Cdc6) and for

inducing genes involved in reinitiating the cell cycle (Swi4, Cln3) (Spellman et al. 1998;

Pramila et al. 2002). Mcm7, together with Mcm1, may also modulate its own expression

and the expression of other early G1-phase expressed genes (Fitch, Donato, and Tye

2003). Expression of the MCM1 cluster genes is also important for proper induction of

genes expressed in the next cluster or phase (the CLN2 cluster – see below). For

example, SWI4 encodes the DNA binding component of a G1-specific transcription factor

(see below), and if Mcm1 binding elements within the SWI4 promoter are mutated, then

CLN transcripts are low and deregulated (MacKay et al. 2001). Mcm1 binds as a dimer

to a palindromic site within the so-called ECB element (TTtCCcnntnaGGAAA early cell

cycle box) (McInerny et al. 1997), found upstream of MCM1 cluster genes. Like many

CLB2 cluster genes (see below), the promoters of MCM1 cluster genes also contain a

binding site for the homeodomain repressors Yox1 and Yhp1, which physically bind

Mcm1 (Pramila et al. 2002). YOX1 is expressed from mid-G1 to early S phase (Spellman

et al. 1998; Horak et al. 2002; Pramila et al. 2002), and YHP1 expression occurs slightly

later than YOX1 (Cho et al. 1998; Spellman et al. 1998). Expression of these repressors

16

correlates with inhibition of MCM1 cluster genes, and gene activation requires their

removal, presumably by some activating signal which has yet to be determined.

The second M/G1 cluster includes approximately 25-30 genes (Spellman et al.

1998; Zhu et al. 2000), which are mainly involved in cell separation (Knapp et al. 1996;

Kovacech, Nasmyth, and Schuster 1996) and in distinguishing daughter cells from

mother cells (Doolin et al. 2001). The so-called SIC1 cluster is activated by the Zn-finger

transcription factors Swi5 and Ace2, which have overlapping (Simon, Barnett et al. 2001)

but distinct roles (Dohrmann et al. 1992; Dohrmann, Voth, and Stillman 1996; McBride,

Yu, and Stillman 1999; Doolin et al. 2001; Laabs et al. 2003). For example, the HO

gene, involved in mating type switching, is specifically regulated by Swi5 (Breeden and

Nasmyth 1987; Andrews and Herskowitz 1989), partly due to coregulation of HO by

Pho2, which interacts with Swi5 but not Ace2 (McBride, Yu, and Stillman 1999).

Together, Swi5 and Ace2 regulate genes important for daughter cell-specific gene

expression (Bobola et al. 1996; Sil and Herskowitz 1996; McBride, Yu, and Stillman

1999; Cosma 2004; Paquin and Chartrand 2008). Both Swi5 and Ace2 become

functional only in late anaphase after relocalization to the nucleus (Nasmyth et al. 1990;

Moll et al. 1991; Shirayama et al. 1999).

1.2.2 G1/S transition

During the G1/S transition, more than 200 genes are upregulated (Spellman et al.

1998; Cho et al. 1998; Wittenberg and Reed 2005; Orlando et al. 2008; Granovskaia et al.

2010). These genes form the CLN2 cluster and include the G1 cyclins CLN1, CLN2,

PCL1, and PCL2 and the S phase cyclin genes CLB5 and CLB6. At the G1/S transition,

17

activation of transcription largely depends on two dimeric transcription factors, SBF

(SCB-Binding Factor) and MBF (MCB-Binding Factor) (Breeden 1996). Both

transcription complexes contain Swi6 as a common component, which interacts

separately with two DNA binding subunits, Swi4 and Mbp1, to form the heterodimers

SBF and MBF, respectively. The optimal binding site for SBF is called the SCB (Swi4

Cell cycle Box), and the optimal binding site for MBF is called the MCB (MluI Cell

cycle Box) (Breeden 1996; Lee et al. 2002; Kato et al. 2004), The SBF complex

activates transcription of G1 cyclin genes (Ogas, Andrews, and Herskowitz 1991;

Espinoza et al. 1994; Measday et al. 1994), the HO endonuclease gene involved in yeast

mating-type switching (Kostriken and Heffron 1984; Andrews and Herskowitz 1989),

and various cell wall biosynthesis genes (Igual, Johnson, and Johnston 1996). The MBF

complex activates transcription of S-phase cyclin genes (Koch et al. 1993) and many

DNA synthesis genes (de Bruin et al. 2006). Cln3-Cdc28 is the primary activator of

SBF- and MBF-mediated G1/S-specific transcription (Tyers, Tokiwa, and Futcher 1993;

Dirick, Bohm, and Nasmyth 1995; Stuart and Wittenberg 1995). Cln3, in association

with Cdc28, activates SBF by removing the repressor Whi5 from SBF (Costanzo et al.

2004; de Bruin et al. 2004), and it activates MBF by regulating the activity of the MBF-

associated protein Stb1 (de Bruin, Kalashnikova, and Wittenberg 2008).

1.2.3 S phase

During S phase, approximately 180 genes are upregulated (Pramila et al. 2006),

including FKH1 and FKH2, which encode transcription factors that function later during

the G2/M transition (Spellman et al. 1998) (see below). Among the many genes

18

expressed in S phase are the MET gene cluster and the histone gene cluster (Cho et al.

1998; Spellman et al. 1998). Many of the MET cluster genes encode proteins involved in

microtubule dynamics, the kinetochore, and sister-chromatid cohesion (Pramila et al.

2006), and their proper expression is important to prevent chromosome loss (Daniel et al.

2006). Although Clb-associated activity may have a minor regulatory role in S phase

(Richardson et al. 1992), transcription of MET cluster genes during S phase depends

largely on Hcm1, a transcription factor of the forkhead family (Zhu et al. 1993). The

HCM1 promoter is bound by SBF and MBF (Iyer et al. 2001), and both HCM1 mRNA

(Cho et al. 1998) and Hcm1 protein (Pramila et al. 2006) peak at the G1/S transition

(Rowicka et al. 2007). Hcm1 is also important for the S phase transcription of other

genes, such as FKH2, NDD1, YHP1 and WHI5, which have critical transcriptional

regulatory roles at different points in the cell cycle (Figure 1-2B) (Pramila et al. 2006).

Fkh2 may also have a role in S phase transcription, because it localizes to histone gene

promoters (Simon, Barnett et al. 2001). Histone genes are specifically activated during S

phase. Their regulation is clearly dependent on a number of histone chaperone

complexes, and histone promoters have binding sites for SBF (Osley et al. 1986; Simon,

Barnett et al. 2001; Iyer et al. 2001; Lee et al. 2002; Kato et al. 2004). The HIR histone

chaperone complex requires a specific sequence in histone gene promoters and works

with other histone chaperones, including Rtt106 (Fillingham et al. 2009), to repress

histone gene expression outside of S-phase (Sherwood, Tsang, and Osley 1993; Spector

et al. 1997; Dimova et al. 1999). Recently, restriction of histone gene transcription to S

phase has been shown to require phosphorylation of the chromatin boundary protein Yta7

by S phase-specific forms of both Cdc28 and Casein Kinase 2 (Kurat et al. 2011).

19

1.2.4 S/G2 transition

During the S/G2 transition, approximately 120 genes are upregulated (Spellman et

al. 1998). The S/G2 gene cluster includes the first wave of B-type cyclins, CLB3 and

CLB4 (Fitch et al. 1992). Surprisingly little is known about how the S/G2 gene cluster is

regulated, but Fkh1 localizes to the CLB4 promoter (Simon, Barnett et al. 2001), and the

promoters of other genes expressed at the S/G2 transition are enriched in Fkh2 binding

sites (Kato et al. 2004), suggesting that Fkh proteins may be important activators of this

cluster. Clb3 and Clb4 induce a Cdc28-dependent kinase activity from mid-S phase until

mitosis (Grandin and Reed 1993), which is required in the subsequent G2/M phase to

establish mitotic spindle orientation (Segal et al. 2000). In contrast to CLB3 and CLB4,

which peak early in S phase and decline near the end of nuclear division, the transcripts

of the mitotic cyclins CLB1 and CLB2 peak later on at the time of nuclear division

(Ghiara et al. 1991; Surana et al. 1991; Fitch et al. 1992). Consistent with the timing of

their transcription, CLB3 and CLB4 appear dedicated to roles in DNA replication and

spindle assembly, whereas the later-transcribed CLB1 and CLB2 genes seem to be

specialized for regulation of later mitotic events (see next section). Other members of the

S/G2 cluster (Spellman et al. 1998) are CWP1 and CWP2 (Caro et al. 1998), which

encode proteins involved in cell wall biosynthesis (van der Vaart et al. 1995). Because

the expression of cell wall biosynthesis genes is not confined to S/G2, CWP1 and CWP2

seem to have evolved specialization in the constitution of the wall of the growing buds in

G2, rather than the small buds of G1 (Smits et al. 2006).

20

1.2.5 G2/M transition

During the G2/M transition, approximately 35 genes are upregulated, including

CLB1 and CLB2, and the genes SWI5 and ACE2 encoding factors required for the next

wave of transcription at the M/G1 transition (Zhu et al. 2000; Kumar et al. 2000; Koranda

et al. 2000; Pic et al. 2000). Mcm1 forms a complex on CLB2 gene cluster promoters

with the forkhead transcription factor Fkh2 and Ndd1 (Lydall, Ammerer, and Nasmyth

1991; Loy, Lydall, and Surana 1999; Koranda et al. 2000; Kumar et al. 2000; Pic et al.

2000; Zhu et al. 2000; Hollenhorst, Pietz, and Fox 2001). This complex is regulated by

cell cycle-dependent kinases that cooperate to give maximal activation through

phosphorylation of Fkh2 and Ndd1 (Darieva et al. 2003; Reynolds et al. 2003; Pic-Taylor

et al. 2004; Darieva et al. 2006). Ndd1 is phosphorylated sequentially by Clb2-Cdc28

and the polo-like kinase Cdc5, which is required for the normal temporal expression of

the CLB2 cluster (Darieva et al. 2006). Clb2 and Cdc5 seem to be important activators of

their own transcription through Ndd1 phosphorylation (Amon et al. 1993; Reynolds et al.

2003), suggesting that the CLB2 cluster of genes is regulated by positive feedback.

1.2.6 Complexity of cell cycle promoters

Although key regulators of most waves of cell cycle transcription have been

identified, it is clear that periodic gene expression is under combinatorial control by

multiple promoter elements and regulators (Lee et al. 2002; Kato et al. 2004; Wang 2007;

Tuch et al. 2008; Li and Zhan 2008; Eser et al. 2011). Regulation by several promoter

elements means that the periodicity of gene expression may be maintained even when a

key regulator is compromised. For example, SPC110 periodicity is not lost in hcm1Δ

21

cells, even though maximal levels of SPC110 transcripts are reduced (Zhu and Davis

1998). The maintenance of periodicity is attributed to the presence of an MCB element

in the SPC110 promoter (Zhu and Davis 1998). Similarly, the CLB2 cluster gene CDC20

contains Fkh2-binding sites, but maintains periodicity in a strain lacking FKH1 and

FKH2, indicating that other factors, such as Hcm1 and Fhl1, might bind the CDC20

promoter (Zhu et al. 2000). Interestingly, the CDC20 promoter contains a binding motif

for the repressor Yox1 (Pramila et al. 2002). Thus, the G2/M expression of CDC20 is

likely due to activation of Mcm1 by Fkh2 with Ndd1, but gene expression is repressed

until M phase because its promoter also contains a Yox1 repressor site. The periodicity

of CDC20 is shifted to an M/G1 pattern in a fkh1Δfkh2Δ strain, since Yox1 repression

constrains transcription to the M/G1 window in the absence of the primary physiological

activator (Zhu et al. 2000).

Multiple promoter elements may also allow coordination of cell cycle-dependent

gene expression with sensitivity to other signals. For instance, RNR1 encodes a subunit

of ribonucleotide reductase and is cell cycle regulated with a transcriptional peak at the

G1/S transition (Elledge et al. 1993; de Bruin et al. 2006). RNR1 is also induced by DNA

damage (Elledge and Davis 1990), since ribonucleotide reductase is required for DNA

repair (Elledge et al. 1993). Similarly, AGA1, which encodes a cell wall protein (Roy et

al. 1991; Zhao et al. 2001), is cell cycle regulated with a transcriptional peak at the M/G1

transition and also induced by mating pheromone (Roy et al. 1991). Aga1 is inducible

and cell cycle regulated because the activity it contains is required for the formation of

both mating projections and buds (Gehrung and Snyder 1990; Chenevert, Valtz, and

Herskowitz 1994).

22

1.3 CDK-dependent regulation of the G1/S phase transition

As noted earlier, accumulation of the G1 cyclins (CLN1, CLN2, and CLN3) is rate-

limiting for the G1/S transition (Nash et al. 1988; Cross 1988; Hadwiger et al. 1989;

Wittenberg, Sugimoto, and Reed 1990; Gallego et al. 1997; Polymenis and Schmidt

1997; MacKay et al. 2001; Newcomb, Hall, and Heideman 2002), and factors that control

G1 cyclin accumulation determine the proper timing of G1/S. In this section, I discuss the

molecular bases of the events that lead to the G1/S transition (Figure 1-3), with an

emphasis on two important aspects of cell cycle entry at G1/S – positive feedback and

Cln3 abundance as an activating trigger.

23

Figure 1-3. CDK-dependent regulation of the G1/S phase transition. Accumulation of G1 cyclins

(CLN1, CLN2, CLN3) is rate-limiting for the G1/S transition. The molecular events of the G1/S transition

begin with SBF and MBF bound to the promoters of CLN2 cluster genes in an inactive form. SBF (Swi4-

Swi6) and MBF (Mbp1-Swi6) are bound to SCB and MCB elements early in G1 phase in complexes

containing Stb1 and Whi5. Cln3-Cdk1 activity emerges in early G1 phase and antagonizes the repressive

activity of Whi5 and Stb1. Once Cln3-Cdk1 induces CLN1 and CLN2 expression through activation of

SBF/MBF, the newly formed Cln1-Cdk1 and Cln2-Cdk1 CDK complexes activate their own production

through a positive feedback loop, which causes the remaining genes of the CLN2 cluster to become

activated. The hyper-phosphorylation of Stb1 and Whi5 allows the CLN2 cluster to be further activated.

The emergence of Cln1-Cdk1 and Cln2-Cdk1 also causes increased phosphorylation of Sic1, which is a

repressor of Clb-Cdk1 activity. Phosphorylation of Sic1 targets it for degradation. The liberation of Clb1-

Cdk1 and Clb2-Cdk1 from Sic1 turns off SBF-dependent transcription until the next G1 phase. MBF-

dependent transcription is turned off when the MBF-specific repressor Nrm1 binds MBF.

OFF ON OFF

Swi6

SCB

Whi5

Stb1

Swi6

SCB

Stb1

SCB

Whi5 Swi6

P P

Cdk1

Cdk1

Cln1, 2

P P

P P

P

Swi6

Stb1

MCB

Swi6

Mbp1

Stb1

MCB

Swi6

Mbp1

Nrm1 Stb1 P

P P

G0

MCB

Mbp1

Early G1 Early S START

Sic1

P

P Cln3

Swi4 Swi4

Swi4

Cdk1

Clb1,2 M

P

P

24

1.3.1 Cln3-dependent activation of SBF/MBF

SBF (Harrington and Andrews 1996; Koch et al. 1996) and MBF (de Bruin et al.

2006) are bound to SCB and MCB elements early in G1 phase in complexes containing

Stb1 (SBF- and MBF-specific) (de Bruin, Kalashnikova, and Wittenberg 2008) and Whi5

(SBF-specific) (Costanzo et al. 2004; de Bruin et al. 2004) (Figure 1-3). Timely

activation of SBF- (Dirick, Bohm, and Nasmyth 1995; Stuart and Wittenberg 1995) and

MBF-containing (de Bruin, Kalashnikova, and Wittenberg 2008) complexes depends on

Cln3-Cdk1 activity, which functions largely by antagonizing the repressive activity of

Whi5 (Costanzo et al. 2004; de Bruin et al. 2004) and Stb1 (de Bruin, Kalashnikova, and

Wittenberg 2008). Once Cln3-Cdc28 kinase activates CLN1 and CLN2 expression

through activation of SBF/MBF (Nasmyth and Dirick 1991; Ogas, Andrews, and

Herskowitz 1991; Tyers et al. 1992; Tyers, Tokiwa, and Futcher 1993; Cvrckova and

Nasmyth 1993), newly-formed Cln1-Cdc28 and Cln2-Cdc28 complexes activate their

own production through positive feedback and also activate expression of the remaining

genes in the CLN2 cluster (see next section) (Eser et al. 2011). Cln1-Cdc28 and Cln2-

Cdc28 phosphorylate and target Sic1, a repressor of Clb-Cdc28 activity, for degradation

(Schwob et al. 1994; Schneider, Yang, and Futcher 1996; Verma et al. 1997; Nash et al.

2001). Liberated Clb2-associated kinase then inactivates SBF (Amon et al. 1993;

Siegmund and Nasmyth 1996), turning off SBF-dependent transcription until the next G1

phase. The MBF-specific repressor Nrm1 inactivates MBF (de Bruin et al. 2006), turning

off MBF-dependent transcription until the next G1 phase.

25

1.3.2 A positive feedback loop -- G1/S

As noted above, a considerable body of evidence suggests that proper timing and

coherent activation of CLN2 cluster gene transcription results from the activity of a

positive feedback loop (Cross and Tinkelenberg 1991; Dirick and Nasmyth 1991; Price,

Nasmyth, and Schuster 1991; Ogas, Andrews, and Herskowitz 1991; Skotheim et al.

2008). In WT cells, the initial activation of the loop depends on Cln3-Cdc28 activity in

early G1, which precedes Cln1- and Cln2-associated activity (Tyers, Tokiwa, and Futcher

1993). Two studies challenged the positive feedback model, because mutant cln1Δcln2Δ

cells cannot initiate positive feedback, and yet Cln3 can adequately activate SBF- and

MBF-dependent transcription in these cells (Stuart and Wittenberg 1995; Dirick, Bohm,

and Nasmyth 1995). However, time-lapse microscopy of single cells revealed that

activation of SBF-regulated genes is clearly delayed in the absence of Cln1 and Cln2

(Skotheim et al. 2008), indicating that the positive feedback model is in fact correct. The

earlier studies (Stuart and Wittenberg 1995; Dirick, Bohm, and Nasmyth 1995) failed to

find positive feedback because of the greater heterogeneity in expression of CLN2 cluster

genes in cln1Δcln2Δ cells (Skotheim et al. 2008). Interestingly, CLN2pr-GFP expression

in cln1Δcln2Δ cells persists for a longer period of the cell cycle and at higher levels once

it is activated (Skotheim et al. 2008), likely because the inhibitor of SBF, Clb2, is not

produced as promptly in the cln1Δcln2Δ mutant (Carey, Leatherwood, and Futcher

2008). Therefore, in cln1Δcln2Δ cells that happen to trigger CLN2 transcription early,

the strength of this activation is likely masked at a population level by lack of activation

in the remaining cells (Carey, Leatherwood, and Futcher 2008), making the cln1Δcln2Δ

cells seem like WT cells in the earlier studies (Stuart and Wittenberg 1995; Dirick,

26

Bohm, and Nasmyth 1995). In cln3Δ cells (Cross 1988) the coherence of START is

likely maintained by positive feedback involving CLN1 and CLN2 (Bean, Siggia, and

Cross 2006). Because passage through START is coincident with peak transcription of

the CLN1 and CLN2 genes, the timing of START depends on when the positive feedback

loop is triggered (Eser et al. 2011).

The positive feedback loop at the G1/S transition is an effective way to convert a

relatively gradual increase in Cln3-associated kinase activity in early G1 into a switch-

like signal in late G1. The switch-like nature of cell cycle commitment has properties

similar to ultra-sensitivity (Ferrell 1996). For example, because multiple

phosphorylations of Swi6 (Sidorova, Mikesell, and Breeden 1995; Wijnen, Landman, and

Futcher 2002; Ubersax et al. 2003; Geymonat et al. 2004) and Whi5 are required for

activating G1/S transcription (de Bruin et al. 2004; Costanzo et al. 2004), activation of

SBF-regulated genes may be ultra-sensitive to the concentration of gradually

accumulating Cln3-Cdc28 complexes. Similar features are likely present in the activation

of G2/M-regulated CLB2 cluster genes (Amon et al. 1993) and in the embryonic cell

cycle, where a positive feedback loop has been hypothesized to convert the gradual

accumulation of cyclin into a sharp activation of the MPF cdc2/cyclin kinase complex,

which triggers entry from interphase into mitosis (Solomon et al. 1990; Pomerening,

Sontag, and Ferrell 2003).

1.3.3 Cln3 abundance as a threshold for cell cycle commitment

Cln3 abundance has been proposed to set the rate of CLN1 and CLN2 mRNA

accumulation and hence the timing of START, relative to nutrient status (Tyers, Tokiwa,

27

and Futcher 1993). In cells released from nutritional arrest, accumulation of CLN2

mRNA begins only after peak CLN3 expression (Hubler, Bradshaw-Rouse, and

Heideman 1993). In particular, CLN3 transcription is induced rapidly upon glucose

treatment of nutrient-limited cultures (Barbet et al. 1996; Newcomb et al. 2003) and in

media containing rich nitrogen sources (Parviz and Heideman 1998). Interestingly,

translation of CLN3 mRNA is also repressed 8-fold under nitrogen deprivation conditions

(Gallego et al. 1997), providing an additional constraint on Cln3 protein levels in

starvation conditions. Because the Cln3 protein is very low in abundance and unstable

(Tyers et al. 1992; Cross and Blake 1993), the levels of Cln3 might also reflect the

general rate of protein synthesis.

Translation of CLN3 is exquisitely sensitive to decreases in the abundance of

translational initiation complexes (Barbet et al. 1996; Polymenis and Schmidt 1997;

Gallego et al. 1997; Hall et al. 1998). In particular, if the translation initiation factor eIF3

is inactivated, Cln3 translation is reduced by approximately half (Polymenis and Schmidt

1997). A three nucleotide long region of the CLN3 mRNA 5’ leader sequence has been

identified as a translational control element, which represses CLN3 expression during

periods of diminished protein synthesis or slow growth (Polymenis and Schmidt 1997).

Moreover, overexpression of CLN3 or the translation initiation factor-encoding gene

EIF5A can bypass the G1 arrest of cells engineered with a hyperactive pheromone

signalling pathway (Edwards et al. 1997), indicating that EIF5A, like eIF3, is important

for proper CLN3 translation (Kang and Hershey 1994). Consistent with the ability of

multiple translation factors to impinge on CLN3 translation, overexpression of the

translation initiation factor-encoding gene EIF4E increases CLN3 expression and causes

28

premature entry into S phase (Anthony, Zong, and De Benedetti 2001). Moreover,

expression of hyperstable or eIF4E-independent alleles of CLN3 restores G1/S

progression in strains mutated for eIF4E (Danaie et al. 1999) and suppresses rapamycin-

induced G1 arrest (Barbet et al. 1996). Strikingly, overexpression of eIF4E transforms

mammalian cells (Lazaris-Karatzas, Montine, and Sonenberg 1990) and results in

increased levels of cyclin D1 (Rosenwald et al. 1993), the analog of budding yeast CLN3

(Hatakeyama et al. 1994).

29

1.4 A CDK-dependent oscillator

Work on the G1/S phase transition (see above) and other cell cycle regulatory

events has led to the description of the cell cycle as a negative feedback oscillator (Qiao

et al. 2007) controlled by the periodic activity of a single master CDK regulator (Futcher

2002) (Figure 1-4). According to the oscillator model, the periodic activity of Clb2-Cdk1

is responsible for cell cycle progression. As described in the previous section, the

oscillator begins with activation of the transcription factors SBF/MBF by active Cln3-

Cdk1 (Tyers, Tokiwa, and Futcher 1993; Stuart and Wittenberg 1995; Dirick, Bohm, and

Nasmyth 1995), which leads to expression of the CLN2 gene cluster (Spellman et al.

1998). The sudden appearance of Cln1- and Cln2-Cdk1 accelerates expression of the

CLN2 cluster by positive feedback (Skotheim et al. 2008) and causes degradation of Sic1,

the inhibitor of Clb-Cdc28 kinase (Schwob et al. 1994; Schneider, Yang, and Futcher

1996; Tyers 1996; Verma et al. 1997). The sudden appearance of Clb2-Cdk1 then

inactivates SBF/MBF (Amon et al. 1993; Siegmund and Nasmyth 1996) and

simultaneously activates a transcriptional complex comprised of Mcm1, Fkh2, and Ndd1.

The Mcm1, Fkh2, Ndd1 complex drives transcription of the CLB2 cluster of genes, which

includes genes that encode activators of the SIC1 cluster (Amon et al. 1993; Koranda et

al. 2000; Kumar et al. 2000; Pic et al. 2000; Zhu et al. 2000). The expression of the SIC1

cluster increases production of Sic1, which inactivates residual Clb2-Cdk1 (Schwob et al.

1994; Toyn et al. 1997). Ultimately, the inactivation of Clb2 allows SBF/MBF to once

again become available for activation by Cln3-Cdk1. Consistent with the opposite effects

of Cln3 and Clb2 on SBF/MBF activity within this model, genome-wide transcriptional

30

Figure 1-4. A CDK-dependent oscillator. Top panel: In S. cerevisiae, six main clusters of genes are

synchronously expressed at distinct points in the cell cycle. Different cyclin-CDKs complexes drive the

synchronous expression of these gene clusters. Representative members of a gene cluster are shown within

the curves. Bottom panel: A CDK-dependent oscillator. The Cln3-Cdk1 CDK complex activates

SBF/MBF transcription factors bound to the promoters of CLN2 cluster genes, causing both the activation

of CLN2 cluster gene expression and the initiation of a positive feedback loop. Activation of the CLN2

gene cluster eventually leads to activation of the Clb2-Cdk1 CDK complex, causing both inhibition of

CLN2 cluster gene expression through inhibition of SBF/MBF activity, and the activation of a Mcm1-Fkh2-

Ndd1 transcriptional complex on the promoters of CLB2 cluster genes. Activation of the CLB2 cluster

eventually leads to activation of the SIC1 gene cluster, which includes genes encoding many proteins

including Sic1, an inhibitor of Clb2-Cdk1 activity. Inhibition of Clb2-Cdk1 allows reactivation of

SBF/MBF by the re-emergence of Cln3-Cdk1 activity in G1 phase.

Cln3 SBF MBF

Clb2 Mcm1 Fkh2 Ndd1

CLB2 cluster

I II

IV V

VI

CLN2 cluster

SIC1 cluster

III

CLB1 CLB2 SWI5 ACE2

CLN1 CLN2 PCL1 PCL2 CLB5 CLB6 NDD1

HTA2 HTB2 CLN3

SWI4

CLB3 CLB4

SIC1 ASH1 RME1

G1 S G2 M

Cdk1 Cdk1 Cdk1 Cdk1 Cdk1

Cln3 Clb3,4 Clb1,2 Clb5,6 Cln1,2

31

changes caused by CLB2 induction are almost the opposite of those caused by CLN3

overexpression (Spellman et al. 1998).

Although the CDK-dependent oscillator model illustrates the basic logic behind

cell cycle progression, many individual proteins that drive the oscillator are not essential

for cell viability. For example, deletion of CLB2 is not lethal to cells (Surana et al. 1991;

Fitch et al. 1992), probably because Clb1 can fulfill the essential functions of Clb2, while

clb1Δclb2Δ cells are nearly inviable (Fitch et al. 1992). Similarly, deletion of other

oscillator components, such as CLN3, is not lethal, while cells lacking all three G1 cyclins

arrest at START (Richardson et al. 1989; Cross 1990). The essential function of Cln-

CDK1 kinases appears to be degradation of SIC1, since a triple cln deletion mutant

lacking SIC1 is viable (Tyers 1996; Schneider, Yang, and Futcher 1996). In this genetic

situation, Clb-associated kinase activity is able to execute functions related to S and M

progression. Therefore, genetic redundancy is a prevalent feature of important cell cycle

regulators and may provide plasticity in the budding yeast CDK-dependent oscillator.

Consistent with this idea, if cln1Δcln2Δcln3Δ cells overexpress CLB5, then viability is

restored (Epstein and Cross 1992; Schwob and Nasmyth 1993), indicating that if Clb5

levels are high enough it is possible to overcome repression due to Sic1. Interestingly,

CLB5 is one of only two genes in the budding yeast genome whose overexpression

confers viability to a cln1Δcln2Δcln3Δ strain. The other gene is BCK2, which I discuss

in more detail below.

32

1.5 CDK-independent regulation of G1/S and cell cycle progression

The first indication that a CDK-independent oscillator exists was the discovery of

several CDC mutants that displayed synchronous and periodic rounds of budding in the

absence of DNA replication (Hartwell 1971a) and cytokinesis (Hartwell 1971b) such as

CDC3, CDC10, CDC11, and CDC12. Later, cells disrupted for all S-phase and mitotic

cyclins (clb1,2,3,4,5,6) were constructed that arrest at the G1/S transition with

unreplicated DNA (Lew and Reed 1993; Haase and Reed 1999; Haase, Winey, and Reed

2001), and yet these cells displayed synchronous rounds of budding, Cln2-associated

kinase activity (Haase and Reed 1999) and synchronous gene expression (Orlando et al.

2008), in a manner virtually identical to that seen in WT cells (Simon, Barnett et al. 2001;

Lee et al. 2002; Pramila et al. 2006). These findings were consistent with the existence

of a CDK-independent oscillator. Consistent with the ability of CLB-less cells to

promote budding, cells capable of producing hyper-stable Sic1 retain the ability to bud

synchronously (Mathias, Steussy, and Goebl 1998). The CDK-independent oscillator

embodies the extent of regulatory integration underlying the cell division cycle.

Strikingly, although Cln3-Cdc28 absolutely requires intact SBF/MBF (Wijnen, Landman,

and Futcher 2002; Ferrezuelo et al. 2010), only 25% of gene promoters expressed at

START actually bind SBF and MBF (Iyer et al. 2001) by the method of ChIP-chip (Buck

and Lieb 2004), implying that a large fraction of genes periodically expressed at START

might be regulated in a Cln3-independent manner.

The major CDK-independent pathway for activating G1-specific transcription

involves proteins with specific roles in ion homeostasis (Figure 1-5). Although the links

between ion homeostasis and cell cycle progression are not fully delineated, it is clear

33

Figure 1-5. CDK-independent regulation of the G1/S phase transition. The major CDK-independent

pathways that regulate the G1/S transition contain proteins with roles in intracellular ion homeostasis.

These proteins include various plasma membrane cation transporters, phosphatases, and phosphatase

regulators. Proteins in these CDK-independent pathways coordinate internal K+ and H

+ levels with cell

cycle initiation at START. The Sit4 and Ppz1 phosphatases are representative members of the two major

pathways that coordinate ion homeostasis with START. Sit4 with either Sap155 or Sap185 regulates the

activity of Nha1, a membrane protein that extrudes metallic cations as it imports protons. As a

consequence of their effect on Nha1 activity, Sit4-Sap185 promotes K+ efflux, whereas Sit4-Sap155

represses K+ efflux. Another major CDK-independent pathway is defined by the Ppz1 phosphatase, which

inhibits the activity of the K+ transporters Trk1, Trk2, and the cation efflux pump Ena1. Hal3 and Vhs3 are

repressors of Ppz1 phosphatase activity. Hal4 and Hal5 are partially redundant activators of K+ transport

through positive modulation of Trk1 and Trk2 transporter activity. Ptk2 is a Hal5-like protein involved in

activation of the Pma1 proton efflux pump. How all of these proteins interface with the G1/S

transcriptional machinery is not known. Bck2 is an enigmatic protein of unknown molecular function that

can activate the G1/S transition to a large extent in the absence of the CDC28 (CDK1) gene and may

explain how CDK-independent pathways transmit upstream signals to the cell cycle machinery.

Bck2

?

CLN1 CLN2 CLB5 SWI4

G1 S

K+

Trk1,

Trk2

Na+/Li

+/K

+ Na

+/Li

+/K

+

Nha1

H+

Pma1

ATP

ADP

ATP

ADP

Ena1

Hal4

Ptk2

Hal3/

Sis2 Ppz1

Vhs3

Hal5

Sit4

H+

Sap155 Sap185

+ +

Cdk1

Cln3

SBF MBF

34

that transit through START is heavily impacted by intracellular levels of important ions.

Plasma membrane transporters and regulatory phosphatases are two important classes of

proteins involved in cell cycle progression. In G1 phase, these proteins act together to

balance internal K+ and H

+ levels and allow START to occur (Arino, Ramos, and

Sychrova 2010). The balance between internal K+ and H

+ levels is key for maintaining

proper cellular pH and to maintain electrical balance -- K+ flow in one direction must be

replaced by H+ flow in the opposite direction (Arino, Ramos, and Sychrova 2010). K

+ is

important for many physiological functions, whereas Na+ is toxic and must be extruded

from the cell (Arino, Ramos, and Sychrova 2010). Alterations in intracellular ions also

alters internal turgor pressure, (Lew et al. 2006), which must be maintained at proper

levels to allow processes such as cell wall expansion and division to occur (Arino,

Ramos, and Sychrova 2010). Therefore, the commitment to START requires that the

integrity of the cell wall is also maintained.

35

1.5.1 Sit4 & Ppz1 pathways

The SIT4 and PPZ1 genes encode phosphatases necessary for the proper

execution of START and define two pathways distinct from the CDK-dependent pathway

(Sutton, Immanuel, and Arndt 1991; Fernandez-Sarabia et al. 1992; Clotet et al. 1999).

Sit4 is a type 2A-related Ser-Thr protein phosphatase (Arndt, Styles, and Fink 1989)

homologous to human PP6 phosphatases (Bastians and Ponstingl 1996). Sit4 functions in

a pathway genetically independent of Cln3-Cdc28, because sit4 mutants are synthetic

lethal with cln3 (Fernandez-Sarabia et al. 1992; Clotet et al. 1999) and cdc28 (Sutton,

Immanuel, and Arndt 1991) mutants. Physical association of Sit4 with the co-factors

Sap155 or Sap185 is important for transit through the G1/S transition (Luke et al. 1996).

The G1/S transition defects of sit4 mutants can be partly cured by constitutive CLN2

expression, which allows sit4 cells to replicate their DNA but not to form buds

(Fernandez-Sarabia et al. 1992). In contrast, overexpression of SWI4 is sufficient to

rescue both the DNA replication and budding defects of sit4 mutants (Fernandez-Sarabia

et al. 1992). PPZ1 encodes a negative regulator of G1 cyclin expression and acts in

opposition to Sit4 at START (Posas et al. 1992; Clotet et al. 1999). Ppz1 likely functions

in a pathway independent of Cln3-Cdc28 because the slow-growth phenotype of PPZ1-

overexpressing cells is exacerbated in a cln3Δ strain (Clotet et al. 1999), indicating that

the two pathways are additive. The HAL3 (SIS2) gene product encodes a negative

regulatory subunit of the Ppz1 Ser-Thr protein phosphatase (de Nadal et al. 1998; Clotet

et al. 1999). High-copy HAL3 can rescue the slow-growth defect of sit4 mutants (Sutton,

Immanuel, and Arndt 1991; Di Como, Bose, and Arndt 1995). Combined loss of SIT4

and HAL3 arrests cells in G1 phase due to defective expression of CLN1, CLN2, CLB5,

36

and SWI4 (Di Como, Bose, and Arndt 1995; Simon, Clotet et al. 2001). However, the

lethality of a hal3Δsit4Δ strain can be rescued by deletion of PPZ1 (Clotet et al. 1999),

suggesting that removal of Ppz1 inhibitory activity is sufficient for activating START

even in the absence of the START-promoting activity of Sit4.

1.5.2 Ion homeostasis in G1 phase

Sit4 and Ppz1 activate START through regulation of cation transporter activity at

the plasma membrane (Figure 1-5). High Na+, Li

+, and K

+ levels outside of the cell tend

to increase K+ efflux and also induce SIT4 transcription (Masuda et al. 2000). The effect

of Sit4 on K+ efflux requires the presence of the Nha1 membrane transporter

(Manlandro, Haydon, and Rosenwald 2005). NHA1 encodes an antiporter that increases

Na+ (Prior et al. 1996) and Li+ tolerance when the external pH is acidic (Banuelos et al.

1998; Sychrova, Ramirez, and Pena 1999). The association of Sit4 specifically with

Sap185 is necessary for activating K+ efflux, whereas association with Sap155 is

necessary for repressing K+ efflux (Manlandro, Haydon, and Rosenwald 2005).

Intracellular K+ levels are also controlled by the Hal3/Ppz1 pathway, which requires the

high-affinity K+ transporters encoded by the TRK1 and TRK2 genes (Yenush et al. 2002).

HAL3 was originally identified by its ability to confer tolerance to high salinity when

present at high-copy (Ferrando et al. 1995). Consistent with antagonism between Hal3

and Ppz1 activities (de Nadal et al. 1998; Clotet et al. 1999), ppz1 mutants are resistant to

Na+ and Li

+ (Posas, Camps, and Arino 1995) and display increased intracellular K

+ levels

and sensitivity to high extracellular KCl concentrations (Merchan et al. 2004). Ppz1-

deficient strains also show increased induction of ENA1 (Posas, Camps, and Arino 1995),

37

which encodes the ATPase pump responsible for efficient efflux of Na+ (Platara et al.

2006). Conversely, ENA1 induction is significantly decreased in hal3 mutants (Ferrando

et al. 1995). Collectively, the Nha1, Trk1, Trk2, and Ena1 plasma membrane transporters

control cation concentrations inside the cell by reacting to the activities of Sit4 and Ppz1

phosphatases in G1 phase.

The G1 arrest of a strain lacking SIT4 and HAL3 (Di Como, Bose, and Arndt

1995; Simon, Clotet et al. 2001) has been exploited to find other CDK-independent

activators of START. In one study, fourteen genes that bypass the G1 arrest phenotype of

a sit4 hal3 double mutant strain were identified (Munoz et al. 2003). These genes can be

binned into three functional categories: (1) ion homeostasis (HAL4, HAL5, PTK2, NHA1,

YAP7, VHS1, VHS2, and VHS3); (2) PP2A-related phosphatases (PPH21, PPH22,

PTC2); and (3) cell cycle activators (SWI4, CLN3, BCK2). Hal4 and Hal5 are partially

redundant activators of K+ transport that act by positively modulating Trk1 and Trk2

(Madrid et al. 1998; Mulet et al. 1999; Perez-Valle et al. 2007). The HAL5 family

member, Ptk2, activates the membrane transporter Pma1 (Goossens et al. 2000), which is

an H+-ATPase that can be activated by glucose (Eraso, Mazon, and Portillo 2006). YAP7

encodes a transcription factor of the bZIP family (Fernandes, Rodrigues-Pousada, and

Struhl 1997). The means by which Yap7 activates START is not presently known,

although another member of the YAP family, Yap6 (Hal7), can induce ENA1 expression

(Mendizabal et al. 1998). ENA1 expression is also strongly increased in ppz1 mutant

strains (Posas, Camps, and Arino 1995), indicating that cation extrusion might be an

important aspect of Yap7 function. Finally, although little is known about Vhs1, the

Vhs2 protein effects polarization of the actin cytoskeleton (Gandhi, Goode, and Chan

38

2006), and Vhs3, like Hal3, is another inhibitory subunit of Ppz1 (Ruiz et al. 2004;

Munoz et al. 2004).

The remaining six genes that suppress the lethality of the sit4 hal3 mutant strain

encode proteins with previously ascribed roles in cell cycle regulation. The genes PPH21

and PPH22 encode the two isoforms of the catalytic subunit of the type 2A protein

phosphatases (PP2Ac) in S. cerevisiae (Stark 1996). The slow-growth defect of sit4

mutants can be partially suppressed by high-copy PPH22 (Sutton, Immanuel, and Arndt

1991), indicating that PP2Ac can replace Sit4 function in vivo. However, PP2Ac usually

functions at the G2/M transition (Lin and Arndt 1995; Yang et al. 2000), so its role at

START is ambiguous. PTC2 encodes one of the five members of the type 2C Ser-Thr

phosphatase (PP2C) family in budding yeast (Stark 1996). The ability of CLN3 to

suppress the G1 arrest of a strain lacking SIT4 and HAL3 likely reflects the ability of

CLN3 to induce SWI4 transcription (Tyers, Tokiwa, and Futcher 1993; Foster, Mikesell,

and Breeden 1993), which activates many genes required for cell wall integrity.

Collectively, all of the genes discussed above, which suppress the G1 arrest of a

strain lacking SIT4 and HAL3, seem to function in processes much earlier than START.

Accordingly, although overexpression of CLN2 can rescue swi4Δmbp1Δ cells that are

blocked for START transcription (Wijnen and Futcher 1999), CLN2 was not isolated as a

high-copy suppressor of the sit4 hal3 strain, although this has not been tested directly.

However, one high-copy suppressor of the sit4 hal3 arrest phenotype, the enigmatic

BCK2 (“Bypass of C Kinase mutation”) gene, also rescues the lysis defects of various

mutants in the PKC1 pathway (Lee, Hines, and Levin 1993), which has roles in cell wall

integrity. More remarkably, overexpression of BCK2 also rescues the G1 arrest

39

phenotype of a strain lacking CLN1, CLN2, and CLN3 (Richardson et al. 1989; Cross

1990). These results suggest that Bck2 is an important integrator of signals during the

G1/S phase transition, and my thesis work has focused on understanding the molecular

function of Bck2 as a means of exploring CDK-independent regulation of the cell cycle.

1.5.3 The BCK2 gene

As noted above, BCK2 (Epstein and Cross 1994) and CLB5 (Epstein and Cross

1992; Schwob and Nasmyth 1993) are the only genes whose overexpression is known to

bypass the lethality of a cln1Δcln2Δcln3Δ strain (Richardson et al. 1989; Cross 1990),

although highly engineered CLB2 alleles can also suppress cln1Δcln2Δcln3Δ lethality

(Amon, Irniger, and Nasmyth 1994). The only reported mutational bypass of CLN

function (i.e. cln1Δcln2Δcln3Δ) occurs as a result of loss of function mutations at the

BYC1 locus (bypass of CLN requirement) (Epstein and Cross 1994) later shown to be

SIC1, an inhibitor of Clb-Cdc28 kinases (Tyers 1996). SIC1 requires the presence of

CDC28, SWI4, and SWI6 in order to overcome the G1 arrest of cln1Δcln2Δcln3Δ cells

(Epstein and Cross 1994), indicating that it is in a CDK-dependent pathway for START

activation (Figure 1-3). Because Clb5 is capable of performing all START-related

processes (Cross 1995; Oehlen, Jeoung, and Cross 1998), CLB5 overexpression likely

allows Cln3-independent activation of SBF/MBF target genes in the absence of CLNs.

Interestingly, overexpression of BCK2 stimulates CLB5 expression in a SWI4-

independent manner (Di Como, Chang, and Arndt 1995), indicating that BCK2 might

suppress the lethality of a cln1Δcln2Δcln3Δ strain partly by increasing CLB5 dosage.

The aforementioned increase in CLB5 expression by overexpressed BCK2 might also be

40

MBF-independent as proper transcription of another MBF-regulated gene, RNR1, is

restored by overexpressed BCK2 in a swi6Δ mutant (Di Como, Chang, and Arndt 1995).

Curiously, CLB5 transcription is also known to increase in strains overexpressing HAL3

(Di Como, Bose, and Arndt 1995), indicating that other proteins with known roles in ion

homeostasis (Figure 1-5) might also contribute to CLN bypass.

Several lines of evidence suggest that BCK2 functions in the SIT4 pathway,

because BCK2 shares many genetic interactions with SIT4. For example, like

bck2∆cln3∆ and bck2∆swi6∆ strains, sit4∆cln3∆ and sit4∆swi6∆ strains are synthetic

inviable (Fernandez-Sarabia et al. 1992; Di Como, Chang, and Arndt 1995), whereas a

bck2∆sit4∆ mutant has similar growth rates as sit4∆ alone (Di Como, Chang, and Arndt

1995). Moreover, overexpression of PPZ1 aggravates the slow-growth defect of bck2Δ

cells (Clotet et al. 1999), indicating that Bck2 is in a pathway distinct from Hal3/Ppz1.

Consistent with this idea, hal3Δbck2Δ cells grow much slowly than hal3Δ cells (Clotet et

al. 1999), again indicating that Bck2 functions in a pathway distinct from Hal3/Ppz1.

However, because overexpression of PPZ1 also aggravates the slow-growth defect of

cln3Δ cells (Clotet et al. 1999), definitive assignment of Bck2 to any single pathway is

difficult.

1.5.4 Bck2 – the problem of SBF/MBF-dependent transcription

Several lines of evidence suggest that Bck2 promotes the G1/S phase transition

(Epstein and Cross 1994; Wijnen and Futcher 1999) through SBF and MBF (Wijnen and

Futcher 1999; Di Como, Chang, and Arndt 1995). First, overexpressed BCK2 activates

CLN2 transcription through a region that contains three SCB and two MCB elements,

41

termed UAS1 (Stuart and Wittenberg 1994; Di Como, Chang, and Arndt 1995). Second,

overexpressed BCK2 can also activate transcription from artificial promoters constructed

purely of multiple synthetic SCB or MCB elements (Di Como, Chang, and Arndt 1995),

which rely exclusively on heterodimeric SBF and MBF for their activation. As a

consequence, the original isolation of BCK2 as a high-copy suppressor of the cell lysis

phenotype of PKC1 pathway mutants (Lee, Hines, and Levin 1993) has been proposed to

reflect the ability of Bck2 to activate SBF/MBF-dependent cell wall biosynthesis genes,

since both PKC1 (Mazur et al. 1995; Zhao et al. 1998) and SBF/MBF (Spellman et al.

1998) induce cell wall genes. Third, both bck2Δ and cln3Δ cells are defective in CLN2

transcription and cln3Δbck2Δ cells are dead (Di Como, Chang, and Arndt 1995), but can

be rescued by high-copy CLN2 (Wijnen and Futcher 1999). These data are consistent

with the idea that Cln3 and Bck2 function in parallel pathways to activate SBF and MBF.

In addition to a function that may depend on SBF/MBF, other evidence suggests

that Bck2 also activates the G1/S phase transition as part of a CDK-independent pathway

that does not require SBF/MBF. As described above, the CDK-dependent activation of

the G1/S transition requires removal of the repressor Whi5 from SBF (Swi4-Swi6) and

removal of Stb1 from MBF (Mbp1-Swi6) by Cln3-Cdc28 activity (Figure 1-3). Cln3

cannot function in the absence of Swi6 (Wijnen, Landman, and Futcher 2002), Swi4 or

Mbp1 (Ferrezuelo et al. 2010), and both Whi5 and Stb1 require Swi6 for association with

SBF and MBF (de Bruin et al. 2004; de Bruin, Kalashnikova, and Wittenberg 2008). In

contrast to the CDK-dependent pathway, activation of SCB/MCB-regulated genes by

Bck2 does not require SWI6 (Ferrezuelo, Aldea, and Futcher 2009). In fact, Bck2 can

activate several natural SBF/MBF target gene promoters in the absence of either the

42

SWI4 and MBP1 genes (Wijnen and Futcher 1999) or the elements that SBF/MBF bind

(Di Como, Chang, and Arndt 1995). Accordingly, although WHI5 overexpression causes

lethality in cln3Δ or cdc28-4 cells, WHI5 overexpression does not cause lethality in

bck2Δ cells (Costanzo et al. 2004) indicating that Bck2 does not antagonize the

repressive activity of Whi5. As expected for a CDK-independent activator of G1/S,

overexpression of BCK2, but not CLN3, can activate the SBF target PCL1 and the MBF

target RNR1 in a cdc28 mutant (Wijnen and Futcher 1999). Moreover, the UAS2 region

of the CLN2 promoter, which completely lacks SCB or MCB elements, has

transcriptional activity in a cdc28 mutant (Stuart and Wittenberg 1994), and this region

also responds to BCK2 overexpression (Di Como, Chang, and Arndt 1995).

1.6 Thesis rationale

The goal of my work was to explore mechanisms of G1-specific transcription and

how gene expression is linked to the intra- and extracellular environments. To better

understand how multiple regulatory pathways control the activity of the G1-specific

transcription factors SBF/MBF, I began my project by using affinity chromatography to

discover SBF/MBF binding proteins. In Chapter 2, I describe in vivo reconstitution of

purified heterodimeric SBF and MBF for use in biochemical assays. To choose proteins

to test for SBF/MBF-binding, I used the findings of a functional genomic screen that

identified genes important for proper transcription from SBF- and MBF-dependent

promoters. Out of seventeen candidates tested, I found four proteins that physically

interacted with both SBF and MBF: Elm1, Rtf1, Ykr077w, and Bck2. I focused on Bck2

because genetic studies strongly suggested a role for Bck2 in the activation of SBF and

43

MBF target genes, but the mechanism of Bck2 action was obscure. By ChIP, I show that

Bck2 localizes to two genes known to be activated at the G1/S transition by SBF/MBF.

The in vivo localization of Bck2 to promoters is also consistent with the ability of Bck2

fusion proteins to activate general transcription in reporter assays. The aggregate data in

Chapter 2 suggest that Bck2 might interact in vivo with SBF and MBF on promoter DNA

in order to activate transcription of genes whose expression peaks at G1/S.

In Chapter 3, I looked more closely at the property of Bck2 to auto-activate

transcription, and explored the possibility that Bck2 functions through promoter-bound

proteins other than SBF/MBF. I first asked whether the Bck2 auto-activation phenotype,

which occurs when a protein fused to a heterologous DB domain activates transcription

of a cognate reporter gene, is necessary for Bck2 in vivo function. I generated twenty

fusions of fragments of the BCK2 gene to the Gal4 DB domain and discovered that the

region of Bck2 which contained the auto-activation activity was not required for in vivo

function. I then used the largest Bck2 fragment lacking auto-activation but retaining in

vivo function as a bait protein in a Y2H screen. I discovered six novel binding partners

for Bck2 including the DNA-binding proteins Mot3 and Mcm1. The physical interaction

between Bck2 and Mot3 suggested a plausible explanation for suppression of cell wall

defects by high-copy BCK2.

My two-hybrid screen also identified Mcm1 as a Bck2-interacting protein, and I

explore the significance of this interaction in Chapter 4. Mcm1 regulates early G1 phase

transcription through binding ECB elements in the promoters of a set of genes that

include CLN3 and SWI4. Because early G1 phase expression of CLN3 and SWI4 delays

the G1/S transition, the physical interaction with Mcm1 suggested to me that Bck2 might

44

actually function much earlier than the G1/S transition point, when SBF and MBF are

activated by CDK activity. Consistent with an early G1 phase role of Bck2, I show that

BCK2 is required for the proper timing of CLN3 and SWI4 expression. Furthermore, the

localization of Bck2 to the promoters of CLN3 and SWI4 specifically required intact ECB

elements where Mcm1 also binds. Bck2 also localized to the CLN2 promoter, which is

predominantly activated by SBF. However, because the CLN2 promoter also has Mcm1

binding sites, my work suggests that Bck2 may function through Mcm1 sites rather than

SBF/MBF-binding sites in the CLN2 promoter. I propose that Bck2 performs a

predominant regulatory role early in G1 phase rather than at the G1/S transition as

previously thought. In contrast to the SBF-regulated HO gene, most G1/S-regulated

genes such as CLN2 contain multiple distinct promoter motifs, which reflects the extent

to which CDK-independent pathways impinge on cell cycle control. The roster of Bck2-

interacting proteins that I identified in my two-hybrid screen also implicates Bck2 in

coordinating production of G1/S phase regulatory factors with environmental stress

signals.

45

Chapter 2

A Biochemical Approach Identifies Bck2 as a

Binding Partner of the G1 transcription factors

SBF and MBF

46

Chapter 2: A biochemical approach identifies Bck2 as a binding partner of the G1

transcription factors SBF and MBF

2.1 Abstract

Protein-protein interactions in vitro often indicate an important functional interaction in

vivo. It has been difficult to produce purified SBF and MBF heterodimers for

biochemical analyses. To address this problem, I used a baculovirus-mediated expression

system to produce purified SBF and MBF in insect cells and tested the purified proteins

for DNA-binding in an in vitro assay. I then developed an affinity capture assay to ask if

putative activators of G1 phase gene expression bound purified SBF and MBF. To

choose putative activators, I used the findings of a functional genomic screen that

identified genes important for proper transcription from SBF- and MBF-dependent

promoters. In that screen, SCB-lacZ and MCB-lacZ reporter plasmids were introduced

into all of the viable single gene deletion mutants of budding yeast in order to identify

genes defective in SBF- and MBF-dependent transcription. I used TAP-tagged versions

of putative SBF/MBF regulators identified in the screen to test binding to SBF and MBF

in my affinity capture assay. Out of seventeen candidates tested, I found four proteins

that interacted with both SBF and MBF: Elm1, Rtf1, Ykr077w and Bck2. I focused on

Bck2 because genetic studies strongly substantiated the role of Bck2 in the activation of

SBF and MBF target genes. I observed that Bck2 not only interacts with the promoters

of two SBF- and MBF-regulated genes, but it also auto-activates transcription, consistent

with its role in transcriptional activation. These experiments suggest that Bck2 regulates

G1 transcription by binding to SBF and MBF on the promoters of late G1-expressed

genes.

47

2.2 Introduction

To understand how the G1/S phase transition occurs, it is necessary to discover

how the transcription factors SBF and MBF are regulated. Of the ~200 genes that exhibit

peak expression at the G1/S transition, most contain SCB and MCB elements (Cho et al.

1998; Spellman et al. 1998), which bind Swi4-Swi6 and Mbp1-Swi6 complexes (Primig

et al. 1992; Koch et al. 1993). DNA binding is not sufficient for activation, however,

because SBF (Harrington and Andrews 1996; Koch et al. 1996) and MBF (de Bruin et al.

2006) bind promoter elements early in G1 phase, yet gene activation does not occur until

late G1 phase. The timely activation of SBF (Dirick, Bohm, and Nasmyth 1995; Stuart

and Wittenberg 1995) and MBF (de Bruin, Kalashnikova, and Wittenberg 2008) depends

on Cln3-Cdk activity. However, because a cln3Δ strain is still viable and SBF/MBF-

dependent transcription still occurs in a cln3Δ strain, albeit slightly later, alternative

mechanisms must exist to activate SBF/MBF (Nasmyth and Dirick 1991; Stuart and

Wittenberg 1995). For example, one alternate regulator of SBF and MBF activity is Stb1

(de Bruin, Kalashnikova, and Wittenberg 2008). A stb1Δcln3Δ strain has more

pronounced G1-transcription defects than either stb1Δ or cln3Δ, which suggests that Stb1

and Cln3 act in non-redundant pathways (Ho et al. 1999). However, a stb1Δcln3Δ strain

is still alive, indicating that there are other SBF and MBF activators.

The regulatory mechanisms controlling SBF and MBF activity are complex,

genetically redundant, and not well understood. For instance, CLN1 expression is

regulated by SBF binding to MCB promoter elements (Partridge, Mikesell, and Breeden

1997), and isolated DNA binding domains from Swi4 or Mbp1 bind to both SCB and

MCB sequences in vitro (Koch et al. 1993; Partridge, Mikesell, and Breeden 1997;

48

Taylor et al. 2000). Moreover, Swi4 and Mbp1 bind overlapping sets of promoters in

vivo (Iyer et al. 2001; Simon, Barnett et al. 2001), and the absence of either Swi4 or

Mbp1 results in an increase in binding of the other factor to many SBF- and MBF-

specific promoters (de Bruin et al. 2006). Deletion of both DNA binding components of

SBF and MBF, Swi4 and Mbp1, leads to lethality (Koch et al. 1993), while single

mutants are viable, again supporting ‘cross-talk’ between SBF and MBF in vivo. Also,

deletion of SWI6 is not lethal, which is thought to reflect alternative mechanisms for

activating Swi4. Whether Mbp1 also has residual activity in the absence of SWI6 is not

clear, but Mbp1 can bind Skn7, and SKN7 is required for MBF-dependent transcription in

the absence of SWI6 (Bouquin et al. 1999; Lee et al. 2002). MBF sites in the RNR1

promoter are responsive to mutation of MBP1, but not of SWI4 (Dirick et al. 1992), and

when the same sequences are assayed in a reporter construct, mutation of SWI4 affects

expression (Verma et al. 1992) even though Swi4 does not localize to the endogenous

RNR1 promoter (de Bruin et al. 2006). Finally, for some genes, deletion of Mbp1 or

Swi4 enhances transcription, suggesting that these factors may also function as repressors

of transcription (Bean, Siggia, and Cross 2005).

To better understand the complex network of signalling in the activation of SBF

and MBF, a genomic screen was performed to identify novel regulators of SBF and MBF

dependent transcription (H. Friesen, M. Costanzo; unpublished). With the exception of

the HO gene, whose expression is completely dependent on SBF (Breeden and Nasmyth

1987; Breeden and Mikesell 1991), only synthetic SCB and MCB-driven reporter genes

exhibit a specific dependence on SBF and MBF, respectively (Andrews and Herskowitz

1989; Verma et al. 1992). To identify regulators that may be specific to SBF and MBF,

49

the Synthetic Genetic Array (SGA) (Tong et al. 2001) method was used to introduce

MCB-lacZ and SCB-lacZ plasmids into an array of approximately 5000 S. cerevisiae

deletion mutants (Winzeler et al. 1999), and the resultant strains were scored for

defective reporter activity. To identify G1-specific regulators amongst the candidate

genes identified in the screens I created a ‘short-list’ of genes with some biological

connection to cell cycle regulation. Like STB1, these candidate genes may encode

proteins that activate G1 transcription by physically associating with SBF and MBF (Ho

et al. 1999), an idea I tested in the experiments described in this Chapter.

2.3 Materials and methods

2.3.1 Yeast strains and plasmids

Yeast strains (Table 2-1) and plasmids (Table 2-2) were constructed using standard

methods.

Table 2-1

S. cerevisiae strains

Strainsa BY# Relevant genotype Source

BY4741 1623 MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 (Brachmann et al. 1998)

BY3015 3015 MATa cln3Δ::HphR bck2Δ::NatR This study

his3Δ1 leu2Δ0 met15Δ0 ura3Δ0lys2Δ0

pGAL-CLN3 (URA3)

Y8930 4889 MATα trp1-901 leu2-3,112 his3-200 ura3-52 (Yu et al. 2008)

gal4Δ gal80Δ LYS2::GAL1-HIS3 GAL2-ADE2

met2::GAL7-lacZ cyhR

BCK2-TAP 2719 MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 (Ghaemmaghami et al. 2003)

SWD1-TAP 2720 MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 (Ghaemmaghami et al. 2003)

RTF1-TAP 2721 MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 (Ghaemmaghami et al. 2003)

ELM1-TAP 2722 MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 (Ghaemmaghami et al. 2003)

MON2-TAP 2723 MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 (Ghaemmaghami et al. 2003)

SWD3-TAP 2724 MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 (Ghaemmaghami et al. 2003)

RTS1-TAP 2725 MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 (Ghaemmaghami et al. 2003)

BRE1-TAP 2726 MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 (Ghaemmaghami et al. 2003)

LGE1-TAP 2728 MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 (Ghaemmaghami et al. 2003)

SRL3-TAP 2729 MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 (Ghaemmaghami et al. 2003)

YKR077W-TAP 2730 MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 (Ghaemmaghami et al. 2003)

SKN7-TAP 2732 MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 (Ghaemmaghami et al. 2003)

MSN5-TAP 2735 MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 (Ghaemmaghami et al. 2003)

SLT2-TAP 2736 MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 (Ghaemmaghami et al. 2003)

CLN3-TAP 2742 MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 (Ghaemmaghami et al. 2003)

50

YOL131W-TAP 2744 MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 (Ghaemmaghami et al. 2003)

YBR174C-TAP 2745 MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 (Ghaemmaghami et al. 2003)

STB1-TAP 2747 MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 (Ghaemmaghami et al. 2003)

SWI4-TAP 4161 MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 (Ghaemmaghami et al. 2003)

a Strain backgrounds are isogenic to S288C

Table 2-2

Plasmids

Plasmid BA# Description Source

pBA1725 1725 pDEST8-FLAG-MBP1 This study

pBA1719 1719 pDEST8-FLAG-SWI4 This study

pBA1720 1720 pDEST8-SWI6 (native) This study

pBA 253 253 Xmn, Nru fragment of URS2

Sma site of pUC18; SCB-HO

pGAL-FLAG 350V LEU2 CEN GAL1pro-Gal4 DB-ORF-FLAG (Ho et al. 2002)

pGAL-BCK2-FLAG 2412 LEU2 CEN GAL1pro-Gal4 DB-ORF-FLAG (Ho et al. 2002)

pSH18-34 1971 (Estojak, Brent, and Golemis

1995)

pEG202 403V HIS3 2 μm ADH1pro-LexA DB-ORF (Gyuris et al. 1993)

pEG202-LexA-BCK2 2413 HIS3 2 μm ADH1pro-LexA DB-ORF This study

pDEST32 401V LEU2 CEN ADH1pro-Gal4 DB-ORF Invitrogen

pDEST32-Gal4-BCK2 2414 LEU2 CEN ADH1pro-Gal4 DB-ORF This study

2.3.2 Recombinant protein expression and purification

The Bac-N-Blue baculovirus system (Invitrogen) was used to express Swi6, Mbp1 and

Swi4 derivatives in insect cells. Plasmids pBA1725, pBA1719 and pBA1720, encoding

FLAG-Mbp1, FLAG-Swi4 and native Swi6, respectively were transformed into

DH10Bac E. coli cells (Invitrogen). Bacmid was isolated from each transformant and

recombinant viruses were generated and amplified over four rounds in Sf9 cells. Optimal

infection ratios were determined empirically for each virus. For expression of

recombinant complexes, flasks with 70% confluent Hi5 cells covering a 500 cm2 surface

were co-infected with the corresponding viruses and incubated for 45 hours. Cells were

then harvested and washed once with cold PBS and then resuspended in 0.75 ml lysis

buffer H (50 mM Tris-Cl, pH 7.5, 150 mM NaCl, 5 mM EDTA, 0.1% Nonidet-P40, 5

mM NaF, and one CompleteTM

Mini-tab (Boehringer) per 5 ml buffer). Following

51

incubation for 1 hour on ice, debris was removed by centrifugation at 14,000 x g for 8

minutes. Supernatants were incubated with 200 μl anti-FLAG M2-coupled agarose beads

(Sigma) in 1 ml TBS (10 mM Tris-Cl, pH 7.4, 150 mM NaCl) for one hour at room

temperature. Following three washes with TBS, bound protein was eluted twice with 100

μl TBS + 100 μg/ml FLAG peptide (Sigma) for 10 minutes at room temperature.

2.3.3 EMSA analyses

The SCB-HO probe was prepared by digesting plasmid pBA253 with EcoRI and BamHI.

The digestion products were then incubated with the Klenow fragment of E. coli DNA

polymerase I, [α32

P] dATP, and unlabeled dTTP, dCTP, and dGTP to label the HO

promoter fragment at both the EcoRI and BamHI ends. The labelled fragment was

purified by native PAGE electrophoresis on a 12% polyacrylamide gel followed by

excision of the DNA band, which was crushed and soaked in elution buffer C (0.5 M

NH4OAc pH 8, 1 mM EDTA) then incubated at 37°C overnight. The sample was

centrifuged at 14,000 x g for 5 minutes at 4°C before removal of supernatant for

subsequent purification by ethanol precipitation. The MCB-TMP1 (CDC21)

oligonucleotide is a 72-bp fragment containing a 54-bp region from the TMP1 promoter

(-163 to -110 5’of the initiation codon) (McIntosh et al. 1991), with the sequence 5’-

TGGTGACGCGTTAAATAGAAAAAATGAAAAAGACCTTAATTGACGCGTTTCC

TG-3’ flanked by 5’ BamHI and 3’ EcoRI linkers. Underlined are the two MluI sites,

which are MCB elements. Mutant MCB-TMP1 oligonucleotides were the same as the

WT, except that two nucleotides within the MluI site were mutated. The 5’ MCB was

mutated from ACGCGTTA to ACTCATTA and the 3’ MCB mutated from ACGCGTTT

52

to ACTCATTT, respectively (sense-strand sequences) (Dirick et al. 1992).

Complementary sets of oligonucleotides were synthesized to make dsDNA. Sense strand

oligonucleotide was end-labelled using [γ32

P] ATP and T4 polynucleotide kinase.

Unincorporated label was removed using a G50 microcolumn (General Electric). The

labelled oligonucleotide was combined with a 5-fold molar excess of cold antisense

oligonucleotide, and the two were annealed by placing in boiling water and allowing the

mixture to cool slowly to room temperature. Unlabelled competitor binding site probes

were made by annealing equimolar amounts of sense and antisense oligonucleotides.

Oligonucleotides were purified as described above using a PAGE gel. Binding reactions

were set up in 20 μl volumes containing 9 μl Buffer D, 1 μl poly dI-dC competitor DNA,

1 μl radiolabelled probe, and in some cases 1 μl MBF cell extract and/or 1 μl competitor.

Binding reactions were incubated at room temperature for 15 minutes, and then were

loaded onto a 5% polyacrylamide gel that had been pre-electrophoresed at 180V for 30

minutes in 0.5x TBE buffer. The electrophoresis was continued for a further 2 hours, at

which time the gel was fixed, dried, and exposed to film.

2.3.4 SBF/MBF affinity resins

Elutions of FLAG-purified SBF or MBF were dialyzed in buffer K (100 mM NaCl, 20

mM HEPES, 10% glycerol, 0.1 mM EDTA, 1 mM DTT) with stirring overnight at 4°C.

Affi-gel 15 (Bio rad) was washed once in cold ddH2O and then washed in buffer K before

addition of ~3 ml recombinant protein solution from pooled elutions. Approximately

1200 μg of SBF or MBF were combined with 1200 μl of Affi-gel 15 at 4°C to generate a

53

1 mg/ml affinity resin. Coupling was performed by incubation at 4°C for 3.5 hours on a

nutator mixer.

2.3.5 Affinity capture binding assays

TAP-tagged yeast strains were grown in YPD, harvested in mid-log phase by

centrifugation, and the resultant cell pellet was frozen before processing. Extracts were

prepared from frozen pellets by resuspending in lysis buffer E (100 mM HEPES-NaOH

pH 7.4, 1 mM EDTA, 10% glycerol, 10 mM MgCl2, 100 mM NaCl, 20 mM NaF, 50 mM

β-glycerophosphate, 1 mM DTT, 1 mM PMSF, and one CompleteTM

Mini-tab

(Boehringer) per 5 ml buffer). 1 ml lysis buffer E was added to 1 g cell pellet in a 14 ml

round-bottom polypropylene tube (Sarstedt), and 0.5 mm glass beads (BioSpec) were

added up to 2 mm below the meniscus. Samples were vortexed 5 x 1 min at 4°C with 1

min rests on crushed ice in between. A 21G 1½ needle (Becton Dickinson) was heated

on a flame before puncturing the bottom of the polystyrene tube and placing it into a

lidless 50 ml conical tube (Falcon) to allow the extract to flow out. Extracts were

standardized to a concentration of 30 μg/μl, after which 300 μl was added to 60 μl of 50%

slurry of each one of the three different versions of Affi-gel 15 resin: Affigel-SBF,

Affigel-alone, Affigel-MBF. Following incubation for 1 hour at 4°C on a nutator mixer,

samples were centrifuged at 500 x g at 4°C for 1 minute. Samples were washed three

times with 1 ml cold lysis buffer E, resuspended in loading buffer, incubated at 95°C for

2 min and then loaded on PAGE gels. Binding of the TAP-tagged protein to resins was

assessed by Western blot analysis using PAP (Sigma) antibody, which recognizes the

TAP tag.

54

2.3.6 ChIP analysis

Strains were grown in either YPD or minimal medium as appropriate, subjected to in vivo

cross-linking, then harvested for subsequent chromatin immunoprecipitation assays.

Quantitative real-time PCR was carried out by using a dual fluorogenic reporter TaqMan

assay in an ABI 7500 system. Target gene probes were labeled with 6-carboxyfluorescin

(FAM) and black hole quencher (BHQ), and an internal control probe for a

transcriptionally inert region of chromosome 2 was labeled with VIC and BHQ. Probes

and flanking primers were designed using primers against the SCB-containing upstream

regions of CLN2 and PCL1. Capture efficiency was calculated as the ratio of

immunoprecipitated versus total DNA for probe signal divided by the signal from the

transcriptionally inert region of chromosome 2.

2.3.7 Bck2-DB constructs, complementation and auto-activation assays

To construct the LexA DB-Bck2 fusion protein, BCK2 was cloned into the plasmid

vector pEG202. The LexA DB-BCK2 construct was co-transformed into WT yeast cells

with the reporter plasmid pSH18-34, in which the lacZ gene is driven by LexA operator

elements. Transformants were spotted at equivalent optical density onto plasmid

selective medium, incubated for 48 hr at 30°C, and subjected to β-galactosidase overlay

assays (Barral, Jentsch, and Mann 1995). LexA DB-BCK2 fusion constructs used in

transcription assays were tested for functionality by testing for complementation of the

growth defect of a cln3Δbck2Δ GAL-CLN3 strain (BY3015) when grown on glucose-

containing medium. Transformants were streaked onto YPGal to suspend plasmid

selection, then streaked onto medium containing glucose + 5’FOA and incubated for 48

55

hr at 30°C. To construct the Gal4 DB-Bck2 fusion protein, BCK2 was cloned into the

plasmid vector pDEST32 (LEU2) by GatewayTM

recombinational cloning. A yeast-two-

hybrid bait strain (Y8930) (where Gal4 UAS promoters drive expression of ADE2, HIS3,

and lacZ reporter genes) bearing either a Gal4 DB-BCK2 fusion plasmid or vector

(plasmid pDEST32) were spotted at equivalent optical density onto plasmid selection

medium. β-galactosidase overlay assays were performed as described (Barral, Jentsch,

and Mann 1995).

2.4 Results

2.4.1 Purification of recombinant SBF and MBF from insect cells

Similar to Cln3, the Swi4 protein is an unstable protein (Foster, Mikesell, and

Breeden 1993; Taylor et al. 2000). Although Swi6 can be purified from E. coli (Ho et al.

1999), attempts to produce Swi4 in bacteria have been met with limited success (Taba et

al. 1991; Sidorova and Breeden 1993). To circumvent this problem, the baculovirus-

mediated expression system (Jarvis 2009) was used to produce Swi4 in insect cells (Baetz

and Andrews 1999). Specifically, coexpression of Swi4- and Swi6-encoding

baculoviruses allows purification of in vivo reconstituted SBF and increases the stability

of Swi4 (Baetz and Andrews 1999). In the original study (Baetz and Andrews 1999),

SBF was partially purified by heparin affinity chromatography, which exploits the

property of a high-capacity cation exchanger to separate groups of proteins. The major

disadvantage of heparin affinity chromatography is its broad specificity, which means

that it is not possible to obtain a homogenous protein preparation in a single step.

56

In order to purify SBF and MBF in one step, Swi4 and Mbp1 were separately

tagged with FLAG (Hopp et al. 1988), but Swi6 was left untagged so to allow co-

purification with the tagged DNA-binding subunits. Because Swi4 is expressed at low

amounts in insect cells (Baetz and Andrews 1999), FLAG-tagged Swi4-encoding virus

was titrated against a constant amount of native Swi6-encoding virus, to determine the

optimal amount of virus solution to use for infecting insect cells (O. Schub; data not

shown). Because MBF had never been produced in insect cells, I performed a titration of

FLAG-Mbp1-encoding virus against a constant amount of Swi6-encoding virus (Figure

2-1A), as Mbp1 is also expressed at lower amounts than Swi6. Cultured insect cells

(Hi5) were co-infected with baculovirus encoding FLAG-tagged Mbp1 and baculovirus

encoding native, untagged Swi6. The amount of baculovirus encoding the native Swi6

subunit was kept constant, while the volume of baculovirus encoding the FLAG-tagged

Mbp1 subunit was progressively increased in order to optimize the desired 1:1

stochiometry of MBF. Using my protocol, and that previously established for SBF, I was

able to reconstitute highly purified SBF and MBF for use in in vitro assays (Figure 2-1B).

57

Figure 2-1. Purification of recombinant MBF and SBF from insect cells. (A) Cultured insect cells

(Hi5) were co-infected with two baculoviruses encoding separately (1) native, untagged Swi6, and (2)

FLAG-tagged Mbp1. The infection volume of baculovirus encoding the native Swi6 subunit was kept

constant, while the infection volume of baculovirus encoding the FLAG-tagged Mbp1 subunit was

progressively increased in order to optimize the desired 1:1 stochiometry. Infected cells were harvested,

crude lysate prepared and the MBF heterodimer was purified using an anti-FLAG resin in pulldown assays.

After several washing steps, purified MBF was eluted in sample buffer, boiled and loaded onto a

polyacrylamide gel and subjected to SDS-PAGE. Gels were subsequently stained with Coomassie dye for

visualization. (Heavy chain = the immunoglobulin, originally conjugated to the anti-FLAG resin, which

becomes solubilized in the elution buffer, PPM = Pre-stained Protein Marker). (B) Cultured insect cells

(Hi5) were co-infected to produce either MBF or SBF. To produce MBF, insect cells were co-infected with

two baculoviruses encoding separately (1) native, untagged Swi6, and (2) FLAG-tagged Mbp1. To

produce SBF, insect cells were co-infected with two baculoviruses encoding separately (1) native, untagged

Swi6, and (2) FLAG-tagged Swi4. Infected cells were harvested, crude lysate prepared and enriched by

anti-FLAG pulldown for purification of the heterodimer from the total protein lysate. After several

washing steps, purified SBF and MBF were eluted using FLAG-peptide, dialyzed, boiled and loaded onto a

polyacrylamide gel and subjected to SDS-PAGE. Gels were subsequently stained with Coomassie dye for

visualization. (PPM = Pre-stained Protein Marker, CE = Crude Extract, SUP = Supernatant of the CE after

the anti-FLAG pulldown step, E1, E2 = Elution 1, Elution 2).

Native Swi6 FLAG-Mbp1

Heavy Chain

175

83

62

48

kDa FLAG-Mbp1 baculovirus Native Swi6

FLAG-Swi4

Native Swi6 FLAG-Mbp1

175

83

62

48

kDa

PPM CE SUP CE SUP E1 E2 E1 E2

F-MBF F-SBF

A

B

PPM

58

2.4.2 Purified SBF and MBF bind DNA specifically in vitro

In order to ask if my preparations of insect cell-purified SBF and MBF retained

DNA-binding activity, I used an established electrophoretic mobility shift assay (EMSA)

to test whether SBF and MBF were able to bind DNA promoter fragments containing

either SCB or MCB elements in vitro (Baetz and Andrews 1999). My purified SBF

preparation showed specific binding to an HO probe containing SCB sites (Harrington

and Andrews 1996), but no binding activity was seen when a TMP1 (CDC21) probe

containing only MCB elements was used in the assay (Figure 2-2A). Conversely, my

purified MBF preparation bound specifically to the MCB sites in the TMP1 promoter

(Dirick et al. 1992) (Figure 2-2B). The interaction of FLAG-purified MBF with the

MCB-containing probe was specific because the MBF-DNA complex was effectively

competed by titration with unlabeled WT, but not mutant TMP1. I conclude that co-

expression of FLAG-SWI4 + untagged SWI6, as well as FLAG-MBP1 + untagged SWI6,

in insect cells can functionally reconstitute pure SBF and MBF, respectively.

59

Figure 2-2. DNA-binding by purified SBF and MBF. (A) Purified SBF was incubated with a

radiolabeled fragment of the HO promoter (lane 1; lane 2 is free probe), which contains SCB elements

known to bind SBF (left panel), or a radiolabeled fragment of the TMP1 promoter (lane 1; lane 2 is free

probe), which contains MCB elements known to bind MBF (right panel). Binding reactions were loaded

onto a native 5% polyacrylamide gel to visualize protein-DNA complexes. Arrows denote the location of

the SBF-HO probe complex or free probe (left panel), or free probe (right panel). (B) Purified MBF was

incubated with a radiolabeled fragment of the TMP1 promoter, which contains MCB elements (lanes 2, 8)

and an increasing concentration (125x, 250x, 500x, 1000x) of wild-type ʺcoldʺ competitor (lanes 3, 4, 5, 6)

or an increasing concentration (125x, 250x, 500x, 1000x) of mutant ʺcoldʺ competitor (lanes 9, 10, 11, 12).

Binding reactions were loaded onto a native 5% polyacrylamide gel in order to visualize protein-DNA

complexes retarded in the gel. Arrows denote the location of the MBF-TMP1 probe complex or free probe.

FLAG-MBF +

TMP1 Probe

Free Probe

WT “cold”

competitor

MUT “cold”

competitor

FLAG-SBF +

HO Probe

Free HO

Probe

Free TMP1

Probe

A

B

60

2.4.3 Novel binding partners of SBF and MBF – Elm1, Rtf1, Ykr077w, Bck2

Affinity chromatography has been successfully used for many years to discover

new binding partners for known transcriptional regulators (Greenblatt and Li 1981b,

1981a; Burton et al. 1988). To test whether candidate G1-phase transcriptional activators

physically interacted with SBF and MBF heterodimers, I devised an in vitro affinity

capture binding assay. I coupled purified SBF and MBF heterodimers to Affigel-15 resin

in order to generate two affinity resins. I then prepared crude extracts from yeast strains

expressing TAP-tagged derivatives of putative SBF/MBF-binding partners at their

endogenous loci (Rigaut et al. 1999; Ghaemmaghami et al. 2003). The extracts were

incubated with SBF and MBF affinity resins, and eluates were probed with an antibody

that specifically recognizes the TAP epitope on Western blots. Known SBF and MBF

binding proteins, Stb1 (Ho et al. 1999) and Rad53 (Sidorova and Breeden 1997), bound

my affinity resins but not the resin alone (Figure 2-3), confirming the utility of the assay.

To choose candidate proteins to test for SBF or MBF binding, I used a list of

genes identified in a genome-wide screen for regulators of SCB or MCB reporter gene

expression. Several candidate proteins tested, including a reported Swi6-interacting

protein Yol131w (Ito et al. 2001), either bound non-specifically to the resin, probably

because it is positively charged, or were undetectable by Western blot analysis.

However, four of the seventeen proteins tested interacted with SBF and MBF: Elm1,

Rtf1, Ykr077w, and Bck2. Elm1 functions at the G2/M phase transition by inhibiting

Swe1 (Sreenivasan and Kellogg 1999), which is an inhibitor of Cdc28 (Harvey et al.

2005). Inhibition of Elm1 kinase activity in a strain lacking CLN1 and CLN2 is lethal

61

Figure 2-3. Affinity capture analysis using SBF/MBF affinity resins. (A) SBF and MBF were

reconstituted in insect cells, purified by anti-FLAG pulldown assays, eluted with FLAG peptide, dialyzed

and covalently coupled to an Affigel matrix. Crude extracts were prepared from yeast strains that had the

endogenous copy of a given candidate gene TAP-tagged at its C-terminus and incubated with SBF or MBF

affinity resins. Binding was determined by Western blot using a PAP (Peroxidase Anti-Peroxidase)

antibody directed against the TAP-tag. High density bands in the Western autoradiograms were called

“+++”, whereas an absence of any bands received the designation “-”. Intermediate intensity bands in the

autoradiogram are denoted by “++” or “+”. (B) Western autoradiograms of positive interactions in the

affinity capture assay. Rad53 and Stb1 are positive controls. Rad53 has been reported to physically

interact with the components of SBF (Swi4, Swi6) and MBF (Mbp1, Swi6), and Stb1 interacts with Swi6

(the common regulatory subunit of SBF and MBF).

A B

SBF Resin MBF

Stb1

Elm1

Rtf1

Ykr077w

Bck2

Rad53

62

(Sreenivasan et al. 2003; Zou et al. 2009), indicating that Elm1 and Cln3 could have

redundant functions. My in vitro assay suggests that a G1 phase function for Elm1 may

be partially mediated through binding to G1 transcription factors. I also identified Rtf1 as

a protein that interacts with SBF and MBF in vitro. Rtf1 is a component of the

Paf1/RNA polymerase II complex which contains Paf1, Ctr9, Cdc73, Leo1, and Rtf1

(Betz et al. 2002; Mueller and Jaehning 2002; Krogan et al. 2002). Deletion of Paf1

complex components affects transcript levels of SBF/MBF target genes (Betz et al.

2002). I also tested the Rtf1-binding proteins Swd1 and Swd3 (Krogan et al. 2003) for

interaction with SBF and MBF affinity resins (Figure 2-3), but the results were

inconclusive.

Finally, I discovered two proteins, Msa2 (Ykr077w) and Bck2, that bound SBF

and MBF resins and had been previously linked to G1 transcription (Epstein and Cross

1994; Di Como, Chang, and Arndt 1995; Wijnen and Futcher 1999; Ashe et al. 2008).

Overproduction of MSA2 increases mating pheromone resistance, rescues the

temperature-sensitivity of a cdc28-4 strain at the restrictive temperature, and partially

suppresses the lethality of a cln1Δcln2Δcln3Δ strain by restoring G1 transcription (M.

Costanzo; unpublished). Although the mechanism by which Msa2 regulates SBF and

MBF remains unclear, I chose to focus on the interaction that I discovered between Bck2

and SBF/MBF. Because of the large body of evidence suggesting that BCK2 is important

for G1 transcription, I decided explore the mechanism by which Bck2 activates G1

transcription (Epstein and Cross 1994; Di Como, Chang, and Arndt 1995; Wijnen and

Futcher 1999; Munoz et al. 2003; Ferrezuelo, Aldea, and Futcher 2009).

63

2.4.4 Localization of Bck2 to promoters of genes transcribed at the G1/S transition

The physical interaction of Bck2 with SBF and MBF affinity resins suggested that

Bck2 might function near promoter DNA. SBF and MBF localize to the CLN2 and PCL1

promoters (Iyer et al. 2001; de Bruin et al. 2006; Ashe et al. 2008; Takahata, Yu, and

Stillman 2009) and activate their transcription (Espinoza et al. 1994; Dirick, Bohm, and

Nasmyth 1995; Baetz et al. 2001). As noted earlier, the BCK2 gene is also an activator of

CLN2 (Di Como, Chang, and Arndt 1995) and PCL1 (Wijnen and Futcher 1999)

transcription, and at least one BCK2-responsive element lies within a CLN2 promoter

region that contains two SCB-like sequences and one intact MCB-like sequence (Di

Como, Chang, and Arndt 1995). However, association of Bck2 with promoter regions

had not been previously reported.

I tested whether a TAP-tagged version of the Bck2 protein could be detected at

CLN2 and PCL1 promoters using a chromatin immunoprecipitation (ChIP) assay (Hecht

and Grunstein 1999). In this strain, BCK2-TAP was expressed from its chromosomal

locus under its own promoter. Bck2 is a very low abundance protein and cannot be

detected by Western blotting in extracts from asynchronous cells (Ghaemmaghami et al.

2003) (Figure 2-3A). Nevertheless, Bck2-TAP immunoprecipitates from asynchronous

cultures were enriched for CLN2 and PCL1 promoter fragments 2-fold above

background, whereas the same promoter fragments were enriched more than 10-fold

above background in Swi4 immunoprecipitates (Figure 2-4A). I next attempted to

enhance the Bck2 ChIP signal by using a strain carrying a plasmid-borne inducible copy

of tagged BCK2 in my ChIP experiment. Under inducing conditions, I observed 3-fold

enrichment of CLN2 promoter DNA fragments relative to a vector control (Figure 2-4B).

64

Figure 2-4. Localization of Bck2 to the promoters of G1/S transcribed genes. (A) WT (BY4741;

untagged), BCK2-TAP and SWI4-TAP strains were grown to mid-log phase, then subjected to in vivo cross-

linking for subsequent chromatin immunoprecipitation assays. Enrichment of CLN2 or PCL1 promoter

DNA was determined by Q-PCR using primers against the SCB-containing upstream regions of CLN2 and

PCL1. ChIP efficiency (Y-axis) measures enrichment of promoter DNA for the target gene indicated

relative to enrichment of non-promoter DNA from an untranscribed region of yeast chromosome II. (B) A

WT (BY4741) strain was transformed with pGAL-BCK2-FLAG or pGAL-FLAG (empty vector). Final

transformants were grown to saturation in plasmid selective medium, subcultured in YPRaff or YPGal to

mid-log phase, subjected to in vivo cross-linking for subsequent chromatin immunoprecipitation assays.

Enrichment of CLN2 promoter DNA was determined by Q-PCR using primers against the SCB-containing

upstream region of CLN2. ChIP efficiency (Y-axis) was measured as described in (A). Error bars reflect

standard deviations from multiple Q-PCR runs from the same experiment.

65

These results suggest that Bck2 may be a part of SBF and MBF complexes that form on

the promoters of G1/S transcribed genes. However, it is possible that Bck2 may localize

to G1 promoters by interacting with other proteins or with non-SCB/MCB regions.

66

2.4.5 Bck2 DB fusion proteins can activate transcription when tethered to DNA

Bck2 is a potent transcriptional activator (Titz et al. 2006), which may be linked

to its G1-specific role. Thus, I asked if Bck2 could activate transcription when tethered to

promoters via the LexA DNA binding domain. I introduced a plasmid expressing a

LexA-Bck2 fusion protein into a strain carrying a reporter plasmid with LexA operator

sequence driving the expression of lacZ. The LexA DB-Bck2 fusion protein effectively

activated transcription of the reporter gene (Figure 2-5). The LexA DB-BCK2 fusion

construct used in this assay was functional because it could complement the inviability

phenotype of a cln3Δbck2Δ GAL-CLN3 (URA3) strain (Figure 2-5). In order to

substantiate the finding that Bck2 could activate transcription in a promoter-bound

context, I also constructed a Gal4 DB-Bck2 fusion which also activated transcription of a

cognate reporter gene (Figure 2-5). This construct was also functional because it

complemented the inviability phenotype of a cln3Δbck2Δ GAL-CLN3 (URA3) strain (data

not shown). The finding that Bck2 is a promoter-proximal activator of late G1 phase

transcription is consistent with: (1) the genetic phenotypes associated with mutation of

BCK2; (2) the physical interaction of Bck2 with SBF/MBF; (3) the localization of Bck2

to the CLN2 and PCL1 promoters.

67

Figure 2-5. Bck2 auto-activates transcription when fused to a DNA-binding domain. (A) Yeast cells

(BY1623) were co-transformed with a reporter plasmid where the lacZ gene is driven by LexA operator

(LexAOp) elements (pSH18-34 = LexAOp-lacZ, URA3), in addition to either LexA DB-BCK2 (plasmid

pEG202, HIS3) or LexA DB (plasmid pEG202, HIS3). Four independent isolates of each were spotted at

equivalent optical density onto plasmid selective medium, incubated for 48 hr at 30°C, overlaid with a top

agar solution containing X-Gal, and incubated further at 30°C until blue color was seen (β-galactosidase

overlay assay). (B) LexA DB-BCK2 fusion constructs used in transcription assays are functional as

determined by complementation of a cln3Δbck2Δ GAL-CLN3 (URA3) strain (BY3015) (sections 1-3 in

black). Three independent transformants of LexA DB-BCK2 (plasmid pEG202, HIS3) and one

transformant of LexA vector (plasmid pEG202, HIS3) in strain cln3Δbck2Δ GAL-CLN3 (URA3) were

streaked onto YPGal to temporarily suspend plasmid selection, then streaked onto medium containing

glucose + 5’FOA, and incubated for 48 hr at 30°C. A ura3Δ strain (BY1623) was used as a control

(section 5 in red). (C) Four isolates of a yeast-two-hybrid bait strain (Y8930) (where Gal4 UAS elements

drive expression of ADE2, HIS3, and lacZ reporter genes) bearing either a Gal4 DB-BCK2 fusion (LEU2)

plasmid (within plasmid pDEST32), or vector (plasmid pDEST32) were spotted at equivalent optical

density onto: (i) plasmid selective medium that would subsequently be assayed for lacZ transcription using

the β-galactosidase overlay assay (shown completed); (ii) medium lacking histidine to visualize growth

proportional to transcription from the HIS3 gene; or (iii) plasmid selection medium to visualize

development of red pigment indicative of defects in ADE2 transcription.

+ LexA

DB-BCK2

+ LexA DB

+ LexA DB-BCK2

SD – U – H + X-Gal

SC + 5-FOA

1 2

3

+ LexA DB

WT

+ Gal4 DB

+ Gal4 DB-

BCK2

+ Gal4 DB

+ Gal4 DB-

BCK2

+ Gal4 DB

+ Gal4 DB-

BCK2

SD – H

SD – L

SD – L + X-Gal

1 2 3

C A

B

WT + LexAOp-lacZ

4

5

cln3Δbck2Δ GAL-CLN3 (URA3)

(i)

(ii)

(iii)

68

2.5 Discussion

2.5.1 SBF and MBF affinity resins for high-throughput screening of G1-specific

regulators

In this Chapter, I describe the use of an affinity capture assay to identify SBF- and

MBF-binding proteins. The use of heterodimeric protein affinity resins for detecting

protein-protein interactions is advantageous for several reasons. First, transient

interactions can often be better identified using affinity resins rather than co-purification

(Formosa et al. 1991), because many protein complexes are too fragile to allow for co-

purification of the interacting species from cellular extracts. Such relatively weak

binding is to be expected, since components that must bind to one another reversibly in

the concentrated environment of the cell may fall apart in a dilute extract (Formosa et al.

1991). Both the Clb6-Swi6 (Geymonat et al. 2004) and Hrr25-Swi6 (Ho et al. 1997)

physical interactions, for instance, are biologically relevant but were discovered using

purified proteins, rather than co-purification from cell extracts. Second, some

interactions require heterodimeric SBF and MBF, rather than their components. For

example, for Cln3 to bind SBF, both Swi6 and Swi4 have to be present (Wang et al.

2009). Both Whi5 (Costanzo et al. 2004; de Bruin et al. 2004) and Nrm1 (de Bruin et al.

2006) also require the heterodimer forms of SBF and MBF, respectively.

I designed my affinity capture strategy to allow rapid screening of proteins

identified in genetic or other screens as potential regulators of G1-specific transcription. I

chose this approach since the biochemical test alone cannot be used to infer that an

interaction occurs in vivo and whether the interaction is direct and involves transcription

activation. For example, although Kap120 (Kim et al. 2010), Clb6 (Basco, Segal, and

Reed 1995; Geymonat et al. 2004), and Msn5 (Queralt and Igual 2003) all physically

69

associate with Swi6 in a biologically meaningful manner, the effect on SBF/MBF

dependent transcription is indirect because these proteins control the localization of Swi6

at a time unrelated to the activation of promoter-bound SBF, which occurs in late G1

phase. Another example of indirect activation of SBF is in cell wall integrity (CWI)

signalling, wherein the signal is transduced through sequential activation of Pkc1, Bck1,

Mkk1, Mkk2 and Slt2, leading to the association of SBF with Slt2 (Madden et al. 1997)

on the FKS2 promoter (Kim, Truman, and Levin 2008). Slt2 is the only member of this

pathway known to directly interact with SBF. However, overexpression of MKK1

activates FKS2 expression (Zhao et al. 1998), presumably through indirect activation of

SBF. Thus, the distinction between a genetic activator and a physical activator must be

considered when using biochemical results to build models about G1 regulation.

2.5.2 Identification of known SBF and MBF transcriptional regulators

The specificity of SBF and MBF affinity resins was established by determining

whether known SBF and MBF binding proteins could be detected. I detected three

known SBF/MBF binding proteins in my assay: Stb1, Rad53, and Msa2. Stb1 was first

identified as a Swi6-binding protein in an affinity chromatography assay (Ho et al. 1999;

Costanzo, Schub, and Andrews 2003) and was subsequently shown to interact with both

SBF and MBF to regulate G1 transcription (de Bruin, Kalashnikova, and Wittenberg

2008). Rad53 phosphorylates recombinant Swi6 in vitro (Sidorova and Breeden 1997)

and in vivo (Sidorova and Breeden 2003b). Moreover, Swi6, Mbp1, Swi4, and Whi5

interacted with Rad53 in a large-scale proteomic study (Smolka et al. 2006), suggesting

the interaction of Rad53 with Swi6 occurs in the context of SBF and MBF. Msa2 was

70

identified as an interactor of Swi4 (Ashe et al. 2008) and Swi6 (Krogan et al. 2006).

Consistent with a role in transcriptional activation, Msa2 localizes to the nucleus (Huh et

al. 2003) and was identified in a screen for strong transcriptional activators in a yeast

two-hybrid experiment (Titz et al. 2006). In our lab, we found that overproduction of

MSA2 increased mating pheromone resistance, rescued the temperature-sensitivity of a

cdc28-4 strain at the restrictive temperature, and suppressed the lethality of a

cln1Δcln2Δcln3Δ strain by restoring G1 transcription (M. Costanzo; unpublished). The

physical interaction of Msa2 with SBF and MBF (Figure 2-3) is consistent with these

genetic phenotypes. The Msa2 homolog, Yor066w (Msa1), shares 28% amino acid

identity and 43% similarity with Msa2. Like Msa2, Msa1 physically interacts with SBF

and MBF in a biologically relevant way (Ashe et al. 2008).

2.5.3 Identification of novel SBF and MBF transcriptional regulators

In addition to proteins known to affect SBF and MBF transcription, I identified

novel proteins for which a significant role in G1/S-specific transcription was not

suspected. I identified two proteins that may bind SBF and MBF in vivo -- Elm1 and

Rtf1. Elm1 is a serine/threonine protein kinase that regulates cellular morphogenesis,

septin behaviour, and cytokinesis (Garrett 1997; Koehler and Myers 1997; Bouquin et al.

2000). Elm1 was originally identified by a mutation that causes elongated cell

morphology and affects pseudohyphal development (Blacketer et al. 1993). Elm1 has

roles in the control of bud growth and cytokinesis (Sreenivasan and Kellogg 1999;

Bouquin et al. 2000). Elm1 regulates the activity of the Kin4 checkpoint kinase during

activation of the spindle position checkpoint (Moore et al. 2010; Caydasi et al. 2010).

71

Thus, an important function of Elm1 is the coordination of spindle positioning with cell

cycle progression. Elm1 may also regulate the activity of Cln3/Cdc28 complexes,

because inhibition of Elm1 kinase activity causes a delay in CLN2 transcription, and loss

of Elm1 function is lethal in a cln1Δcln2Δ background (Sreenivasan et al. 2003; Zou et

al. 2009). Elm1 has also been implicated in cellular energy balance (Hong et al. 2003;

Sutherland et al. 2003) through interaction with Snf1 kinase in vivo and in vitro (Hardie,

Carling, and Carlson 1998; Kemp et al. 2003). Overproduction of hyperactive Elm1

derivatives produces phenotypes, such as pseudohyphal invasive growth, that all require

the function of Snf1 (Sutherland et al. 2003). Given that Snf1 physically interacts with

Swi6 (Pessina et al. 2010), it is possible that Elm1 interacts with SBF/MBF through

regulation of Snf1 activity.

The physical interaction of Rtf1 with SBF/MBF is another novel interaction that

may indicate how transcription complexes form following SBF/MBF-binding (Figure 2-

3). Rtf1 is a subunit of the RNAPII-associated Paf1 complex that also includes Paf1,

Cdc73, Leo1 and Ctr9 (Mueller and Jaehning 2002). CTR9 was isolated as a gene that is

required in the absence of CLN3 (Di Como, Chang, and Arndt 1995). Consistent with a

shared role with CLN3 in G1 transcriptional activation, mutation of CTR9 causes

defective CLN2 transcription (Koch et al. 1999). There are a number of other intriguing

genetic connections between Rtf1 and late G1 phase transcription. Loss of Paf1 disrupts

transcription of cell wall biosynthetic genes controlled by the PKC pathway (Chang et al.

1999), and paf1Δ mutants are sensitive to cell wall-damaging agents such as sodium

dodecyl sulphate and caffeine (Shi et al. 1996; Chang et al. 1999). Nearly 30% of genes

whose expression decreases in a paf1Δ strain are regulated during the cell cycle (Porter et

72

al. 2002), including many SBF- and MBF-regulated genes. For example, expression of

CLN1, RNR1 and HO are diminished in a paf1Δ strain (Porter et al. 2002; Mueller and

Jaehning 2002). Genetically, elimination of Paf1 in strains with already compromised G1

transcription, like a swi4Δswi6Δ strain, paf1Δswi6Δ (Porter et al. 2002), paf1Δcln3Δ and

ctr9Δcln3Δ (Koch et al. 1999) strains, results in lethality. Finally, overexpression of

MBP1 or SWI4 suppresses the sensitivity to HU of a paf1 mutant (Porter et al. 2002).

Rtf1 is important for TATA site selection during transcription initiation (Stolinski,

Eisenmann, and Arndt 1997), which suggests that Rtf1 could affect start-site selection for

SBF/MBF-regulated genes. In a swi6Δ strain, the normal length RNR1 RNA is reduced

in abundance, while a longer RNR1 RNA is present (Di Como, Chang, and Arndt 1995).

In a swi4Δ strain, the pattern of CLN2 mRNA start site selection is also altered (Stuart

and Wittenberg 1994). Taken together, these findings suggest that Rtf1 may physically

link SBF/MBF to the Paf1 complex to allow the general transcriptional apparatus to

initiate G1/S-specific transcription from appropriate transcription start sites.

73

Chapter 3

Dissection of Bck2 Protein Domains and

Identification of Bck2 Binding Partners using a

Yeast Two-Hybrid Screen

74

Chapter 3: Dissection of Bck2 protein domains and identification of Bck2 binding

partners using a yeast two-hybrid screen

3.1 Abstract

Full-length Bck2 protein fused to the Gal4 DNA-binding domain (DBD) activates

transcription of a cognate reporter gene, a phenomenon called Yeast 2-hybrid (Y2H)

auto-activation. In some cases, auto-activation reflects an important in vivo function of

the protein in the recruitment of RNAPII to promoters. For other proteins, such as Cln3,

the auto-activation phenotype does not appear central to protein function. The auto-

activation phenotype is common for transcription factors such as Bck2 and makes it

impossible to screen for binding proteins using the Y2H system. I sought to use the Y2H

system to expand our view of Bck2-binding proteins and generated 20 fusions of

fragments of the BCK2 gene to the Gal4 DBD. I discovered that the C-terminal third of

Bck2 contained the Y2H-auto-activation activity and that this region was not required for

in vivo function. I then used the largest Bck2 fragment lacking auto-activation but

retaining in vivo function as a bait protein in a Y2H screen. I discovered six novel

binding partners for Bck2: Mcm1, Mot3, Yap6, Tpd3, Std1, and Mth1.

3.2 Introduction

Strictly speaking, a transcriptional activator is composed of a DNA-binding

domain (DBD) and an activation domain (AD). The DBD portion targets the whole

protein to a specific binding site in the promoter of a gene and the AD portion mediates

transcription initiation through recruitment of basal transcriptional machinery (Kadonaga

2004). This type of protein is a physiological transcriptional activator. While DBDs are

75

functionally and structurally well characterized (Kadonaga 2004), ADs in general do not

share easily recognizable motifs or structures (Triezenberg 1995), and there is no

consensus on what constitutes a typical AD (Giniger and Ptashne 1987; Sadowski et al.

1988; Courey et al. 1989; Mermod et al. 1989; Minter, Brennan, and Mapp 2004).

A physiological transcriptional activator, however, differs from a Y2H auto-

activator, which activates transcription on promoters in artificial systems. A Y2H auto-

activator is tethered to promoter DNA because it is fused to the DBD of Gal4. Auto-

activation may reflect the physico-chemical properties of protein fusion which enable

recruitment of RNAPII to the reporter gene promoter. For example, random fusion

proteins derived from fragments of the E. coli genome (Ruden et al. 1991), small

synthetic peptides (Seipel, Georgiev, and Schaffner 1994), and small non-protein

molecules (Minter, Brennan, and Mapp 2004) can auto-activate transcription, suggesting

that auto-activation alone is a poor determinant of whether a protein is a physiological

transcriptional activator. Also, the ability to auto-activate in the Y2H system is estimated

to be present in approximately 5% of all proteins (Van Criekinge and Beyaert 1999) yet

may be dispensable for the normal function of the protein. For example, although Cln3

has been reported to be a strong transcriptional activator, the Y2H auto-activation activity

of Cln3 is dispensable for its normal cell cycle function (Wijnen, Landman, and Futcher

2002).

Nevertheless, a Y2H auto-activator is more likely than a random protein to be a

physiological transcriptional activator. Of 451 budding yeast Y2H auto-activators tested,

20% have the GO annotation (Ashburner et al. 2000) ‘transcriptional activator’, in

comparison to Y2H non-activators (~3%) (Titz et al. 2006). Y2H auto-activators are also

76

more likely to be nuclear localized (41%) relative to Y2H non-activators (22%) (Titz et

al. 2006). Furthermore, analyses of the properties of Y2H auto-activators shows that they

possess a lower isoelectric point, lower hydrophobicity, higher molecular weight,

enrichment of asparagines clusters, and tend to specifically interact with components of

the transcription machinery (Titz et al. 2006).

The BCK2 gene encodes a 2,553-bp open reading frame corresponding to a

polypeptide with a predicted length of 851 amino acids and a calculated molecular size of

94 kDa (Lee, Hines, and Levin 1993). The predicted BCK2-encoded protein is rich in

serine and threonine residues (24% Ser plus Thr) but it is not closely related to any

known protein based on primary sequence similarity (Lee, Hines, and Levin 1993). The

N-terminal third of the protein is probably not essential for biological function, because

these sequences were absent from the original clones of BCK2 that suppressed the lysis

phenotype of a mpk1Δ strain (Lee, Hines, and Levin 1993). Bck2 is post-translationally

modified, as at least four sites are phosphorylated in vivo: Ser317, Ser334, Ser 757, and

Ser 761 (Chi et al. 2007; Albuquerque et al. 2008). The significance of these potential

post-translational modifications is not known, but likely reflects some control over Bck2

activity or localization.

Several properties of Bck2 suggest that it may be a physiological transcriptional

activator. Bck2 is (1) low abundance (Ghaemmaghami et al. 2003); (2) partly nuclear

(Huh et al. 2003); (3) high molecular weight (Lee, Hines, and Levin 1993); and (4) quite

acidic [pI 4.5] in the C-terminal region (529 to 851). All of these properties are found in

transcriptional activators (Ma and Ptashne 1987; Sadowski et al. 1988; Ptashne and Gann

1990; Titz et al. 2006). The significance of the strong Y2H auto-activation by Bck2

77

(Chapter 2) (Titz et al. 2006) to in vivo function is not known. In this chapter, I show that

Bck2 auto-activation is not central for in vivo function because this activity is not

required to sustain growth. Using the remaining N-terminal two-thirds of the protein as a

bait in the Y2H system, I identified six novel Bck2-binding proteins: Mcm1, Mot3, Yap6,

Tpd3, Std1, and Mth1. I discuss how well these interactions fit into our current

understanding of how the G1/S transition is coordinated with environmental sensing.

3.3 Materials and Methods

3.3.1 Yeast strains and plasmids

Yeast strains (Table 3-1) and plasmids (Table 3-2) were constructed using standard

methods.

Table 3-1

S. cerevisiae strains

Strainsa BY# Relevant genotype Source

BY4741 1623 MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 (Brachmann et al. 1998)

BY3015 3015 MATa cln3Δ::HphR bck2Δ::NatR his3Δ1 leu2Δ0 This study

met15Δ0 ura3Δ0 lys2Δ0 pGAL-CLN3 (URA3)

Y8930 4889 MATα trp1-901 leu2-3,112 his3-200 ura3-52 (Yu et al. 2008)

gal4Δ gal80Δ LYS2::GAL1-HIS3 GAL2-ADE2

met2::GAL7-lacZ cyhR

Y8800 4890 MATa trp1-901 leu2-3,112 his3-200 ura3-52 (Yu et al. 2008)

gal4Δ gal80Δ LYS2::GAL1-HIS3 GAL2-ADE2

met2::GAL7-lacZ cyhR

T487 4891 containing plasmids EV (DB) x EV (AD) (Vidal et al. 1996)

T488 4892 containing plasmids Rb (DB) x E2FΔF1 (AD) (Vidal et al. 1996)

T489 4893 containing plasmids c-fos (DB) x c-jun (AD) (Vidal et al. 1996)

T490 4894 containing plasmids Gal4 (DB) x Gal4 (AD) (Vidal et al. 1996)

T491 4895 DP1 (DB) x E2F (AD) (Vidal et al. 1996) a Strain backgrounds are isogenic to S288C

Table 3-2

Plasmids

Plasmids BA# Description Source

pDONR201 346V Gateway donor vector Invitrogen

RPL39-lacZ 67V RPL39-lacZ This study

CYC1-lacZ 347V CYC1-lacZ (Guarente and Mason 1983)

pDEST DB 2415 LEU2 CEN ADH1pro-Gal4 DB-ORF

(Gateway equivalent of pPC97)

78

pDEST DB-Gal4-BCK2 2416 LEU2 CEN ADH1pro-Gal4 DB-BCK2 (F1) This study

pDEST DB-Gal4-BCK2 2417 BCK2 aa81-851 (Fragment 2) This study

pDEST DB-Gal4-BCK2 2418 BCK2 aa178-851 (Fragment 3) This study

pDEST DB-Gal4-BCK2 2419 BCK2 aa250-851 (Fragment 4) This study

pDEST DB-Gal4-BCK2 2420 BCK2 aa529-851 (Fragment 5) This study

pDEST DB-Gal4-BCK2 2421 BCK2 aa1-766 (Fragment 6) This study

pDEST DB-Gal4-BCK2 2422 BCK2 aa81-766 (Fragment 7) This study

pDEST DB-Gal4-BCK2 2423 BCK2 aa178-766 (Fragment 8) This study

pDEST DB-Gal4-BCK2 2424 BCK2 aa250-766 (Fragment 9) This study

pDEST DB-Gal4-BCK2 2425 BCK2 aa529-766 (Fragment 10) This study

pDEST DB-Gal4-BCK2 2426 BCK2 aa1-662 (Fragment 11) This study

pDEST DB-Gal4-BCK2 2427 BCK2 aa81-662 (Fragment 12) This study

pDEST DB-Gal4-BCK2 2428 BCK2 aa178-662 (Fragment 13) This study

pDEST DB-Gal4-BCK2 2429 BCK2 aa250-662 (Fragment 14) This study

pDEST DB-Gal4-BCK2 2430 BCK2 aa529-662 (Fragment 15) This study

pDEST DB-Gal4-BCK2 2431 BCK2 aa1-610 (Fragment 16) This study

pDEST DB-Gal4-BCK2 2432 BCK2 aa81-610 (Fragment 17) This study

pDEST DB-Gal4-BCK2 2433 BCK2 aa178-610 (Fragment 18) This study

pDEST DB-Gal4-BCK2 2434 BCK2 aa250-610 (Fragment 19) This study

pDEST DB-Gal4-BCK2 2435 BCK2 aa529-610 (Fragment 20) This study

3.3.2 Cloning and construction of BCK2 truncations

Fragments of the BCK2 gene were amplified from genomic DNA by PCR using primers

complementary to the BCK2 gene at various positions. PCR primers were designed to be

compatible with the Gateway system of recombinational cloning (Table 3-3). The hybrid

primers contained sequences homologous to BCK2 in continuity with sequences

containing attB sites, which are required for the in vitro recombination reaction. All PCR

products were recombined into the donor vector pDONR201 using the BP clonase II

system and transformed into DH5α E. coli cells. Plasmid DNA was isolated and

diagnosed by restriction digest using BsrGI to liberate the BCK2 insert. Positive clones

were fully sequence-confirmed. BCK2 truncations within pDONR201 were recombined

into the destination vector pDEST-DB using the LR clonase II system and transformed

into DH5α E. coli cells. Plasmid DNA was isolated and diagnosed by restriction

digestion using BsrGI to liberate the BCK2 insert.

79

Table 3-3

Oligonucleotides

Primera Sequence 5’ to 3’

BCK2F-Start GGGGACAAGTTTGTACAAAAAAGCAGGCTTAATGCCGAAGAATAGTCACCACCA

BCK2 aa81F GGGGACAAGTTTGTACAAAAAAGCAGGCTTAATGACTAAGGCGAAGAAGAGTAGTAGA

BCK2 aa178F GGGGACAAGTTTGTACAAAAAAGCAGGCTTAATGGACGCCTCGTCACTAACAACCAAA

BCK2 aa250F GGGGACAAGTTTGTACAAAAAAGCAGGCTTAATGTTCACTGAAAGCGAAACAAATTCT

BCK2 aa529F GGGGACAAGTTTGTACAAAAAAGCAGGCTTAATGTCAACTCCAAATGTGCTTGAAACAC

BCK2 aa610R noTAG GGGGACCACTTTGTACAAGAAAGCTGGGTCTTAATTCGGAATATCTTGTACTAGGAC

BCK2 aa662R noTAG GGGGACCACTTTGTACAAGAAAGCTGGGTCTTAGTTGCTTGTTGTAGTGGATATTGAGTT

BCK2 aa766R noTAG GGGGACCACTTTGTACAAGAAAGCTGGGTCTTAGTTGATGGCGCTGTTGCTGTTG

BCK2R-Stop noTAG GGGGACCACTTTGTACAAGAAAGCTGGGTCTTAGTTGCTATTATCAAAATAAAAAGACTG a Primers are shown 5’ to 3’

3.3.3 β-Galactosidase assays

Quantitative β-Galactosidase assays were performed as follows. Exponentially growing

cells at an optical density at 600 nm (OD600) of 0.2 to 0.25 were harvested. Extracts were

prepared by vortexing the cells in 1 ml Z-buffer (0.1 M NaPO4 [pH 7.0], 0.01 M KCl, 1

mM MgSO4, 4 mM 2-mercaptoethanol) + 20 μl 0.1% SDS + 40 μl chloroform and then

vortexing them for 45 seconds. After 5 min, 0.2 ml of o-nitrophenylgalactoside (Sigma

N-1127; at 4 mg/ml in Z buffer) solution was added and incubated at 30°C until a slight

yellowing was observed in the samples. The reactions were stopped by the addition of 0.5

ml of 1 M Na2CO3 at that time, and the samples were centrifuged for 3 minutes at 13,000

x g. The A420 of the supernatant was determined. β-Galactosidase units were calculated

using the following formula: units = (A420)*(1000) / (time of reaction, in

minutes)*(volume of extract in assay = 1 ml)*(cellular concentration in OD600 values).

For each time point, the assays were performed on three separate cultures, and the

average is reported. Qualitative β-galactosidase overlay assays were performed as

described (Barral, Jentsch, and Mann 1995).

80

3.3.4 Complementation analysis

Transformants of a cln3Δbck2Δ pGAL-CLN3 (URA3) strain (BY3015) bearing ADH1-

GAL4 DBD-BCK2 (1-20) plasmids (LEU2) were first grown in plasmid selective medium

followed by growth in YPGal, then prepared at equivalent optical density and spotted in

serial 10-fold dilutions onto plasmid selective medium containing either galactose –

5’FOA or glucose + 5’FOA, and incubated for 48 hr at 30°C.

3.3.5 Genome-wide Y2H screen

The Y2H ORFeome method (Xin et al. 2009) was used to screen for Bck2-interacting

proteins by mating a prey strain with a bait strain. The AD ORFeome (prey strain) is a

pooled collection of AD-ORF plasmids (Gelperin et al. 2005; Yu et al. 2008) in a 96-well

format. The single 96-well plate covers the ORFeome twice. There are ~200 AD-ORF

plasmids per well, and the bottom 1/3 of the plate is redundant with the top 2/3. A Y2H

pooled AD ORFeome (Gelperin et al. 2005; Yu et al. 2008) was grown in a 96-well

culture block containing 600 μl per well of SD – Trp media at 30°C for 48 hours. Five μl

of AD ORFeome culture were spotted from the 96-well culture block onto large YPD

plates and allowed to dry. Next, 5 μl of a DBD ORF strain culture (bait strain

transformed with a single DBD-ORF plasmid) were dispensed directly onto the

ORFeome strain spots. Five diploid control strains were spotted onto the same plate at an

empty location in order to ensure the quality of selection plates and to help evaluate the

phenotype of interactions. The monoplate was incubated at 30°C for more than 1 day

before replica plating onto a monoplate containing SD – Leu – Trp media and grown for

3 days at 30°C in order to select only diploid yeast. This monoplate was replica plated

81

onto a final selection plate containing SD – Leu – Trp – Ade and grown for 5 days until

foci were observed. At least 3 foci per spot were picked, and subjected to colony PCR

using primers homologous to the sequences flanking the ORF within the AD plasmid

(AD-F GTGTGTCGCGTTTGGAATCACTACAGGG; AD-R

GTGTGTGGAGACTTGACCAAACCTCTGGCG). Yeast cells were scraped from

plates into 30 μl of lysis solution (2.5 mg zymolyase in 1 ml 1 M sorbitol) and incubated

at 37°C for 15 minutes, then 95°C for 5 minutes, before addition of 120 μl of ddH2O.

PCR was performed in 25 μl volumes containing 5 μl of the yeast cell preparation

described. PCR was performed using 5 minute extension times in order to ensure that

large ORF inserts were isolated. PCR products were electrophoresed on agarose gels,

purified using the PureLink kit (invitrogen) and sent for sequencing analysis using

specific primers (XF075_AD_5' CGCGTTTGGAATCACTACAGGG and

XF076_Term_3' GGAGACTTGACCAAACCTCTGGCG). Sequences were processed

using Chromas Lite v. 2.01 and uploaded in FASTA format for subsequent BLAST

analysis in order to identify the relevant ORF.

3.3.6 Direct pair-wise Y2H assays

Yeast transformants carrying ADH1-GAL4 DBD (vector; LEU2) or ADH1-GAL4 DBD-

BCK2 Fragment 11 (Bck2) in a two-hybrid bait strain (Y8930) were mated to yeast

transformants of a two-hybrid prey strain (Y8800) bearing specific gene ORFs fused to

the N-terminal GAL AD (activation domain; TRP1), i.e. ADH1-GAL4 AD-ORF plasmid.

Diploids were selected by streaking cultures onto double plasmid selection medium (SD

– Leu – Trp). Diploids, of equivalent optical density, were spotted in serial 10-fold

82

dilutions on double plasmid selection medium (SD – Leu – Trp), or medium where

growth was proportional to transcription of the ADE2 gene (SD – Leu – Trp - Ade), and

incubated for 48 hr at 30°C. Six diploid strains carrying different combinations of AD

and DBD ORF-fusions (Gelperin et al. 2005; Yu et al. 2008) were used as a spectrum of

positive and negative controls.

83

3.4 Results

3.4.1 Truncation analysis of the BCK2 gene

To interrogate the relevance of a putative activation domain to Bck2 molecular

function, I created 20 truncations of the BCK2 gene and fused them to the Gal4 DBD

(Figure 3-1). First, I assessed the ability of each truncation construct to activate

transcription of two reporter genes in which the Gal4 UAS is upstream of lacZ (Figure 3-

2) or ADE2 (Figure 3-3). Bck2 residues 662 to 851 were required for transcriptional

activation (fragments 11-20), while the Bck2 N-terminal region had some apparent

repressive activity (Figure 3-2), since deletions of the N-terminus resulted in significant

increases in reporter gene expression. A construct containing fragment 5 lacking the first

529 amino acids of Bck2 was the most potent Y2H auto-activator in the lacZ reporter

assay (Figure 3-2).

To explore the relationship between Y2H auto-activation and biological function,

I assessed the ability of each Bck2 fragment to complement the synthetic lethal

phenotype of a cln3Δbck2Δ strain (Figure 3-4). I discovered that fragments of Bck2

containing residues 250 to 662 are essential for complementation of the lethality of the

cln3Δbck2Δ strain (Figure 3-1; 3-4). Consistent with the observation that the first 178

residues of Bck2 are not necessary for suppression of the pkc1 lysis phenotype by high-

copy BCK2 (Lee, Hines, and Levin 1993), a derivative of Bck2 lacking the N-terminal

178 residues was able to complement the inviability of the cln3Δbck2Δ strain (Figure 3-

4). Bck2 residues 529 to 851 alone failed to complement the cln3Δbck2Δ strain but were

sufficient for Y2H auto-activation. This region is also insufficient to suppress the lysis

84

Figure 3-1. Truncation analysis of the BCK2 gene. Fragments of the BCK2 gene (black bar) were

amplified from genomic DNA using primer positions shown and designated as “amino acid + F (Forward),

or R (reverse)”. PCR products were cloned into a yeast two-hybrid vector to create BCK2 fragments fused

to the N-terminal GAL4 DBD (DNA Binding Domain). High density growth spots (in either the ADE2

transcription activation assay or the complementation assay) were called “+” or “++” depending on extent

of growth. A complete absence of growth was called “-”. Numbers in the β-Gal column represent

averaged quantities (per fusion protein) in Miller Units (U).

Fragment # - Ade

1 +

β-Gal

0.43

3

Rescue

cln3Δ

bck2Δ

+

1 851 BCK2

Start-F 81F 178F 250F 529F 610R

662R 766R Stop-R

+ 0.40

0

+ 2 1.62

2

+++ + 3 + 2.38

8

+++ 4 + 3.84

4

- 5

+ 0.40

0

++ 6 + 0.22

2

++ 7 + 1.76

6

+++ 8 + 1.52

2

+++ 9 +

-

2.69

9 0.09

9

-

+

10

11 - 0.14

4

+ 12 - - 0.16

6

13 - - 0.20

0

14 - - 0.02

2

15

- 0.26

6

+ 16 - 0.13

3

+ 17 - 0.31

1

- 18 - 0.11

1

- 19 - 0 - 20

85

Figure 3-2. Transcriptional activation by BCK2 truncations in lacZ reporter assays. A yeast-two-

hybrid bait strain where Gal4 UAS elements drive expression of ADE2, HIS3, and lacZ reporter genes

(Y8930) bearing truncated versions of the BCK2 gene, ADH1-GAL4 DB-BCK2 (F1-20) (LEU2) plasmids

were spotted onto plasmid selection medium, incubated for 48 hr at 30°C, overlaid with a top agar solution

containing X-Gal, and incubated at 30°C until blue color was seen (inset). To quantify the relative

differences in transcriptional activity observed by the overlay assay, three independent isolates from a

single transformation reaction (per plasmid, of which there are 21), were grown to mid-log phase in

plasmid selective medium and subjected to quantitative β-galactosidase liquid assays to measure lacZ

expression. Y-axis values are expressed in relative Miller Units. Error bars reflect standard deviation of

values obtained from 3 independent transformants in separate experiments.

86

Figure 3-3. Transcriptional activation by BCK2 truncations in ADE2 reporter assays. A yeast-two-

hybrid bait strain where Gal4 UAS elements drive expression of ADE2, HIS3, and lacZ reporter genes

(Y8930) bearing ADH1-GAL4 DB-BCK2 (1-20) (LEU2) plasmids were spotted in serial 10-fold dilutions

on plasmid-selection medium or medium where growth is proportional to transcription of the ADE2 gene.

Plates were incubated for 48 hr at 30°C.

SD-L-Ade

SD-L-Ade

SD-L

SD-L

2

3

4

5

6

7

8

9

10

1

12

13

14

15

16

17

18

19

20

11

Vector

87

Figure 3-4. Complementation of a cln3Δbck2ΔpGAL-CLN3 strain growth defect by BCK2 truncation

derivatives. Transformants of a cln3Δbck2Δ GAL-CLN3 (URA3) strain (BY3015) bearing ADH1-GAL4

DBD-BCK2 (1-20) plasmids (LEU2) were grown in plasmid selective medium, grown to equivalent optical

density and spotted in serial 10-fold dilutions onto plasmid selective medium containing either galactose –

5’FOA or glucose + 5’FOA. Plates were incubated for 48 hr at 30°C.

2

3

4

5

6

7

8

9

10

1

12

13

14

15

16

17

18

19

20

11

Vector

SC+5`FOA

SC+5`FOA

SG-U-L

SG-U-L

88

phenotype of pkc1 mutants (Lee, Hines, and Levin 1993). I conclude that the central

region (250 to 662) of Bck2 lacking the N- and C-terminal ends is necessary but not

sufficient to complement essential in vivo functions of Bck2 and that this essential

function is unrelated to the Y2H auto-activation activity of the Bck2 protein.

89

3.4.2 A yeast-two-hybrid screen using Gal4 DB-Bck2 as bait identifies six novel

interacting proteins

Thus far, no known protein interaction partners of Bck2 easily explain the cell

cycle transcription phenotypes associated with deletion of BCK2. The Y2H auto-

activation property of Bck2 has precluded identification of Bck2 binding partners using

the Y2H screening method. In order to carry out a yeast two-hybrid screen, I decided to

use the largest Bck2 construct that did not auto-activate the ADE2 Y2H reporter gene, but

complemented the inviability of the cln3Δbck2Δ strain (fragment 11 (Figure 3-1)). I

chose this fragment for my screen since complementing regions are often important

protein-protein interaction domains. For example, the minimal region of the Ada2

protein required for complementation is the same region required for physical interaction

with Gcn5 and Ada3 (Candau and Berger 1996).

I used the ORFeome Y2H screening method (Rual et al. 2005) to discover

potential Bck2-interacting proteins. I identified 6 proteins that interacted with Bck2:

Mcm1, Yap6, Tpd3, Std1, Mth1, and Mot3 (Figure 3-5A). With the exception of Tpd3,

all of these proteins are transcriptional regulators that physically bind DNA. Mcm1 is a

member of a class of MADS box transcription factors that are found in all eukaryotic

organisms, including humans (Wynne and Treisman 1992; Treisman 1994; Shore and

Sharrocks 1995), and I describe my detailed analysis of the Mcm1-Bck2 interaction in

Chapter 4. Yap6 is a basic leucine zipper (bZIP) transcription factor that activates a

number of genes involved in sodium and lithium tolerance (Fernandes, Rodrigues-

Pousada, and Struhl 1997; Mendizabal et al. 1998). Yap6 is part of the yAP family, the

yeast homologs of AP1-like factors in mammals (Moye-Rowley, Harshman, and Parker

1989) that contain DNA-binding properties identical to the product of the c-jun proto-

90

Figure 3-5. Bck2-interacting proteins identified in a genome-wide yeast two-hybrid screen. (A) Positive clones that represent Bck2-interacting proteins are shown next to the number of times they were

identified in the genomic screen. (B) Yeast transformants carrying ADH1-GAL4 DBD (vector; LEU2) or

ADH1-GAL4 DBD-BCK2 Fragment 11 (Bck2) in a two-hybrid bait strain (Y8930) were mated to yeast

transformants of a two-hybrid prey strain (Y8800) bearing specific gene ORFs fused to the N-terminal

GAL AD (activation domain; TRP1, i.e. ADH1-GAL4 AD-ORF plasmid). Diploids were selected by

streaking on double plasmid selection medium (SD – Leu – Trp). Strains were grown to equivalent optical

density, and spotted in serial 10-fold dilutions on double plasmid selection medium (SD – Leu – Trp) or

medium where growth is proportional to transcription of the ADE2 gene (SD – Leu – Trp - Ade). Plates

were incubated for 48 hr at 30°C. Six diploid strains carrying different combinations of AD and DBD

ORF-fusions were used as a spectrum of positive and negative controls for determination of relative

physical interaction strength.

EV (DB) x EV (AD)

Rb (DB) x E2FΔF1 (AD)

fos (DB) x jun (AD)

Gal4 (DB) x Gal4 (AD)

DP1 (DB) x E2F (AD)

SD - L - Trp SD - L – Trp - Ade

Bck2

Vector

Bck2

Vector

Bck2

Vector

Bck2

Vector

Bck2

Vector

Bck2

Vector

Mcm1

Mbp1

Swi4

Yap6

Mot3

Stb1

B

A

Negative regulator of the glucose-sensing signal transduction

pathway, required for repression of transcription by Rgt1 10 Mth1

Transcription factor involved in control of glucose-regulated

gene expression 2 Std1

Regulatory subunit of the heterotrimeric protein phosphatase 2A

(PP2A), which contains Cdc55 and either Pph21 or Pph22 1 Tpd3

Transcription factor with a putative role in regulation of

expression of genes involved in carbohydrate metabolism 9 Yap6

Transcriptional regulator involved in repression of hypoxic

genes by Rox1 17 Mot3

Transcription factor involved in cell-type specific transcription,

pheromone response, and cell cycle progression 1 Mcm1

Description # Clones Prey Protein

91

oncogene (Bohmann et al. 1987). Tpd3 is the scaffold subunit A of the heterotrimeric

protein phosphatase 2A (PP2A) (van Zyl et al. 1992), which is involved in the TOR

pathway for nutrient sensing (Jiang and Broach 1999). Std1 and Mth1 are both

controllers of glucose-regulated gene expression (Schmidt et al. 1999; Sabina and

Johnston 2009), which are required for transcriptional repression of HXT (hexose

transport) genes (Lakshmanan, Mosley, and Ozcan 2003; Kim and Johnston 2006). The

destruction of Std1 and Mth1 results in derepression of HXT gene transcription, leading

to a rapid influx of glucose into cells (Kaniak et al. 2004). Mot3 is a Zn-finger

transcription factor that activates a number of cell wall genes (Abramova, Sertil et al.

2001) and represses transcription of the DAN/TIR group of genes that encode cell wall

mannoproteins during anaerobic growth (Abramova et al. 2001).

92

3.4.3 Bck2 and Mot3 activate CYC1 and RPL39 transcription

BCK2 was originally isolated as a high-copy suppressor of the cell wall lysis

phenotype of mpk1Δ and pkc1Δ mutants (Lee, Hines, and Levin 1993). This phenotype

may reflect Bck2-dependent activation of cell wall genes, many of which contain

SCB/MCB elements and are also regulated by SBF/MBF at the G1/S transition. Although

Bck2 activates transcription from SCB and MCB elements in a SWI4 or SWI6-dependent

manner (Di Como, Chang, and Arndt 1995), deletion of SWI4 or SWI6 has little effect on

the ability of overexpressed BCK2 to induce expression of several natural SBF/MBF

target genes (Di Como, Chang, and Arndt 1995). Natural promoters usually have binding

sites for multiple factors, leaving open the possibility that Bck2 activates other DNA-

bound factors on the promoters of cell wall genes. As noted above, Mot3 is a

transcription factor that activates a number of cell wall genes such as CWP2 (Abramova

et al. 2001). In mot3Δ cells, levels of CWP2 mRNA are strongly decreased (Abramova,

Sertil et al. 2001), while CWP2 expression is increased in cells overexpressing BCK2 in a

SWI6-independent manner (Ferrezuelo, Aldea, and Futcher 2009). MOT3 overexpression

also suppresses the cell wall integrity defect of mpk1Δ mutants (Grishin et al. 1998). My

Y2H screen identified Mot3 as a Bck2-interactor, suggesting that association of Bck2

with Mot3 may promote induction of cell wall genes.

As a first test of whether Bck2 activates Mot3-regulated genes, I assessed

activation of a CYC1-lacZ reporter gene in cells lacking either BCK2 or MOT3. Mot3

binds CYC1 promoter DNA in vitro and activates CYC1 transcription in vivo (Grishin et

al. 1998). Both bck2Δ and mot3Δ cells were defective in CYC1 lacZ reporter activity

(Figure 3-6A). Next, I performed a similar test with an RPL39-lacZ reporter gene, since

93

Figure 3-6. Bck2 and Mot3 activate CYC1 and RPL39 transcription. (A) WT (white bars), bck2Δ

(black bars), or mot3Δ (grey bars) yeast transformants carrying CYC1-lacZ were assessed for lacZ

expression. Asynchronous cells were grown to mid-log phase in selective medium and subjected to

quantitative β-galactosidase assays. Y-axis values are expressed in relative Miller units. Error bars reflect

values obtained from 3 independent transformants in separate experiments. (B) WT (white bars), bck2Δ

(black bars), or mot3Δ (grey bars) yeast transformants carrying RPL39-lacZ were assessed for lacZ

expression. Asynchronous cells were grown to mid-log phase in selective medium and subjected to

quantitative β-galactosidase assays to measure lacZ expression. Y-axis values are expressed in relative

Miller units. Error bars reflect standard deviation of values obtained from 3 independent transformants in

separate experiments.

94

levels of RPL39 mRNA are substantially increased (5.6-fold) in cells overexpressing

BCK2 (Martin-Yken et al. 2002). Again, both bck2Δ and mot3Δ cells were defective in

RPL39 lacZ reporter gene expression (Figure 3-6B). These transcriptional effects are

likely not general defects in transcription because both bck2Δ (Di Como, Chang, and

Arndt 1995) and mot3Δ (Grishin et al. 1998) strains express normal levels of ACT1

mRNA. These results suggest that Bck2 and Mot3 may function to co-regulate the CYC1

and RPL39 promoters, and perhaps other genes, in vivo.

95

3.5 Discussion

The ability of Bck2 to act as a genetic activator of G1/S-expressed genes,

physically interact with SBF/MBF, auto-activate transcription, and localize to the

promoters of G1/S-regulated genes led to the hypothesis that Bck2 activates SBF/MBF-

regulated genes by associating with their promoters in order to recruit the general

transcription machinery and lead to their transcriptional activation. In this study, I

characterized the in vivo activity of various truncated fragments of Bck2 in order to

identify the region responsible for the property of auto-activation. The most striking

result of these experiments was that the Bck2 auto-activation domain was not required for

viability. During the course of these studies, I was able to isolate an allele of BCK2 that

would allow me to use Bck2 as a bait protein in a Y2H assay to identify potentially novel

Bck2-interacting proteins. Five of the six proteins I isolated are DNA-binding proteins

with roles in ion homeostasis and nutrient sensing. One of these proteins, Mot3, has been

implicated in cell wall biosynthesis and cell cycle progression. I determined that Mot3

and Bck2 both are activators of CYC1 and RPL39 transcription.

3.5.1 Bck2 activity under hyper-osmotic conditions

The association of Bck2 with Yap6, Std1 and Mth1 suggests that Bck2 may be

required under hyper-osmotic conditions for coordinating ion homeostasis with G1/S

progression, processes similar to those coordinated by the Ppz1/Hal3 and Sit4 pathways

(Section 1.5). Increased expression of ENA1, which encodes a P-type ATPase sodium

pump that promotes cation efflux (Ruiz and Arino 2007) positively correlates with

activation of G1/S progression and rapamycin resistance and may be an important read-

96

out for the activity of proteins that co-regulate ion homeostasis and cell cycle. For

example, a halotolerant Ppz1-deficient strain has a high level of ENA1 expression (Posas,

Camps, and Arino 1995) and an accelerated G1/S transition (Posas et al. 1992; Clotet et

al. 1999). Similarly, Sit4 is an activator of ENA1 expression (Crespo et al. 2001) and

accelerates the G1/S transition (Sutton, Immanuel, and Arndt 1991; Fernandez-Sarabia et

al. 1992). ENA1 expression is induced by high-copy YAP6 (HAL7) (Mendizabal et al.

1998) and STD1 (Rios, Ferrando, and Serrano 1997; Ganster, McCartney, and Schmidt

1998), indicating that activation of ENA1 by BCK2 is an important event in the activation

of G1/S progression.

Several lines of evidence suggest that poor carbon or nitrogen conditions may

activate a pathway defined by Bck2, Yap6, Std1 and Mth1 to activate genes involved in

halotolerance. First, cells lacking BCK2 have a growth defect (Smith et al. 1996) that is

aggravated by growth in minimal complete media (Giaever et al. 2002). Second,

expression of ENA1 is higher on derepressing carbon sources, such as galactose or

raffinose, than on glucose (Rios, Ferrando, and Serrano 1997; Alepuz, Cunningham, and

Estruch 1997). Third, inhibition of the TOR nutrient sensing pathway by rapamycin

treatment (Crespo and Hall 2002) or starvation (Garciadeblas et al. 1993) increases ENA1

expression (Crespo et al. 2001). Fourth, loss of Snf1, which is required for proper

adaptation to poor glucose, results in enhanced sensitivity to Na+ or Li

+, likely due to

deficient induction of ENA1 by high salt concentrations (Alepuz, Cunningham, and

Estruch 1997). Fifth, Std1 physically interacts with Snf1 (Hubbard, Jiang, and Carlson

1994), TBP (Tillman et al. 1995), and Bck2. Finally, among the preferred promoter

targets of Yap6 are the ribosomal protein genes (Ni et al. 2009), which are activated by

97

the TOR signalling pathway (Klein and Struhl 1994; Neuman-Silberberg, Bhattacharya,

and Broach 1995; Cardenas et al. 1999; Hardwick et al. 1999; Powers and Walter 1999;

Wang et al. 2004).

3.5.2 Bck2 activity under hypo-osmotic conditions

Under hypo-osmotic conditions (i.e. cell swelling or increased turgor), Bck2

activity may be required to coordinate ion homeostasis with cell cycle progression

through activation of Mcm1-regulated genes. In particular, hypo-osmotic conditions may

dictate that H+ be effluxed because cells maintain electrical balance through the exchange

of H+ with cations (Arino, Ramos, and Sychrova 2010). Thus, the slow growth

phenotype of bck2Δ cells in basic pH (Giaever et al. 2002) is consistent with a defect in

H+ efflux. The PMA1 gene encodes a H

+ ATPase efflux pump that can be activated by

glucose (Eraso, Mazon, and Portillo 2006) or Ptk2 (Goossens et al. 2000) (Section 1.5).

Interestingly, overexpression of BCK2 induces expression of the PMA1 gene (Ferrezuelo,

Aldea, and Futcher 2009), whose promoter binds Mcm1 (Kuo and Grayhack 1994).

Consistent with the requirement for proper PMA1 expression for cell cycle progression,

overexpression of both PTK2 and BCK2 can bypass the G1 arrest of a strain lacking SIT4

and HAL3 (Munoz et al. 2003). Interestingly, overexpression of SWI4 and CLN3 also

bypass the G1 arrest of a strain lacking SIT4 and HAL3 (Munoz et al. 2003), and SWI4

and CLN3 are rapidly induced upon refeeding nutrient-deprived cells (Gray et al. 2004;

Radonjic et al. 2005), indicating that their induction could be due to carbon source

regulation of PMA1 expression.

98

Several lines of evidence suggest that Bck2 is activated by the Sln1 osmosensing

pathway (Ketela et al. 1998; Li et al. 1998) to activate various Mcm1-regulated cell wall

genes, as part of processes required for cell cycle progression under hypo-osmotic

conditions. Sln1 encodes a protein activated by hypo-osmolarity (Posas et al. 1996;

Posas and Saito 1998) and repressed by hyper-osmolarity (Maeda, Wurgler-Murphy, and

Saito 1994; Posas et al. 1996; Fassler et al. 1997). In addition to its role in the G1 phase

activation of CLN3, SWI4, CDC6 and CDC47 (McInerny et al. 1997; MacKay et al.

2001), Mcm1 activates several cell wall genes, including GFA1 (Kuo and Grayhack

1994) and AGA1 (Oehlen, McKinney, and Cross 1996). Similarly, Bck2 activates GFA1

(Martin-Yken et al. 2002) and AGA1 transcription in a SBF/MBF-independent manner

(Ferrezuelo, Aldea, and Futcher 2009). Moreover, high-copy BCK2 stimulates the

expression of the Mcm1-dependent reporter gene, P-lacZ, which is also stimulated by

high-copy SLN1 gene (Yu, Deschenes, and Fassler 1995; Li et al. 1998). Interestingly,

the induction of CLN3 and SWI4 transcription as a consequence of SLN1 overexpression

is abrogated in a bck2Δ strain (N. Bastajian; unpublished observations), indicating that

activation of Bck2 by activated Sln1 allows cell cycle progression to occur under hypo-

osmotic conditions.

A downstream effector of Sln1 activity is the SKN7 gene product (Li et al. 2002),

which may act together with Bck2 to control various aspects of cell wall biosynthesis and

cell cycle progression. Similar to BCK2 (Lee, Hines, and Levin 1993), high-copy SKN7

suppresses the inviability of cells deficient in cell wall integrity (Brown, North, and

Bussey 1993) and bypasses the requirement for Swi6 in transcription from MCB

elements to which MBF binds (Morgan et al. 1995; Wijnen and Futcher 1999).

99

Moreover, overexpression of either SKN7 (Williams and Cyert 2001) or BCK2

(Ferrezuelo, Aldea, and Futcher 2009) increases FKS1 expression, and activates a Mcm1-

dependent reporter gene (Li et al. 1998). Thus, Skn7 functions like Bck2 in acting to

restore cell wall integrity and in acting without the requirement for Swi6. Whether high-

copy BCK2, like SKN7 (Brown, North, and Bussey 1993), is able to suppress the cell wall

defects of a kre9Δ strain is not clear, but high-copy BCK2 suppresses the cell wall defects

of a pkc1Δ strain (Lee, Hines, and Levin 1993), indicating that Bck2 may induce genes

similar to Skn7. Pkc1, which is known to become activated in hypo-osmotic conditions

(Davenport et al. 1995), may be part of a distinct pathway because there is limited

overlap in target genes when comparing the PKC1 (Jung and Levin 1999; Roberts et al.

2000) and BCK2 pathways (Martin-Yken et al. 2002; Ferrezuelo, Aldea, and Futcher

2009). Consistent with Bck2 acting in a distinct pathway from Pkc1, both Skn7 (Brown,

Bussey, and Stewart 1994) and Bck2 (Lee, Hines, and Levin 1993) have additive genetic

interactions with Pkc1. Interestingly, Yap6 in association with Skn7 binds the promoters

of many genes encoding constituents of the cell wall (Ni et al. 2009), indicating that

Yap6 and Skn7 may act as effectors of a Bck2-specific pathway.

3.5.3 Tpd3 and Bck2 in nutrient sensing

Under nutrient-rich conditions, Bck2 might be activated by a PP2A complex

containing Tpd3. PP2A (Jiang and Broach 1999) and the PP2A-like protein Sit4 (Rohde

et al. 2004) positively act in the TOR pathway for nutrient sensing, which can be

inhibited by lack of nutrients or exposure to the drug rapamycin leading to cell cycle

arrest in G1 (Barbet et al. 1996). Tpd3 is the scaffold subunit A of the heterotrimeric

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protein phosphatase 2A (PP2A) (van Zyl et al. 1992), which binds one of the two B

regulatory subunits, Cdc55 or Rts1, and one of the two C catalytic subunits, Pph21 and

Pph22 (Yang et al. 2000). Both sit4Δ (Cutler et al. 2001) and tpd3Δ (Banuelos et al.

2010) strains are rapamycin sensitive, and cells that overexpress SIT4 are rapamycin

resistant (Munoz et al. 2003). Similar to Bck2, the Sit4 protein is required for the normal

accumulation of SWI4, CLN1 and CLN2 mRNAs in late G1 (Fernandez-Sarabia et al.

1992). Interestingly, high-copy PPH21 or PPH22 rescues (Munoz et al. 2003) the

growth defect of sit4 mutants on glycerol (Arndt, Styles, and Fink 1989; Sutton,

Immanuel, and Arndt 1991), indicating that a PP2A complex containing Tpd3 can

substitute for Sit4 in G1 phase. Accordingly, induction of SWI4 transcription, as a

consequence of PPH22 overexpression, is abrogated in a bck2Δ strain (N. Bastajian;

unpublished observations). In addition to the major PP2A species in the cell (Dobrowsky

et al. 1993), Tpd3 is also part of a ceramide-activated protein phosphatase (CAPP)

comprised of Tpd3, Cdc55 and Sit4 (Nickels and Broach 1996), which is important for

arresting budding yeast cells (Fishbein et al. 1993; Nickels and Broach 1996) and

mammalian cells (Okazaki et al. 1990; Bielawska, Linardic, and Hannun 1992; Obeid et

al. 1993) in G1. The findings outlined above suggest that activation of Bck2 likely

involves the normal PP2A complex containing Tpd3.

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Chapter 4

Bck2 Interacts with the MADS box protein Mcm1

on Cell Cycle-Regulated Promoters to Activate

Early G1 Phase Transcription in Budding Yeast

102

Chapter 4: Bck2 interacts with the MADS box protein Mcm1 on cell cycle-regulated

promoters to activate early G1 phase transcription in budding yeast

4.1 Abstract

The protein Bck2 has been posited to act at the G1/S transition as a late G1 phase activator

of periodically-expressed genes in budding yeast. Although Bck2 is a potent genetic

regulator of early cell cycle progression, the mechanism of gene activation by Bck2

remains obscure. To date, most experiments have focused on assessing a potential role

for Bck2 in activation of the late G1-specific transcription factors SBF (containing Swi4,

Swi6) and MBF (containing Mbp1, Swi6). I discovered that Bck2 interacted with Mcm1,

an essential protein which binds to early G1 promoters such as CLN3 and SWI4 through

Early Cell cycle Box (ECB) elements (Chapter 3). Mcm1 is inhibited by association with

two repressors, Yox1 and Yhp1, and gene activation ensues once repression is relieved

by an unknown activating signal. Here, I show that Bck2 interacts physically with Mcm1

to activate ECB-containing genes. I used chromatin immunoprecipitation (ChIP)

experiments to show that Bck2 localized to the promoters of the CLN3 and SWI4 genes,

provided functional ECB elements exist. The Bck2-Mcm1 interaction required valine 69

on Mcm1, a residue known to be involved in the interaction with Yox1 at G2/M-regulated

promoters. Moreover, overexpression of BCK2 decreased Yox1 localization to the CLN3

promoter. Finally, overproduction of BCK2 rescued the lethality caused by

overexpression of YOX1. I envision a model whereby Yox1 and Bck2 compete for

access to the Mcm1-ECB scaffold. As early G1 phase is a critical time in assessing

nutrient status, the cell may use nutrient-regulated dosage/activity of Bck2 as a means of

co-ordinately activating the initial suite of genes required for cell cycle commitment.

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4.2 Introduction

How Bck2 activates the G1/S phase transition (Epstein and Cross 1994; Wijnen

and Futcher 1999) is not known, but appears to involve activation of the SBF/MBF

transcription factors in late G1 phase. The CLN1, CLN2, and PCL1 genes are part of a

co-ordinately regulated cluster of genes that peak sharply at the G1/S transition

(Spellman, Sherlock et al. 1998) due to activation of SBF (Costanzo, Nishikawa et al.

2004; de Bruin, McDonald et al. 2004) and MBF (de Bruin, Kalashnikova, and

Wittenberg 2008). Both CLN1 (Wijnen and Futcher 1999) and CLN2 (Di Como, Chang,

and Arndt 1995), which contain SCB and MCB elements in their promoters, are activated

by overexpressed BCK2. These observations imply a role for Bck2 in promoting the

G1/S transition through activation of SBF and MBF on the promoters of genes such as

CLN2. However, the presence of SBF/MBF or the elements to which they bind is not

necessary for Bck2 to function as a transcriptional activator. For instance, the SBF-

regulated gene PCL1 (Dirick, Bohm et al. 1995; Wijnen, Landman et al. 2002) is induced

by overexpression of BCK2 both in a WT and a swi4Δmbp1Δ strain (Wijnen and Futcher

1999). Moreover, the UAS2 region of CLN2, which completely lacks SCB or MCB

elements (Stuart and Wittenberg 1994), is activated by BCK2 overexpression (Di Como,

Chang, and Arndt 1995), and the UAS2 region remains transcriptionally active in cdc28-

arrested cells (Stuart and Wittenberg 1994). These findings collectively suggest that

Bck2 activity may operate through an unidentified DNA-binding factor whose activity is

Cln3-Cdc28-independent. Thus, despite the clear importance of upstream Cln3-Cdc28

activity in the activation of CLN2 transcription in late G1 phase (Cross and Tinkelenberg

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1991; Breeden and Mikesell 1994), a Bck2-specific pathway for CLN transcription also

exists that appears to involve an unknown DNA-bound factor.

The MADS box transcription factor Mcm1 has an important regulatory function

at two points in the cell cycle – M/G1 and G2/M. During M/G1, Mcm1 functions as a

critical constituent of a complex that forms on ECB elements (Early Cell cycle Box) in

promoters of genes expressed at the M/G1 phase transition such as CLN3 and SWI4 (Mai,

Miles, and Breeden 2002). Two related homeodomain proteins Yox1 and Yhp1 act as

repressors of Mcm1 on ECB elements by physically interacting with Mcm1 and with

DNA binding sites next to the Mcm1 site in the ECB element (Pramila et al. 2002).

Mcm1 is also a critical constituent of complexes that form during the G2/M phase

transition to control the CLB2 cluster of genes, which includes CDC20, CLB2, and

SPO12 (Althoefer et al. 1995). The CLB2 gene cluster is activated by the Clb-Cdc28 and

Cdc5 kinases (Reynolds et al. 2003; Darieva et al. 2003; Pic-Taylor et al. 2004; Darieva

et al. 2006), which regulate a promoter-bound complex of Mcm1, Fkh2 and Ndd1 (Loy,

Lydall, and Surana 1999; Koranda et al. 2000; Kumar et al. 2000; Pic et al. 2000; Zhu et

al. 2000). Many CLB2 cluster genes contain hybrid elements composed of an Mcm1-

binding site flanked by Yox1- and Fkh2-binding sites (Zhu and Davis 1998). In CLB2

cluster genes that contain such elements, Yox1 and Fkh2 compete for binding to DNA-

bound Mcm1 despite the spatial separation of their DNA recognition elements (Darieva

et al. 2010). Interestingly, no such juxtaposed binding motifs are obvious in the vicinity

of ECB elements and no other binding partners for Mcm1 that positively regulate these

genes have been identified.

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In this chapter, I show that Bck2 activates CLN3 and SWI4 expression through an

interaction with Mcm1 on ECB elements in the promoters of these genes. Moreover,

removal of Yox1 correlates with increased Bck2 dosage, indicating that Bck2 might

activate CLN3 and SWI4 expression by displacing Yox1 from Mcm1. Consistent with

this hypothesis, mutation of a key residue on Mcm1 known to prevent interaction with

Yox1 also prevents interaction with Bck2. Finally, overproduction of BCK2 rescues the

lethality caused by overexpression of YOX1, indicating that Bck2 and Yox1 may compete

for access to Mcm1 on promoters. The experiments described in this Chapter suggest

that Bck2 co-ordinately regulates the expression of key genes necessary for the

subsequent switch-like transition from G1 to S phase.

4.3 Materials and Methods

4.3.1 Yeast strains and plasmids

All yeast strains used in this study (Table 4-1) were derivatives of either S288C or W303.

Yeast plasmids are described in Table 4-2. Yeast cultures were grown in YEP (1% yeast

extract, 2% bacto peptone) supplemented with 2% glucose. Synthetic minimal medium

supplemented with the appropriate nutrients was used to select for plasmid maintenance

and gene replacements. Yeast transformation, tetrad analysis and general manipulation

of yeast cells were performed using standard techniques.

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Table 4-1

S. cerevisiae strains

Strainsa BY# Relevant genotype Source

BY4742 1624 MATα his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 (Brachmann et al. 1998)

BY3015 3015 MATa cln3Δ::HphR bck2Δ::NatR his3Δ1 This study

leu2Δ0 met15Δ0 ura3Δ0 lys2Δ0

pGAL-CLN3 (URA3)

Y8930 4889 MATα trp1-901 leu2-3,112 his3-200 ura3-52 (Yu et al. 2008)

gal4Δ gal80Δ LYS2::GAL1-HIS3 GAL2-ADE2

met2::GAL7-lacZ cyhR

Y8800 4890 MATa trp1-901 leu2-3,112 his3-200 ura3-52 (Yu et al. 2008)

gal4Δ gal80Δ LYS2::GAL1-HIS3 GAL2-ADE2

met2::GAL7-lacZ cyhR

Y8890 4896 MATa cdc20-3 KanR his3Δ1 leu2Δ0 (Li et al. 2011)

met15Δ0 ura3Δ0

4897 MATα cdc20-3 bck2Δ (#2) (Y8890 x bck2Δ) This study

BY2125 4898 MATa ade2-1 his3-11, 15 leu2-3, 112 trp1-1 (MacKay et al. 2001)

ura3 can1-100 ssd1-d

BY2680 4899 MATa ade2-1 his3-11, 15 leu2-3, 112 trp1-1 (MacKay et al. 2001)

ura3 can1-100 ssd1-d cln3ecb5 swi4ecb

YOX1-TAP 4900 MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 (Ghaemmaghami et al. 2003)

YHP1-TAP 4901 MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 (Ghaemmaghami et al. 2003)

MCM1-TAP 4902 MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 (Ghaemmaghami et al. 2003)

Y7092 4903 MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 (Costanzo et al. 2010)

bck2Δ 4904 bck2Δ::NatR (Costanzo et al. 2010)

yox1Δyhp1Δ 4905 yox1ΔKanRyhp1ΔLEU2(#4α1) This study

T487 4891 EV (DB) x EV (AD) (Vidal et al. 1996)

T488 4892 Rb (DB) x E2FΔF1 (AD) (Vidal et al. 1996)

T489 4893 c-fos (DB) x c-jun (AD) (Vidal et al. 1996)

T490 4894 Gal4 (DB) x Gal4 (AD) (Vidal et al. 1996)

T491 4895 DP1 (DB) x E2F (AD) (Vidal et al. 1996) a Strain backgrounds are isogenic to S288C, except BY2125 and BY2680, which are isogenic to W303.

Table 4-2

Plasmids

Plasmids BA# Description Source

pCLM771 2436 4xP-lacZ URA3 2μ (Li et al. 1998)

pBD1790 2437 CLN3-lacZ URA3 (McInerny et al. 1997)

pBD1867 2438 CLN3(ecb)-lacZ URA3 (McInerny et al. 1997)

pBD1577 2439 SWI4-lacZ URA3 (McInerny et al. 1997)

pBD1637 2440 CDC6-lacZ URA3 (McInerny et al. 1997)

pBD1951 2441 CDC47-lacZ URA3 (McInerny et al. 1997)

ACT1-lacZ 2442 ACT1-lacZ URA3 (Hughes et al. 2001)

pGAL-FLAG 350V LEU2 CEN GAL1pro-ORF-FLAG (Ho et al. 2002)

pGAL-BCK2-FLAG 2412 LEU2 CEN GAL1pro-ORF-FLAG (Ho et al. 2002)

pGAL-CLN3-FLAG 2443 LEU2 CEN GAL1pro-ORF-FLAG (Ho et al. 2002)

pGAL-YOX1 \ 2444 URA3 (FLEX collection) (Hu et al. 2007)

pDEST32 401V LEU2 CEN ADH1pro-Gal4 DB-ORF Invitrogen

pDEST32-Gal4 DB-BCK2 2414 LEU2 CEN ADH1pro-Gal4 DB-ORF This study

pDEST22-Gal4 AD- MCM1WT 2445 TRP1 CEN ADH1pro-Gal4 AD-ORF This study

pDEST22-Gal4 AD-MCM1V69E 2446 TRP1 CEN ADH1pro-Gal4 AD-ORF This study

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4.3.2 β-Galactosidase assays

Exponentially growing cells at an optical density at 600 nm (OD600) of 0.2 to 0.25 were

harvested. Extracts were prepared by vortexing the cells in 1 ml Z-buffer (0.1 M NaPO4

[pH 7.0], 0.01 M KCl, 1 mM MgSO4, 4 mM 2-mercaptoethanol) + 20 μl 0.1% SDS + 40

μl chloroform and then vortexing them for 45 seconds. After 5 min, 0.2 ml of o-

nitrophenylgalactoside (Sigma N-1127; at 4 mg/ml in Z-buffer) solution was added and

incubated at 30°C until a slight yellowing was observed in the samples. The reactions

were stopped by the addition of 0.5 ml of 1 M Na2CO3, the samples were centrifuged for

3 minutes at 13,000 x g and the A420 of the supernatant was determined. β-Galactosidase

units were calculated by the following formula: units = (A420)*(1000)/(time of reaction,

in minutes)*(volume of extract in assay)*(cellular concentration in OD600 values). For

each time point, the assays were performed on three separate cultures, and the average is

reported.

4.3.3 Cell synchronization

The cdc20-3 temperature-sensitive strain was grown in YPD medium at 21°C, arrested in

M phase by incubation at 37°C for 3.5 hours, and released into the cell cycle by

transferring the culture back to 21°C. Arrest was determined by visualization of large

budded cells under a light microscope.

4.3.4 mRNA purification and RT-qPCR

Total RNA was isolated by phenol-chloroform extraction and resuspension in DEPC

water. Samples were further purified using the RNeasy kit (Qiagen). RNA was

108

transcribed into cDNA using the Superscript II Reverse Transcriptase kit (Invitrogen) and

RNA was then removed by addition of NaOH. Reactions were run on the ABI 7500

system (Applied Biosystems) using standard Q-PCR conditions. Data were analyzed

using ABI7500 system software. VIC and FAM labelled fluorogenic primers (ABI), used

to detect CLN2, ALG9, CLN3, SWI4, BCK2, CLB2 and ACT1 cDNA, are described in

Table 4-3.

Table 4-3

Oligonucleotidesa

ORF Forward Primer Reverse Primer Fluorogenic Probe

CLN2 GAGTACCTTAATGAACGGCATTG GGGTAGAACACCATTGACCG TCCTAACTCCTTGATGGAAGTG ALG9 CTGCCGTTGCCATGTTG GAAGTAGACCCAGTGGACAG GGTGCCACCAGACACTC CLN3 GGTCCATCTGTCAGTCGG CCAAAGGGGCAGAAAGGAC CTACGTCCCCGTTATCGC SWI4 AGGGAACACTCCACTGC ACGCAGGCGATTCGTTATC CTGGTATATCTTGGTGCGTC BCK2 CCGTTCTCTGGAAACTTGTCC GGCGGTTTTGCTAGTGAAG CCGAGAAAGAAGTCGTTCCG CLB2 CCAAGACCCAAGTAGTCAGC CTTCGCTGAGGAGGATTCTTG GTGCGCTAACTTCTATAAAGGAG ACT1 GAAACTTTCAACGTTCCAGCC CCAGTAGTTCTACCGGAAGAG CGTTTCCATCCAAGCCG a Oligonucleotides are shown 5’ to 3’

4.3.5 Yeast two-hybrid assays

Yeast transformants carrying ADH1-GAL4 DBD (vector; LEU2) or ADH1-GAL4 DBD-

BCK2 Fragment 11 in a two-hybrid bait strain (Y8930) were mated to yeast

transformants of a yeast two-hybrid prey strain (Y8800) bearing specific gene ORFs

fused to the N-terminal GAL4 AD (activation domain; TRP1; ADH1-GAL4 AD-ORF

plasmid). Diploids were selected by streaking cells onto double plasmid selection

medium (SD – Leu – Trp). Diploids, of equivalent optical density, were spotted in serial

10-fold dilutions on double plasmid selection medium (SD – Leu – Trp), or medium

where growth was proportional to transcription of the ADE2 gene (SD – Leu – Trp -

Ade), and incubated for 48 hr at 30°C. Six diploid strains carrying different

109

combinations of AD and DBD ORF-fusions (T487-T491) were used as a spectrum of

positive and negative controls.

4.3.6 Mutagenesis

To construct the Mcm1V69E

yeast two-hybrid prey plasmid, the Mcm1WT

prey plasmid

was subject to in vitro mutagenesis using the QuikChange site-directed mutagenesis kit

(Stratagene). Approximately 60 ng of plasmid DNA was added to a 50 μl reaction mix

containing 20 μM forward (5'-TCAGAAACAGGTTTGGAATATACTTTCAGCACG-

3') and 20 μM reverse (5'-CGTGCTGAAAGTATATTCCAAACCTGTTTCTGA-3')

primers.

4.3.7 ChIP assays

Yeast strains transformed with pGAL-flag plasmids were grown in raffinose-containing

minimal media overnight to prevent gene induction, and then grown separately in

raffinose- (non-inducing conditions) or galactose-containing (inducing conditions) to

mid-log phase. Cultures were harvested and anti-FLAG ChIPs were analyzed for CLN2,

CLN3 and SWI4 promoter DNA by TaqMan Q-PCR (Applied Biosystems) using primers

with homology to target gene promoter DNA (Table 4-4). Enrichment of promoter DNA

was determined relative to non-promoter DNA from an untranscribed region of

chromosome II.

Table 4-4

Oligonucleotidesa

ORF Forward Primer Reverse Primer Fluorogenic Probe CLN2 ATCAATTCATGCGCGCTTTA GACAAATTTCGCGATGATTTCC CCGGCTCCATCTTTCCGAAA

CLN3 GATGTCCTAGTGCAGCCAC CCATTTGACTGGCAGACTCAG GGTGCACCTTTATCGTGTGC

SWI4 GCATTGTGACTGCGTACTTTAAACG CGTTTGAAAAAGCGACGACCAATGC GTAAAAGGCTGTCGAAATCTCTC a Oligonucleotides are shown 5’ to 3’

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4.4 Results

4.4.1 Bck2 activates early G1 phase-expressed genes that contain Mcm1-binding

sites in their promoters

Several observations implicate Bck2 in the activation of Mcm1 target genes in

early G1 phase: (1) SWI4 mRNA accumulates much more slowly in bck2Δ than WT cells

in synchronized cultures, whereas SWI4 is upregulated in cells that overexpress BCK2

(Di Como, Chang, and Arndt 1995); (2) high-copy BCK2 stimulates the expression of the

Mcm1-dependent reporter gene, P-lacZ (Li et al. 1998); (3) overexpression of BCK2

causes increased transcription of CLN3 and SWI4 by microarray analysis (Martin-Yken et

al. 2002); and (4) Mcm1 and Bck2 interact in a 2-hybrid assay (Chapter 3). To evaluate

the significance of the Mcm1-Bck2 physical interaction in vivo, I first assessed the effect

of BCK2 deletion on expression of lacZ reporter genes whose expression was dependent

on either multiple Mcm1-binding sites (4 x P-sites) or the upstream activating sequences

of four Mcm1-regulated genes expressed in early G1 phase – CLN3, CDC6, CDC47, and

SWI4 (MacKay et al. 2001) (Figure 4-1A). In these plasmid reporter assays, deletion of

BCK2 had no effect on expression of a control ACT1-lacZ reporter gene. However, I saw

a pronounced reduction in expression of the CLN3-lacZ, CDC6-lacZ, CDC47-lacZ,

SWI4-lacZ, and P-lacZ reporter genes in the bck2Δ strain (Figure 4-1B). These results

suggest a role for BCK2 in early G1 gene expression.

To verify the results of the reporter gene assays, I next examined endogenous

levels of Mcm1 target gene expression in a bck2Δ strain (Figure 4-2). Since Mcm1

controls CLN3 and SWI4 transcript accumulation at a very early point in G1, I

synchronized cultures in mitosis with a cdc20-3 temperature-sensitive allele and released

them into the subsequent G1 phase. Cells in this experiment were slow growing and

111

Figure 4-1. Bck2 activates Mcm1-driven lacZ reporter constructs. (A) Diagrams of plasmid reporter

constructs used to assess the effect of BCK2 deletion or overexpression. Constructs containing either

multiple synthetic Mcm1-binding sites upstream of the lacZ gene (4 x P-site, pCLM771) or endogenous

promoters that contain ECB elements (CLN3 pBD1790, SWI4 pBD1577, CDC6 pBD1637, CDC47

pBD1951) are shown. Black boxes represent distinct Mcm1-binding sites such as Mcm1-binding P-site

elements or ECB elements, whereas white boxes represent MCB elements or GRE elements. The 4 x P-site

construct (pCLM771) contains 4 copies of a minimalist palindromic Mcm1-binding site. Boxes marked

with an “X” represent mutation of an element. Dashed boxes represent promoter regions contained within

a specific reporter construct. (B) WT (grey bars) or bck2Δ (black bars) yeast transformants carrying P-

lacZ, CLN3-lacZ, SWI4-lacZ, CDC6-lacZ, CDC47-lacZ, and ACT1-lacZ were assessed for lacZ expression

level. Asynchronous cells were grown to mid-log phase in selective medium and subjected to quantitative

β-galactosidase assays to measure lacZ expression. Y-axis values are expressed in relative Miller units.

Error bars reflect values obtained from 3 independent transformants in separate experiments.

GREs ECBs CLN3

ECB MCBs SWI4

ECBs MCBs CDC6

ECBs CDC47

4 x P-site

GREs ECBs CLN3 (ecb)

lacZ

lacZ

lacZ

lacZ

lacZ

lacZ X X X X

A

B

X X

112

exhibited highly periodic CLN2 transcription in both WT and bck2Δ cells, indicating that

these cultures were synchronously released from the mitotic block. Consistent with

previous reports, CLN2 transcript was reduced in the bck2Δ strain (Epstein and Cross

1994; Di Como, Chang, and Arndt 1995), while levels of a control transcript (ALG9)

were unaffected. Strikingly, the accumulation of CLN3 and SWI4 mRNAs was

significantly reduced in bck2Δ cells, and peak expression was also delayed, at least for

the CLN3 transcript. In wild-type cells, CLB2 mRNA peaked after G1 transcripts as

expected. However, in bck2Δ cells CLB2 transcripts were reduced and began to

accumulate near the end of the time-course, after the peak of CLB2 expression seen in

wild-type cells. Thus, BCK2 is required for the appropriate expression of CLN3, SWI4,

and CLB2 mRNA.

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Figure 4-2. Effect of BCK2 deletion on CLN2, ALG9, CLN3, SWI4, BCK2 and CLB2 mRNA

accumulation during the cell cycle. WT (cdc20-3; blue) and bck2Δ cultures (cdc20-3; pink) were grown

to log-phase, arrested at M/G1 by incubating for 3.5 hours at 37°C (block), then released into the cell cycle

by re-incubation at 21°C. Samples were harvested every 15 minutes and mRNA levels (Y-axis) quantified

by Q-PCR using ACT1 mRNA levels as a normalizing control. In WT cells, the peak of CLN2

transcription marks the G1/S transition, the peak of CLN3 and SWI4 marks M/G1, and the peak of CLB2

marks G2/M.

0 15 30 45 60 75 90

WT bck2Δ

ALG9 CLN2

CLB2

CLN3 SWI4

BLOCK LOG

BCK2

0 15 30 45 60 75 90 BLOCK

LOG

0 15 30 45 60 75 90 BLOCK

LOG 0 15 30 45 60 75 90 BLOCK

LOG

0 15 30 45 60 75 90 BLOCK

LOG 0 15 30 45 60 75 90 BLOCK

LOG

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4.4.2 Bck2 requires ECB elements for transcriptional activation

The observation that Bck2 is required for proper transcription of early G1 genes

that contain Mcm1-binding sites suggested that Bck2 may function through early cell

cycle box (ECB) elements. To test this hypothesis, I assayed the effect of BCK2 deletion

on expression of lacZ reporter plasmids containing either a WT CLN3 promoter that has

intact ECB elements or a mutated CLN3 promoter that lacks functional ECB elements

(McInerny et al. 1997). Deletion of BCK2 in combination with mutation of ECB

elements caused a level of reporter gene expression similar to that seen with either

perturbation alone (Figure 4-3A), indicating that Bck2 acts through ECB elements in

order to control expression of early G1 genes.

To gather more evidence that Bck2 works through ECB elements, I next tested the

effects of BCK2 overexpression on ECB-containing reporter gene expression. For this

experiment, I used a lacZ reporter gene driven by a version of the CLN3 promoter in

which ECB elements were mutated (McInerny et al. 1997). As previously seen

(McInerny et al. 1997) when ECB elements were mutated, expression of CLN3-lacZ in

wild-type cells was substantially reduced (Figure 4-3B). Overexpression of BCK2 could

not maximally induce expression of a CLN3-lacZ reporter gene when intact ECB

elements were mutated (CLN3ecb-lacZ) (Figure 4-3B). I conclude that ECB elements are

required for Bck2 to maximally activate transcription through the CLN3 promoter.

To substantiate the requirement of ECB elements in transcriptional activation of

M/G1-regulated genes by overproduced Bck2, I next assayed the effects of BCK2

overexpression on expression of the endogenous CLN3 and SWI4 genes. I compared the

115

Figure 4-3. Bck2 requires intact ECB elements in the CLN3 promoter for transcriptional activation. (A) WT (grey bars) or bck2Δ (black bars) yeast transformants harboring a CLN3-lacZ or CLN3ecb-lacZ

reporter plasmid were grown to mid-log phase in selective medium and subjected to quantitative β-

galactosidase assays. (B) WT yeast strains containing a CLN3-lacZ, CLN3ecb-lacZ or ACT1-lacZ reporter

plasmid were co-transformed with pGAL-BCK2-FLAG (black bars) or vector (grey bars) and grown to

mid-log phase in selective medium containing galactose (inducing conditions) and subjected to quantitative

β-galactosidase assays to measure lacZ expression. Y-axis values are expressed in relative Miller units.

Error bars reflect values obtained from 3 independent transformants in separate experiments.

116

expression of CLN3 and SWI4 in a wild-type strain to a strain where ECB elements in the

promoters of both CLN3 and SWI4 were mutated (cln3(ecb)swi4(ecb)) (MacKay et al.

2001). Consistent with my reporter gene assays, overproduction of Bck2 increased CLN3

and SWI4 transcript levels in a WT strain, but not the cln3(ecb)swi4(ecb) mutant strain

(Figure 4-4), indicating that Bck2 functions through ECB elements in endogenous early

G1 promoters. As previously seen, overproduction of Bck2 also increased CLN2

expression in WT cells (Di Como, Chang, and Arndt 1995; Ferrezuelo, Aldea, and

Futcher 2009), but that induction (Figure 4-4) was entirely independent of the ECB

elements in the CLN3 and SWI4 gene promoters. This result suggests that the induction

of CLN2 transcription by overexpressed BCK2 is not an indirect consequence of

increased CLN3 and SWI4 expression. Recently, Cross and colleagues showed that

Mcm1 binds to the CLN2 promoter (Bai, Ondracka, and Cross 2011), suggesting that the

localization of Bck2 to the CLN2 promoter may be mediated by a Bck2-Mcm1

interaction at the same sites Mcm1 was shown to bind.

Similarly, the induction of CLB2 expression by overexpressed BCK2 (Di Como,

Chang, and Arndt 1995; Ferrezuelo, Aldea, and Futcher 2009) was also independent of

the ECB elements in the CLN3 and SWI4 promoters (Figure 4-4), again suggesting that

the CLB2 induction is not an indirect effect of defects in early G1 phase gene expression.

Finally, overproduced BCK2 did not alter ALG9 expression, a non-ECB containing gene,

indicating that Bck2 is not a general activator of global transcription. Together, my

analyses of the effects of BCK2 deletion and overexpression show that Bck2 activates

CLN3 and SWI4 transcription in an ECB-dependent manner, while promoting expression

117

Figure 4-4. Bck2 requires intact ECB elements in the CLN3 and SWI4 promoters for transcriptional

activation of CLN3 and SWI4. (A) WT (BY2125) and a strain lacking functional ECB elements in the

CLN3 and SWI4 promoters (BY2680; cln3(ecb)swi4(ecb)) were transformed with pGAL-BCK2-FLAG

(hatched bars in WT; black bars in mutant) or vector (white bars in WT; grey bars in mutant).

Transformants were grown to saturation in plasmid selective medium and subcultured in YPGal to mid-log

phase before harvesting for quantification of mRNA levels by Q-PCR analysis using ACT1 mRNA levels

as a normalizing control. Relative enrichment of CLN3, SWI4, CLN2, ALG9 and CLB2 mRNA normalized

against ACT1 mRNA is shown on in the left panel. The Y-axis represents the normalized mRNA levels.

Relative enrichment of BCK2 mRNA from the same samples normalized against ACT1 mRNA is shown in

the right panel.

GREs ECBs

GREs ECBs

CLN3

CLN3 (ecb) X X X X

A

B

X X

SWI4

SWI4 (ecb)

cln3(ecb)swi4(ecb)

WT

cln3(ecb)swi4(ecb)

WT

ECB MCBs

ECB MCBs

X

118

of late G1- (CLN2) and G2/M-regulated (CLB2) genes by a mechanism that is not yet

known.

119

4.4.3 Bck2 localizes to early and late G1 phase promoters

Since Bck2 functions through ECB elements (see above) and physically interacts

with Mcm1 (Chapter 3), I next asked if Bck2 localized to the promoter regions of G1-

phase genes. I first used ChIP and a strain carrying a TAP-tagged allele of Bck2 to assess

association of Bck2 with the CLN2, CLN3 and SWI4 promoters. I detected a reproducible

enrichment of promoter DNA in the Bck2-IP, but the signal was very low relative to

Swi4 or Mcm1 IPs. To improve my assay, I repeated the ChIP experiment using a strain

in which a Flag-tagged derivative of Bck2 was conditionally overproduced (Figure 4-5).

Under inducing conditions (galactose), Bck2-flag IPs were enriched in CLN2, CLN3 and

SWI4 promoter DNA relative to non-inducing conditions (raffinose) (Figure 4-5) or

vector control (data not shown). The enhanced enrichment of CLN3 promoter DNA

compared to SWI4 promoter DNA in Bck2 IPs likely reflects the presence of more ECB

elements in the CLN3 promoter (6 versus 1). Association of Bck2 with the CLN3 and

SWI4 promoters was entirely dependent on the presence of ECB elements, while

association with the CLN2 promoter was unaffected, consistent with my gene expression

analysis (Figure 4-4). My findings are supported by a recent study that identified Bck2 as

a constituent of DNA-bound complexes containing Mcm1 (Lambert et al. 2010). I

conclude that Bck2 localizes to the promoters of CLN3 and SWI4 in a manner that

depends on ECB elements. Bck2 also localizes to the CLN2 promoter through an

unknown binding element, which may also require Mcm1 (Bai, Ondracka, and Cross

2011). Mcm1-dependence is difficult to assess directly in these experiments since

MCM1 is an essential gene.

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Figure 4-5. Bck2 localizes to the promoters of early and late G1 phase-expressed genes. A WT strain

(BY2125; W303) or a strain containing mutated ECB elements in the CLN3 and SWI4 promoters (BY2680;

cln3(ecb)swi4(ecb)) was transformed with a pGAL-BCK2-FLAG plasmid and grown separately in

raffinose- (i.e. RAF; non-inducing conditions; white in WT, grey in mutant) or galactose-containing

medium (i.e. GAL; inducing conditions; black in WT, lined in mutant) to mid-log phase. Cultures were

harvested and anti-FLAG ChIPs were analyzed for CLN2, CLN3 and SWI4 promoter DNA by Q-PCR. The

Y-axis measures enrichment of promoter DNA for the target gene indicated relative to enrichment of non-

promoter DNA from an untranscribed region of chromosome II. Error bars reflect values obtained after

multiple Q-PCR runs from the same experiment.

121

4.4.4 Bck2 competes with Yox1 for access to Mcm1 on ECB elements

As noted earlier, the closely related homeodomain proteins Yox1 and Yhp1 act as

repressors of Mcm1 by interacting directly with Mcm1 at DNA binding sites adjacent to

the actual Mcm1 DNA binding site within the ECB (Pramila et al. 2002). Activation of

Mcm1 on ECB elements correlates with removal of Yox1 from ECB elements, while

deletion of Yhp1 has little effect on the level or periodicity of gene expression and is not

part of the predominant complex at ECB elements (Pramila et al. 2002). These

observations suggest that Bck2 may activate CLN3 and SWI4 transcription through ECB

elements by promoting the removal of Yox1. To test this hypothesis, I first assessed

Yox1 binding to ECB elements within the CLN3 promoter when Bck2 levels were

elevated. Overexpression of BCK2 significantly reduced the amount of Yox1 associated

with the CLN3 promoter (Figure 4-6A; left panel), implicating Bck2 in Yox1 removal.

Yhp1 was not localized to the CLN3 promoter to the same extent as Yox1, nor was the

association affected by BCK2 overproduction, consistent with a secondary role for Yhp1

(Pramila et al. 2002). Consistent with previous reports that Mcm1 remains localized to

promoters throughout the cell cycle (Pramila et al. 2002), I observed that Mcm1

localization was not significantly affected by BCK2 dosage (Figure 4-6A; right panel).

These experiments suggest that Bck2 overproduction leads to the displacement of Yox1

repressor from the CLN3 promoter, which is correlated with activation of M/G1-regulated

genes (Pramila et al. 2002).

To further test the model that Bck2 may compete with Yox1 binding to Mcm1 at

ECB elements, I asked if the toxicity caused by YOX1 overexpression was suppressed by

concurrent overexpression of BCK2. Previous work on the CLB2 gene cluster, which is

122

Figure 4-6. Bck2 competes with Yox1 for interaction with Mcm1. (A) YOX1-TAP, YHP1-TAP and

MCM1-TAP strains carrying a pGAL-BCK2-FLAG plasmid (black bars) or vector (grey bars), were

individually grown in plasmid selective medium containing galactose (inducing conditions) to mid-log

phase. Cultures were harvested and anti-TAP ChIPs were performed using IgG sepharose resin and

analyzed for CLN3 promoter DNA using Q-PCR. The Y-axis represents enrichment of CLN3 promoter

DNA. Error bars reflect values obtained after multiple Q-PCR runs from the same experiment. (B) A WT

strain (Y7092) was co-transformed with a GAL-YOX1 (URA3) plasmid and Vector, GAL-CLN3, or GAL-

BCK2 (LEU2) plasmid. Transformant cultures of equivalent optical density were spotted in serial 5-fold

dilutions on selective medium that was either non-inducing (glucose) or inducing (galactose), and

incubated for 72 hr at 30°C. (C) A yeast two-hybrid bait strain (Y8930) carrying either plasmid ADH1-

GAL4 DB (vector) or plasmid ADH1-GAL4 DB-BCK2 Fragment 11 (Bck2) were mated to yeast two-hybrid

prey strain (Y8800) carrying either ADH1-GAL4 AD-MCM1WT

or ADH1-GAL4-MCM1V69E

to create

diploid yeast strains. Diploids were spotted in serial 10-fold dilutions on double plasmid selection medium,

or medium where growth is proportional to transcription of the ADE2 gene, and incubated for 48-72 hr at

30°C.

+ Vector

+ GAL-CLN3

+ GAL-BCK2

WT + GAL-YOX1

SD - U - L SGal - U - L

Mcm1-TAP Yox1-TAP Yhp1-TAP

+ pGAL-FLAG

+ pGAL-BCK2-FLAG

A

B

Vector + Mcm1V69E

Bck2 + Mcm1V69E

Vector + Mcm1WT

Bck2 + Mcm1WT

SD - L - Trp SD - L – Trp - Ade

C

123

expressed at the G2/M phase transition, has shown that Mcm1 acts as a common scaffold

for recruitment of Yox1 and the forkhead protein Fkh2 (Darieva et al. 2010). These

physical interactions with Mcm1 are mutually exclusive and are mediated by distinct

Yox1 and Fkh2 DNA binding sites that flank a central Mcm1 DNA binding site. When

bound to Mcm1 on promoters, Fkh2 recruits the positively acting co-regulator Ndd1 in

order to activate the CLB2 cluster genes (Koranda et al. 2000). Constitutive

overexpression of YOX1, which is toxic to cells (Pramila et al. 2002), inhibits Ndd1

binding to the Yox1-regulated SPO12 promoter (Darieva et al. 2010), consistent with a

mechanism based on mutual exclusivity between an activator and repressor. My

observation that overproduction of Bck2 leads to reduced Yox1 on the CLN3 promoter

(Figure 4-6A) suggested that the YOX1 dose-lethality phenotype might be suppressed by

concurrently overproducing BCK2. Indeed, I observed that overexpression of BCK2 was

able to significantly suppress the lethality caused by overexpressing YOX1 (Figure 4-6B).

Overexpression of CLN3 failed to rescue the YOX1 overexpression phenotype, suggesting

that the BCK2 rescue does not simply reflect an indirect effect of accelerated G1 phase.

This genetic observation is consistent with a competitive relationship between Bck2 and

Yox1 for interaction with the Mcm1 scaffold.

Mcm1 contains a hydrophobic pocket found on the surface of the MADS DNA-

binding domain, and mutation of this pocket by the introduction of a V69E mutation

disrupts interaction with Fkh2 (Tan and Richmond 1998; Boros et al. 2003; Darieva et al.

2010), and also prevents binding of Yox1 to Mcm1 (Darieva, Clancy et al. 2010). I

hypothesized that the competitive binding mechanism that allows Mcm1 to activate genes

transcribed at the G2/M transition also functioned for genes transcribed at the M/G1

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transition. Specifically, I wondered whether Bck2 might activate Mcm1 at ECB elements

through competition with Yox1 for binding to Mcm1. Consistent with my hypothesis, I

found that the two-hybrid-based interaction between Bck2 and Mcm1 was abolished in

the Mcm1V69E

mutant (Figure 4-6C). I conclude that Bck2 may act to remove Yox1 by a

competitive binding mechanism similar to that seen at G2/M phase promoters (Darieva,

Clancy et al. 2010).

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4.5 Discussion

My work identifies a previously unappreciated role for Bck2 in the regulation of

early G1-specific transcription in budding yeast. In contrast to the CLB2 gene cluster, no

positively acting partner protein had yet been found that cooperates with Mcm1 to

regulate M/G1-expressed genes. I describe several observations that suggest that Bck2

functions to control Mcm1 activity on promoter elements to ensure the proper regulation

of early G1 events. First, the ability of Bck2 to activate expression of SWI4, CDC6, and

CDC47 is consistent with the ability of Bck2 to promote the G1/S transition through

activation of genes in an SBF/MBF-independent manner. Second, the requirement for

intact Mcm1-binding ECB elements within the promoters of CLN3 and SWI4 is

consistent with physical interaction between Bck2 and Mcm1 by yeast two-hybrid

analysis. Third, dosage-dependent competition between Bck2 and Yox1 for a common

binding pocket on Mcm1 suggests that M/G1- and G2/M-regulated genes may be

controlled by similar mechanisms. Because Bck2 also activates CLB2 expression, I

discuss the possibility that Bck2 regulates other Mcm1-dependent genes. These findings

are summarized in Figure 4-7A. At the CLN2 promoter, Bck2 may also function through

Mcm1, although the possibility remains that Bck2 may further function through Swi4

with or without Swi6 (Figure 4-7A; right panel). However, the specific mechanism by

which Bck2 might directly activate CLN2 is not known.

4.5.1 SBF/MBF-independent activation by Bck2

My finding that Bck2 activates Mcm1-regulated genes in early G1 phase is

consistent with previous observations that Bck2 activates G1 phase gene expression in a

126

Figure 4-7. How Bck2 activates the G1/S phase transition. (A) Possible mechanism by which Bck2

activates M/G1- and G1/S-regulated promoters. In the case of MCM1 cluster genes activated at the M/G1

transition, such as CLN3 and SWI4 (left panel), Yox1 inhibits Mcm1 activity on ECB promoter elements

(containing Yox1 and Mcm1 (P) sites) throughout the cell cycle by binding to a hydrophobic pocket on

Mcm1 that precludes binding by Bck2. In early G1 phase, Bck2 displaces Yox1 from the hydrophobic

pocket on Mcm1 to activate gene expression. In the case of CLN2 cluster genes activated at the G1/S

transition (right panel), Bck2 activates an unknown promoter-bound protein that may be Mcm1 or Swi4.

(B) Activation of the G1/S transition by Bck2. A CDK-independent pathway for activation of the G1/S

transition is defined by Bck2 activity (red arrows). Bck2, in pre-START cells, initiates expression of SWI4

and CLN3, which encode important constituents of the CDK-dependent positive feedback ʻswitchʼ for

START that activate CLN2 (and CLN1) expression predominantly through activation of promoter-bound

SBF/MBF. Bck2 also induces CLN2 expression directly through physical association with other factors,

likely Mcm1 or Swi4. Bck2 also induces at least one other gene, XXX, which is not CLN1.

A

B Bck2

Mcm1

SWI4

?

CLN2

CLN3

S

G1 CDK-

Independent

CDK-Dependent

XXX

M/G1 G1/S

Yox1

Yox1 P

Mcm1 ++

P

Mcm1 ++

SCB

Swi4

?

ECB

SBF/MBF

Bck2

Bck2

127

pathway distinct from that involving Cln-Cdc28 activation of promoter-bound SBF/MBF.

First, there is no evidence that CDK activity is required for activation of M/G1-expressed

genes. For example, many genes with peak expression at the M/G1 transition are induced

in a cdc28-13 mutant (Cho et al. 1998), and some of these genes (SWI4, FAR1, ASH1,

TIP1, TEC1, CDC6 and CDC47) are also activated by Bck2 (Chapter 4) (Martin-Yken et

al. 2002) in a SBF/MBF-independent manner (Ferrezuelo, Aldea, and Futcher 2009).

Second, high-copy BCK2 activates SBF/MBF target genes in a cdc28-4 mutant (Wijnen

and Futcher 1999) and suppresses the G1-arrest of a cln1Δcln2Δcln3Δ strain (Epstein and

Cross 1994). These activities likely reflect increased SWI4 expression, because Bck2

activates SWI4 transcription (Di Como, Chang, and Arndt 1995; Martin-Yken et al.

2002), and high-copy suppression by BCK2 of the G1-arrest of a cln1Δcln2Δcln3Δ strain

requires SWI4 (Epstein and Cross 1994). Third, Bck2 can activate several natural

SBF/MBF target gene promoters in the absence of either SWI4 or MBP1 (Wijnen and

Futcher 1999), or the elements that SBF/MBF bind (Di Como, Chang, and Arndt 1995).

For example, the UAS2 region of CLN2, which completely lacks SCB or MCB elements

(Stuart and Wittenberg 1994), responds to BCK2 dosage (Di Como, Chang, and Arndt

1995) and is induced in a cdc28 mutant (Stuart and Wittenberg 1994), indicating that at

UAS2 Bck2 likely acts through another DNA-bound factor distinct from SBF/MBF.

Moreover, even a point-mutated version of UAS1 lacking SCB/MCB elements or UAS2

is sufficient for proper expression of CLN2 in a strain lacking Cln-Cdc28 activity (Stuart

and Wittenberg 1995), suggesting that Bck2 might activate the same factor at UAS1 and

UAS2. Interestingly, Mcm1 binds to the CLN2 promoter in two regions (Bai, Ondracka,

128

and Cross 2011) that fall within UAS1 and UAS2 (Stuart and Wittenberg 1994), which

are both responsive to BCK2 dosage (Di Como, Chang, and Arndt 1995).

4.5.2 Bck2 as an activator of G2/M genes

Despite evidence suggesting that Bck2 may be an activator of CLB2 expression

(Figure 4-2, 4-4), the possibility that Bck2 directly regulates CLB2 cluster genes in vivo is

unlikely for several reasons. First, Mcm1 alone can protect the entire ECB sequence

from nuclease digestion (Kuo and Grayhack 1994), suggesting that Mcm1 can be the only

DNA-binding component necessary for ECB activation. In contrast, G2/M-specific

promoter elements within CLB2 cluster genes require at least one other protein to bind a

site adjacent to Mcm1 (McInerny et al. 1997). Accordingly, at least for M/G1-regulated

genes, Yox1 can repress Mcm1 activity in the absence of an adjacent Yox1 site on DNA

(Pramila et al. 2002). In contrast, Yox1 DNA-binding sites are present in many G2/M-

regulated genes in the CLB2 cluster, such as SPO12 but not CLB2 itself, and mutation of

Yox1 DNA-binding sites alters the precise timing of gene expression (Pramila et al.

2002). Second, unlike the CLB2 cluster, M/G1 promoters of the MCM1 cluster (Spellman

et al. 1998) do not contain Fkh1 or Fkh2 DNA binding sites, and deletion of FKH1 or

FKH2 does not affect expression of the M/G1 cluster (Zhu et al. 2000). Third, Mcm1, but

not Fkh1, Fkh2 or Ndd1, can be localized to promoters of M/G1-regulated genes such as

MCM3, CDC46, MCM6, and CDC6 (Simon, Barnett et al. 2001), indicating that

induction of M/G1-regulated genes by Mcm1 (Pramila et al. 2002) depends on other

positively-acting proteins besides Fkh1, Fkh2 or Ndd1. Fourth, artificial overproduction

of Clb2 can activate genes expressed at M/G1, in addition to its normal targets at G2/M

129

(Spellman et al. 1998), indicating that overexpression of CLB2 can lead to off-target

effects. Thus, induction of G2/M genes due to overexpression of BCK2 may also be an

off-target effect outside of normal activation of M/G1 genes (Ferrezuelo, Aldea, and

Futcher 2009).

4.5.3 How Bck2 activates the G1/S transition

The findings of this study, in combination with the expansive literature as it

relates to cell cycle commitment at START, are consistent with the model shown in

Figure 4-7B. Bck2, in pre-START cells, initiates expression of gene products that

constitute the positive feedback ‘switch’ for START (Skotheim et al. 2008). The raw

materials for the ‘switch’, such as Swi4 and Cln3, are steadily produced by activated

Bck2 on ECB elements, while certain components (Swi4 and Cln3) are likely prevented

from prematurely triggering the ‘switch’ because they are buffered by inhibitory factors

such as active Far1 and inactive Whi5/Stb1, which require removal by the CDK-

dependent pathway (Figure 1-3). Indeed, Cln3 has to overcome Far1 to trigger Cln-

Cdc28 activation, which then turns on SBF/MBF (Alberghina et al. 2004). Because this

model is predicated on Bck2 acting on Mcm1 rather than SBF, certain genes expressed

precisely at the G1/S transition, such as HO, would not be responsive to Bck2 activity in

early G1 phase. Unlike CLN2, however, both the HO promoter or isolated SCB elements

strictly depend upon Swi4 and Swi6 for their activity (Breeden and Nasmyth 1987), and

are therefore inactive until just at the Cdc28-dependent point of START (Nasmyth 1985;

Breeden and Nasmyth 1985). In contrast to the HO promoter, the Mcm1-binding sites in

the CLN2 promoter could allow a large amount of inactive Cln2 protein to accumulate,

130

which could be buffered until late G1 phase when the positive feedback ‘switch’ is

triggered. In particular, the nucleolytic activity of the HO protein might not be buffered

adequately – a problem that can be circumvented by tying the expression of HO to the

activity of SBF by the CDK-dependent pathway, exclusively. Indeed, HO expression is

the most tightly regulated and START-dependent gene in S. cerevisiae (Kostriken and

Heffron 1984). Importantly, because BCK2 overexpression bypasses the lethality of a

cln1Δcln2Δcln3Δ strain (Epstein and Cross 1994), Bck2 likely induces expression of at

least one other gene, XXX, which is not CLN1 (Figure 4-7B). In summary, the

importance of Bck2 in the cell likely reflects an ill-defined environmental checkpoint that

links production of the cell-cycle-commitment machinery to an acceptable cocktail of

proliferation signals.

131

Chapter 5

Summary and Future Directions

132

Chapter 5 - Summary and future directions

5.1 Thesis summary

The CDK-dependent model for cell cycle progression posits that cyclin-dependent

kinases drive the molecular events necessary for cell division. A key driver of CDK-

mediated processes is periodic transcription, wherein groups of genes are co-ordinately

regulated at specific points during progression of the cell cycle. Underlying periodic

transcription itself is a regulatory circuit comprised of various transcriptional factors that

are sometimes direct targets of CDKs. Periodic transcription plays a prominent role in

the G1/S phase transition, a key regulatory period during which cell cycle exit or

commitment can occur. The G1/S transition has been intensively studied and it is clear

that Cdk1-dependent activation of promoter-bound SBF/MBF transcription factors is

necessary for expression of genes required for cell cycle entry. SBF/MBF activation is a

complex process, and cells can still transit G1/S in the absence of SBF/MBF activity (i.e.

swi6Δ), indicating that there are alternate methods of passing START. Moreover, several

CDK-independent pathways involved in ion homeostasis impinge upon G1 phase

progress, suggesting that integration of environmental signals is very important for entry

into the cell cycle. One important protein involved in CDK-independent cell cycle

regulation is Bck2, an enigmatic activator of the G1/S transition that might partly function

through SBF/MBF. The major objective of my thesis was to gain insight into how Bck2

activates START.

As noted earlier, the Cln3-Cdc28 form of CDK is important for the activation of

the related transcription factors SBF (Dirick, Bohm, and Nasmyth 1995; Stuart and

Wittenberg 1995; Wijnen, Landman, and Futcher 2002) and MBF (Wijnen, Landman,

133

and Futcher 2002; de Bruin, Kalashnikova, and Wittenberg 2008). However, a cln3Δ

strain is still viable, and SBF/MBF-dependent transcription still occurs in a cln3Δ strain,

which suggests alternative mechanisms must exist to activate SBF/MBF (Nasmyth and

Dirick 1991; Stuart and Wittenberg 1995). Indeed, a genomic screen aimed at identifying

activators of SBF and MBF dependent transcription identified many genes, often with no

known connections to CLN3 or CDC28, which strongly impacted SBF and MBF activity

(H. Friesen, M. Costanzo; unpublished). In Chapter 2, I asked if genes identified in this

screen encoded proteins that physically associate with SBF and MBF. Out of seventeen

candidates tested, I found four novel SBF- and MBF-interacting proteins: Elm1, Rtf1,

Ykr077w, and Bck2. I focused on Bck2 because genetic studies strongly implicated

Bck2 as a CDK-independent activator of SBF and MBF for which a clear mechanism was

lacking. I observed that Bck2 not only interacted with the promoters of SBF- and MBF-

regulated genes, but that Bck2 had the property of transcriptional auto-activation,

consistent with its role as a genetic activator of the G1/S phase transition.

Whether the localization of Bck2 to SBF/MBF-specific promoters (Chapter 2)

required the presence of SBF and MBF was not pursued, which left open the possibility

that Bck2 might interact with G1-specific promoters through some other DNA-binding

factors. Because the identification of Bck2-interacting proteins using biochemical assays

has met with limited success, I decided to perform a yeast-two-hybrid screen to find

novel Bck2-interacting proteins. In Chapter 3, I describe a systematic truncation analysis

of Bck2 which uncovered a region of Bck2 sufficient to perform its essential function but

that did not auto-activate transcription in a two-hybrid bait strain. I used this truncated

version of Bck2 as a bait protein in a systematic two-hybrid screen and discovered six

134

novel Bck2-interacting proteins: Mcm1, Mot3, Yap6, Tpd3, Std1, and Mth1. I present

evidence that both Mot3 and Bck2 activate several genes involved in metabolism,

suggesting a shared role in integrating metabolic signals with cell cycle progression.

Mcm1 is an essential protein which binds to early G1 promoters such as CLN3 and

SWI4 through Early Cell cycle Box (ECB) elements. The physical interaction of Bck2

with Mcm1 (Chapter 3) suggested that the requirement for Bck2 in activation of genes at

the G1/S transition may reflect a primary defect in CLN3 and SWI4 expression in early G1

phase, which subsequently impairs transit through G1/S. In Chapter 4, I show that Bck2

is required for the proper expression of early G1 phase genes and Bck2 localizes to the

promoters of early G1 phase genes in an ECB-dependent manner. In early G1 phase,

Mcm1 is inhibited by association with two repressors, Yox1 and Yhp1, and gene

activation ensues once repression is relieved by an unknown activating signal. Therefore,

in Chapter 4 I ask whether Yox1-mediated repression is relieved by Bck2. I provide

evidence that Bck2 removes Yox1 from the CLN3 promoter. In addition, I show that

BCK2 overexpression can suppress the toxicity of YOX1 overexpression, which suggests

that Bck2 might compete with Yox1 for access to Mcm1. Consistent with this model, I

show that the Bck2-Mcm1 interaction requires valine 69 on Mcm1, a residue known to be

involved in the interaction with Yox1 at G2/M-regulated promoters. I envision a model

whereby Yox1 and Bck2 compete for access to the Mcm1-ECB scaffold on early G1

phase promoters. As early G1 phase is a critical time for assessing environmental

conditions, the cell may use nutrient-regulated dosage/activity of Bck2 as a means of co-

ordinately activating the initial suite of genes required for cell cycle entry.

135

5.2 Future directions

My discovery that Bck2 interacts with regulators of metabolic genes (Chapter 3)

and also activates M/G1-regulated genes (Chapter 4) suggests that Bck2 may play a role

in integrating signals required for re-entry of quiescent cells into the cell cycle (G0/G1

transition). The M/G1 transition of cycling cells is analogous to the G0/G1 transition of

quiescent cells because environmental signals are assessed in both cases. For example,

mating pheromone in the extracellular environment causes G1-arrest whether cells are

released into G1 phase from nocodazole-induced mitotic arrest or from rapamycin-

induced arrest in early G1 phase (Barbet et al. 1996). The pheromone-induced G1 arrest

pathway of S. cerevisiae is controlled by a mating MAPK cascade (Herskowitz 1995) that

leads to sequential activation of a kinase cascade (Ste20, Ste11, Ste7, Fus3 and Kss1)

(Elion 1995), which ultimately leads to activation of Far1, a Cln/Cdc28-inhibitor, by the

Fus3 kinase (Elion, Satterberg, and Kranz 1993; Peter et al. 1993; Tyers and Futcher

1993; Henchoz et al. 1997; Jeoung, Oehlen, and Cross 1998; Breitkreutz, Boucher, and

Tyers 2001). Importantly, Far1 can repress only Cln-Cdc28 (Kurjan 1993; Bardwell et

al. 1994), but not Clb-Cdc28 complexes (Peter and Herskowitz 1994), suggesting that

high Clb-Cdc28 activity may substitute for Cln-Cdc28 activity in overcoming the G1

arrest of cln1Δcln2Δcln3Δ cells (Richardson et al. 1989; Cross 1990). Indeed,

overexpression of either CLB5 (Epstein and Cross 1992; Schwob and Nasmyth 1993) or

certain CLB2 alleles (Amon, Irniger, and Nasmyth 1994) allows cells to pass START in

the absence of CLN function. Interestingly, Bck2 activates CLB5 (Di Como, Chang, and

Arndt 1995) and CLB2 (Di Como, Chang, and Arndt 1995; Ferrezuelo, Aldea, and

Futcher 2009) expression in an SBF-independent manner, suggesting that high-copy

136

BCK2 may overcome the G1 arrest of a cln1Δcln2Δcln3Δ mutant (Epstein and Cross

1994) through induction of CLB expression. In contrast, degradation of the Clb-Cdc28

repressor, Sic1, is triggered by CLN cyclins (Schwob et al. 1994; Schneider, Yang, and

Futcher 1996; Verma et al. 1997), suggesting that the pheromone-resistance seen in

strains overexpressing CLNs may reflect precocious Sic1 degradation. Intriguingly, high-

copy BCK2 can overcome the G1-arrest of a mutant with a hyperactive pheromone

pathway (Edwards et al. 1997), which suggests that Bck2 can promote proliferation under

conditions where Far1 is inhibiting Cln-Cdc28 and Sic1 is inhibiting Clb-Cdc28 --

perhaps because pheromone activates Bck2.

As noted in Chapter 1, several lines of evidence suggest that nutrients regulate

Bck2 activity in G1 phase. Cells lacking BCK2 grow slowly (Smith et al. 1996) in

minimal media (Giaever et al. 2002) and are defective in SWI4 and CLN3 expression

(Chapter 4). Interestingly, 85% of genes induced by overexpressed BCK2, such as CLN1,

CLN3, SWI4 and CDC28 (Martin-Yken et al. 2002), are downregulated under conditions

of glucose, nitrogen or phosphate deprivation (Klosinska et al. 2011), suggesting that

nutrients may signal through Bck2 to activate expression of these genes. Moreover,

greater than 25% of genes induced by overexpressed BCK2 (Martin-Yken et al. 2002) are

also induced upon exit from quiescence, including RPA135, TYE7, RPS3, AAH1,

RPL22A, RPL22B, CLN3, RPL25, and RPL38 (Radonjic et al. 2005). In contrast to Bck2

target genes, BCK2 expression is induced by nutrient limitation (Klosinska et al. 2011) or

in stationary phase (Gasch et al. 2000), consistent with a role for Bck2 in nutrient-

regulated gene expression at G0/G1. Intriguingly, the activation of Mcm1 by Bck2 in S.

cerevisiae appears similar to the activation of SRF by p62TCF

in human cells (Treisman

137

and Ammerer 1992). For example, Bck2 might activate promoter-bound Mcm1 by

binding to it only when nutrients are rich, mimicking the behaviour of certain SRF

coregulatory proteins, such as p62TCF

. Before my work identifying Bck2 as a Mcm1 co-

factor, no positively-acting partner of Mcm1 was known for activation of M/G1 genes in

S. cerevisiae. Collectively, my findings suggest that Bck2 and Mcm1 may function

together in nutrient-regulated transcription in ways analogous to specific activators of

SRF in humans.

5.2.1 Does Bck2 activate MAT cluster genes at M/G1?

The MAT cluster of genes (Spellman et al. 1998) are induced by mating

pheromone and encode transcription factors that induce genes required for cell-type

determination and mating. In the absence of pheromone, MAT genes are expressed

periodically at the M/G1 transition in a manner dependent on both the transcription factor

Ste12 binding to pheromone-response elements (PRE) (Oehlen, McKinney, and Cross

1996) and Mcm1 binding to ECB elements (Wittenberg and Reed 2005). For example,

the gene encoding the pheromone receptor, STE2 (Hwang-Shum et al. 1991), and the

gene encoding the inhibitor of Cln-Cdc28 complexes, FAR1 (Oehlen, McKinney, and

Cross 1996), contain hybrid promoters with PRE elements next to ECB elements.

However, the requirement for Ste12 in the periodic expression of MAT cluster genes is

not well understood. Some mating genes are induced in response to pheromone by Ste12

alone (Dolan, Kirkman, and Fields 1989; Sengupta and Cochran 1990; Hagen,

McCaffrey, and Sprague 1991; Hwang-Shum et al. 1991), whereas other mating genes

are regulated by Ste12 together with Mcm1 (Errede and Ammerer 1989; Primig, Winkler,

138

and Ammerer 1991; Kirkman-Correia, Stroke, and Fields 1993; Oehlen, McKinney, and

Cross 1996). In some cases, even basal expression of several MAT cluster genes, such as

FUS1 and AGA1, is strongly Ste12-dependent, although Mcm1 also regulates these genes

(Oehlen, McKinney, and Cross 1996).

Many genes that are induced by BCK2 overexpression in an SBF/MBF-

independent manner (Ferrezuelo, Aldea, and Futcher 2009) are genes of the MAT cluster,

such as STE12, FAR1, TEC1, KAR4, ASH1, STE2, STE6, AGA1, FUS1 and SST2.

Induction of many of these MAT cluster genes by BCK2 overexpression is Ste12-

dependent (Ferrezuelo, Aldea, and Futcher 2009), indicating that Bck2 might generally

require Ste12 for activation of all MAT cluster genes. Ste12 neither binds to nor activates

ECB elements (Oehlen, McKinney, and Cross 1996), suggesting that Bck2/Ste12-

coregulation requires both a PRE and ECB sequence. In addition to PRE and ECB

sequences, Mot3 binding sites may also be important. Deletion of MOT3, which I

identified as a Bck2-interacting protein by Y2H analysis (Chapter3), induces the MAT

cluster gene FUS1, while MOT3 overexpression decreases FUS1 expression (Grishin et

al. 1998). Likewise, deletion of the pheromone-response pathway kinase KSS1 induces

FUS1 expression, while overexpression of KSS1 decreases FUS1 expression (Bardwell et

al. 1998). In contrast, overexpression of BCK2 induces the expression of the MAT cluster

gene FUS1 (Edwards et al. 1997; Ferrezuelo, Aldea, and Futcher 2009).

These observations implicate Kss1 and Mot3 as repressors of MAT cluster genes,

although they may function via distinct mechanisms (Bardwell et al. 1998; Grishin et al.

1998). In particular, in kss1Δ cells, active Fus3 likely compensates for the absence of

Kss1 to phosphorylate and activate Ste12, allowing induction of FUS1 and other Ste12

139

target genes. However, in mot3Δ cells, active Ste12 is sufficient for basal and

pheromone-induced activation of FUS1 expression, because the pheromone signalling

pathway is intact in mot3Δ cells. Consistent with this idea, deletion of MOT3 fails to

increase FUS1 expression in strains mutated for upstream components of the pheromone-

signalling pathway (Grishin et al. 1998). Curiously, MOT3 (Grishin et al. 1998) or BCK2

(Wijnen and Futcher 1999) overexpression decreases sensitivity to pheromone, whereas

mot3Δ cells (Grishin et al. 1998) or bck2Δ cells (Di Como, Chang, and Arndt 1995)

increase sensitivity to pheromone (Grishin et al. 1998). One testable model stemming

from these data is that pheromone activates a promoter-bound complex of Bck2 and

Mot3, causing MAT cluster genes to be induced.

Collectively, my work and the observations summarized above underscore two

related puzzles. First, how does Bck2 simultaneously promote cell cycle progression and

arrest? Here the answer may be complex. As noted earlier, overexpression of Bck2

induces expression (Ferrezuelo, Aldea, and Futcher 2009) of cell cycle arrest factors such

as FAR1 but also genes required for recovery from cell cycle arrest, such as SST2 (Kurjan

1993; Bardwell et al. 1994; Dohlman et al. 1996; Dohlman and Thorner 1997; Parnell et

al. 2005). However, increased production of Far1 due to BCK2 overexpression need not

lead to cell cycle arrest, because overexpression of FAR1 in the absence of pheromone is

not sufficient to inhibit Cln2-Cdc28 activity (Chang and Herskowitz 1992), suggesting

that abundant inactive Far1 may be inconsequential to cell cycle progression. The second

puzzle involves understanding how Bck2 and Mot3 act antagonistically to regulate MAT

cluster gene expression. My work suggests that Bck2 and Mot3 have supporting roles in

the regulation of metabolic genes, such as CYC1 and RPL39 (Chapter3) (Grishin et al.

140

1998), but may have opposing roles at MAT cluster genes. For example, basal

transcription of the pheromone-inducible FUS1, SST2 and AGA1 promoters is decreased

in a MOT3-overexpressing strain (Grishin et al. 1998), while it is increased in a BCK2-

overexpressing strain (Ferrezuelo, Aldea, and Futcher 2009). However, as noted earlier,

BCK2 and MOT3 mutants are phenotypically similar with respect to pheromone

sensitivity. Cells overexpressing BCK2 or MOT3 (Grishin et al. 1998) increase

pheromone resistance, while bck2Δ and mot3Δ (Grishin et al. 1998) mutants are

pheromone sensitive. Thus, the roles of Bck2 and Mot3 may be different at FUS1

relative to other promoters.

In order to better understand how Bck2 promotes expression of both arrest and

proliferation genes, which MAT cluster genes are activated by Bck2 and whether this

occurs co-ordinately with Ste12, Kss1 and Mot3 on promoters should be determined

using a multi-pronged approach: [1] To ask whether Bck2 activates all MAT cluster

genes, it will be important to observe the expression of STE12, FAR1, TEC1, KAR4,

ASH1, STE2, STE6, AGA1, FUS1 and SST2 as a function of BCK2 overexpression or

deletion in the presence and absence of pheromone; [2] next, the induction of MAT

cluster genes by overexpressed BCK2 in ste12Δ, kss1Δ or mot3Δ mutants should be

assessed, which will indicate whether Ste12, Kss1 or Mot3 function in a Bck2-specific

pathway; [3] does Bck2 physically interact with Ste12 and Kss1? Using a series of ChIP

assays, Bck2 localization to MAT cluster promoters can be tested in ste12Δ and kss1Δ

strains, and the potential interaction of Bck2 with Ste12 or Kss1 can be tested by the Y2H

assay method; [4] if the extent of MAT cluster gene expression in bck2Δste12Δ,

bck2Δkss1Δ, and bck2Δmot3Δ strains is tested, it would help determine the genetic

141

contributions of these genes to MAT cluster activation; [5] testing the pheromone

resistance caused by overexpressed KSS1 (Ma, Cook, and Thorner 1995) in a ste12Δ,

mot3Δ or bck2Δ strain, would help establish epistatic relationships amongst these genes;

[6] the transcriptional repression of FUS1 by overexpressed MOT3 should be tested in

ste12Δ, kss1Δ and bck2Δ strains to see if Ste12, Kss1, and Bck2 are important in Mot3

function; [7] using a ChIP assay to examine localization of Bck2 to the FUS1 promoter in

ste12Δ, kss1Δ and mot3Δ strains in the presence and absence of pheromone can be

performed to reveal whether the amount of promoter-associated Bck2 is increased by

pheromone and whether other proteins are required for this localization; [8] Ste12 (Chou,

Lane, and Liu 2006), Kss1 (Pokholok et al. 2006), and Mot3 (Montanes, Pascual-Ahuir,

and Proft 2011) have all been localized to promoters in vivo using the method of ChIP.

Thus, the ChIP method should be used to assess whether Ste12, Kss1 or Mot3

localization to the FUS1 promoter is altered under conditions of BCK2 overexpression,

which will reveal how promoter-bound complexes are influenced by Bck2 dosage.

142

5.2.2 Bck2 functional analogs in humans.

S. cerevisiae Mcm1 and human SRF are homologous DNA-binding proteins,

which regulate nutrient-dependent transcription using similar mechanisms (Figure 5-1).

Serum-starved mammalian cells re-enter the cell cycle when subjected to serum

stimulation (Cooper 2003) and induce serum-responsive genes called immediate early

genes (IEG) (Cochran et al. 1984) required for re-entry into the cell cycle (Treisman

1985). Similarly, S. cerevisiae quiescent G0 cells stimulated with nutrients exit from

quiescence into the G1 phase of the cell cycle (Werner-Washburne et al. 1993; Gray et al.

2004) by expressing many genes required for cell cycle re-entry, such as SWI4 and CLN3

(Gray et al. 2004; Radonjic et al. 2005). The promoters of IEGs contain serum response

elements (SRE) bound by the serum response factor (SRF) (Treisman 1987; Norman et

al. 1988). Similarly, the promoters of early G1-expressed genes contain early cell-cycle

box elements (ECB) bound by Mcm1 (McInerny et al. 1997). Serum modulates the

activities of SRF coregulators rather than causing SRF to bind DNA (Messenguy and

Dubois 2003; Cen, Selvaraj, and Prywes 2004). For instance, in vivo footprinting at the

c-fos SRE indicates that SRF is bound to DNA before, during, and after growth factor

stimulation and target induction (Herrera, Shaw, and Nordheim 1989; Konig 1991).

Similarly, nutrients modulate the activity of Mcm1 coregulators rather than causing

Mcm1 to bind DNA (Yuan, Stroke, and Fields 1993; Mai, Miles, and Breeden 2002). For

example, Mcm1 is bound to ECB elements of CLN3 and SWI4 before, during, and after

activation (Pramila et al. 2002).

143

Figure 5-1. Early G1 phase signaling in human cells and budding yeast. (A) SRF bound to SREs can

be activated by either p62TCF

or myocardin/MKL proteins, but their binding to SRF is mutually exclusive.

p62TCF

is activated by ERK1/2 through a MAPK phosphorylation cascade. A second SRF activation

pathway involves RhoA, which is required for TCF-independent activation by serum. Serum activates

RhoA, which activates G protein coupled receptors, which causes release of MKL1 from a G-actin complex

and subsequent translocation of MKL1 to the nucleus. Inducible phosphorylation of MKL1 has been

observed, suggesting that RhoA induction of an unknown MKL1-kinase may be a critical regulatory step.

(B) The analogous pathway in budding yeast. Mcm1 bound to P-sites can be activated by either Fkh2-

Ndd1 or Bck2, and their binding may be mutually exclusive. Fkh2-Ndd1 is activated by Clb-Cdk1 and

Cdc5 phosphorylation. The activators of Bck2 are not known but could include the kinases Kss1 and Sln1.

Like myocardin/MKL, Bck2 is a phosphoprotein that might also translocate to the nucleus.

HUMAN

Ets SRE IEG

c-fos c-jun c-myc

srf

ELK1 SAP1

NET

ERK1/2

P

TCFp62

Myocardin MKL1

MKL2 MKL

P

Raf

Ras

MEK1

EGF

BUDDING

YEAST

Fkh2 P ECG

SWI4 CLN3 MCM1

Clb2

Cdk1

Fkh2

P

Ndd1

Skn7

Sln1

?

Bck2

P Cdc5

Foxo4

Yox1

MKL (nuclear)

Actin

?

RhoA

Ste7

Ste11

Kss1

SRF

Mcm1

?

144

The general regulatory similarities between Mcm1 and SRF are substantiated by

functional similarities between the two proteins at the amino acid sequence level. Mcm1

binds the human c-fos SRE in vitro and increases c-fos SRE activity in vivo (Passmore,

Elble, and Tye 1989), suggesting a high level of functional similarity. Indeed, the portion

of Mcm1 with the highest identity to SRF (amino acids 18-98) is sufficient for DNA

binding and viability (Christ and Tye 1991; Darieva et al. 2010). The same region within

SRF is essential for both DNA-binding and dimerization (Norman et al. 1988).

Moreover, the regions Mcm1 and SRF use to physically interact with co-factors are

similar (Schroter et al. 1990; Primig, Winkler, and Ammerer 1991; Mueller and

Nordheim 1991; Treisman and Ammerer 1992) – specifically, Mcm1 and SRF contain a

hydrophobic pocket on the surface of their DNA-binding domains (Tan and Richmond

1998; Hassler and Richmond 2001) that is required for interaction of Fkh2 or α2 with

Mcm1, and for interaction of SAP-1 or ELK1 with SRF (Boros et al. 2003). Finally,

analogous residues are also important for processes such as DNA bending, DNA binding

affinity, transcriptional activation and cell growth (Acton et al. 2000). Residues in SRF

required to bind DNA when complexed with SAP-1 are similar to the residues in Mcm1

required to bind DNA when complexed with α1 (Mo et al. 2001).

In humans, two major signalling pathways control the activity of SRF target gene

expression in response to serum. Activation of the Ras-Raf MAPK pathway leads to

phosphorylation of ternary complex factor (TCF), also known as p62TCF

(Janknecht et al.

1993; Buchwalter, Gross, and Wasylyk 2004), encoded by ELK1, SAP1 (ELK4), or NET

(ELK3, SAP2, and ERP) (Lee, Vasishtha, and Prywes 2010), which are Ets-family

proteins that share similar domain structure (Yang, Whitmarsh et al. 1998; Yang, Yates et

145

al. 1998; Ducret et al. 2000; Hassler and Richmond 2001). ELK1 binds constitutively to

SRE target sites and is associated with the transcriptional coactivators CREB-binding

protein and/or p300 (Janknecht and Nordheim 1996; Nissen, Gelly, and Hipskind 2001;

Li et al. 2003).

Serum-dependent induction of SRE reporter genes occurs even in the absence of

the p62TCF

pathway (Johansen and Prywes 1995). The other major pathway for activation

of SRF/SRE is through activation of receptors that signal to the RhoA GTPase to induce

actin polymerization (Hill, Wynne, and Treisman 1995). RhoA signalling then causes

activation of one of the three SRF coregulatory proteins of the megakaryoblastic

leukemia (MKL) family (Miralles et al. 2003): myocardin, MKL1 and MKL2 (Wang et

al. 2001; Sasazuki et al. 2002; Wang et al. 2002; Selvaraj and Prywes 2003; Cen et al.

2003; Miralles et al. 2003). Whereas myocardin is expressed in cardiac and smooth

muscle cells, MKL1 and MKL2 are ubiquitously expressed (Wang et al. 2001; Wang et

al. 2002; Du et al. 2003; Cen et al. 2003). An important aspect of MKL1/2 regulation is

translocation from the cytoplasm to the nucleus, where it forms a complex with SRF to

activate gene expression (Sotiropoulos et al. 1999; Miralles et al. 2003).

Although both Fkh2 and Bck2 physically bind and activate Mcm1, several lines

of evidence suggest that Bck2 is the functional analog of myocardin/MKL proteins rather

than Fkh2, which is more analogous to p62TCF

factors. First, whereas Mcm1-Fkh2 acts in

G2/M, the SRF-TCF complex acts at G1 (Treisman 1994), consistent with the proposed

role of Bck2 at the M/G1 and G0/G1 transitions. Second, the CLB2 promoter (G2/M) has a

Fkh2-binding site adjacent to the Mcm1-binding site (Pramila et al. 2002; Darieva et al.

2010), similar to the c-fos SRE, which has a p62TCF

binding site adjacent to the SRF-

146

binding site (Treisman, Marais, and Wynne 1992; Shore et al. 1996; Sharrocks et al.

1997). In contrast, M/G1-regulated promoters of the MCM1 cluster (Spellman et al.

1998) do not have obvious Fkh1 or Fkh2 binding sites adjacent to the Mcm1-binding site

(Zhu et al. 2000), similar to myocardin/MKL-regulated promoters, which lack obvious

ternary complex factor binding sites (Selvaraj and Prywes 2003). Third, both Bck2

(Figure 3-2) and myocardin/MKL proteins contain strong transcriptional activation

domains in their C-terminal regions, and deletion of their N-terminal regions increases

activation by GAL4 fusion proteins (Wang et al. 2001; Cen et al. 2003; Selvaraj and

Prywes 2003; Du et al. 2004). Fourth, both Bck2 (Martin-Yken et al. 2002; Ferrezuelo,

Aldea, and Futcher 2009) and MKL1/2 (Miralles et al. 2003; Cen et al. 2003) activate

expression of the same factors (i.e. MCM1 and srf) they physically associate with on

target gene promoters (i.e. Mcm1 and SRF). Fifth, like the increased binding of MKL to

target promoters in serum-rich media (Lee, Vasishtha, and Prywes 2010), my preliminary

evidence suggests that binding of Bck2 to target promoters may also be increased in

nutrient-rich media (Figure 5-2). In contrast to Bck2, binding of Mcm1 to ECB elements

is higher in the presence of rapamycin (Figure 5-2), which is consistent with its increased

localization to promoters in cells grown on poor nutrient sources (Mai, Miles, and

Breeden 2002). Finally, both ELK1 p62TCF

and MKLs bind to the same hydrophobic

pocket in the SRF DNA binding domain such that their binding to SRF is mutually

exclusive (Miralles et al. 2003; Wang et al. 2004; Zaromytidou, Miralles, and Treisman

2006), a mode of regulation that likely extends to other SRF-binding proteins (Cen,

Selvaraj, and Prywes 2004). In Chapter 4, I provide evidence that Bck2 might also bind

147

Figure 5-2. Localization of Bck2 to early G1 phase promoters in the presence of rapamycin. Mcm1-

TAP and Bck2-TAP strains were grown 30°C to early-log phase at in YPD (rich medium), after which to

one set of cultures for a given strain was added rapamycin (RAP). Cultures were allowed to grow for an

additional 3 hours, after which they were harvested for ChIP analysis. Anti-TAP ChIPs were analyzed for

SWI4 and CLN3 promoter DNA by Q-PCR. The Y-axis measures enrichment of promoter DNA for the

target gene indicated relative to enrichment of non-promoter DNA from an untranscribed region of

chromosome II.

148

the same region of Mcm1 that binds Fkh2 and Yox1 in a mutually-exclusive manner

(Darieva et al. 2010).

To better understand the role of Bck2 in nutrient-sensing, I will first determine the

levels of Bck2 protein relative to transcript levels in cells released from stationary phase

or grown in media containing various sources of carbon (glycerol, acetate, ethanol),

nitrogen, and phosphate. These experiments will tell me if Bck2 abundance is

significantly regulated by specific nutrient conditions BCK2 expression is induced by

nutrient limitation (Klosinska et al. 2011) or in stationary phase (Gasch et al. 2000),

suggesting that Bck2 protein abundance may be similarly influenced by nutrient quality.

Second, I will ask if nutrient-stimulation of bck2Δ cells in stationary phase blocks SWI4

and CLN3 expression, similar to the blockage of target gene expression in serum-

stimulated cells depleted of MKL1/2 (Lee, Vasishtha, and Prywes 2010), which would

indicate if Bck2 is required for transducing the nutrient signal to the transcription

machinery. Third, I will ChIP Bck2, Fkh2 and Yox1 to the SWI4 and CLN3 promoters in

a timecourse experiment, where cultures are allowed to enter and exit stationary phase in

a strain where Bck2, Fkh2 and Yox1 are all epitope-tagged. This experiment will tell me

if the association of Bck2 with Mcm1 on promoters positively correlates with target gene

expression, and whether association of Yox1 with Mcm1 is negatively correlated. The

promoter-localization of Fkh2, if any, will be useful in assessing the specificity of control

over Mcm1-dependent gene expression in early G1 phase. In some instances, p62TCF

binding to the c-fos SRE increases after serum induction from undetectable levels (Zhou,

Hu, and Herring 2005). Thus, I am also interested in testing if Fkh2 can bind to ECB

elements in the SWI4 and CLN3 promoters after growth in rich media. Interestingly,

149

Fkh2-Mcm1 has been shown to interact with the PHO5 promoter by the ChIP assay

(Pondugula et al. 2009), suggesting that Fkh2-Mcm1 may activate other genes of the

PHO regulon at the M/G1 transition (Cho et al. 1998; Spellman et al. 1998) in order to

scavenge limiting phosphate (Lenburg and O'Shea 1996). Fourth, because Bck2 has been

localized to both the nucleus and cytoplasm (Huh et al. 2003), I am curious to know if

Bck2 translocates to the nucleus in rich media, mimicking the translocation of MKL

proteins in serum (Lee, Vasishtha, and Prywes 2010). Fifth, I am interested in assaying

the phosphorylation status of Bck2 as a function of various alterations in nutrient

conditions, because MKL1 phosphorylation is induced by serum induction (Miralles et al.

2003; Selvaraj and Prywes 2003). If Bck2 is phosphorylated in rich nutrients, then I will

mutate residues previously reported to be phosphorylated in the central region of Bck2

(Chi et al. 2007; Albuquerque et al. 2008) and ask if any phospho-mutants phenocopy a

bck2Δ strain. Collectively, the experiments outlined will help me better understand the

role of Bck2 in nutrient sensing. Most mutations resulting in human cancer are in genes

encoding proteins involved in processes that occur in G1 phase (Sidorova and Breeden

2003a). If Bck2 activity turns out to be intimately related to external nutrient conditions,

then S. cerevisiae stands to be an ideal system to understand how aberrant proliferation

can begin inside the living, undividing cell.

150

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