Understanding mechanisms of zinc homeostasis in Schizosaccharomyces pombe
DISSERTATION
Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy in the Graduate School of The Ohio State University
By
Sang‐Yong Choi
The Ohio State University Nutrition Graduate Program
The Ohio State University
2015
Dissertation Committee:
Amanda J. Bird, Ph.D. (Advisor)
Earl H. Harrison, Ph.D.
Daren L. Knoell, Pharm.D.
Kichoon Lee, Ph.D.
Copyrighted by
Sang‐Yong Choi
2015
Abstract
Zinc is essential for cell growth, but can be toxic when in excess. As a consequence, intracellular zinc levels are tightly controlled by complex mechanisms that maintain zinc homeostasis in all biological creatures. In eukaryotic cells, factors that affect zinc homeostasis include zinc transporters, zinc buffering molecules, and zinc‐ regulatory factors. The levels of these factors are precisely regulated to maintain
optimal intracellular zinc levels. To extend the growing literature in the roles of genes involved in zinc homeostasis, this dissertation investigates the role of the above
executants in maintaining cytosolic zinc levels in Schizosaccharomyces pombe.
In S. pombe, Loz1 plays a specific role in repressing gene expression when zinc is
in excess. In zinc‐replete conditions, Loz1 down‐regulates zrt1, a gene encoding a high affinity zinc uptake transporter, and indirectly up‐regulates zym1 expression, a gene which encodes a zinc metallothionein. Deletion of Loz1, thus, causes constitutive zrt1 expression and low zym1 expression. In addition, cells lacking a functional loz1 gene
hyperaccumulate zinc in zinc‐rich medium. In chapters 2 and 3, I take advantage of the
loz1 mutation, and other yeast mutants to determine the roles of specific zinc
transporters and zinc buffering molecules in maintaining zinc levels in the cytosol.
ii
Specifically, I utilize genetically encoded zinc‐responsive FRET‐based sensors, which
allow changes in the labile pool of cytosolic labile zinc to be measured. My results show
that Zhf1, a zinc transporter in endoplasmic reticulum membrane, and Zrg17 and Cis4,
which reside in Golgi membrane, play a role in maintaining the cytosolic labile zinc pools
upon a zinc shock. In addition, my works describe how Loz1 controls zinc homeostasis in
zinc‐replete conditions and reveal that phytochelatins, small molecules that have a well‐
known role in the detoxification of toxic heavy metals, also have an important zinc
buffering role.
In chapters 4 and 5, I investigate the molecular mechanisms by which zinc‐
responsive transcription factors are regulated by zinc. Using chimeric proteins containing Loz1, this study examines the accessory domain adjacent to a double zinc finger, and shows that it is necessary for Loz1‐mediated zinc‐responsive changes in gene expression.
This dissertation provides a platform for the understanding of zinc homeostasis
mechanisms in fission yeast by examining the role that specific zinc transporters, zinc
buffering molecules, and the zinc‐regulated factor Loz1 play in regulating cellular zinc
levels. The results suggest that specific zinc transporters and Loz1 control the labile zinc
pool in cells. Also, the roles of phytochelatins are highlighted as zinc buffering molecules.
This discovery extends the current knowledge of how zinc buffering molecules influence
metal homeostasis. While a number of zinc transporters have been identified in various
iii
eukaryotic cells, the zinc buffering molecules that modify the labile zinc pool remain to
be further investigated.
iv
Acknowledgments
My sincere gratitude goes to my advisor and mentor, Dr. Amanda Bird, for all her
guidance and support throughout my doctoral studies. I would not have been able to
complete all this without her kind assistance and optimistic view. I am thankful to all the
members in Dr. Bird’s lab for their help as well. My dissertation committee members, Dr.
Kichoon Lee, Dr. Daren Knoell, and Dr. Earl Harrison, have been a great support for my
research. Also, my studies would not have been possible without the ZapCY1 and
ZapCY2 constructs kindly provided by Dr. Amy Palmer at University of Colorado. I would
like to thank Dr. Hyeyoung Kim at Yonsei University, who has been a warm and
supportive teacher throughout the years.
Finally, I would like to thank my mother, Seon‐Sook Yi, and father, Joong‐Suk
Choi, for all their love and support from home. Also, I am grateful to Janice Jung for
countless things.
v
Vita
2006 ...... B.S. Food & Nutrition and Biology, Yonsei
University, South Korea
2008 ...... M.S. Food & Nutrition, Yonsei University,
South Korea
2009 to present ...... The Ohio State University Nutrition Program,
The Ohio State University, USA
Publications
KM Ehrensberger, ME Corkins, S Choi, and AJ Bird. The double zinc finger domain and
adjacent accessory domain from the transcription factor Loss of zinc sensing 1 (Loz1) are necessary for DNA binding and zinc sensing. J Biol Chem 2014 (289) 18087‐18096.
S Choi and AJ Bird. Zinc'ing sensibly: controlling zinc homeostasis at the transcriptional
vi
level. Metallomics 2014 (6) 1198‐1215.
YC Chan, J Banerjee, SY Choi, and CK Sen. miR‐210: the master hypoxamir.
Microcirculation 2012 (3) 215‐223.
SY Choi, JW Lim, T Shimizu, K Kuwano, JM Kim, and H Kim. Reactive oxygen species
mediate Jak2/Stat3 activation and IL‐8 expression in pulmonary epithelial cells
stimulated with lipid‐associated membrane proteins from Mycoplasma pneumoniae.
Inflamm Res. 2012 (5) 493‐501.
SY Choi, JH Yu, and H Kim. Mechanism of α‐lipoic acid‐induced apoptosis of lung cancer
cells: Involvement of Ca2+. Ann NY Acad Sci. 2009 (1171) 149‐155.
Fields of Study
Major Field: The Ohio State University Nutrition Graduate Program
vii
Table of Contents
Abstract ...... ii
Acknowledgments ...... v
Vita ...... vii
Table of Contents ...... viiii
List of Tables ...... xiv
List of Figures ...... xv
CHAPTER 1. Zinc in life ...... 1
1.1 Introduction ...... 1
1.2. Zinc in public health ...... 2
1.2.1. Occurrence of zinc deficiency ...... 2
1.2.2. Nutritional intervention of zinc deficiency ...... 3
1.3. Biochemical functions of zinc ...... 5
1.3.1. Functions of zinc in metalloproteins ...... 6
1.3.2. Zinc signaling ...... 7
viii
1.4. Zinc homeostasis in mammals ...... 7
1.4.1. Zinc transporters ...... 8
1.4.2. Metallothioneins (MTs) ...... 14
1.4.3. MTF‐1 ...... 15
1.5. Zinc homeostasis in S. cerevisiae ...... 20
1.5.1. Zinc transporters ...... 21
1.5.2. Zap1 ...... 25
1.6. Zinc homeostasis in Schizosaccharomyces pombe ...... 28
1.6.1. Zinc transporters ...... 29
1.6.2. Metal storage proteins/peptides ...... 32
1.6.3. Loz1 ...... 34
1.7. Overview ...... 38
CHAPTER 2. Monitoring intracellular zinc distribution using FRET‐based genetic zinc sensors ...... 40
2.1. Introduction ...... 40
2.2. Materials and methods ...... 43
2.2.1. Yeast strains and growth conditions ...... 43
2.2.2. Plasmid construction ...... 43
ix
2.2.3. Yeast transformation ...... 47
2.2.4. Crossing strains and tetrad analysis ...... 47
2.2.5. RNA extraction and Northern blot ...... 48
2.2.6. Protein isolation and Western blot ...... 49
2.2.7. LacZ reporter assay ...... 51
2.2.8. FRET measurement ...... 52
2.2.9. Atomic Absorption Spectroscopy ...... 53
2.3. Results ...... 53
2.3.1. FRET‐based zinc sensors can monitor intracellular zinc distribution in the
fission yeast ...... 53
2.3.2. The effects of loss of specific zinc transporters on cytosolic labile zinc
levels ...... 65
2.4. Discussion ...... 79
CHAPTER 3. Phytochelatins influence the cytosolic labile zinc pool in fission yeast ...... 87
3.1. Introduction ...... 87
3.2. Materials and methods ...... 89
3.2.1. Yeast strains and growth conditions ...... 89
3.2.2. Plasmids ...... 90
x
3.2.3. Crossing strains and tetrad analysis ...... 90
3.2.4. Serial dilution growth assays ...... 90
3.2.5. FRET measurement ...... 91
3.2.6. Northern blot ...... 91
3.3. Results ...... 91
3.3.1. Loz1 knock‐out strains are resistant to zinc toxicity ...... 91
3.3.2. Loz1 controls cytosolic zinc distribution ...... 96
3.3.3. Overexpression of zrt1 is required to accumulate more zinc and modulate
the kinetics of labile zinc in loz1Δ cells ...... 108
3.3.4. Zym1 reduction is not responsible for zinc hyperaccumulation and
tolerance of loz1Δ cells ...... 112
3.3.5. Phytochelatins play a role for zinc buffering in loz1Δ cells ...... 117
3.4. Discussion ...... 121
CHAPTER 4. The double zinc finger and adjacent accessory domain are critical in controlling Loz1 regulons ...... 125
4.1. Introduction ...... 125
4.2. Materials and methods ...... 126
4.2.1 Site‐directed mutagenesis ...... 126
xi
4.2.2 RNA isolation and Northern blot ...... 126
4.3. Results ...... 127
4.3.1. The Loz1 double zinc finger domain and adjacent accessary domain are
necessary for zinc‐dependent gene expression ...... 127
4.3.2. The adjacent accessory domain in Loz1 includes histidine residues ...... 132
4.4. Discussion ...... 135
CHAPTER 5. Understanding the zinc‐regulated mammalian cell proliferation ...... 137
5.1. Introduction ...... 137
5.2. Materials and methods ...... 138
5.2.1. Cell culture ...... 138
5.2.2. MTT assay ...... 139
5.2.3. Reverse Transcription PCR (RT‐PCR) and quantitative real‐time PCR (qRT‐
PCR)...... 139
5.2.4. Western blot ...... 139
5.2.5. siRNA Transfection ...... 140
5.2.6. Plasmid construction ...... 140
5.2.7. Dual luciferase reporter assay ...... 141
5.3. Results ...... 141
xii
5.3.1. HiNF‐P contains a ZF pair with similar amino acid residues ZF pair from
Zap1 ...... 141
5.3.2. Cell viability upon TPEN‐induced zinc deficiency ...... 146
5.3.3. Histone H4 mRNA expression in TPEN‐induced zinc deficient cells ...... 149
5.3.4. Regulation of H4 mRNA expression by HiNF‐P ...... 152
5.3.5. Critical role of the ZF pair in transcriptional activity and zinc
responsiveness of HiNF‐P ...... 154
5.3.6. Cell proliferation in zinc‐deficient medium with chelated‐FBS ...... 156
5.3.7. H4 mRNA level in cells grown in medium containing different zinc
concentrations ...... 158
5.4. Discussion ...... 160
CHAPTER 6. Concluding remarks ...... 163
REFERENCES ...... 166
APPENDIX A. Additional tables ...... 181
xiii
List of Tables
Table 1.1. MTF‐1 genes in various organisms ...... 16
Table A.1. S. pombe strains used in this study ...... 181
Table A.2. Primers and oligos used in this study...... 185
Table A.3. Plasmids used in this study ...... 187
xiv
List of Figures
Figure 1.1. Domain structure of human MTF‐1 ...... 19
Figure 1.2. Schematic diagram of Zap1 in S. cerevisiae ...... 26
Figure 1.3. Schematic diagram of Loz1 ...... 37
Figure 2.1. Construction of ZapCY1 and ZapCY2 sensors ...... 45
Figure 2.2. ZapCY1 and ZapCY2 expressions are stable in the fission yeast cells ...... 55
Figure 2.3. Both ZapCY1 and ZapCY2 sensors do not significantly interfere zinc
homeostasis ...... 59
Figure 2.4. Zinc uptake of WT cells upon zinc shock ...... 63
Figure 2.5. Zinc uptake of zrt1Δ cells upon zinc shock ...... 67
Figure 2.6. Zinc uptake of zhf1Δ cells upon zinc shock ...... 71
Figure 2.7. FRET responses of ZapCY1 or ZapCY2 in zrg17Δ or cis4Δ cells upon zinc shock
...... 75
Figure 2.8. FRET responses of ZapCY1 or ZapCY2 in zip3Δ cells upon zinc shock ...... 78
Figure 2.9. FRET responses of ZapCY1 in zrg17Δ cells upon pyrithione and zinc shock .. 84
Figure 3.1. Cells lacking loz1 are resistant to zinc toxicity ...... 94
Figure 3.2. Zinc uptake of loz1Δ cells upon zinc shock ...... 98
xv
Figure 3.3. ZapCY1 sensor in loz1Δ cells may still be saturated upon TPEN treatment in
short‐term ...... 102
Figure 3.4. ZapCY1 sensor in loz1Δ cells becomes an apo form by pyrithione ...... 107
Figure 3.5. Zrt1 is required for loz1Δ cells to have zinc tolerance, but is not sufficient to
confer the tolerance to WT cells ...... 110
Figure 3.6. FRET responses of ZapCY1 or ZapCY2 in loz1Δ zym1Δ cells upon zinc shock .....
...... 113
Figure 3.7. Overexpression of metallothioneins does not nullify the zinc tolerance of
loz1Δ cells ...... 115
Figure 3.8. Phytochelatin may play a role for zinc buffering in the loz1Δ cells that were
exposed to zinc shock ...... 119
Figure 4.1. The adjacent accessory domains are important for zinc‐dependent repression
of Loz1 ...... 130
Figure 4.2. The adjacent accessary domain in Loz1 includes histidine residues ...... 134
Figure 5.1. Schematic structures and amino acid comparisons of ZF1/2 pair in Zap1 and
HiNF‐P ...... 144
Figure 5.2. Cell viabilities in different zinc levels...... 148
Figure 5.3. Histone H4 and MT1 mRNA expressions in different zinc levels ...... 150
Figure 5.4. H4 and HiNF‐P genes expressions in HiNF‐P deficient cells ...... 153
Figure 5.5. Importance of ZF1/2 for transcriptional activity and zinc responsiveness of
HiNF‐P ...... 155
xvi
Figure 5.6. Cell viability in zinc‐normal (ZN) or ‐deficient (ZD) medium ...... 157
Figure 5.7. mRNA levels of HiNF‐P target genes and MT1 in HepG2 cells ...... 159
xvii
CHAPTER 1
Zinc in life
1.1 Introduction
Zinc is essential for the growth of all forms of life, including microorganisms,
plants, and animals [1‐3]. It is the second most abundant transition metal ion in humans, where it is primarily found within cells [4]. Given its ubiquitous nature within organisms and relatively copious concentration in cells, zinc has many important functions in both unicellular and multicellular organisms.
When such a fundamental mineral is depleted, living organisms show zinc
deficient symptoms. In contrast, when the intake of zinc is too high, they exhibit toxicity
symptoms. To avoid these undesirable nutritional conditions and maintain an optimal level of zinc, cells have evolved mechanisms to maintain an optimal level of zinc.
This chapter will illustrate the significance of zinc in public health and biochemistry. It will also describe the zinc homeostasis mechanisms in humans and in 2 different yeasts, Saccharomyces cerevisiae and Schizosaccharomyces pombe, that are
widely utilized to better understand the complicated zinc metabolism.
1
1.2. Zinc in public health
Human zinc deficiency was first reported in the Middle East in 1963 in patients
with dwarfism and hypogonadism [5, 6]. At the time, many doctors and nutritionists
doubted the existence of zinc deficiency in humans, since zinc is widely distributed in
the environment [7]. Research, however, revealed that zinc supplementation increases
the rates of growth and sexual development in zinc‐deficient subjects [8, 9]. Since these
initial observations, the health effects associated with zinc deficiency have been
extensively studied, especially in infants and younger children of both developing and
developed countries [10]. Despite the relatively narrow range of zinc levels in the body,
treating zinc deficiency has become an important strategy to improve public health in
various populations.
1.2.1. Occurrence of zinc deficiency
About 2 billion people in the world are classified to be zinc deficient or have inadequate levels in their diet [11, 12]. In developing countries as well as low‐income families in the United States, zinc deficiency has emerged as a major public health problem [13]. Zinc deficiency is chiefly associated with inadequate intake or absorption
of zinc from the diet, as zinc is not stored to any major extent in the human body [10].
Rich sources of zinc include animal products, such as meats and shellfish [14]. In
2
contrast, vegetables and cereals are poor sources of zinc, as plants contain high levels of
phytic acid and dietary fibers that lower bioavailability of zinc. Due to economic and
environmental factors, many people do not have the opportunity to consume a diet
adequate in proteins, and therefore are prone to zinc deficiency.
An increased risk of zinc deficiency is observed in patients with malabsorption
syndromes, diarrhea, parasitic diseases, or sickle‐cell diseases [15]. Another disorder
that results in severe zinc deficiency, unless properly treated, is acrodermatitis
enteropathica. This rare inherited genetic disorder results in symptoms that include skin
lesions, plaques of dry skin, and loss of hair from the scalp and eyebrows. Recent studies
have elucidated that mutations within the zinc transporter gene, SLC39A4 (ZIP4), are responsible for this disease, owing to reduced intestinal absorption of zinc [16, 17].
1.2.2. Nutritional intervention of zinc deficiency
The effects of zinc supplementation on health—both therapeutic and preventive—have been evaluated in a number of randomized controlled trials, and
meta‐analyses that combine results from multiple clinical studies [18‐22]. Some studies
on zinc supplementation [19, 20, 22] have investigated its therapeutic effects on
diarrhea, glucose homeostasis, and the common cold. In children with diarrhea,
especially in low‐ and middle‐income populations, oral zinc supplementation has been shown to reduce diarrhea duration, stool output, and frequency and length of hospitalization, and to decrease diarrhea mortality [19, 21, 23]. 3
Zinc supplementation affects glucose homeostasis by lowering the concentration
of fasting glucose [22, 24]. This effect has been significant in patients with chronic metabolic diseases, including diabetes and obesity. According to a recent meta‐analysis, there is an inverse association between the level of fasting glucose and total zinc intake, including food sources as well as supplements, in individuals with the SLC30A8 (ZnT8)
variant. This gene variant is therefore thought to be a risk allele for high fasting glucose
levels and type 2 diabetes risks [24].
Zinc is also thought to influence the common cold. Oral zinc supplies, in
particular, have been associated with a shorter duration of the common cold in healthy
people [20]. When administrated within 24 hours of the symptom onset, orally taken zinc shortened a day of cold duration. This effect of zinc is suggested to be due to the
antiviral activity against rhinoviruses. The efficacy is dependent on the amount of
positively‐charged ionic zinc that can be effectively released from zinc compounds, including zinc acetate and zinc gluconate [25].
Some other studies have examined the preventive effects of zinc supplementation, aiming to diminish the incidence of many common diseases, especially in the children of low‐income populations. The supplementation lowers the occurrences of diarrhea by 18%, pneumonia and lower respiratory tract infections by
41%, and malaria by 38% [26, 27]. Furthermore, preventive zinc supplementation is associated with a 6% reduction in total child mortality, including the reduction of mortalities from diarrhea and pneumonia [18, 28].
4
Regarding both its therapeutic and preventive effects, zinc supplementation clearly has the potential to be recommended to many individuals with zinc deficiency or the aforementioned health issues. A major limitation of zinc nutriture in humans, however, is that there is yet no accurate method for measuring zinc deficiency. To
better understand the interaction of zinc nutriture and human health problems, it is
necessary to investigate the mechanisms of how zinc levels are sensed and maintained
in the body [29]. This information will help nutritionists make recommendations for
various populations in different environments and extend our knowledge of the
connections between zinc metabolisms and diseases.
1.3. Biochemical functions of zinc
According to proteomic analysis, 10% of the entire human proteome is
estimated to bind zinc ions [30]. The majority of zinc‐bound proteins are proposed to be
involved in the regulation of gene expression. In addition to its importance in transcription factors, zinc ions can function as a catalyst and help to maintain structural
domains within many enzymes. Another noteworthy role of zinc is intra‐ and
intercellular signaling.
5
1.3.1. Functions of zinc in metalloproteins
Metalloproteins are defined as proteins that are capable of binding metal ions, in
which the bound metal ion facilitates the biological function of other cellular proteins,
support the structure of these proteins, or regulate their activities [30]. Numerous
metalloproteins in nature are known to require zinc ions. The predicted number of zinc‐ proteins in humans has increased since zinc finger structures were first identified in the early 1990s. The current number of proteins that contain a zinc‐finger like domain is
now estimated to be approximately 2800 [30].
Zinc functions as a catalytic cofactor in over 300 enzymes. Carbonic anhydrase was the first enzyme to be identified in which zinc ions were required for a catalytic
reaction of carbon dioxide and water to form bicarbonate [31]. In this enzyme, the zinc
ion is located in an active site where it facilitates the hydrolysis of water so that carbon
dioxide can be rapidly converted to bicarbonate.
Zinc also stabilizes the overall conformational structure of proteins and certain
protein domains. One of the most common classes of structural zinc domain is the C2H2 type zinc finger, which is composed of 2 cysteines and 2 histidines as ligands for a zinc ion. This zinc finger is found in a wide variety of transcription factors and is responsible for stabilizing the quaternary structure of the metalloproteins [32].
In some proteins, zinc can also have a regulatory role. Examples of these proteins
include zinc‐responsive activator protein (Zap1) and Metal regulatory Transcription
Factor‐1 (MTF‐1) [33, 34].
6
1.3.2. Zinc signaling
Zinc has been shown to be both extracellular and intracellular signaling
molecules [35, 36]. Similar to how neurotransmitters are transferred, zinc ions in
neurons are released from the nerve terminal into the synaptic cleft by exocytotic stimulation [37]. The zinc ions in the extracellular microenvironment are then co‐ translocated to postsynaptic neurons through neurotransmitter receptors or channels.
The roles of zinc in postsynaptic neurons are not yet known, but are suggested to be involved in the intracellular signaling pathway [38].
Zinc also functions as an intracellular second messenger. In mast cells, the intracellular free zinc ion levels increase upon stimulation in a manner that is dependent on calcium influx and mitogen‐activated protein kinase (MAPK) activation [36]. While this rapid boost in zinc level is analogous to what is observed with other secondary
messengers, the direct targets of the zinc messenger are currently unknown. Suggested
possible downstream targets include the zinc ions that regulate phosphatase activity
and cytokine production [36].
1.4. Zinc homeostasis in mammals
The mechanisms of zinc homeostasis have been widely examined in mammals
[39‐41]. In humans, zinc is absorbed at the apical surface of the gastrointestinal tract.
7
The regulation of this absorption process into and across the enterocyte is a crucial
factor to maintain zinc homeostasis in the whole body. Another factor that influences
zinc homeostasis in human bodies is the intestinal excretion of endogenous zinc
responding to both recent absorption of zinc and zinc status.
In addition to the systemic regulation of zinc levels, intracellular zinc levels are tightly regulated. As zinc cannot solely diffuse across cellular membranes, zinc needs a
group of proteins called zinc transporters, which are dedicated to its transport across the membrane. In the following section, I will describe our current knowledge of zinc transporters, storage proteins, and how intracellular zinc levels are sensed. I will also discuss how these homeostasis executants are regulated by zinc.
1.4.1. Zinc transporters
Before the first mammalian zinc transporter gene, ZnT1, was identified in 1995,
zinc transport in animals was thought to involve co‐transport of zinc with other ions or
amino acids [42]. The discoveries of various zinc transporters have allowed zinc metabolism to be investigated in terms of the regulation and interrelation of the
transporters. The zinc transporter genes in humans fall into 2 main families: SLC30A
(ZnT) and SLC39A (ZIP). The human genome encodes 10 ZnT members (ZnT1‐ZnT10) and
14 ZIP members (ZIP1‐ZIP14). The ZnT members generally engage in the efflux of zinc from the cytosol to the extracellular medium or intracellular organelles, such as
Endoplasmic Reticulum (ER), Golgi apparatus, and mitochondria; whereas the ZIP 8
proteins transport zinc from the extracellular fluid or intracellular organelles into the
cytoplasm. As these 2 transporter families are the primary means by which zinc enters
or leaves a cell or cellular compartment, the regulation of their expression and/or
activity results in dynamic changes of intracellular zinc level.
The ZnT transporters belong to the Cation Diffusion Facilitator (CDF) family, a
group of intermembrane transporters that function in the homeostasis of various
divalent metal cations [43]. The ZnT transporters, except for ZnT5 which contains 15
predicted membrane‐spanning domains, share partial sequence homology within their 6 transmembrane domains as well as their N‐ and C‐termini on the cytoplasmic side of membranes [44, 45]. In contrast, the amino acid sequences upstream of the first
transmembrane domains are more varied in sequence and often contain subcellular
targeting signals that assign diverse subcellular localization to each ZnT member.
As a CDF family member, ZnT1 is located on the plasma membrane and takes on
a fundamental role in exporting zinc to the extracellular environment [42]. The
transporter is expressed in a wide range of tissues and cell types. In enterocytes, ZnT1 is
specifically located on the basolateral surface. In addition, dietary zinc levels regulation
affect its levels, which in turn, affects zinc transfer into the circulation [46, 47]. Mice with homozygous knockout of ZnT1 gene do not survive embryonic development [48].
This finding indicates the essential roles of ZnT1 in transporting maternal zinc into the embryonic environment [48].
9
Some ZnT members, namely, ZnT2, ZnT3, ZnT4, and ZnT8, are expressed on the
membranes of transport/secretory vesicles in the cells of specific tissues [49‐52]. These
transporters pump zinc into the vesicles in order to deliver zinc to key proteins in the
respective compartment or to enable zinc to be secreted through exocytosis. For
example, ZnT2 and ZnT4 are each highly expressed in the mammary gland epithelial cells of humans and mice, respectively [49, 51]. During lactation, the mammary epithelial cells accumulate zinc in the secretory vesicles via the zinc transporters and secrete the
mineral with other components into milk. Genetic mutations in the genes encoding
ZnT2 and ZnT4, hence, lead to zinc‐deficient milk [49, 51]. As for the mammalian ZnT3,
this protein is expressed at a high level in the brain. Here, ZnT3 transports zinc into
presynaptic vesicles where it can be released into synaptic cleft upon stimulation [50].
Although the role of synaptic zinc is not yet known, ZnT3 knockout mice show learning
and memory impairment [53]. Predominantly found in pancreatic beta cells, ZnT8 plays
a key role in transferring zinc into insulin secretary vesicle [52]. A recent in vivo study
using beta cell‐specific ZnT8 knockout mice revealed lower zinc accumulation in the beta
cell, which caused an abnormal morphology and reduced islet insulin processing [54].
ZnT5 shows unusual characteristics depending on its isoform [55, 56]. Its splice variants, ZnT5A and ZnT5B, contain different amino acid sequences in the N‐ and C‐ terminal regions. These variants exhibit nonidentical subcellular localizations as 2 GFP‐ tagged isoforms, ZnT5A and ZnT5B were shown to be localized to the Golgi apparatus and plasma membrane, respectively [56]. Together with ZnT6 and ZnT7, ZnT5A is
10
expressed on the membrane of the Golgi apparatus where it delivers zinc into the intracellular compartment and secretory pathway [57]. In a study measuring the activity of alkaline phosphatases (ALPs), which require 2 zinc ions to become holoenzyme in the secretory pathway, the hetero‐dimer complex of ZnT5A and ZnT6 and the homo‐dimer complex of ZnT7 are both considered to supply zinc to the ALPs as they mature in the secretory pathway [57]. The other splice variant ZnT5B has been found to reside in the apical membrane in polarized human intestine Caco‐2 cells and human enterocytes [58,
59], and the membrane of ER in HeLa cell lines [60].
The transporters in the ZIP family are responsible for zinc influx into the cytosolic space. Depending on its subcellular localization, each ZIP transporter delivers zinc from the extracellular environment or intracellular compartments to the cytoplasm. Most ZIP members are predicted to contain the following: 8 transmembrane domains, a long loop variable region with histidine‐rich sequences between transmembrane 3 and 4, and
both N‐ and C‐terminal ends facing towards the outside of cytoplasm. Depending on
their degree of sequence conservation, the 14 ZIP proteins are classified into 4
subgroups: ZIP subfamily I, ZIP subfamily II, gufA subfamily, and LIV‐1 subfamily [61].
Unlike the ZIP subfamily I (ZIP9) and gufA subfamily (ZIP11), the functions of ZIP
subfamily II and LIV‐1 subfamily have been extensively investigated.
The ZIP subfamily II members—ZIP1, ZIP2, and ZIP3—are widely expressed
throughout the human body [62]. These proteins have been reported to be zinc uptake
transporters on plasma membrane of mammalian cells [63‐65]. In overexpression and
11
knockdown experiments using human erythroleukemia K562 cells, ZIP1 and ZIP2 levels
show a positive correlation with Zn65 uptake rate [63, 64]. Also, ZIP3 relocates to the serosal plasma membrane in response to prolactin hormone. This trafficking increases zinc uptake into mammary epithelial cells for them to secrete adequate zinc into milk
[65]. In some cell lines, such as COS‐7 and HepG2, the localization of ZIP1 was observed in the ER membrane [66]. This indicates that ZIP1 may also be in charge of the efflux of
stored zinc within ER. Moreover, mouse homologs of mZIP1 and mZIP3 reside in plasma
membrane under zinc‐limiting conditions, while they are localized to intracellular compartments in zinc‐replete states [67]. Thus, the role of these subfamily II members in zinc trafficking must be studied with careful consideration of cell and tissue types as
well as environmental zinc and hormonal statuses.
In the LIV‐1 subfamily, there are 9 ZIP transporters—ZIP4‐ZIP8, ZIP10, and ZIP12‐
ZIP14—that display sequences homologous to the estrogen‐regulated LIV‐1 protein
whose expression increases in estrogen receptor‐positive breast cancer. Members of
the LIV‐1 subfamily, compared to the other ZIP members, have a longer N‐terminus, as
well as more histidine‐rich repeats and conserved transmembrane sequences [68]. As a
consequence, these zinc transporters are suggested to have broader roles. Recent
studies have also reported on the functions of some LIV‐1 subfamily transporters in
various diseases, suggesting that they may be critical targets to understand the
significance of zinc homeostasis in cancer and inflammation [69‐73].
12
ZIP4 and ZIP5 are 2 LIV‐1 transporters found in the small intestine. ZIP4 has been identified in acrodermatitis enteropathica patients who have mutations in the ZIP4 gene and a defect in the absorption of dietary zinc [74]. Taking into account its abundant expression level in mouse enterocyte and its localization in apical membrane of the cell,
ZIP4 is responsible for zinc absorption in the small intestine [74]. Unlike ZIP4, ZIP5 resides in basolateral surfaces of enterocytes [75]. Because of its expression patterns—
decreasing in periods of dietary zinc deficiency and increasing in sufficiency, ZIP5 is
suggested to remove zinc from blood and help excrete zinc into the intestine when zinc
is in excess.
ZIP6 and ZIP8 are 2 other LIV‐1 subfamily members associated with diverse diseases. ZIP6, also known as LIV‐1 in humans, has been implicated in various cancers,
such as breast, pancreatic, cervical, prostate, and brain. [70‐72, 76, 77]. Expressed in the
plasma membrane of cells, ZIP6 modulates zinc uptake rates [72, 78]. ZIP8 is involved in
inflammatory diseases. Its gene expression is up‐regulated in response to bacterial
endotoxins, cytokines, and sepsis [79, 80]. ZIP8 was identified in the plasma membrane
in lung epithelial cells. However, it was observed in lysosomal membranes in
differentiated macrophages and monocytes where increased expression of ZIP8 induced
zinc transfer into cytoplasm [81, 82]. While studies continue to investigate the roles of
both ZIP8 and zinc ions in the immune response, more recent research have
demonstrated an association between ZIP8‐mediated zinc transport and Nuclear factor‐
kappa B (NF‐κB) pathway that is a key inflammatory signaling pathway in mammals [83].
13
NF‐κB factor transcriptionally modulates ZIP8 expression, and ZIP8‐induced zinc uptake
then represses the NF‐κB activation via a negative feedback loop.
1.4.2. Metallothioneins (MTs)
MTs are a group of small cysteine‐rich proteins (up to 30% of the amino acid residues), which are able to bind specific metals [84, 85]. These proteins can sequester
zinc, cadmium, or copper in cells, potentially protecting the cells from the toxicity of
these metal ions [86]. In humans, there are 4 isoforms of MTs: MT1‐MT4. In addition, at
least 7 genes encode MT1‐type proteins that slightly vary in amino acid sequences. In contrast, only 1 copy of the 3 other isoforms—MT2, MT3 and MT4—is located within genome [87]. With regard to tissue specific expression, MT1 and MT2 are expressed ubiquitously throughout the human body. In contrast, MT3 is predominantly expressed
in the hippocampus of the brain, and MT4 is observed in the stratified squamous epithelial cells in buccal mucosa, esophagus, and uterine cavity [88, 89].
The 3D structure of MTs indicates that all cysteine residues are involved in the
binding of zinc, forming 2 metal‐thiolate clusters: a Zn4Cys11 cluster in the N‐terminal
domain of an MT and a Zn3Cys9 cluster in the C‐terminal domain [90, 91]. Based on
these studies, MT1 in mammals can bind a total of 7 zinc atoms.
Associated with their characteristic sulfur‐rich structures, MTs have been reported to serve a number of physiological functions [92‐94]. The MTs in cells influence zinc homeostasis, detoxification of cadmium, and protection from oxidative stress. 14
These metal binding proteins play a fundamental role especially in zinc metabolism. Zinc is an essential cellular component bound to MTs, and the metal buffering proteins can
modulate the intracellular labile zinc levels, which dynamically fluctuate in very low
ranges [87].
The levels of MT expression are mainly controlled at a transcriptional level [95].
This transcriptional control is mediated by MTF‐1 in response to the exposure of metals,
such as zinc. In zinc‐replete conditions, the MTF‐1 activation rapidly increases MT mRNA
expressions. The cells become capable of sequestering intracellular labile zinc and
maintaining optimal zinc levels in the cytoplasm.
1.4.3. MTF‐1
MTF‐1 is the only well‐characterized zinc‐sensing transcriptional factor in
humans. In addition to being found in mammals, homologs of MTF‐1 are present in
insects, reptiles, and fish (Table 1.1). MTF‐1 was initially identified as a protein that was
required for the metal inducible regulation of MT1 gene expression [95]. However, more
recent studies have demonstrated that MTF‐1 activity is regulated by zinc and that MTF‐
1 regulates the gene expression of crucial proteins in changing labile intracellular zinc
levels. As a consequence, MTF‐1 is now regarded as a key protein in zinc homeostasis.
15
Table 1.1. MTF‐1 genes in various organisms
Gene Response to metal Host organism Reference
Copper‐responsive Drosophila melanogaster (firefly) [96]
Mus musculus (mouse) [97]
Homo sapiens (human) [98]
Fugu rubripes (puffer fish) [99]
Danio rerio (zebrafish) [100]
MTF‐1 Hydrochoerus hydrochaeris Zinc‐responsive [101] (capybara)
Oreochromis aureus (tilapia) [102]
Cyprinus carpio (common carp) [103]
Crassostrea gigas (pacific oyster) [104]
Anguis fragilis (slow worm) [105]
16
MTF‐1 regulates its target gene expression by binding to DNA elements that are
located within the promoter regions of its target genes. These elements are called the
metal response elements (MREs) [106]. In addition to MT1, another important target
gene of MTF‐1 in regard to zinc homeostasis is ZnT1. MTF‐1 is required in the zinc‐
induced ZnT1 expression and its basal expression [107]. MTF‐1 also controls the
expression of ZIP10, which encodes a zinc importer [108]. However, at the ZIP10
promoter, MTF‐1 acts as a transcriptional repressor by binding to MRE elements that are located downstream of the transcriptional start site and interfering with
transcriptional elongation [109].
In humans, MTF‐1 is a protein that is made up of 753 amino acids and contains 6
C2H2 zinc fingers that form the DNA binding domain (Fig. 1.1). Once zinc ions, or other
heavy metal ions, are bound to MTF‐1, the protein is rapidly translocated into nucleus
where it interacts with the MREs in its target genes via the DNA binding domain [110].
This ability of MTF‐1 to bind DNA in vivo is dependent on the zinc concentrations,
suggesting that the DNA binding function may be directly regulated by zinc [111].
Although this is a widely accepted hypothesis, the molecular mechanism behind this
control remains controversial. As it consists of 6 canonical zinc fingers, it has been suggested that zinc binding to these zinc finger domains may control the DNA binding functions of MTF‐1 [34, 112]. Consistent with this hypothesis, constructs with deletions of ZF1 or both ZF5 and ZF6 have impaired‐zinc responsive expression of MT1 gene [113].
Moreover, the ZF1 deletion mutant displays constitutive interaction with MREs after
17
additional zinc treatment [114]. In contrast, linker peptides between the zinc fingers
have also been suggested to be critical for zinc sensing [111]. Amino acid substitutions in
the linker region between ZF1 and ZF2 have resulted in a loss of zinc sensing.
In addition to the regulation of DNA binding activity by zinc, the transcriptional
activation function is regulated by zinc [115]. MTF‐1 contains activation domains that
are classified as acidic, proline‐rich, and serine/threonine‐rich [34]. A reporter assay
with chimeric transcription factors that are composed of the Gal4 DNA binding domain
and different MTF‐1 activation domains provided the evidence that the acidic activation
domain contains a major determinant of zinc inducibility [115].
MTF‐1 has 2 nuclear localization signals (NLSs) and a nuclear export signal (NES)
[111, 115]. A consensus NLS next to ZF1 is auxiliary, while a nonconventional NLS, a
region including ZF1‐ZF3, is sufficient to target the protein into the nucleus [115]. On the
other hand, a classical leucine‐rich NES is embedded in the acidic domain. In resting
cells, mutations in the NES cause nuclear accumulation of MTF‐1 [115]. As zinc‐bound
MTF‐1 migrates into nucleus and binds with MREs, nuclear translocation of MTF‐1 is necessary but not sufficient for zinc‐responsive transcriptional activation.
18
Figure 1.1. Domain structure of human MTF‐1
Six C2H2‐type zinc fingers (ZFs) in the DNA binding domain are represented in numbers and dark blue boxes. Three activation domains—acidic, proline‐rich, and serine/threonine‐rich—are illustrated in yellow, cyan blue, and purple boxes, respectively. The red boxes and letters indicate an auxiliary nuclear localization signal (aNLS) next to ZF1, a nuclear localization signal (NLS) spanning ZF1‐ZF3, and a nuclear export signal (NES) overlapping the acidic domain.
19
1.5. Zinc homeostasis in S. cerevisiae
S. cerevisiae, the budding yeast, is a unicellular eukaryote that has been extensively used as an experimental organism in molecular and cellular biology, and
genetics. As a model organism, the budding yeast has many advantages, which include
versatile genetic modification, short life cycle, a simple and compact genome, and a fully
mapped and sequenced genome database.
With these many advantages for genetic studies, this yeast has been extensively
used as a system to understand genetic and molecular mechanisms of zinc homeostasis.
Due to the evolutionary conservation of genes and their functions, the molecular
mechanisms of zinc homeostasis in yeast have allowed significant advances to be made
in understanding the metal homeostasis in humans. For example, Zrt1, the founding
member of the ZIP family of zinc transporters, was initially discovered in S. cerevisiae
[63, 116].
Multiple genes have been identified to play a crucial role in maintaining zinc
homeostasis in S. cerevisiae. These include zinc transporters from the ZIP and CDF
families, and the zinc‐responsive transcription factor Zap1 [117]. Zap1 modulates the
expression of genes encoding zinc transporters depending on the zinc availability in the
cell.
This section will provide an overview of how S. cerevisiae responds to zinc‐ deficient and zinc‐replete conditions and maintains zinc homeostasis through the
20
transcriptional control of genes encoding zinc transporters. It will also discuss the molecular mechanisms by which the Zap1 factor recognizes intracellular zinc levels.
1.5.1. Zinc transporters
Like those in mammals, most zinc transporters in S. cerevisiae, with a few exceptions, are mainly classified into ZIP and CDF families. In the yeast, there are 4 ZIP
transporters—Zrt1‐Zrt3 and Yke4, and 4 CDF proteins—Zrc1, Cot1, Msc2, and Zrg17
[118].
As the primary transporter for zinc uptake, Zrt1 is required for normal growth in
zinc‐limited environments [116]. In such deficiency, the expression of the ZRT1 gene is
transcriptionally increased; and this regulation is mediated by the Zap1 transcription
factor [33]. The increased expression of ZRT1 leads to an increase in the numbers of Zrt1 proteins that are stably expressed on the plasma membrane, which in turn leads to
increased zinc uptake. In addition to the transcriptional regulation of ZRT1 expression,
Zrt1 is regulated by zinc at a post‐translational level. In response to high levels of extracellular zinc, Zrt1 becomes degraded through endocytosis [119].
Another zinc uptake transporter, Zrt2 has a lower affinity for zinc ions than Zrt1
[120]. Due to its lower affinity, Zrt2 contributes to zinc acquisition under mildly zinc‐ deficient or zinc‐replete states [121]. Although this protein is a target gene of the Zap1
transcription activator, its mRNA expression is suppressed in severe zinc‐limited cells
and remains high upon zinc supplementation. This unexpected regulation profile is 21
known to occur due to a weak Zap1 binding site in the ZRT2 promoter which is located
downstream of the TATA box, and 2 strong Zap1 binding elements that are found in the
upstream promoter region [121]. In severe zinc deficiency, the Zap1 expression is
induced by autoregulation; thus, the high level of Zap1 results in it being able to bind to the weak Zap1 binding site in the ZRT2 promoter, which inhibits the expression of ZRT2.
This unique mechanism enables the cell to solely use Zrt1, the high efficient zinc transporter, when it needs zinc from its surrounding.
Found in the vacuole membrane, Zrt3 plays the role to “offset” deficiency in zinc‐ depleted cells [122]. Similar to other mammalian intracellular ZIP proteins, Zrt3 pumps
stored zinc out of the vacuole into the cytoplasm when there is not much extracellular
zinc available. In such zinc deficiency, the Zap1 transcriptional activator up‐regulates the
expression of ZRT3, which ensures that zinc is released from the vacuolar store.
Yke4 transporter has been identified as a yeast homolog gene of the mouse/human Ke4, which is also known as Zip7 [123, 124]. While Ke4 in mammals is known to reside at the ER or Golgi membrane, Yke4 was found to be localized specifically to the ER using subcellular membrane fractionation and
immunofluorescence assays [123‐125]. On the ER membrane, the transporter was
suggested to function for bidirectional transport of zinc depending on cellular zinc status
[124]. According to the study, Yke4 may increase cytosolic zinc, delivering zinc out of ER in zinc‐limited cells. When there is enough zinc in the environment, however, the
transporter may pump and store zinc into ER. Also, the expression of mouse Ke4 was
22
able to complement Yke4 knockout cells grown in high zinc medium. This result is interesting in that the mouse Ke4, which only releases zinc from the secretory pathway, substituted the bidirectional Yke4 in zinc‐replete cells. Further studies, therefore, need
to investigate how Yke4 modulates intracellular zinc distribution and what molecular
aspects contribute to the differences between Yke4 and mammalian Ke4.
Two members of the CDF family, Zrc1 and Cot1 are transporters that reduce the
cytosolic zinc level, transferring zinc into the vacuole [122]. The vacuole in S. cerevisiae
is known as a major organelle for zinc storage [117]. Under periods of zinc limitation, zinc can be released from the vacuole by Zrt3 for use in the cytosol.
Due to their role in vacuolar zinc storage, Zrc1 and Cot1 are important for
cellular tolerance to excessive zinc levels. Zrc1, in particular, has been shown to be
critical for a “proactive” mechanism to help protect zinc‐limited cells from a sudden
exposure to high zinc [126]. As a Zap1 target gene, ZRC1 expression increases in zinc‐ limited cells. As zinc‐limited cells express high levels of the zinc uptake transporter Zrt1 on their plasma membrane, when they are resupplied with zinc, a very high level of zinc comes into the cytosol and the cells experience a condition known as “zinc shock”. To resist against this shock, it is essential for the Zrc1 expression to be proactively induced
[126]. Consistent with this proactive protective role, ZRC1‐deleted cells display
hypersensitivity to zinc shock. In contrast, the ZRC1 mutant cells pre‐grown in zinc‐
replete conditions do not show such levels of sensitivity when inoculated into high zinc
medium.
23
Two other CDF members, Zrg17 and Msc2 form a heteromeric complex which
transports zinc into the ER [127]. Many proteins, which are either retained in the ER or
moved through it, require zinc as a cofactor. As transporters responsible for zinc entry
into ER, Zrg17 and Msc2 are therefore both important for ER functions and protein
processing [127‐129]. In zinc‐limited cells, zinc transfer into ER is maintained by Zap1.
Under these conditions, Zap1 directly binds to the promoter region of ZRG17 and induces its expression [129]. Strains that contain a chromosomal mutation within the
ZRG17 promoter or that lack ZRG17 exhibit higher ER stress responses under zinc‐ deficient conditions compared to wild type cells. These results indicate that zinc delivery into ER, which is both Zap1‐dependent and Zrg17‐mediated, is crucial in order for ER functions to be maintained in these cells during periods of zinc limitation.
Apart from the zinc transporters in the ZIP and CDF families, S. cerevisiae
contains 2 additional transporters on plasma membrane, Fet4 and Pho84, which play a
role in zinc uptake [130, 131]. Although it was originally identified as an iron transporter
[132], Fet4 has been demonstrated to mobilize zinc into cells. Also, its mRNA level is up‐
regulated by Zap1 in response to zinc deficiency [130]. PHO84 expression, in contrast, is
not modified according to zinc level. Nevertheless, this high affinity phosphate
transporter has been suggested to transfer zinc as ZnPO4 from extracellular
environments [117, 131].
24
1.5.2. Zap1
Zap1 was identified in S. cerevisiae in a genetic screen to isolate mutants that
induced the expression of genes required for zinc uptake in zinc‐replete cells [33]. In zinc‐deficient cells, Zap1 becomes active and binds to zinc responsive elements (ZREs) that are located in the promoter region of Zap1 target genes [133]. To date, studies have reported over 80 target genes of Zap1 whose expressions are associated with
homeostasis and adaptive responses against mild and severe zinc‐deficient conditions, respectively [134, 135].
As it contains a ZRE within its promoter, Zap1 is transcriptionally up‐regulated by the Zap1 factor in zinc‐limited cells [136]. This positive autoregulatory mechanism
amplifies the zinc‐responsive activity of Zap1 factor. Moreover, it enables cells to react
differently to severe or mild zinc deficiency as seen in the Zap1‐mediated Zrt2 regulation.
25
A
B
Figure 1.2. Schematic diagram of Zap1 in S. cerevisiae A. Zap1 consists of 880 amino acids and includes 2 activation domains (ADs) and a DNA binding domain (DBD). AD1 does not contain any zinc finger (ZF) while AD2 and DBD contain 2 and 5 zinc fingers, respectively. The 7 ZFs are represented in numbered boxes. The loop structures found between ZF1 and ZF2, and ZF3 and ZF4 indicate the unique ZF pair structures of ZF1/2 and ZF3/4. Other traditional ZFs are illustrated in light blue boxes. B. The amino acid sequences of ZF1/2 (578‐644 a.a.) in AD2. The key amino acid
sequences for ZF pair structure (CWCH2) and zinc binding are displayed in the red letters.
26
The functional domains of the Zap1 protein have been extensively studied [137‐
139] (Fig. 1.2). Zap1 is composed of 880 amino acids that include a DNA binding domain
and 2 activation domains (AD1 and AD2), all of which are regulated by zinc [138, 140].
The DNA binding activity is mediated by 5 C2H2 type zinc fingers: ZF3‐ZF7. Each of these
zinc fingers tightly binds zinc, even during severe zinc starvation. For this reason, the
DNA binding domain maintains its structure and ability to bind to ZREs under severely zinc‐limited conditions. AD1 does not contain any classical C2H2 type zinc finger motif
but has multiple cysteine and histidine residues that coordinate zinc [33]. It is not clear
how this domain responds to zinc deficiency. Studies have demonstrated that AD1 can
activate most of Zap1 target genes expressions [141], while AD2 plays a vital role during
periods of severe zinc limitation [138, 140]. These results imply that both AD1 and AD2,
in an independent manner, function to overcome the zinc deficiency and contribute to cell growth responding to low zinc.
The zinc‐responsive mechanism of AD2 has been relatively well studied [102,
138, 139, 142]. As presented in Fig. 1.2, this domain contains 2 zinc finger motifs (ZF1 and ZF2), which potentially play a key role in the zinc‐sensing mechanism of Zap1. ZF1
and ZF2 form a unique paired structure due to the existence of a tryptophan residue
that is located between the 2 canonical cysteine residues in each zinc finger (Fig. 1.2.B).
This ZF1 and ZF2 pair structure (ZF1/2) is hypothesized to exist as a tight closed form in
the presence of zinc ion. In zinc‐limited cells, the zinc bound to the ZF1/2 is lost, which
in turn leads to a conformational change of the domain [102, 142]. As a result, the ZF1/2
27
becomes an opened structure, and acidic residues, which are widely found in most
transcription activators, are exposed to basic transcriptional machinery. This
conformational change, in turn, causes Zap1 activation and target genes expression.
The zinc ion bound to the Zap1 ZF1/2 is labile in nature and hence relative to the
zinc bound to other ZF pairs [2, 102, 139]. These pairs include another ZF pair from Zap1
that is composed of ZF3 and ZF4 (ZF3/4). ZF3/4 is known to be critical to the DNA
binding function of Zap1 protein. Since ZF1/2 releases zinc ions rapidly under zinc‐ limiting states whereas ZF3/4 sustains the ability to bind zinc in both zinc‐deplete and replete conditions, this observation is consistent with the response of Zap1 to a low zinc level—in that AD2, where the ZF1/2 pair is embedded, is active, while the DNA binding domain containing ZF3/4 maintains an interaction with the ZREs. Finally, the critical role of ZF1/2 in the zinc responsiveness of Zap1 has been demonstrated in ZF1/2 mutants with disrupted zinc binding sites (mutated from C2H2 to C2Q2) [138]. The ZF1/2 mutants
have been shown to have defects in Zap1 activity in response to certain ranges of zinc
level compared to wild type.
1.6. Zinc homeostasis in Schizosaccharomyces pombe
S. pombe, the fission yeast, is a rod‐shaped unicellular eukaryote. It grows by tip
extension and divides by medial fission. In addition to its versatility of genetic manipulation, the fission yeast contains various characteristics that have rendered it as
an attractive model organism for zinc homeostasis research. 28
The fission yeast S. pombe is evolutionarily divergent from the budding yeast.
While Zap1 serves as a transcription factor that is central for zinc homeostasis in the budding yeast, homologs of Zap1 have not been found in the fission yeast. S. pombe, nonetheless, regulates its own growth and the expression of many genes in response to the zinc availability in the environment. It is evident that this fission yeast develops
distinct mechanisms to maintain zinc homeostasis.
Like other organisms, the fission yeast requires zinc for optimal growth, and its
zinc acquisition from the environment is mediated by zinc transporters. Unlike the
budding yeast, the fission yeast produces an MT (named Zym1) that sequesters zinc,
which plays roles in zinc homeostasis. S. pombe also synthesizes phytochelatins (PCs), a
second putative zinc‐buffering molecule. In spite of the absence of a Zap1 homolog in S. pombe, the expression of genes encoding the Zrt1 zinc uptake transporter and Zym1 are controlled by cellular zinc status [143, 144]. A recent study has identified a zinc‐ responsive transcription factor that alters these genes in the fission yeast, named Loss
Of Zinc sensing 1 (Loz1) [145]. Loz1 has been shown to regulate a range of zinc
homeostasis‐associated genes in response to zinc availability.
1.6.1. Zinc transporters
S. pombe contains 3 members in each of the ZIP family and the CDF family. The 3
ZIP members are Zrt1, Zip2 (SPCC126.09), and Zip3 (SPAP8A3.03); and the CDF proteins
29
are Zhf1, Cis4, and Zrg17. Most transporters are known to follow the rule of zinc transfer in that the ZIP increases cytosolic zinc while the CDF decreases it.
Zrt1 is the only zinc uptake system among the ZIP transporters in S. pombe [143].
Loss of the Zrt1 function severely reduces intracellular zinc accumulation, and especially
in zinc‐depleted medium, causes impaired proliferation of the cell [102, 143]. Thus, Zrt1 is essential for the growth of fission yeast during zinc deficiency. Similar to the Zrt1 in S. cerevisiae, Zrt1 in S. pombe is a high‐affinity zinc transporter and its expression is strongly responsive to zinc availability; under zinc deficiency both mRNA and protein
levels of Zrt1 increase while they decrease upon zinc repletion [143]. This zinc‐ responsiveness is controlled by Loz1, a transcription factor that represses zrt1 gene
expression in zinc‐replete cells [145].
As another member in the ZIP family in S. pombe, Zip2 is encoded by the
SPCC126.09 gene [143]. This protein is categorized in the gufA subfamily which includes
ZIP11 in mammals and Zrt3 in S. cerevisiae; ZIP11 is localized in Golgi, and Zrt3 is in the
vacuolar membrane [122, 146]. The Zip2 protein in S. pombe has been shown to be
localized in ER membranes. Its knockout mutant cells, interestingly, accumulate more zinc than wild type cells under zinc deficiency [143]. Despite these findings, the role of
Zip2 in zinc homeostasis needs to be further examined.
The third ZIP protein, Zip3 is encoded by the SPAP8A3.03 gene [143]. Although it was proposed to be similar with Yke4 in S. cerevisiae in that it belongs to the LIV‐1
30
subfamily [143, 147], the roles of the Zip3 transporter in the fission yeast have not been fully described.
Within the CDF family in the fission yeast, the most characterized transporter is
Zhf1. The zhf1 gene was identified from a mutant whose growth was severely inhibited
in zinc‐supplemented medium [148]. Additional studies revealed that Zhf1 is localized in
ER membranes and its expression affects zinc accumulation in ER, the main storage
organelle of zinc in the fission yeast [148]. The strong growth phenotypes associated
with the zhf1 mutant are consistent with Zhf1 being crucial for zinc tolerance and/or the
removal of excessive zinc from cytosol [144, 148]. In spite of its importance in zinc‐ replete cells, mRNA and protein levels of Zhf1 transporter are not regulated by zinc
availability [148]. The possibility of zinc regulation in Zfh1, nevertheless, still remains at
the post‐translational regulation level. Future studies need to illustrate whether the
transporter activity is altered in response to zinc levels.
Two other CDF family members, cis4 and zrg17 are homologs of MSC2 and
ZRG17 in S. cerevisiae, respectively [149]. The Cis4 transporter resides in cis‐Golgi
apparatus and influences Golgi membrane trafficking by mobilizing zinc into the lumen
of Golgi. It is found to physiologically bind with Zrg17, indicating that both Cis4 and
Zrg17 may form a heteromeric complex to function for zinc transport [149]. However, studies have not yet demonstrated how these transporters impact on the intracellular zinc distribution or balance. Unlike the case with Zhf1, which delivers zinc into the ER compartment, cis4 mutants do not show any growth defect in high zinc medium [147].
31
1.6.2. Metal storage proteins/peptides
S. pombe possesses 2 principal groups of metal‐binding proteins: MT (referred to as Zym1 in S. pombe) and PCs. Zym1 sequesters zinc ions in the cell [144]. For this
reason, unlike the MTs in S. cerevisiae, such as Cup1 and Crs5, which preferentially bind
with copper, Zym1 takes part in zinc homeostasis [150, 151]. In the zhf1‐deleted strain,
which is hypersensitive to zinc, overexpression of zym1 alleviates the hypersensitivity of the cell [144]. In zinc‐supplemented medium, zym1‐knockout cells accumulate less zinc
than wild types. The knockout mutants also show slight impairment of zinc tolerance,
indicating that Zym1 plays a minor role in zinc detoxification [144].
Although the protein structure of Zym1 is not yet available, the zinc binding property of Zym1 has been estimated. Compared to the 20 cysteines found in mammalian MT1, Zym1 contains 12 cysteines. This smaller number of cysteines implies that the metallothionein in S. pombe may have a lower capability to sequester zinc ions.
Consistent with this prediction, research using a cysteine‐specific reagent that releases
zinc ion from metal‐thiolate clusters revealed that Zym1 chelates 4 zinc atoms [144].
Similar to that of mammalian MTs, the level of Zym1 proteins increases
according to zinc supplementation [144, 145]. This is one of the mechanisms by which S. pombe can sequester more zinc and maintain intracellular zinc balance under zinc‐ replete conditions. The zinc‐regulated Zym1 expression is mediated by the Loz1 transcription factor [145]. 32
PCs are well‐known as cadmium‐binding peptides in the fission yeast and plants
[152, 153]. S. pombe and Arabidopsis thaliana significantly increase PC accumulations
upon cadmium exposure. The metal‐binding peptides confer the tolerance to the toxic
heavy metal in S. pombe [154]. A recent report, however, suggested the role of PC in the
tolerance of essential metal zinc in zhf1‐knockout cells [155]. It found that more PCs
accumulate in the zhf1 mutants, a strain that potentially retains higher levels of zinc in
the cytoplasm than wild type cells [155]. This finding corresponds with the increase of phytochelatin synthase (PC synthase) activity in response to greater zinc availability
[156]. These results propose that PCs may be involved in maintaining zinc homeostasis
against excessive zinc in the fission yeast.
PCs are oligopeptides that are composed of polymers of glutamate and cysteine
dipeptides with a terminal glycine, as expressed in (Glu‐Cys)n‐Gly (n=2 to 11). In contrast
to the gene‐encoded MTs, PCs are enzymatically synthesized from glutathione [157].
The enzyme, known as PC synthase, acts as a dipeptidyl transpeptidase, transferring the
Glu‐Cys moiety to glutathione or the growing PC chains [156]. In the fission yeast, PC synthase is encoded by the pcs2 gene whose homologs are found in Arabidopsis thaliana and Caenorhabditis elegans, but not in S. cerevisiae [154]. A mutant lacking
pcs2 does not synthesize PCs in response to cadmium [154, 155]. In S. pombe, therefore,
PC formations are dependent on PC synthase.
33
1.6.3. Loz1
Although S. pombe does not contain Zap1 homolog, the cell is able to maintain
zinc homeostasis. This, therefore, leads to the question of how S. pombe sustains
intracellular zinc levels against fluctuating zinc availability. While this question has been
deeply investigated [102, 148], a recent study has successfully described a vital factor
that regulates zinc homeostasis‐associated genes depending on zinc levels: Loz1 (Loss Of
Zinc sensing 1).
When mutants lacking a functional loz1 were grown in zinc‐rich medium, they displayed the typical patterns of gene expressions of zinc‐deficient cells: up‐regulation of zrt1 gene encoding zinc uptake transporter and down‐regulation of zym1 encoding
zinc metallothionein [145]. Consistent with the up‐regulation of zrt1 gene expression,
loz1Δ cells were also found to hyperaccumulate zinc [145]. This led to the hypothesis that Loz1 acts as a transcriptional repressor specifically in zinc‐replete cells.
One of the Loz1 target genes is zrt1, which encodes a high‐affinity zinc transporter on plasma membrane [145]. This Loz1‐mediated Zrt1 regulation is a critical
mechanism for the fission yeast to maintain zinc homeostasis. When the cell is exposed
to zinc‐replete conditions, the Loz1 factor represses the zrt1 expression to inhibit
further zinc uptake and preserve optimal levels of intracellular zinc. In contrast, under zinc‐deplete conditions, Loz1 is inactive and zrt1 expression is derepressed. The factor that activated zrt1 gene expression is currently unknown.
34
Loz1 also plays an indirect role in regulating zym1 expression. In contrast to the
regulation of zrt1, zym1 is expressed in zinc‐replete cells, and deletion of loz1 leads to a
decrease in zym1 expression. A potential explanation for this reciprocal regulation is
that Loz1 represses an intergenic transcript at the zym1 promoter [145]. The contingent
mechanism was illustrated by RNA blot analysis, using a probe that hybridizes to the
upstream regulatory region of zym1. The Loz1‐modified intergenic transcript is
conversely expressed with zym1 mRNA levels, suggesting a regulatory association [145].
The mutant lacking loz1 also displays constant low expressions of zym1 in either zinc
condition—deplete or replete. These results indicate that Loz1 up‐regulates the zinc
metallothionein under zinc‐abundant conditions potentially through a non‐coding RNA‐
mediated mechanism.
Loz1 also transcriptionally auto‐regulates its own expression [145]. This auto‐
regulation was observed when activity was measured in wild type cells expressing a loz1
promoter‐driven lacZ reporter. β‐galactosidase activity was controlled by zinc in the
presence of Loz1, but not in its absence. Thus, Loz1 controls its own synthesis by reducing the mRNA and protein levels of itself in zinc‐replete cells. Although its
expression level remains low in these cells, the Loz1 factor suppresses target gene transcription in the nucleus.
As pointed out in previous research, Loz1 regulates the expression of a number
of genes in a zinc‐dependent manner; however, the mechanisms by which zinc regulates
the functions of Loz1 need to be further investigated. To better understand these
35
mechanisms, it is critical to examine the functional roles of the protein domains of Loz1.
Loz1 contains a cysteine/histidine domain and a double zinc finger domain in the N‐ terminus and the extreme C‐terminus, respectively (Fig. 1.3) [145, 158]. The N‐terminal cysteine/histidine cluster is conserved in the Loz1 homologs in Schizosaccharomyces
species; however, its function is currently unknown. This domain may potentially be
related to the zinc‐regulated functions of Loz1, considering that the AD1 in S. cerevisiae
contains many cysteine/histidine residues that are important for zinc‐responsiveness of
AD1 [159].
The double zinc finger domain located in the extreme C‐terminus of Loz1 is constituted of a tandem arrangement of general C2H2 type zinc fingers. The fact that this
domain is highly conserved in Loz1 homologs suggests its important role in Loz1
functions. From an in vitro DNA binding study, the double zinc finger domain binds to a
GNNGATC cis‐acting element that is found in target genes of Loz1 [145]. This binding
capability was disrupted by a mutation in a cysteine residue, which is predicted as a
direct zinc binding site, in the first zinc finger of Loz1. These findings indicate that the double zinc finger domain mediates the zinc‐regulated DNA‐binding activity of Loz1.
36
Figure 1.3. Schematic diagram of Loz1 The Loz1 protein consists of 522 amino acids. It also contains a cysteine/histidine domain and a double zinc finger domain, which are each displayed in yellow and dark blue boxes. The numbers 1 and 2 represent the first and second zinc finger, respectively.
37
1.7. Overview
The eukaryotic cells’ repertoire to maintain zinc homeostasis includes the post‐
translational regulation of zinc transporters, as well as the ability to modulate the zinc
buffering capacity or zinc sequestration in cells. Zinc‐regulatory transcription factors are
another means by which cells can balance intracellular zinc levels by regulating the
expression of zinc transporters and/or zinc buffering agents. In this study, S. pombe was
chosen as a model organism to investigate zinc homeostasis. Using this cell as a versatile tool for genetic modification, my goal is to provide insights into the role of genes involved in zinc homeostasis.
Another advantage of utilizing S. pombe in a study for zinc homeostasis is the
presence of Loz1, a zinc‐responsive transcription factor. Most of the knowledge about zinc‐regulatory transcription factors in eukaryotic cells is derived from studies of MTF‐1 and Zap1, which are involved in regulating the expression of zinc buffering proteins
and/or zinc transporters. The fission yeast does not have a homolog of MTF‐1 or Zap1.
Instead, S. pombe expresses Loz1, which controls both the expression of genes encoding
zinc buffering proteins and zinc transporters. Thus, examining the mutants lacking Loz1
may corroborate critical information of zinc homeostasis in eukaryotic cells and provide
new information about other novel pathways regulated by zinc.
Taking the advantages of the fission yeast, this study demonstrates how various
zinc transporters play a key role in regulating cytosolic zinc levels. In chapters 2 and 3, I
utilized the ZapCY1 and ZapCY2 genetically encoded FRET‐based sensors to detect the
38
kinetics of cytosolic labile zinc in cells lacking Loz1, a specific zinc transporter, or specific zinc buffering molecules. These studies provide information which helps us to
understand how Loz1 controls zinc homeostasis. They also reveal that phytochelatins, in
addition to their well‐known role in the detoxification of toxic heavy metals, have an
important role as a zinc buffering molecule.
Chapters 4 and 5 focus on the molecular mechanisms by which zinc‐responsive transcription factors are regulated by zinc. Specifically, I examine the minimal domain that confers zinc responsiveness in the Loz1 protein. In addition, based on the current knowledge in molecular structures of Zap1, I used a novel strategy to search for zinc‐ regulatory transcription factor in humans. This research may provide insights to
understand zinc homeostasis, extending the current knowledge of zinc‐sensing mechanisms in unicellular organisms to complicated multicellular system.
39
CHAPTER 2
Monitoring intracellular zinc distribution using FRET‐ based genetic zinc sensors
2.1. Introduction
Zinc transporters in humans belong to the ZIP and CDF family proteins. The ZIP
family members generally transport zinc into the cytosol, whereas the CDF proteins play
a role in transporting zinc out of the cytosol. The S. pombe genome encodes 3 members
of the ZIP family—Zrt1, Zip2, and Zip3—and 3 members of the CDF family—Zhf1, Cis4,
and Zrg17 (See Chapter 1 for additional information). So far, a number of studies have
shown that cells lacking Zrt1 and Zhf1 have significant growth phenotypes related to
changes in zinc supply [143, 144, 148]. Additional methods that have been used to
examine the role of each of the zinc transporters include utilizing a radioactive isotope
Zn65. Atomic absorption spectroscopy (AAS) and inductively coupled plasma‐optical
emission spectrometer (ICP‐OES) have also been used to measure zinc uptake rates or cell‐associated zinc levels [143]. Studies using these methods have shown that Zrt1 directly functions in high affinity zinc uptake and that Zhf1 is necessary for transport of
40
zinc into the ER [143, 148]. Although it is clear that zinc transport genes are required for zinc transport into or out of a specific organelle, no study has examined the role of each zinc transporter in maintaining cytosolic zinc homeostasis.
In addition to the above methods, a number of tools have been developed to measure changes in the subcellular levels of zinc. These tools include small molecule fluorescent probes, such as Zinquin and FluoZin‐3 [7, 105]. One advantage of these latter molecules is that multiple zinc‐specific probes are available with varying affinities for zinc. Limitations of the small molecule fluorescent sensors include that their concentrations and localizations are difficult to control.
ZapCY1 and ZapCY2 are alternative zinc‐responsive reporters that function based
on the principles of fluorescent resonance energy transfer (FRET) [160]. The FRET‐based
sensors contain ZF1/2 from the transcription factor Zap1, flanked by a cyan fluorescent
protein (CFP) and a yellow fluorescent protein (YFP). The ability of these reporters to sense zinc results from a unique pair structure between zinc finger 1 and zinc finger 2
(ZF1/2) of Zap1. As the metal ion bound to ZF1/2 is kinetically labile, the zinc finger pair only forms when zinc is in excess and both zinc fingers bind zinc. [102]. As a consequence, when ZapCY1 is converted from the apo‐form to the holo‐form by
available zinc, the distance between CFP and YFP becomes closer, which induces FRET.
Moreover, this conversion is reversible in a manner that is dependent upon the
availability of labile zinc. ZapCY2 is a low affinity derivative of ZapCY1, in which 2 cysteines in ZF1/2 were replaced by histidines [160].
41
The ZapCY1 and ZapCY2 genetically encoded zinc sensors have several advantages compared to the chemically synthetic probes including FluoZin‐3 [104]. Both
ZapCY1 and ZapCY2 are produced by the cell itself, which allows its expression level and
localization in the cell to be controlled. Its controllable expression also allows
monitoring the kinetics of cellular zinc dynamics. The ZapCY1 FRET sensor has a high
affinity and specificity for zinc by virtue of the natural metal binding domain of Zap1. In
addition, its affinity can be adjusted by point mutations in the zinc‐binding domain,
creating a reporter with a different sensing range for zinc. For example, the lower
affinity sensor ZapCY2 is used to measure cytosolic zinc levels in mammalian cells, while
ZapCY1 has been used to measure zinc in organelles that have a relatively small pool of
the metal ion in mammalian cells [160].
To elucidate the roles that individual zinc transporters play in maintaining
cytosolic zinc levels in fission yeast, I developed ZapCY1 and ZapCY2 sensors that could
be stably expressed from the genome. I also established a system—zinc shock—to
monitor dynamic changes of cytosolic labile zinc in the cells. Using the advantages of the
sensors, in this chapter, I investigate the roles of zinc transporters Zrt1, Zhf1, Zrg17, Cis4, and Zip3 in modulating cytosolic labile zinc while zinc shock occurs. These studies will help to elucidate the role that individual zinc transporters play in maintaining the metal
balance in cells.
42
2.2. Materials and methods
2.2.1. Yeast strains and growth conditions
All of the S. pombe strains used in this study are shown in Table A of the
Appendix. Two different mediums were used for yeast cultures; yeast extract plus supplements (YES) medium (0.5% yeast extract, 3% glucose, and 750 μg/ml of histidine,
uracil, leucine, and adenine), and Zinc‐Limited Edinburgh minimal medium (ZL‐EMM), a
derivative of EMM that lacks the zinc supplement [38]. To generate zinc‐deficient cells
for zinc shock experiments, each strain was pre‐grown in YES medium to exponential
phase at 31oC, before they were washed 3 times in ZL‐EMM. The cells were then inoculated at approximately an optical density (OD595) of 0.2 into ZL‐EMM and grown for
17‐19 hours before a zinc shock was performed. For other experiments, cells were pre‐
grown in YES medium, washed, inoculated into ZL‐EMM (OD595=0.5), and grown for 17‐
19 hours in ZL‐EMM with or without the indicated zinc supplementation.
2.2.2. Plasmid construction
The plasmids, except for pJK148, pTN‐lacZ, and pTN‐zrt1‐lacZ, were generated in
this study; pTN‐lacZ and pTN‐zrt1‐lacZ were previously produced using the pTN215
vector (Table B in Appendix) [96].
To create the vector that constitutively expresses ZapCY1 and ZapCY2 in S.
pombe, a 1 kb promoter region of pgk1 gene was PCR‐amplified and introduced into the
43
KpnI and EcoRI restriction sites of JK148 (JK‐pgk1). Terminator sequences of adh4 were
also amplified using primers that contained BamHI/SacI restriction sites. PCR products
were digested with BamHI/SacI before being inserted into similar sites in the JK‐pgk1
(JK‐pgk1‐adh4T).
The original ZapCY1 and ZapCY2 in mammalian vectors were obtained from Dr.
Amy E. Palmer of the Department of Chemistry and Biochemistry at the University of
Colorado. To generate pZapCY1, the plasmid encoding ZapCY1 in S. pombe, the DNA
sequences that encode Zap1 ZF1/2 and flanking 2 proteins, CFP and YFP, were amplified
using the primers containing EcoRI and BamHI restriction sites. The PCR product was
digested with EcoRI and BamHI and cloned into similar sites in JK‐pgk1‐adh4T (pZapCY1).
The pZapCY2 for S. pombe was also produced with the same primers, enzymes, and
vectors, but the mammalian ZapCY2 vector was used as a template. All the primers that were used in this study were custom‐designed (Sigma‐Aldrich, St. Louis, MO) and are
shown in Table C of the Appendix.
44
Figure 2.1. Construction of ZapCY1 and ZapCY2 sensors
A. Schematic illustration of ZapCY1 sensor. The zinc finger 1 and 2 pair (ZF1/2 pair) is flanked by a cyan fluorescent protein (CFP) and a yellow fluorescent protein (YFP). Once ZapCY1 is converted from the apo‐form to the holo‐form by available zinc, the distance between CFP and YFP becomes closer, which induces FRET. B. Schematic diagram of the pZapCY1 sensor and the amino acid sequences of zinc finger 1 and zinc finger 2. Key amino acids required for pair formation or the coordination of zinc in the ZF1/2 pair are shown in red and blue. C. Diagram of pZapCY2. In ZapCY2, a key cysteine residue that coordinates zinc in each zinc finger has been converted to a histidine residue.
45
Fig.2.1
A
B
C
46
2.2.3. Yeast transformation
To transform the plasmids into the fission yeast, a modified lithium acetate method was performed [97]. All plasmids created from the JK148 backbone were linearized with NruI (New England Biolabs, Beverly, MA) prior to integration at the leu1 locus [98]. The lacZ reporters derived from pTN215 were digested with AatII and then transformed into ade6 locus.
2.2.4. Crossing strains and tetrad analysis
To create zinc transporter knockout strains containing an integrated‐FRET sensor gene (zrt1Δ pZapCY1, zrt1Δ pZapCY2, cis4Δ pZapCY1, cis4Δ pZapCY2, zip3Δ pZapCY1, and
zip3Δ pZapCY2), strain lacking the respective zinc transporter gene and the WT strain containing ZapCY1 or ZapCY2 were crossed. Strains crossed were of the opposite mating types so that they could produce transient diploids on malt extract plates supplemented with adenine, histidine, uracil, and leucine [99]. Following an overnight incubating on
ME, diploids were selected on EMM agar plates lacking adenine by complementing
adenine markers. Diploids were then allowed to sporulate on YES plates. Individual spores were dissected apart using a microscope (Nikon Eclipse 50i, Nikon, Melville, NY)
attached with a tetrad dissection micro‐manipulator. Once each spore formed a colony,
they were struck on both YES + G418, and EMM without leucine plates. This procedure
enabled a strain with complementing markers, KanMX6 and leu1, to be selected.
47
2.2.5. RNA extraction and Northern blot
To isolate total RNAs, a modified hot acid phenol method was used [100]. Cell pellets were collected by centrifugation at 3500 rpm for 5 min. The pellets were then resuspended in 650 μL of TES (10 mM Tris‐Cl pH 7.5, 10 mM EDTA pH 8.0, 0.5% SDS). An equal amount of acidic phenol/chloroform (phenol:chloroform:isoamyl alcohol
(125:24:1) pH 4.3; Fisher Scientific) was then added before cells were incubated at 65oC.
After 1 h, the samples were centrifuged at 13,000 rpm for 5 min. In a new tube, the upper aqueous layer from the centrifuged sample was carefully collected, and 500 μL of chloroform was added. This solution was then mixed and centrifuged at 13,000 rpm for
5 min. The upper layer was collected in a new tube and then precipitated with 50 μL 3 M
sodium acetate (pH 5.2) and 900 μL of 100% molecular biology grade ethanol. After
centrifugation at 13,000 rpm for 10 min, RNA pellets were obtained and air dried for 5
min. The pellets were resuspended in ddH2O, and their RNA concentrations were
measured with the NanoDrop 1000 spectrophotometer (Thermo Fisher Scientific,
Waltham, MA).
For the Northern blot assay, an agarose gel (1.2% Agarose/37% formaldehyde) and 1X MOPS buffer (20 mM MOPS pH 7.0, 2 mM sodium acetate, 1 mM EDTA) were prepared. 10 μg of total RNA with an equal volume of 2X loading dye (90% formamide,
1X MOPS buffer, bromophenol blue) was denatured in 65oC for 15 min. The RNA
samples were then loaded on the gel, and run overnight at 23V in the 1X MOPS buffer.
48
On the next day, the gel was stained with ethidium bromide, and ribosomal RNAs on the
gel were visualized on UV and saved as an image.
RNAs on the gel were transferred to a positively charged nylon membrane
(BrightStar‐Plus, Ambion, Austin, TX) using a capillary method. The capillary blotting
system was set up with an alkaline transfer solution (3 M NaCl and 0.01 M NaOH), the
nylon membrane, Whatman 3 mm filter paper, a stack of paper towels and a weight.
After 7‐10 h of the transfer, RNAs were fixed to the membrane using a UV crosslinker
(Stratagene, La Jolla, CA).
Membranes were hybridized with strand‐specific RNA probes, which include α‐
32P labeled dCTPs (Perkin‐Elmer, Waltham, MA). The probes were created from DNA
templates, which were amplified with gene‐specific primers (Table C in the Appendix)
using a MAXIscript T7 kit (Ambion). Probe hybridization was facilitated using UltraHyb
buffer (Ambion) or a custom hybridization buffer (0.5 M sodium phosphate pH 7.2, 7%
SDS, 1 mM EDTA) at 60oC overnight. The membrane blots were then washed in 2X SSC
(0.316 M NaCl, 0.03 M sodium citrate dihydrate) with 0.1% SDS and exposed to a
phosphor screen (Amersham Biosciences/GE Healthcare, Piscataway, NJ). The screen
was imaged with the Typhoon Phosphoimager (Amersham Biosciences/GE Healthcare).
2.2.6. Protein isolation and Western blot
Total protein extracts were prepared utilizing a modified trichloroacetic acid
(TCA) protocol [101]. Cell pellets were collected and resuspended in 0.5 mL of cold TCA 49
buffer (0.02 M Tris‐Cl pH 8.0, 50 mM ammonium acetate, 2 mM EDTA). 1 volume of 0.5
mm glass beads and 500 mL of cold 20% TCA were added to cells on ice. The cell
suspensions were vortexed twice for 1 min and cooled down on ice for 2 min in
between. Supernatants from the cell suspensions were then collected in a fresh tube and centrifuged at 13,000 rpm for 5 min. TCA precipitates were resuspended in 200 μL
Laemmli buffer and boiled for 10 min to denature proteins.
Sodium dodecyl sulfate (SDS)‐polyacrylamide gel electrophoresis (PAGE) was carried out with Mini‐PROTEAN Tetra Cell system (Bio‐Rad Laboratories, Hercules, CA). 5
μL of protein samples were loaded on an acrylamide gel (8% acrylamide/bis, 0.4 M Tris
pH 8.8, 0.1% SDS, 0.1% APS, 0.1% TEMED), and run in SDS‐PAGE running buffer (25 mM
Tris‐base, 0.2 M glycine, 0.02% SDS).
Proteins on the gel were transferred to 0.45 µm pore size PVDF membrane
(Millipore, Billerica, MA) using a wet‐transfer system (Bio‐Rad). The protein transfer was run with 70V for 2 h in the standard Towbin buffer containing 20% methanol.
After transfer, membranes were incubated for 2 h in Odyssey Blocking Buffer (LI‐
COR Biosciences, Lincoln, NE) and probed with primary antibodies overnight on rocking
platform at 4oC. As a primary antibody to detect CFP and YFP, anti‐GFP (G1544, Sigma‐
Aldrich) was chosen based on the sequence homologies of the proteins. Anti‐Act1 (Actin
1) (ab3280‐500, Abcam, Cambridge, MA) was used as a loading control. Membranes were washed 3 times with TBST (0.05% Tween‐20, NaCl 140 mM, Tris 10 mM, pH 7.5) and probed with fluorescent secondary antibodies, IRDye 800CW‐conjugated anti‐
50
mouse IgG and IRDye 680RD‐conjugated anti‐rabbit IgG (LI‐COR Biosciences) for 2 h.
Protein bands on the membranes were then imaged using the Odyssey Classic infrared
imaging system (LI‐COR Biosciences).
2.2.7. LacZ reporter assay
β‐galactosidase assays were performed using cells expressing the lacZ reporter
gene from the indicated promoter. Cells were grown in ZL‐EMM with or without zinc
supplementation overnight. Cells were spun down at 3500 rpm for 3 minutes and the
resulting pellets resuspended in lacZ buffer (60 mM Na2HPO4, 40 mM NaH2PO4, 10 mM
KCl, 1 mM MgCl2). Cell numbers of the suspension were measured using a UV‐Vis spectroscopy (Agilent Technologies, Santa Clara, CA) at an absorbance of 595 nm and recorded for later calculation.
1.5 mL of cell suspensions was mixed with 100 µL of chloroform and 100 µL 0.1%
SDS, and vortexed for 10 seconds. 3 aliquots of 500 µL mixture were placed into fresh
tubes to allow triplicate measurements. To start β‐galactosidase reaction, 100 µL O‐
nitrophenyl‐β‐D‐galactoside (ONPG; 4 mg/mL) was added to each sample and the time
was recorded. The reaction was stopped by adding 250 μL 1 M Na2CO3 once the ONPG
mixture turned light‐yellow. The time periods of the reaction till being stopped were
also saved. Cell debris from the yellow‐colored samples was spun down, and the
supernatants were examined to measure at OD420.
51
β‐galactosidase activity was calculated by the following formula: (OD420 /
(Reaction time (min) x OD600)) x 1000. An average of the triplicate samples was regarded as an experiment. The presented data displays an average of values from 3 independent experiments.
2.2.8. FRET measurement
For zinc shock, zinc‐limited cells were generated by pre‐grown in ZL‐EMM for 17‐
19 h. Cells were briefly vortexed, and equal volumes of the cells in ZL‐EMM placed in 96‐
well plates with black walls and clear bottoms (Corning Life Sciences, Corning, NY). Using
a multi‐mode microplate reader (Synergy H1, Biotek, Winooski, VT), OD595 of the cells on the plate were determined; all strains were in the similar range.
For FRET analysis, 3 different fluorescent intensities (FIs)—FI from CFP
(excitation/emission wavelengths, 434/475 nm); FI from YFP (515/535); and FI from
FRET (434/535)—were evaluated by the multi‐mode microplate reader. For the kinetics observation, 2 FI readings from CFP and FRET were recorded due to the limitation of the microplate reader. The 2 FIs were determined every 90 sec with vigorous plate shaking
steps in between. After marking the FIs in the zinc‐deficient cells for 5 min, the indicated
concentrations of zinc supplements were immediately added to each well. The plate
was the re‐loaded and read again under the same protocol for another 60 min. All the
FIs were collected in an Excel file, and the FRET ratios in each time frame (90 sec) were
calculated by dividing the FI from FRET by FI from CFP [102]. 52
2.2.9. Atomic Absorption Spectroscopy
Zinc‐deficient cells to induce zinc shock were prepared as indicated in the
previous sections. At time 0, 1 mL of cell solutions was collected and quickly spun down.
The cell pellets were washed twice with 10 mM EDTA and twice with ddH20, and frozen
down for further nitric acid digestion. While the remaining cells were treated with diverse levels of zinc supplement at time 0, the cell pellets were collected in the same
way at the indicated time points. The cell pellets were then digested by boiling them in
150 μL of metal‐free nitric acid (Fisher Scientific) for 30 min in the fume hood.
Acid‐digested samples were diluted with ddH20, and zinc contents in the samples
were measured using an AAS (Varian, Agilent Technologies). Cell numbers at each time
point were measured at OD595 using the UV‐Vis spectroscopy. The cell‐associated zinc levels were calculated by normalizing the zinc contents to the cell numbers.
2.3. Results
2.3.1. FRET‐based zinc sensors can monitor intracellular zinc distribution in the fission yeast
In mammalian cells, FRET‐based zinc sensors ZapCY1 and ZapCY2 have been used to monitor the cytosolic labile zinc dynamics upon zinc supplementation [160]. To create derivatives of pZapCY1 and pZapCY2, which could be integrated into the genome of
53
yeast, plasmid vectors were generated that expressed each sensor from the pgk1
promoter. The pgk1 promoter was used as its expression is not effected by zinc, or gene
deletions that affect zinc uptake [37, 145]. Each vector was transformed into wild type
(WT) cells to generate the strains WT ZapCY1 and WT ZapCY2.
To monitor intracellular zinc levels, it is necessary that the sensors are stably
expressed in cells without their levels being affected by any changes in zinc concentrations. To determine whether the ZapCY1 and ZapCY2 were produced in zinc‐ limited and zinc‐replete cells, WT cells expressing each sensor were examined by fluorescence microscope. This analysis revealed that both ZapCY1 and ZapCY2 reporters
were expressed in the cytosolic space and nucleus, but were excluded from the vacuole
(Fig. 2.1A and B). In addition, changes in zinc status did not alter their localization (Fig.
2.1C).
54
Figure 2.2. ZapCY1 and ZapCY2 expressions are stable in fission yeast cells
A. Cellular localization of ZapCY1 sensor. Wild type (WT) cells bearing pZapCY1 were grown in YES medium and analyzed by fluorescence microscopy. B. Localization of ZapCY2 sensor. C. Cellular localization of ZapCY1 sensor in the zinc‐deplete or replete cells. WT cells expressing pZapCY1 were grown in ZL‐EMM overnight. The cells were then supplied with or without 100 µM zinc for 2 h before all the images were taken. D. Western blot analysis of lysates from cells expressing the indicated plasmids. Cells were grown overnight in ZL‐EMM with or without 200 µM ZnCl2 supplement. Anti‐GFP was used to detect YFP and CFP, while anti‐Actin was used for actin.
55
Fig.2.2
A B
C
D
56
To examine the expression levels of the FRET sensors, Western blot analysis was
performed using an anti‐GFP antibody (Fig. 2.2D). The GFP antibody is known to react against both YFP and CFP due to the high sequence homology between them. As shown in Fig. 2.2D, no GFP band was detected in the vector‐only transformed cells. However, in
cells expressing ZapCY1 or ZapCY2, YFP and/or CFP were strongly and stably expressed
in both zinc‐deplete and ‐replete conditions. These results indicate that both FRET
sensors accumulate at a constant level in cells in a manner that is independent of zinc
status.
As ZapCY1 contains the intact ZF1/2 of Zap1, which has the potential to bind zinc, the reporter’s high‐level expression could potentially perturb zinc homeostasis. To test this hypothesis, zrt1 expression levels were determined using Northern blot analysis
(Fig. 2.3A). The transcript of zrt1 is regulated by zinc status in fission yeast [102, 145]. In the RNA blot analysis, the zrt1 expression was up‐regulated by zinc depletion and down‐ regulated by the zinc supplementation in WT cells regardless of the ZapCY1 sensor. To
quantitatively evaluate the interference of the 2 FRET sensors to zinc homeostasis in
yeast cells, a zrt1 promoter‐driven lacZ reporter construct was generated (Fig. 2.3B). The reporter construct was expressed into WT cells, which were co‐transformed with an
empty vector, pZapCY1, or pZapCY2. β‐galactosidase activity was then measured
following growth in ZL‐EMM medium supplemented with a range of zinc (0‐100 µM zinc)
(Fig. 2.3C). In the reporter assay, β‐galactosidase activity decreased in a manner
57 dependent of zinc level in all 3 strains. These findings highlight that ZapCY1 and ZapCY2 sensors do not significantly alter labile zinc levels and zinc homeostasis.
58
Figure 2.3. Both ZapCY1 and ZapCY2 sensors do not significantly interfere zinc homeostasis
A. Zrt1 gene expression in WT cell expressing pZapCY1 or empty vector. Both cells were grown in ZL‐EMM supplemented with 0, 50, 200, or 500 µM zinc. Total RNA was purified for Northern blot analysis, and the blot was incubated with strand‐specific zrt1 probe. Ribosomal RNAs were stained with ethidium bromide and are shown as a loading control. B. Construction of pTN‐lacZ and pTN‐zrt1‐lacZ. pTN‐zrt1‐lacZ is driven by zrt1 promoter. C. LacZ assay of WT cells expressing vectors, pZapCY1, or pZapCY2. WT cells with empty pJK148 vector, pZapCY1, or pZapCY2 were transformed with pTN‐lacZ (lacZ vector) or pTN‐zrt1‐lacZ (zrt‐lacZ). Each cell was grown in ZL‐EMM supplemented with 0, 1, 10, or 100 µM zinc overnight, and cell pellets were collected to measure β‐ galactosidase activity.
59
Fig.2.3
A
B
C
60
To confirm that the ZapCY1 and ZapCY2 sensors were functional, their activity
was examined by spectrophotometry in WT cells (Fig. 2.4A and B). To avoid using metal
ion chelators that could potentially interfere with zinc buffering, the activity of the FRET
sensors was measured following a zinc shock. In a zinc shock experiment, cells are grown under severely zinc‐limiting conditions, which leads to the up‐regulation of the
Zrt1 zinc uptake transporter. Due to the high levels of Zrt1 in these cells, they rapidly
accumulate zinc when the cells are resupplied with high concentrations of the mineral.
As shown in Fig. 2.4A, when a zinc shock with 0.1 µM zinc was performed on WT cells
expressing ZapCY1, there was a slight increase in the FRET ratio. This induction of FRET
ratio corresponds to an increased level of cytosolic labile zinc that can be recognized by
the FRET sensor [160]. Zinc shocks of 1‐1000 µM zinc immediately induced the FRET intensities to the maximum within 1.5 min. The similarity of the FRET curves induced by varying zinc shock levels suggested that 1 µM zinc was sufficient to saturate the ZapCY1
sensors.
To assess the saturation of ZapCY1 in WT cells, the total cell‐associated zinc after
zinc shock treatments was measured using AAS (Fig. 2.4C). In the WT cells that were pre‐ grown in zinc‐limited medium that were not exposed to additional zinc, there was no
detectable zinc that was accumulated within cells, confirming that the cells were zinc‐ limited. In contrast, zinc shocks with higher levels of zinc resulted in higher zinc
accumulation. Although zinc shock with higher levels of zinc resulted in increased
61
accumulation of zinc, there was no increase in FRET intensities above 1 µM zinc (Fig.
2.4C). This finding shows that the ZapCY1 zinc sensor is saturated by at least 1 µM zinc
shock in WT cells.
As ZapCY2 binds zinc with a lower affinity, it is expected to have a much less tight
ZF1/2 interaction in the presence of zinc. As shown in Fig. 2.4B, the increase in the FRET
ratio on zinc shock is smaller in WT cells expressing ZapCY2 than that of ZapCY1.
However, higher levels of zinc were required to saturate the ZapCY2 FRET sensor
relative to ZapCY1, consistent with it binding zinc with a low affinity. Also, the short response time within which 10 µM and 1000 µM zinc shock induced cytosolic zinc is
indicative of a very rapid delivery of zinc into the cells. Approximately 10 min after zinc
treatment, the cells with 10 µM zinc shock notably started to decrease the FRET ratio,
unlike the traces of 1000 µM; this decrease was maintained throughout the time of
observation. This reduction suggests that some of the zinc ions are released from the
sensor and transferred to another zinc‐binding ligand. The data from ZapCY2 as well as
ZapCY1, therefore, indicate that the sensors provide important information regarding
the kinetics of cytosolic zinc in real time.
62
Figure 2.4. Zinc uptake of WT cells upon zinc shock
A. Representative traces of FRET from ZapCY1 in WT cells. WT pZapCY1 cells were grown in ZL‐EMM overnight. Cells were loaded into a fluorescence microplate reader, and their CFP, YFP, and FRET intensities were observed in the resting state. After a 5 min incubation, the indicated concentrations of zinc were supplied into each well, and the fluorescence intensities were for 1 h. B. Average traces of FRET from ZapCY2 in WT cells. WT pZapCY2 cells grown in ZL‐EMM were exposed to zinc shock, and their FRET ratios were examined. The traces are averages of 3 independent experiments. Error bars represent the mean ± SD. C. Zinc accumulation upon zinc shock in WT cells. WT cells were grown in ZL‐EMM overnight and exposed to a sudden supply of zinc. Cells pellets were collected at the indicated time, and cell‐associated zinc was measured using atomic absorption spectroscopy (AAS).
63
Fig.2.4 A
B
C
64
2.3.2. The effects of loss of specific zinc transporters on cytosolic labile zinc levels
As ZapCY1 and ZapCY2 were able to detect rapid changes of zinc in the
cytoplasm, the zinc‐dependent changes in the FRET ratio in yeast mutants lacking
specific zinc transporter were next investigated. The goal of these studies was to
examine the roles of individual zinc transporters in maintaining cytosolic zinc levels. As
shown in the methods section, various mutant cells lacking different zinc transporters were manipulated to express either the ZapCY1 or ZapCY2 sensor.
Zrt1 is the gene that encodes the major zinc uptake transporter in the fission yeast. Therefore, zrt1 mutants accumulated lower levels of cells [143]. As shown in Fig.
2.5A and B, the kinetics of cytosolic zinc in the zrt1 mutant upon zinc shock were
determined using ZapCY1 or ZapCY2. Although zinc shock with 1 µM zinc led to the
maximum FRET ratio in the WT cells (Fig. 2.4A), in zrt1 mutants it failed to increase the
ratio. Moreover, higher concentrations of zinc were required to saturate the FRET
sensor relative to WT cells. Similarly higher levels of zinc were required to saturate the
low affinity sensor ZapCY2. From the moment of zinc exposure, 10 µM zinc shock gradually increased FRET, yet maintaining the FRET intensity lower than that induced by
100 µM zinc treatment (Fig. 2.5B). Thus, the FRET ratios of the ZapCY1 sensor are consistent with the reduced zinc uptake rates in zrt1 mutants (compare Fig. 5C and Fig.
4C). In addition to confirming that Zrt1 plays a main role for zinc uptake, these findings
demonstrate that zinc supply in the extracellular environment can be delivered into the
65 cytosol by other lower affinity uptake systems, presumably including Fet4 [102], when
Zrt1 is disrupted.
66
Figure 2.5. Zinc uptake of zrt1Δ cells upon zinc shock
A. Representative traces of FRET from ZapCY1 in zrt1Δ cells. Zrt1Δ cells expressing pZapCY1 were grown in ZL‐EMM and exposed to zinc shock. B. Average traces of FRET from ZapCY2 in zrt1Δ cells. The averages from 3 independent experiments are shown. Error bars represent the mean ± SD. C. Total zinc accumulation in zrt1Δ cells upon zinc shock. Cells that underwent zinc shock were collected, and zinc in the cells was examined using AAS.
67
Fig.2.5
A
B
C
68
Another well‐known zinc transporter in the fission yeast is Zhf1, which is
predicted to transfer cytosolic zinc into ER [148]. Although deletion of zhf1 results in
reduced zinc storage, no study has examined whether this mutation also affects
cytosolic zinc levels. To determine if deletion of zhf1 significantly altered cytosolic zinc
levels, the ZapCY1 and ZapCY2 FRET sensors were expressed in zhf1 mutant cells. As
shown in Fig. 2.6A, 1 µM of zinc shock was sufficient to cause immediate uptake of the
metal. However, zinc supplements higher than 1 µM repeatedly and robustly exhibited
unexpected FRET responses. After the initial increase, the FRET ratios of 10‐1000 µM
zinc shocks began to decrease due to yet unknown reasons (see discussion in 2.4).
Considering that the strain lacking zhf1 gene has growth defects in those zinc‐replete conditions in long term observation [143], these FRET responses may be related to
unknown factors associated with cell growth reduction.
The traces for FRET responses of ZapCY2 in zhf1 knockout cells are illustrated in
Fig. 2.6B. 1 µM zinc shock was sufficient to lead to the maximum FRET intensity in zhf1
mutant cells. This is in contrast to WT ZapCY2 cells, which displayed only a slight
increase of FRET with the same concentration of zinc (Fig. 2.4B). The FRET ratio curve of
10 µM zinc supplement also increased and stayed high, whereas in the WT ZapCY2 cells,
it increased but later decreased in the middle of measurement. These results suggest that the cells lacking zhf1 accumulate higher levels of labile zinc in the cytosol than WT
cells upon zinc shocks.
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As shown in Fig. 2.6C, the amount of total zinc uptake in zhf1Δ cells is much less than in WT cells—note the differences in the y‐axis values in Fig. 2.4C. This amount
corresponds with a previous report that found a lower expression of zrt1 in zhf1
knockout cells than WT cells [143]. The low level of total zinc accumulation indicates
that the dynamic changes in both zinc uptake and cytosolic labile zinc levels in the zhf1
mutants occur within low range levels of zinc.
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Figure 2.6. Zinc uptake of zhf1Δ cells upon zinc shock
A. Representative FRET responses of ZapCY1 in zhf1Δ cells upon zinc shock. Zhf1Δ cells containing ZapCY1 were grown in ZL‐EMM, and exposed to zinc shock. B. Average traces of FRET from ZapCY2 in zhf1Δ cells. 3 independent experiments were performed, and their FRET ratios were averaged. Error bars represent the mean ± SD. C. Total zinc accumulation in zhf1Δ cells upon zinc shock. Cell pellets from Zhf1Δ cells exposed to zinc shock were digested in nitric acid. Zinc contents in the acid‐digested samples were measured using AAS.
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Fig.2.6
A
B
C
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In the current study, using ZapCY1 and ZapCY2 sensors, the expected roles of
both Zrt1 and Zhf1 transporters were confirmed—Zrt1 plays a role for zinc uptake from
its extracellular environment, while Zhf1 serves as zinc storage into other cellular
compartments, most likely the ER. To test this whether other intracellular zinc
transporters also affected cytosolic zinc levels, ZapCY1 or ZapCY2 was expressed in yeast mutants lacking specific intracellular zinc transporters. These strains included zrg17Δ
cells with ZapCY1, zrg17Δ ZapCY2, cis4Δ ZapCY1, cis4Δ ZapCY2, Zip3Δ ZapCY1, and Zip3Δ
ZapCY2.
The traces of FRET ratios for ZapCY1 in zrg17 knockout cells are displayed in Fig.
2.7A. Surprisingly, the traces for ZapCY1 were not responsive to any concentrations of zinc shock. The non‐responsiveness of the reporter could be a result of reduced zinc uptake, or that zinc entered the cell but was undetected by the sensor. In regard to the question, the ZapCY2 sensor provided an answer. Zinc shock with both 10 and 100 µM zinc induced zinc influx into the cytosol in the zrg17 mutants (Fig. 2.7B). Although the increase of FRET ratios was not as high as the one seen in WT cells, it occurred
immediately after zinc shocks. This result suggests that the ZapCY1 sensor in the zrg17Δ
cells was not able to detect the zinc influx—perhaps because of already being saturated.
Zrg17 physically interacts with Cis4, and strains lacking either gene share common MgCl2‐sensitive growth defects, which are suppressed by zinc supplements
[149]. Based on these results, the 2 transporters have been suggested to form a
heterodimer complex. As a consequence, if the disruption of Zrg17 in the heteromeric
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form affects the movements of cytosolic labile zinc, the single disturbance of cis4 gene is expected to cause similar impacts on the labile zinc in the cytosol. To test this prediction
(Fig. 2.7C and D), ZapCY1 or ZapCY2 were expressed in cis4Δ cells. Upon zinc shock, the cis4Δ ZapCY1 or ZapCY2 displayed similar trends to those of zrg17Δ cells. In particular,
the FRET responses to 10 µM zinc shock in cis4 mutants slowly became lower than those to 100 µM zinc shock, which is very similar with the FRET responses in zrg17 mutants in terms of time and reduction ranges. These findings indicate that the kinetics of labile zinc in the cytosol are similarly influenced by mutation of either gene—zrg17 or cis4.
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Figure 2.7. FRET responses of ZapCY1 or ZapCY2 in zrg17Δ or cis4Δ cells upon zinc shock
A. Representative FRET responses of ZapCY1 in zrg17Δ cells upon sudden supply of zinc. Zrg17 mutants were pre‐grown in ZL‐EMM, and 6 different levels of zinc supplement were added into the cells. B. Average traces of FRET from ZapCY2 in zrg17Δ cells. 3 independent experiments were performed, and their FRET ratios were averaged. Error bars represent the mean ± SD. C. Representative FRET responses of ZapCY1 in cis4Δ cells upon zinc shock. D. Average traces of FRET from ZapCY2 in cis4Δ cells. To calculate the average, 3 independent experiments were examined. Error bars represent the mean ± SD.
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Fig.2.7
A B
C D
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The next zinc transporter examined was Zip3, a ZIP family member in S. pombe.
Although most ZIP family members are expected to increase the levels of cytosolic zinc,
Zip3 has not been known to serve this function. According to the FRET responses of
ZapCY1 and ZapCY2 in zip3Δ cells (Fig. 2.8A and B), the kinetics of zinc uptake were not interfered by the loss‐of‐function mutation in the genome. The immediate increase of
FRET upon zinc shock demonstrates that the zinc influx from extracellular environment was normal in the cells. This indicates that Zip3 is not involved in the process of zinc uptake, at least when cells are zinc‐deficient.
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A
B
Figure 2.8. FRET responses of ZapCY1 or ZapCY2 in zip3Δ cells upon zinc shock
A. Representative traces of FRET from ZapCY1 in zip3Δ cells. Zip3 mutants expressing ZapCY1 were grown in ZL‐EMM overnight. The cells were transferred into a 96 well‐plate and exposed to the indicated concentrations of zinc while monitoring FRET responses. B. Average traces of FRET from ZapCY2 in zip3Δ cells. Zip3Δ mutant cells expressing ZapCY2 were grown in ZL‐EMM and exposed to 0, 10, or 100 µM zinc shock. FRET ratios from 3 independent experiments were averaged, and the averages are shown. Error bars represent the mean ± SD
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2.4. Discussion
A FRET‐based metal binding probe containing the ZF1/2 of Zap1 was originally created to validate the conformational change of the unique zinc finger pair structure
[102]. The remarkable lability of ZF1/2 in zinc binding resulted in it being developed as an in vivo sensor to monitor labile zinc in cells. This included modifying its fluorescent proteins and linker regions to create a plasmid named ZapCY1. So far, studies with the
ZapCY1 sensor have used transiently transfected mammalian cell lines, which allow
them to monitor the FRET in individual cells or organelles using fluorescent microscopy
[160].
In the present study, the ZapCY1 sensor was permanently integrated into the
genome of fission yeast strains and used to measure labile zinc in the cytosol and nucleus in the cells. By virtue of the integration into the chromosome, the gene encoding FRET sensors are replicated during cell division, and thus the sensors are stably
expressed throughout the homogeneous population of each strain. This advantage
allowed us to determine the FRET responses of whole cell population directly from cell
cultures using the fluorescent microplate reader.
The study also utilized the ZapCY2 sensor, which has a lower affinity to zinc ion
due to amino acid replacements and identified some differences between the results
from the ZapCY1 and ZapCY2 sensors. In WT cells (see Fig. 2.4A and B), changes in
intracellular zinc levels led to dynamic changes in the FRET ratio in WT cells (above 2.5
fold), while the ZapCY2 sensor showed about 1.3 fold changes by the highest
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concentration of zinc shock. These differences in the fold‐responses between ZapCY1
and ZapCY2 result from 2 cysteine mutations in ZapCY2. These mutations alter 2
features of ZapCY2: zinc binding affinity, and conformation of the sensor. ZapCY2, which
contains mutations in critical zinc binding amino acid residues in the zinc fingers, has a
weaker binding affinity for zinc than ZapCY1, which contains 2 intact C2H2 type zinc
fingers. Also, the ZF1mut and ZF2mut in ZapCY2 hinder the tight interaction between them. The loose conformation of the 2 zinc fingers potentially makes the distances
between CFP and YFP longer, which in turn reduces FRET. The amount of FRET
responses of the ZapCY2 sensor, therefore, is limited compared to ZapCY1; however, it
is still proportional to the levels of zinc available to ZapCY2.
ZapCY2 also displays different trace patterns of labile zinc kinetics in WT cells.
This feature is particularly notable during a zinc shock with 10 µM in WT cells (see Fig.
2.4B). The differences in FRET response between the ZapCY1 and ZapCY2 FRET sensors
may have resulted from different zinc binding affinities, which are caused by the
mutations in the zinc binding domain. The lower affinity of ZapCY2 allows faster
dissociation kinetics when there is less zinc available. On the other hand, ZapCY1 has
slow off rates of ion binding, which takes longer to reach the zinc‐free state of the
sensor. The different response kinetics of the 2, thus, makes ZapCY2 the preferred sensor measuring dynamic changes in zinc ion concentrations in mammalian cells [104].
Correspondingly, in WT fission yeast cells, only the ZapCY2 sensor was capable of
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detecting dynamic reductions in cytosolic labile zinc levels. Therefore, ZapCY2 sensor
has the benefit to track the fluctuations of labile zinc levels in the cytosol in S. pombe.
Although zinc transporters in the fission yeast have been defined, with the exception of Zrt1, the roles of the transporters in changing cytosolic labile zinc levels have not been specifically studied. In a previous study, the role of Zrt1 was easily predicted by measuring total zinc accumulation rate in zrt1Δ cells. To better understand
the roles of other transporters, which reside in membranes of intracellular organelles, the present study examined the kinetics of the cytosolic labile zinc in the mutant cells
lacking each transporter gene. Using the features of both ZapCY1 and ZapCY2 sensors, the study provides some clues for the functions of the zinc transporters.
Zhf1 gene knockout cells have higher FRET ratios than WT cells, even when relatively low levels of zinc shock are introduced (see Fig. 2.5B). This finding from the present study supports previous research demonstrating that Zhf1 removes cytosolic
zinc and stores it into the ER, the main zinc storage site in S. pombe [148]. Studies with
the FRET sensors suggest that as zhf1 mutants are unable to transfer cytosolic zinc into
the ER, at least some of this zinc hyperaccumulates in the cytosol. Zinc
hyperaccumulation is known to be potentially toxic to the yeast cells. Zhf1, therefore, offers a tolerance against zinc hyperaccumulation, by delivering zinc into the ER. Such
zinc tolerance mechanisms during zinc shock, in S. cerevisiae, are mediated by the Zrc1
which mobilizes zinc into the vacuole, the main intracellular organelle to store zinc
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[126]. The cells lacking the ZRC1 gene accordingly show higher cytosolic zinc levels—
likewise in the fission yeast cells lacking zhf1.
Another potential reason behind the higher FRET ratios with low levels of zinc zhf1Δ cells could be that the mutants contain less zinc buffering molecules than WT cells. This may lead to more cytosolic zinc available to the ZapCY2 sensor. Both zhf1 mutants and WT cells, however, may similarly contain low levels of zinc buffering
proteins, such as the metallothionein Zym1. The levels are similar because they were pre‐grown in ZL‐EMM before exposed to zinc shock. Nevertheless, there are still many
other candidates of small zinc buffering molecules—citrate, glutamate, and
glutathione—that can affect the cytosolic zinc pool [103]. Whether such molecules exist in low levels in the zhf1Δ cells needs to be examined in future studies.
The experiments examining changes in FRET activity in zrg17Δ and cis4Δ cells suggested that these strains have slightly higher cytosolic labile zinc than WT cells when they are pre‐grown in ZL‐EMM overnight (see Fig. 2.7A and C). The saturation of ZapCY1 before zinc shock in zrg17Δ was verified with the findings that pre‐treatment of a zinc chelating agent made the sensor responsive to zinc shock (Fig. 2.9). As CDF family members, the 2 transporters may play roles to reduce cytosolic zinc, similar to Zhf1. In
this study, the standard zinc‐limited conditions rendered ZapCY1 in the state of apo‐
form only in the zhf1Δ cells. Why did this not occur in the 2 mutants lacking zrg17 or cis4 gene? One reason may be associated with the zinc uptake systems. It has been shown
that the zhf1Δ cells have reduced zrt1 expression and total zinc accumulation [143].
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Although the zrt1 expressions in both zrg17Δ and cis4Δ cells were not determined, my studies revealed that zinc uptake rates in zrg17Δ cells were much faster than those in zhf1Δ cells using AAS (see Fig. 2.6C). This result is consistent with the high affinity zinc uptake system being more highly expressed in the zrg17Δ cells, which may induce a slight difference of cytosolic zinc levels even after they were pre‐grown in ZL‐EMM.
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Figure 2.9. FRET responses of ZapCY1 in zrg17Δ cells upon pyrithione and zinc shock.
Representative FRET ratios of ZapCY1 in zrg17Δ cells treated with pyrithione and/or zinc supplements. The zrg17Δ cells with ZapCY1 were grown in ZL‐EMM overnight. The cells were treated with 50 µM pyrithione before exposed to 0, 10, or 1000 µM zinc.
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In addition to the findings from ZapCY1, studies with the ZapCY2 sensor suggest
that the Zrg17 and Cis4 transporters may also play a role in mobilizing the cytosolic zinc.
According to Fig. 2.7B and D, the FRET trace induced by a 10 µM zinc shock shows a slow
reduction after immediate increase in the beginning. These are in contrast to the FRET
trace in WT cells, which rapidly decrease within 10 and 20 min after zinc shock (see Fig.
2.4B). This comparison indicates that Zrg17 and Cis4 transporters may both contribute to the maintenance of cytosolic zinc levels.
Although the literature has shown that Zrg17 and Cis4 are localized to the Golgi
in fission yeast, it has not been shown whether zinc influx into this organelle is impaired
in the cells lacking both or either transporters [149]. To clearly determine the zinc‐
transferring roles of Zrg17 and Cis4, ongoing studies aim to generate cellular organelle‐
specific FRET sensors. Such sensors will be specifically localized to intracellular
compartments including the ER and Golgi. By expressing the targeted sensors in WT cells and in the zinc transporter knockout cells, these studies will help to establish the impact of Zrg17 and Cis4 on cytosolic zinc homeostasis.
For Zip3, there were no notable differences in FRET responses compared to WT cells. This finding corresponds with existing research about the transporter, which
suggests that it has no effect on zinc uptake rate and reveal that its loss does not lead to
any growth defect in zinc‐deplete or ‐replete conditions [143]. The FRET data indicates
that during zinc shock conditions, the transporter does not significantly affect zinc
uptake into cytosol and mobilization of cytosolic labile zinc.
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These results are also consistent with published studies examining the direction
of zinc transport of Zip3 in S. pombe. In S. cerevisiae, Yke4, a homolog of Zip3, is suggested to be a bidirectional zinc transporter [124]. It has been proposed to transport zinc into the ER in zinc‐replete conditions, while removing the metal from the organelle in zinc‐limited states. The involvement of cellular stress, such as temperature, may also
possibly affect the directions of zinc transfer. If Zip3 functions in a similar way to Yke3, it
is likely that in S. pombe, Zip3 may not function during 1 h period after zinc resupply, i.e.,
zinc delivery in the opposite direction—from organelles to cytosol—could not be
observed in the zinc shock conditions. Culturing the WT and zip3Δ cells in ZL‐EMM
overnight may already deplete the zinc contents in cellular organelles, such as ER. In
order to examine the role of Zip3 in zinc homeostasis using FRET sensors, future studies
should measure FRET responses in WT and zip3Δ cells during the transition from zinc‐ sufficient to ‐deficient conditions. Future studies are also needed to explore whether the cellular stresses, including high temperature, differently affect FRET in the cells.
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CHAPTER 3
Phytochelatins influence the cytosolic labile zinc pool in fission yeast
3.1. Introduction
Most zinc in eukaryotic cells is sequestered in the organelles or is bound to
proteins. In yeast, organelles such as vacuole and ER store zinc, and therefore, separate
the metal from the cytosolic space [122, 148]. Many metalloproteins also bind zinc for
their structure and function [161]. The zinc in the proteins is tightly bound, usually being
regarded as non‐exchangeable.
In addition to the zinc that is bound to protein or is sequestered, an additional pool of zinc is thought to exist in the cytosol. This pool is composed of kinetically labile
zinc that is bound to specific proteins or small molecules. The bound zinc within this
pool rapidly dissociates upon changes of zinc status in the cell or various cellular stresses
(e.g., oxidative stress) [162]. The proteins or molecules that bind zinc in a labile manner are therefore considered to be involved in zinc buffering. One well‐known zinc buffering protein is metallothionein. Metallothioneins are cysteine‐rich small proteins in animal
87
cells that can dynamically coordinate up to 7 zinc ions with different affinities. These properties allow metallothioneins to play an important role in zinc buffering [90].
Specific regulatory proteins have evolved to sense intracellular zinc levels. These factors included Zap1 which directly senses cellular zinc levels to activate target gene expressions during zinc limitation [39, 163]. Zap1 contains multiple zinc sensing domains, including a unique zinc finger pair domain. The zinc within this domain is highly labile so that it easily leaves the factor under zinc‐deficient conditions [164].
The ZapCY1 and ZapCY2 FRET sensors utilize the zinc finger pair domain from
Zap1 protein to detect labile zinc pool in the cytosol [160]. When the level of zinc increases, the high affinity sensor ZapCY1 is readily saturated. However, the low affinity
ZapCY2 sensor remains in an apo or partially apo form [160, 165]. Earlier in Chapter 2,
ZapCY2 was validated to be a better sensor to monitor the kinetics of existing cytosolic
labile zinc in the yeast cells.
Phytochelatins are a group of sulfur‐rich oligopeptides that can bind with various
metals [156]. Upon exposure of heavy metals, such as cadmium, a number of plants
induce the synthesis of phytochelatins, while animal cells up‐regulate metallothioneins.
Thus, in plants, phytochelatins have been known to be important molecules for heavy metals‐detoxification [154]. S. pombe contains a zinc‐metallothionein, known as Zym1, and is able to synthesize phytochelatins using the Pcs2 phytochelatin synthase. Although zym1 and psc2 are both up‐regulated by cadmium [81, 144], a recent study has shown that the activity of the phytochelatin synthase increases in the presence of zinc, in S.
88
pombe and in plants. These results suggest that phytochelatins, like metallothioneins,
may also play a role in zinc homeostasis.
In S. pombe, Loz1 plays a central role in zinc homeostasis by regulating the
expression of zrt1 and zym1 metallothionein. As a consequence, zinc homeostasis is
impaired in cells lacking Loz1. The present chapter investigates the effects of the loz1Δ
mutation on cytosolic zinc levels. The results show that loz1Δ cells constantly reduce zym1 expression regardless of zinc condition, and identify a potential zinc buffering role for phytochelatins in fission yeast. The data obtained from both ZapCY1 and ZapCY2
sensors demonstrates that phytochelatins may affect the cytosolic labile zinc levels.
Evidence is also presented to show that phytochelatin and the Zrt1 transporter allows loz1Δ cells to have an increased tolerance to zinc.
3.2. Materials and methods
3.2.1. Yeast strains and growth conditions
All of the S. pombe strains used in this study are shown in Table A of the
Appendix. Strains expressing ZapCY1 or ZapCY2 were generated either by the transformation of pZapCY1 or pZapCY2 plasmid, or genetic crosses. The cell growth
conditions were the same as described in Chapter 2.
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3.2.2. Plasmids
All plasmids were previously produced using the JK148 and TN215 backbone (see
Table B in Appendix).
3.2.3. Crossing strains and tetrad analysis
To generate pcs2Δ ZapCY1 and pcs2Δ ZapCY2 cells, the opposite mate types of pcs2Δ cells and WT ZapCY1 or ZapCY2 were crossed and were then subject to tetrad dissection analysis. Loz1Δ pcs2Δ ZapCY1 and loz1Δ pcs2Δ ZapCY2 cells were created by crossing pcs2Δ cells with loz1Δ ZapCY1 or loz1Δ ZapCY2. In our studies we found that a
number of gene knockout strains generated by transformation with ZapCY1 or ZapCY2,
showed different expression levels of the sensors. Therefore, in later studies, strains
expressing the FRET sensors were generated by crossing in order to avoid the
discrepancy in expression levels of the sensors.
3.2.4. Serial dilution growth assays
To examine the normal growth rates, the cells were pre‐grown in YES overnight.
The OD of the cells was then measured, and each strain diluted to an OD595=1.0. 5 μL of
the serial diluted cells were then plated on YES agar plate supplemented with the indicated zinc levels. The plate images were taken after 2‐4 d of incubation in 31oC. To compare their growth upon zinc shock, cells in YES were washed 3 times with ZL‐EMM and transferred into ZL‐EMM. After 17‐19 h incubation in ZL‐EMM, the serial dilutions
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from the zinc‐deficient cells were prepared. The next day the growth rates of the cells
were determined and recorded as images.
3.2.5. FRET measurement
The FRET responses upon zinc shock were monitored as previously described in
Chapter 2. To examine the roles of chelators on FRET signals, N,N,N',N'‐tetrakis (2‐
pyridylmethyl) ethane‐1,2‐diamine (TPEN), EDTA, and pyrithione were prepared—250
μM TPEN in DMSO, 250 μM EDTA, and 50 μM pyrithione in ddH2O. Chelators and zinc
were added at the specific times indicated in the figures.
3.2.6. Northern blot
WT and loz1Δ cells were pre‐grown in YES to exponential phase and transferred
into ZL‐EMM supplemented with or without 200 μM ZnSO4. Total RNA extracts were
purified following growth overnight. The northern blot assay was performed as
described in Chapter 2. Zym1 and pgk1 probes were generated from DNA templates
which were amplified with zym1 and pgk1 gene‐specific primers (Table C in the
Appendix).
3.3. Results
3.3.1. Loz1 knock‐out strains are resistant to zinc toxicity
Zrt1 and zym1 are tightly regulated by zinc in a manner that is dependent upon
Loz1 [145]. In WT cells, zrt1 is up‐regulated in zinc‐deficient conditions, while zym1 gene 91
expression decreases in zinc‐deplete cells (Fig. 3.1A). In cells lacking loz1, both genes are not regulated by zinc. Specifically, zrt1 transcript accumulates to high levels in a manner that is independent of zinc availability, and zym1 transcript levels are constantly low. In addition to the above changes in gene expression, loz1Δ cells accumulate higher level of zinc than WT cells under zinc‐replete conditions [145]. Thus, loz1 knockout cells grown
in high zinc‐medium hyperaccumulate zinc, but their gene expressions show patterns
similar to those in zinc‐deficient cells. Based on the findings that loz1Δ cells
constitutively produce the Zrt1 transporter and contain more zinc in the cell, I
hypothesized that they might also be hypersensitive to zinc.
To determine whether loz1Δ cells had growth defects, serial dilution growth assays were performed with the cells. WT, zhf1Δ, and zrt1Δ cells were also plated as controls. All the strains were grown overnight in YES and were then spotted in 10‐fold
serial dilutions onto YES medium with or without various supplements. As expected, the
mutants lacking zrt1, as shown in Fig. 3.1B, exhibited impaired‐growth on YES
supplemented with EDTA. This zinc‐deficient phenotype was suppressed by 200 µM of
zinc supplements. Consistent with published studies, zhf1Δ cells showed growth defects
on zinc‐supplemented medium, indicating that they are hypersensitive to zinc [143,
148]. Although hypothesized to be hypersensitive to zinc, loz1Δ cells were able to grow
on 200 µM zinc‐supplemented medium. Moreover, the mutants lacking loz1 grew well
on YES medium with a 2 mM zinc supplement where WT cells and even zrt1Δ cells
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possessed severe growth defects. Thus, loz1Δ cells have an increased zinc tolerance
despite constitutive expression of zrt1.
To determine whether the unexpected phenotype of loz1Δ cells resulted from a
loss of loz1 gene function, the mutants were transformed with pLoz1 (Fig. 3.1C), a
plasmid containing the loz1 ORF under the control of its native promoter [145], and
growth was examined with the serial dilution growth assay. As shown in Fig. 3.1D, pLoz1
suppressed the phenotype of loz1Δ cells, restoring zinc toxicity of the cells on YES supplemented with 2 mM zinc. This data indicates that the high zinc tolerance was the
effect of loz1 gene deletion from the genome. All in all, the loss of loz1 gene function leads the cells to accumulate high levels of zinc, and increased survival in zinc‐replete conditions.
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Figure 3.1. Cells lacking loz1 are resistant to zinc toxicity A. Zrt1 and zym1 gene expressions in wild‐type (WT) and loz1Δ cells. WT and loz1Δ cells were grown in ZL‐EMM supplemented with or without 200 µM zinc. Total RNA was extracted and subjected to Northern blot analysis. The blot was probed for zrt1, zym1, and pgk1 (phosphoglycerate kinase 1). Ribosomal RNAs in the gel were stained with ethidium bromide. Pgk1 and ribosomal RNAs were shown as loading controls. B. Growth assays showing the different growth rates of WT, loz1Δ, zhf1Δ, zrt1Δ, and zym1Δ strains. The cells grown overnight in YES medium were spotted in 10‐fold serial dilutions onto plates containing 100 µM EDTA, 200 µM, or 2 mM ZnCl2. Plates were incubated for 2 d at 31 °C before photography. C. Schematic diagram of pLoz1. The predicted loz1 5’ UTR (green rectangle) and loz1 ORF (blue rectangle) are indicated. D. Serial dilution growth assays showing the differences of phenotype. WT cells transformed with empty vector and loz1Δ cells introduced with empty vector or pLoz1 were grown in YES. 10‐fold serial dilutions from each strain were plated in YES containing 100 µM EDTA, 200 µM, or 2
mM ZnCl2.
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Fig.3.1
A
B
C
D
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3.3.2. Loz1 controls cytosolic zinc distribution
The paradoxical findings from loz1Δ cells raise the question of where in the cells do they accumulate zinc in order to survive in extremely high zinc environments. This leads to the need to investigate the intracellular zinc distribution in these cells— especially the labile zinc in cytosol, which is known to be critical to the growth of cells
[143, 148]. To examine the kinetics of cytosolic zinc, I created loz1Δ cells expressing
either ZapCY1 or ZapCY2, 2 sensors that are highly suitable for monitoring zinc levels in
cellular compartments.
To determine if there were any differences in the cytosolic labile pool of zinc in
loz1Δ cells, the FRET ratio was measured in loz1Δ cells expressing ZapCY1 and ZapCY2
following a zinc shock (see Chapter 2). In Fig. 3.2A, the FRET traces of ZapCY1 in loz1Δ
cells upon zinc shock with 0.1‐1000 µM zinc revealed very little change in the FRET ratio.
This lack of response in loz1Δ cells contrasts with the response of ZapCY1 in WT cells, in
which the FRET ratio increased over 2.5 fold following a 1‐1000 µM zinc shock. Taking into account that loz1Δ cells stably up‐regulate zrt1 mRNA expression and hyperaccumulate zinc in zinc‐replete medium, the absence of any major increase in the
FRET ratio suggested that the ZapCY1 sensor might be saturated in this background.
As presented in Fig. 3.2C, further experiments were conducted to determine if the ZapCY1 sensor was already saturated with zinc in loz1Δ cells following growth in
zinc‐limited medium. Loz1Δ cells accumulated zinc in a manner dependent of the levels
of zinc supplement. In addition, the total zinc that was accumulated in loz1Δ cells upon
96
zinc shock with 100 µM was higher than in WT cells (see Fig. 2.4C). This finding is
consistent with the increased levels of zrt1 transcript in zinc‐replete conditions.
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Figure 3.2. Zinc uptake of loz1Δ cells upon zinc shock A. Representative traces of FRET from ZapCY1 in loz1Δ cells. Loz1Δ cells containing ZapCY1 were grown in ZL‐EMM overnight and loaded into fluorescence microplate reader. After 5 min incubation, the cells in a 96 well‐plate were supplied with the indicated concentration of zinc. FRET ratio was investigated for 1 h. B. Average traces of FRET from ZapCY2 in loz1Δ cells. Loz1Δ cells expressing pZapCY2 were grown in ZL‐EMM before the addition of 0.1‐1000 µM zinc at t=0. The FRET traces present the averages of 3 independent experiments. Error bars represent the mean ± SD. C. Total zinc accumulation in loz1Δ cells upon zinc shock. Cell pellets from the loz1Δ mutants exposed to zinc shock were collected and digested in nitric acid. Zinc contents in the acid‐ digested samples were measured using AAS.
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Fig.3.2
A
B
C
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The ZapCY1 sensor contains a zinc‐binding domain that has a high affinity to zinc
[160]. As a consequence, if it is occupied with intracellular zinc, the sensor will neither
change its conformation nor produce FRET signals. To determine if the ZapCY1 sensor
was saturated in loz1Δ cells, 2 zinc chelators—TPEN and pyrithione—were used to
examine if they could render ZapCY1 to an apo state.
TPEN is commonly used as the permeable metal chelator in mammalian cells. In
yeast cells, however, its effect has not been rapidly observed [166]. To examine the
effect of TPEN on ZapCY1 in yeast cells, the cells were exposed to TPEN while their FRET
responses were examined (Fig. 3.3A and C). In Fig. 3.3A, the WT cells pre‐grown in ZL‐
EMM overnight were treated with 0, 50, or 200 µM TPEN, and after 20 min, supplied
either with or without 1 mM zinc. Compared to the cells that were not treated with
TPEN, the TPEN‐treated cells display slightly lower and stable FRET ratios. There were no differences in FRET between 50 µM and 200 µM TPEN‐treated cells. After zinc supplement, the TPEN‐treated and untreated cells demonstrated the similar FRET
responses. The FRET ratios reached the maximum because the amount of zinc
supplement was above the 1:1 stoichiometric ratio of TPEN to zinc ion, canceling out the metal chelating effect of TPEN.
In Fig. 3.3B, the ZapCY1 in loz1Δ cells were exposed to TPEN. As in WT cells, no increase in the FRET ratio was observed upon TPEN treatment of loz1Δ cells. Upon zinc shock, however, none of the TPEN‐treated and untreated loz1Δ cells showed a
significant increase in FRET, i.e., 0, 50, or 200 µM TPEN‐treated cells, all of which were
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supplied with 1 mM zinc, exhibited the same FRET ratio as the vehicle‐treated control group. It was also found that the FRET ratio decreased throughout the time of observation in the TPEN‐only treated loz1Δ cells. These findings imply that TPEN did not deplete zinc from the ZapCY1 sensor in loz1Δ cells, but instead, consistent with published studies, preferentially chelates zinc from the extracellular environment.
As a second experiment to examine changes in FRET response following TPEN treatment, WT cells were pre‐grown in zinc‐rich YES medium in order to mimic the
potential conditions of the ZapCY1 sensor in loz1Δ cells before the addition of TPEN. The
data in Fig. 3.3C shows that the holo form of the ZapCY1 sensor—fully saturated by zinc
in YES medium—does not respond to additional zinc supply. Also, 250 µM TPEN, which
is 25 times higher than the level used to unsaturate ZapCY1 in mammalian cells [160],
was not able to dissociate zinc ions from the holo form of the sensor. These results are
consistent with the report that TPEN does not rapidly show chelating effects across
membrane in the yeast cells [166].
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Figure 3.3. ZapCY1 sensor in loz1Δ cells may still be saturated upon TPEN treatment in short‐term A. Representative traces of FRET from ZapCY1 in WT cells that were treated with 0, 50, or 200 µM TPEN, and 0 or 1 mM zinc. WT cells containing ZapCY1 were grown in ZL‐ EMM overnight. While monitoring the FRET responses, 200 µM TPEN or vehicle (DMSO)
was added to the cells, followed by 1 mM zinc or vehicle (ddH2O) treatment. B. Representative traces of FRET from ZapCY1 in loz1Δ cells. The loz1Δ mutants bearing ZapCY1 were grown and examined for FRET under the same conditions as WT cells. C. FRET ratios of ZapCY1 sensor in WT cells grown in YES. WT cells containing ZapCY1 were grown in YES and exposed to 1 mM zinc, 250 µM TPEN, or vehicles.
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Fig.3.3
A
B
C
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As TPEN did not noticeably reduce zinc levels in yeast, an alternative zinc
chelator, pyrithione, was utilized. Pyrithione is a zinc ionophore, which chelates zinc in
the extracellular environment and then diffuses to the intracellular space together with
the metal. On the other hand, in the absence of zinc in the extracellular space, it crosses
the membrane and binds with the metal in the cell [167]. The ZL‐EMM medium that is
used in the zinc shock method does not contain any added zinc. Thus, pyrithione is
acting as a zinc chelator in the studies.
To determine if the ZapCY1 sensor was saturated, cells were pre‐grown in ZL‐
EMM overnight before pyrithione was added to cells. As shown in Fig. 3.4A, when WT cells expressing ZapCY1 were treated with 50 µM pyrithione, there was a slight but rapid decrease in the FRET signal. Upon zinc shock, compared to the FRET‐increase rate in WT
cells without pyrithione (see Fig. 2.4A), the overall FRET responses were slowed down; 1
µM zinc shock did not induce the maximum FRET intensity, while 10 and 100 µM zinc shock took a few minutes to reach the maximum. These findings are consistent with pyrithione entering yeast and chelating zinc.
To further examine the role of pyrithione on ZapCY1, a zinc shock experiment in
the presence of pyrithione was performed with zrt1Δ and zhf1Δ cells expressing the
ZapCY1 FRET sensor. In zrt1Δ cells, the chelator had no effect on the FRET ratio. This may be attributed to the fact that the mutants were already in severe zinc‐deficient
states before adding pyrithione. In contrast, zhf1Δ ZapCY1 cells exhibited similar decreases of FRET compared to WT ZapCY1 cells (Fig. 2.4C). In addition, zhf1Δ ZapCY1
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cells displayed more rapid increases of FRET upon 1 µM zinc shock, which supports the
findings in Chapter 2. Interestingly, in the presence of pyrithione, the atypical FRET
curves by both 10 and 100 µM zinc shock observed in zhf1Δ ZapCY1 cells were not
observed in the mutants treated with pyrithione.
When similar experiments were performed with loz1Δ ZapCY1 cells, a
considerable decrease in the FRET ratio was observed upon pyrithione treatment.
Moreover, after 20 min when zinc was added to the cells, there was a significant increase in the FRET ratio. These results suggest that ZapCY1 sensor was converted from the holo state to apo state by pyrithione. The data underlines that loz1Δ cells accumulate more labile zinc in the cytosol in ZL‐EMM, rendering ZapCY1 to be
saturated.
The results that illustrate ZapCY1 to be saturated in loz1Δ cells suggested that
additional information would be obtained in this strain using the lower affinity ZapCY2
sensor. Due to its lower affinity zinc‐binding domain, ZapCY2 easily releases zinc ions
depending on the kinetics of labile zinc [160]. This characteristic enables the dynamic changes of cytosolic labile zinc to be monitored. The traces of FRET responses of ZapCY2 in the mutant cells lacking loz1 gene are presented in Fig. 3.2B. Followed by zinc shock,
FRET ratios gradually decreased during 60 min. While WT cells maintain certain levels of
cytosolic labile zinc upon zinc shock (see Fig. 2.4B), a similar effect on the FRET ratio was observed with by 10‐1000 µM zinc shocks. This data implies that although zinc is bound
to the sensor upon the initial zinc shock, zinc moves to other molecules in the cytosol or
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other cellular compartments during the 1 h post zinc shock. In this regard, the finding from ZapCY2 may reflect crucial clues to solve the question of where in the cells do they accumulate zinc in order to survive in extremely high zinc environments.
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A B
C D
Figure 3.4. ZapCY1 sensor in loz1Δ cells becomes an apo form by pyrithione
A. Representative FRET ratios of ZapCY1 in WT cells treated with or without pyrithione and zinc supplements. The WT cells with ZapCY1 were grown in ZL‐EMM overnight. The
cells were treated with 50 µM pyrithione or vehicle (ddH2O) before exposed to 0, 1, 10, or 100 µM zinc. B, C, and D. Representative FRET responses of ZapCY1 in zrt1Δ, zhf1Δ, and loz1Δ cells, respectively. All growth conditions and FRET measurement methods were identical with those for WT cells. 107
3.3.3. Overexpression of zrt1 is required to accumulate more zinc and modulate the
kinetics of labile zinc in loz1Δ cells
To start understanding the phenotypes associated with loz1Δ cells, an important
question to address is: which genes are necessary for loz1Δ cells to accumulate more
zinc following growth in zinc‐replete medium? A potential clue to this problem was that loss of loz1 gene function disrupts the regulation of zrt1 expression. Although both WT
and loz1Δ cells up‐regulate zrt1 expression in ZL‐EMM, in YES medium, only loz1Δ cells
up‐regulate the zrt1 and accumulate zinc under zinc‐replete conditions [145]. I,
therefore, hypothesized that overexpression of zrt1 results in higher levels of zinc in
loz1Δ cells.
To demonstrate the role of zrt1 gene expression in the loz1Δ cells, mutants
lacking both loz1 and zrt1 genes were created. The uptake rates and kinetics of cytosolic
labile zinc were then measured using the ZapCY1 and ZapCY2 sensors. In Fig. 3.5A,
ZapCY1 sensor in the double mutant cells responded to the various concentrations of
zinc shocks. This represents that the sensor was in the zinc‐free state indicating that the
loz1Δ zrt1Δ cells contain less zinc in the cytosol. Overall, the loz1Δ zrt1Δ double mutant
has a similar uptake rate for zinc as the single zrt1Δ cells. Unexpectedly, the maximum
FRET ratio of loz1Δ zrt1Δ cells was lower than zrt1Δ cells’. However, when a loz1Δ zrt1Δ
ZapCY1 strain was created by crossing of loz1Δ zrt1Δ cells and WT ZapCY1 cells, the
double knockout cells and the single zrt1Δ cells showed the same values of maximum
FRET ratio. These results suggest that different transformations may express ZapCY1 at
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different levels, and future studies should, therefore, use strains generated via genetic
crosses. In the Fig. 3.5B, the ZapCY2 sensor also displayed similar patterns of FRET ratios
with those in zrt1Δ cells. Moreover, as the FRET ratio remained high in zinc shocked loz1Δ zrt1Δ cells, these results suggest that the ZapCY2 sensor in loz1Δ zrt1Δ cells does
not readily lose zinc. Interestingly, loz1Δ zrt1Δ cells could also not survive in very high
zinc‐containing medium, suggesting that Zrt1 is critical to the increased tolerance of
loz1Δ cells to high zinc (Fig. 3.5C).
The role of zrt1 in regulating the kinetics of cytosolic labile zinc in loz1Δ cells is
underscored in the data from loz1Δ zrt1Δ cells. To further explore the role of zrt1 in the
zinc tolerance of loz1Δ cells, I utilized a plasmid that expresses the zrt1 ORF from the
constitutive pgk1 promoter (Fig. 3.5D). To confirm that this plasmid was function, it was
initially integrated into the genome of zrt1Δ cells, and its ability to transfer zinc into cells
examined using growth assays. Fig. 3.5E shows that the JK pgk1‐zrt1 plasmid rescued
zrt1Δ cells from severe growth defects on the zinc‐limited plate. Zrt1Δ cells expressing the plasmid, however, exhibited less growth on the YES plate.
To investigate the effects of zrt1 overexpression on the growth rate in extremely
high zinc conditions, WT cells transformed with the JK pgk1‐zrt1 or the empty vector
were incubated at 31 °C for 4 d. In the wild‐type background, overexpression of zrt1 had
no effect on growth in YES medium and was inhibitory to growth in the presence of 2 mM zinc. This result indicates that the highly constant expression of zrt1 is not sufficient to enable cells to have zinc tolerance in zinc‐replete conditions.
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Figure 3.5. Zrt1 is required for loz1Δ cells to have zinc tolerance, but is not sufficient to confer the tolerance to WT cells A. Representative FRET traces of ZapCY1 in loz1Δ zrt1Δ cells. The loz1Δ zrt1Δ cells expressing ZapCY1 were generated by transformation of pZapCY1 into loz1Δ zrt1Δ cells. The double knockout mutants containing ZapCY1 were grown in ZL‐EMM and underwent zinc shock. B. Average traces of FRET from ZapCY2 in loz1Δ zrt1Δ cells. 3 independent experiments were performed, and their FRET ratios were averaged. Error bars represent the mean ± SD. C. Growth assay of WT, loz1Δ, zrt1Δ, or loz1Δ zrt1Δ cells. Each strain was grown in YES, and 10‐fold serial dilutions of the cells were spotted onto YES plates supplemented with or without 2 mM zinc. D. Schematic diagram of JK pgk1‐ zrt1 plasmid. The predicted pgk1 promoter (green rectangle) and zrt1 ORF (blue rectangle) were cloned into pJK148 backbone. E. Growth assay showing the differences of growth rates of WT vector, zrt1Δ vector, and zrt1Δ JK pgk1‐zrt1. The cells were grown in YES and plated on YES or YES + 100 µM EDTA medium. F. Growth assay showing phenotypic differences between WT vector and WT JK pgk1‐zrt1. Both cells cultured in YES were spotted onto YES plates supplemented with or without 2 mM zinc.
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Fig.3.5
A B
C
D
E
F
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3.3.4. Zym1 reduction is not responsible for zinc hyperaccumulation and tolerance of
loz1Δ cells
The above results suggest that Zrt1 is necessary for the increased levels of zinc that accumulate in loz1Δ cells and saturate the ZapCY1 sensor, but do not explain why
the ZapCY2 sensor readily loses zinc in this background. One possible explanation is that
the buffering capacity of the cytosol for zinc was altered in the loz1Δ cells. Zym1, a
metallothionein in S. pombe, is a small zinc protein, which sequesters zinc in the cytosol
[144]. In loz1Δ cells, however, zym1 expression is down‐regulated.
To determine the role of Zym1 in loz1Δ cells, the FRET responses from loz1Δ
zym1Δ cells containing ZapCY1 or ZapCY2 were examined (Fig 3.6A and B). Consistent with the low levels of zym1 expression, its absence in loz1Δ cells did not result in any different patterns of FRET from both sensors. This result indicates that the small amount of Zym1 in loz1Δ cells does not play any major role in the zinc buffering capacity in loz1Δ
mutants.
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A
B
Figure 3.6. FRET responses of ZapCY1 or ZapCY2 in loz1Δ zym1Δ cells upon zinc shock A. Representative FRET traces of ZapCY1 in loz1Δ zym1Δ cells upon zinc shock. The loz1Δ zym1Δ cells mutants containing ZapCY1 were grown in ZL‐EMM and underwent zinc shock. B. Representative FRET traces from ZapCY2 in loz1Δ zym1Δ cells upon zinc shock. Loz1Δ zym1Δ cells expressing ZapCY2 were grown in ZL‐EMM and supplied with the indicated concentrations of zinc.
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Since metallothioneins are known to have roles in buffering cytosolic zinc levels,
additional experiments were performed to test the effects of overexpressing Zym1. For
this, strains were generated, which co‐expressed ZapCY1 or ZapCY2 FRET sensor and
overexpressed the metallothionein gene of S. pombe (plasmid TN pgk1‐zym1) or the
human metallothionein1 (MT1) gene (plasmid TN pgk1‐MT1) (Fig. 3.7A). When zinc shock was performed on loz1Δ cells expressing ZapCY2 and TN pgk1‐zym1 or TN pgk1‐
MT1, no significant difference was identified in the FRET signals, compared with the
FRET of loz1Δ cells that contain an empty vector (Fig. 3.7B). As metallothioneins bind zinc, the effects of their overexpression on zinc tolerance were also tested with serial dilution growth assay (Fig. 3.7E). Loz1Δ cells expressing the yeast or human metallothionein genes were resistant in high zinc environments. However, the tolerance for zinc was similar to original loz1Δ cells, which express low basal levels of zym1 (see vector only control). These results indicate that zym1 does not play significant roles in the kinetics of cytosolic labile zinc and zinc tolerance in loz1Δ cells.
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Figure 3.7. Overexpression of metallothioneins does not nullify the zinc tolerance of loz1Δ cells A. Schematic diagrams of TN pgk1‐zym1 and TN pgk1‐MT1 plasmids. The predicted pgk1 promoter (green rectangle) and zym1 ORF (blue rectangle) or human metallothionein1 (MT1) ORF (purple rectangle) were cloned into pTN215 backbone. B. FRET responses of ZapCY2 in loz1Δ cells expressing empty vector. The loz1Δ mutants bearing ZapCY2 and TN pgk1‐zym1 were grown in ZL‐EMM and underwent zinc shock. C. and D. ZapCY2 FRET responses in loz1Δ cells expressing TN pgk1‐zym1 or TN pgk1‐MT1, respectively. E. Growth assay showing phenotypic differences. WT empty vector, loz1Δ empty vector, loz1Δ TN pgk1‐zym1, and loz1Δ TN pgk1‐MT1 cells were grown in YES, and the 10‐fold serial dilutions from each strain were spotted onto YES plates supplemented with or without 2 mM zinc.
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Fig.3.7
A B
C
D
E
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3.3.5. Phytochelatins play a role for zinc buffering in loz1Δ cells
The very limited roles of zym1 in loz1Δ cells suggested that a different metal
buffering system is more important in zinc‐limited cells. As recent study suggested that
phytochelatins may play an important role in zinc detoxification in the fission yeast, I next examined whether PC levels affect the FRET response [155]. In S. pombe, the pcs2
gene encodes a phytochelatin synthase. As a consequence, cells lacking pcs2 are not
able to synthesize phytochelatins [154].
To investigate the zinc buffering capacity of phytochelatins, pcs2Δ cells
expressing ZapCY1 or ZapCY2 were generated. Fig. 3.8A shows the FRET responses of
ZapCY1 in pcs2Δ cells. Surprisingly, the high affinity sensor repeatedly produced high
FRET signals upon a very low level of zinc shock. Whereas it always induced a slight higher FRET than ddH2O in WT cells (see Fig. 2.4A), in pcs2Δ cells, the 0.1 µM zinc supply
caused a rapid increase of FRET reaching close to the maximum intensity. In contrast, the FRET responses of ZapCY2 in pcs2Δ cells were similar to those in WT cells (Fig. 3.8B).
Unlike ZapCY1, the low affinity sensor was not able to detect the loss of zinc buffer between pcs2Δ and WT cells. These findings suggest that the loss of phytochelatin affects the pool of zinc that is available to the high affinity sensor, but not the low affinity sensor in zinc‐limited cells.
The role of phytochelatins in loz1Δ cells, which hyperaccumulate zinc, was investigated with loz1Δ pcs2Δ double mutants. In this background, the ZapCY1 sensor in was saturated, as was observed in loz1Δ single mutants (Fig. 3.8C). On the other hand,
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the ZapCY2 FRET traces in loz1Δ pcs2Δ cells were not similar with those in loz1Δ cells
(Fig. 3.8D). The FRET ratio following a 10 µM zinc shock gradually decreased after the initial maximum obtained at 1.5 min post zinc shock, but was maintained at a slightly higher value than the control. The ratio after a 1000 µM zinc shock also remained higher than that in the loz1 single mutant cells (see Fig. 3.2B). These patterns of FRET responses, hence, appear to take the average values of the patterns in WT and loz1Δ cells. This finding suggests that the phytochelatin synthase is associated with the unique kinetics of cytosolic labile zinc in loz1Δ cells. Taking into account the zinc binding ability of the enzyme’s product, an unknown zinc buffering molecule that chelates the metal upon zinc shock in loz1Δ cells may be phytochelatin. These results are supported by the
growth assay using in Fig. 3.8E. When loz1Δ pcs2Δ cells were transferred from ZL‐EMM
onto zinc‐replete plates, they had growth defects compared to the loz1Δ single mutants.
These results are consistent with PCs having a role in zinc detoxification upon sudden supply of the metal.
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Figure 3.8. Phytochelatin may play a role for zinc buffering in the loz1Δ cells that were exposed to zinc shock A. Representative FRET traces of ZapCY1 in pcs2Δ cells. The mutants containing ZapCY1 were grown in ZL‐EMM and supplied with various levels of zinc supplements. B. Average traces of FRET from ZapCY2 in pcs2Δ cells upon zinc shock. Pcs2Δ cells containing ZapCY2 were grown in ZL‐EMM and underwent zinc shock. The FRET traces present the averages from 3 independent experiments. Error bars represent the mean ± SD. C. Representative FRET traces of ZapCY1 in loz1Δ pcs2Δ cells upon zinc shock. D. Average FRET traces from ZapCY2 in loz1Δ pcs2Δ cells upon zinc shock. E. Growth assay of WT, loz1Δ, pcs2Δ, and loz1Δ pcs2Δ cells underwent zinc shock. Each strain was grown in ZL‐ EMM overnight, and 10‐fold serial dilutions from each were prepared with ZL‐EMM. The cell dilutions were spotted onto YES supplemented with 0, 200 µM, or 2 mM zinc. Images of the plates were taken on the next day.
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Fig. 3.8
A B
C D
E.
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3.4. Discussion
In S. pombe, Loz1 plays an important role in zinc homeostasis by regulating the
expression of genes that are critical for zinc uptake and zinc storage [145]. Key Loz1
target genes include zrt1 and zym1. In zinc‐replete conditions, Loz1 represses zrt1 gene
expression while it indirectly induces zym1 transcript. This regulation helps cells to
balance zinc levels and survive in high zinc medium. Accordingly, loss of Loz1 function was anticipated to have hypersensitivity to zinc, leading severe growth defects in high zinc environments. However, surprisingly, loz1Δ cells grow very well in such conditions where WT cells suffer from zinc toxicity. The ultimate goal of this present study was to investigate the mechanisms underlying these paradoxical characteristics of loz1Δ cells.
Based on monitoring the kinetics of cytosolic labile zinc in loz1Δ cells upon zinc
shock, sudden external zinc supply causes a rapid influx of the metal into the cytosol.
The labile zinc in the cytosol is, however, sequestered by unknown molecules and/or transferred into other compartments.
The present study suggested that phytochelatins take part in chelating the cytosolic labile zinc and controlling its levels. In S. pombe, upon cadmium exposure, Pcs2 synthesizes phytochelatins as a mechanism to protect cells from cadmium toxicity [154].
In addition, high levels of zinc result in increased activity of the enzyme and the levels of phytochelatin [157]. The loss of pcs2 function also decreases the survival rate of zhf1Δ
cells in zinc‐replete conditions [155]. Together these results suggest that phytochelatins
may play an important role in zinc homeostasis. Consistent with this hypothesis, the
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data using FRET sensors indicates that pcs2 is required for loz1Δ cells to lessen the
surplus cytosolic zinc upon zinc shock. The gene is also critical for loz1Δ mutants to grow
when they undergo the sudden supply of zinc. In addition, loss of pcs2 gene function in
WT cells caused the lower capacity of zinc buffering against low concentrations of zinc
supplement.
Why does loss of phytochelatins disrupt the growth of loz1Δ cells under zinc
shock conditions? Tiling array analysis indicates that pcs2 transcript expression is not
regulated by zinc [38]. Phytochelatin formation is, nevertheless, promoted by increasing
the activity of phytochelatin synthase in response to zinc supplements [156, 168]. Taking
into account that high zinc increases phytochelatin synthesis, it is noteworthy that loz1Δ
cells “hyper”accumulate zinc and “hypo”accumulate Zym1. In zinc‐replete medium, loz1
mutants accumulate about 4 times more zinc than WT cells [145]. The presence of a
very limited amount of metallothionein may therefore further enhance the cytosolic
zinc that is available for phytochelatin synthase in loz1 mutants.
Another line of evidence suggesting that phytochelatins have a protective role against zinc toxicity comes from the results generated using the loz1Δ zrt1Δ double mutant. To build zinc tolerance, loz1Δ cells require the zinc uptake transporter Zrt1. This
transporter is also essential for cells to increase cytosolic labile zinc levels. Notably in loz1Δ cells, which contain high zinc and therefore increased PC synthesis, zinc was readily lost from the ZapCY2 sensor. However, in loz1Δ cells lacking zrt1, zinc was not released from the ZapCY2 sensor. These results may indicate that the loz1Δ zrt1Δ cells
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do not contain much phytochelatins, underlining the importance of high zinc levels in phytochelatin synthesis.
In addition to the induction of phytochelatin formation, zinc detoxification in loz1Δ cells seems to occur from other factors as well. As pointed out in the current
study, high zinc influx—hence, expected high contents of phytochelatin—was
insufficient for WT cells to have zinc tolerance. This implies that there exist other
mechanisms under Loz1 regulation. In S. pombe, phytochelatin‐cadmium complexes are
sequestered in the vacuole, and Hmt1 is required to transport the complexes through
the vacuolar membrane [169]. Consequently, down‐regulation of hmt1 induces cadmium sensitivity while its up‐regulation causes the tolerance against the toxic metal
[82]. Although it is currently unknown if Hmt1 can directly transport the phytochelatin‐ zinc complex, its homolog in Caenorhabditis elegans, CeHMT1, was shown to play a role in the detoxification of copper, another essential metal, as well as cadmium [170]. It will, therefore, be worthwhile to further investigate if hmt1 expression is regulated by
Loz1 and influences zinc homeostasis in the fission yeast.
The decline of cytosolic labile zinc shown from ZapCY2 FRET traces in loz1Δ pcs2Δ cells suggest that there are other mechanisms controlling the surplus zinc in loz1Δ cells.
These mechanisms may include unknown transporters that mobilize the cytosolic zinc
into other compartments, and/or other zinc buffering molecules. In S. pombe, 3 CDF
family members—Zhf1, Cis4, and Zrg17—are identified to mobilize the cytosolic zinc
into intracellular organelles (see Chapter 1). The findings from FRET sensors
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demonstrated that the levels of cytosolic zinc can be affected by these transporters (see
Chapter 2). These transporters should be prioritized in further studies that examine
where loz1Δ cells accumulate zinc. Moreover, unknown transporters may also play a
role in limiting the cytosolic zinc. Although zinc export systems have not yet been reported in S. pombe, some bacteria are known to be capable of removing cytosolic zinc out of the cells using the P‐type ATPase transporter family [158, 171].
Other mechanisms that regulate the cytosolic surplus zinc in loz1Δ cells include zinc buffering molecules. In S. pombe, no zinc buffering molecule has been reported
other than Zym1 and phytochelatin. However, in all biological organisms, there are some zinc buffering molecules currently proposed, which include metallothioneins, glutathione, bacillithiol and phytochelatin [172‐174]. These peptide/proteins commonly contain cysteine residues that frequently play a role in zinc ion coordination. To increase the zinc‐buffering capacity under zinc‐replete conditions, cells may need more cysteines
synthesized or imported. According to the study using S. cerevisiae, the genes involved
in sulfate assimilation pathway leading cysteine biosynthesis, are down‐regulated by
Zap1 in zinc‐limited cells [175]. Therefore, the cysteine‐derivatives or the single amino
acid might be worthwhile to be closely examined in further studies that seek to
investigate unknown zinc buffering molecules in S. pombe.
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CHAPTER 4
The double zinc finger and adjacent accessory domain are critical in controlling Loz1 regulons
4.1. Introduction
Zinc is an essential nutrient for cells to grow, but is toxic in excess. All kingdoms
of life, thus, contain zinc‐responsive transcription factors that help cells to maintain an
intracellular zinc balance despite fluctuating zinc levels [176]. In S. pombe, Loz1 is a
critical transcription factor for zinc homeostasis, regulating genes associated with
maintaining zinc homeostasis in response to zinc availability.
Although a number of studies have reported the zinc‐regulatory mechanisms of
Zap1 and MTF‐1 (see Chapter 1), the mechanisms underlying how Loz1 controls gene expressions in response to zinc need to be investigated. Based on the report that C‐ terminal regions of Loz1, composed of double zinc finger and adjacent accessory
domains, are essential for Loz1 function, the present study focuses on examining the
role of the adjacent accessory domain in Loz1. By using site‐directed mutagenesis and
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by generating chimeric protein, here I show that the adjacent accessory domain is essential in the zinc‐regulatory functions of the Loz1.
4.2. Materials and methods
4.2.1 Site‐directed mutagenesis
The method for site‐directed mutagenesis was modified from the manufacturers’ protocol of Quikchange site‐directed mutagenesis (Stratagene, La Jolla,
CA). For the primers, both forward and reverse primers in length of 30 bp both which include mutations in amino acid sequence of interests were designed (Table C in
Appendix). 60 ng template DNA and Pfu Turbo polymerase (Stratagene) were used to
perform the site‐directed mutagenesis PCR according to the manufacturer’s
recommendations. PCR products were digested with DpnI (NEB) to remove the original
temples, and inserted into template plasmid backbones. All plasmids were sequenced to
confirm the mutation.
4.2.2 RNA isolation and Northern blot
RNA isolation and northern blot analysis were performed as described in Chapter
2. Zrt1 and adh4 probes were generated from DNA templates and were amplified with zrt1 and adh4 gene‐specific primers (Table C in the Appendix).
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4.3. Results
4.3.1. The Loz1 double zinc finger domain and adjacent accessary domain are necessary
for zinc‐dependent gene expression
To determine how Loz1 is regulated by zinc, a number of truncations were generated to map a minimal zinc‐responsive domain [96]. Analyses of these truncations revealed that the 96 amino acid sequences in C‐terminus were sufficient to confer zinc‐
dependent regulations of adh4, and partial regulation of zrt1 zinc‐replete conditions
[96]. The C‐terminal regions include a C2H2‐type double zinc finger and a 40 amino acid
region that is adjacent to zinc finger 1. Consistent with it being important for Loz1
function, these regions were sufficient for site specific DNA binding function in vitro, and
mutations within the double zinc finger domain inhibited Loz1 activity in vivo. [145].
Although the additional 40 amino acid residues, which are not included in the double
zinc finger domain, are critical in the minimal zinc‐responsive regulatory region in C‐ terminus, their roles have not been determined in zinc‐responsiveness or gene
repression of Loz1. In a number of other transcription factors, domains that are adjacent to double zinc fingers have been shown to be critical for DNA binding [165‐167].
Therefore, the goal of this chapter is to determine if the adjacent accessory domain was
necessary for zinc‐dependent regulation in Loz1.
MtfA (master transcription factor A) is a transcription factor containing the C2H2 double zinc finger domain and regulates secondary metabolism and morphogenesis in
Aspergillus nidulans. The double zinc finger domains from Loz1 in S. pombe and MtfA in 127
A. nidulans share conserved sequences within the double zinc finger (Fig. 4.1A).
However, significant sequence similarities other than the zinc finger domain were not found. To determine whether the adjacent domain of zinc finger 1 is important for Loz1 functions, plasmids expressing full mtfA ORF from A. nidulans or the chimeras of MtfA and Loz1 were created. All the plasmids were expressed from the zhf1 promoter region
as zhf1 expression is not effected by zinc [38, 145] (Fig. 4.1B).
To determine if MtfA could compensate the Loz1 functions in S. pombe, mtfA was expressed from the zhf1 promoter in loz1Δ cells, and the total RNA was purified for northern blot analysis (Fig. 4.1C). When the blot was probed for zrt1, there was no
regulation of zrt1 expression in response to zinc. Immunoblot analysis detecting GFP tag
in the C‐terminus of the plasmid confirmed that there is no significant change in the
expression level of the protein depending on zinc levels [96]. These results suggest that
the MtfA factor does not bind to the zrt1 gene regulatory region, and/or its activity does not respond to zinc. In contrast, when the chimera that was composed of the N‐terminal region of MtfA and the C‐terminal 96 amino acid sequences of Loz1 was expressed in loz1Δ cells, zrt1 expression was down‐regulated under zinc‐replete conditions (plasmid
#3 in Fig. 4.1B and C). These results are consistent with the C‐terminal region of Loz1
conferring zinc‐dependent repression of zrt1.
The role of the 40 amino acid region that is adjacent to the zinc finger 1 in Loz1
was further investigated using plasmid #4. As shown in Fig. 4.1B, the plasmid contains
the mtfA ORF whose adjacent domain next to zinc finger 1 was replaced by the 40
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amino acid‐adjacent domain of Loz1. Northern blot data in Fig. 4.1C revealed that zrt1 is
robustly regulated by zinc, i.e., the gene expression increased in zinc‐limited medium and decreased in zinc‐replete medium. Similar to other plasmids, the expression levels
of the chimera protein were also not changed by zinc [96]. These results suggest that
the region which is adjacent to the Loz1 double zinc finger is a critical “accessory domain” which is necessary for zinc‐dependent gene repression.
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Figure 4.1. The adjacent accessory domains are important for zinc‐dependent repression of Loz1 A. Alignment of amino acid sequences from the double zinc finger domains of Loz1 and MtfA. Conserved (black), non‐conserved (grey) and predicted key zinc binding amino acid sequences (red) are indicated. B. Schematic diagrams of the chimeras of Loz1 and MtfA. Loz1 is shown in brown while MtfA in light blue. The brown and light blue numbers represent amino acid number in Loz1 and MtfA, respectively. The double zinc fingers in Loz1 and MtfA are shown as rectangles in yellow and dark blue, respectively. The predicted zhf1 promoter and GFP tag were presented at both ends of the chimeras. C. Northern blot analysis of the loz1Δ cells containing respective chimeras. Loz1 cells expressing the chimera constructs or empty vector were grown in ZL‐EMM with or without a 200 µM zinc supplement. Total RNA was extracted and examined for northern blot assay.
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Fig. 4.1
A
B
C
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4.3.2. The adjacent accessory domain in Loz1 includes histidine residues
The rescue with the chimeras for loz1Δ cells produced a question about how the accessory domain plays an important role in zinc‐dependent repression in Loz1. In S.
cerevisiae, Zap1 is a zinc‐sensing transcription factor that contains 7 zinc fingers and
activation domain 1 (AD1) (see Fig. 1.2). Zinc fingers are a type of structural domain that directly binds zinc. In Zap1, they form the DNA binding domain and play a role in zinc sensing in the zinc‐regulatory activation domain2. AD1, in contrast, does not contain any zinc fingers. Despite the absence of zinc fingers, AD1 is known to drive full expressions of certain Zap1 target genes in zinc‐dependent manners [175]. The mechanisms of zinc‐ regulation have not been well understood, but cysteine and histidine residues in the domain potentially serve as zinc ligands.
As the 40 bp accessory domain contains a number of histidine residues (Fig.
4.2A), one possible model for the regulation of Loz1 function by zinc is that zinc binds to
this domain under zinc‐replete conditions, resulting in a conformation in which Loz1 is
able to bind to DNA and repress gene expression. To determine if the zinc finger accessory domain in Loz1 was required for zinc‐dependent repression, the zhf1
promoter‐driven Loz1 plasmid (pZ‐loz1‐GFP) was modified. Specifically, each of the 4
histidines was replaced with alanine (H432A, H439A, H444A, H454A) using site‐directed
mutagenesis. These plasmids were sequenced to confirm the correct mutations and
then were transformed into loz1Δ cells. The loz1Δ cells containing empty vector or expressing the loz1 ORF were also grown for negative or positive control, respectively. In
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Fig. 4.2B, RNA blots of the cells are shown. Consistent with previous findings, both adh4 and zrt1 transcripts were not regulated by zinc in loz1Δ cells expressing the empty
vector, while they were strongly repressed in the presence of the intact loz1 ORF plasmid. Both adh4 and zrt1 gene expressions were also inhibited in the cells containing
each of the 4 histidine‐mutated Loz1 proteins. The data suggests that the mutations did not eliminate Loz1‐mediated gene repression.
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A
B
Figure 4.2. The adjacent accessary domain in Loz1 includes histidine residues. A. Schematic diagram of pZ‐loz1GFP and the amino acid sequences in the adjacent accessary domain in Loz1. pZ‐lozGFP is identical with the plasmid #1 shown in Fig. 4.1B. Red letters indicate the histidine residues in the accessary domain. The amino acid numbers are also presented. B. Northern blot analysis of the loz1Δ cells expressing the Loz1 constructs including each histidine mutation. Amino acid substitutions from 4 each histidine to alanine were indicated. The loz1Δ cells, which were transformed with plasmids for wild‐type Loz1, H432A, H439A, H444A, or H454A, or empty vector, were grown in ZL‐EMM with or without a 200 µM zinc supplement. For northern blot analysis, specific probes against adh4, zrt1 or pgk1 were used.
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4.4. Discussion
Loz1 is a zinc‐responsive transcriptional repressor that contains a C2H2‐type double zinc finger domain [145]. Although in vitro and in vivo experiments show that the double zinc finger in Loz1 function is necessary for DNA binding [96, 145], other structural domains that regulate Loz1 function need to be further understood. The current study was part of this research that has sought to illustrate how Loz1 is regulated by zinc. By using a chimera containing the adjacent region of zinc finger in
Loz1, my studies demonstrate that the adjacent accessory domain is critical to confer the gene regulatory functions to the double zinc finger in Loz1.
Another goal of this study was to determine how the accessory domain affects zinc‐dependent regulation of Loz1. Based on the knowledge of zinc‐sensing domains of
Zap1 in S. cerevisiae, this study investigated the role of histidine residues in the
accessory domain of Loz1. Since the accessory domain is necessary for high affinity DNA
binding of Loz1 [96], it was hypothesized that the histidine residues coordinate zinc ions,
which might help to form the optimal conformation of the protein to bind with DNA.
The result from the site‐directed mutagenesis in each of the histidine residues, however, revealed that the mutations in single histidine do not eliminate the target gene‐ repression by Loz1. However, these results do not eliminate models in which zinc coordination occurs cooperatively by several histidines, or that there may be other unknown amino acids that render the accessory domain to be important.
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Interestingly, Loz1 proteins with H439A or H444A mutation weakly derepressed
adh4 gene expression in zinc‐limited cells. These overcompensations were not found in
zrt1 transcript in the cells with plasmids. One of reasons might be that the mutations
lead other factors to induce adh4 repression. The adh4 promoter includes multiple
binding sites for other transcription factors [145]. According to data from
electrophoretic mobility shift assay analysis, a recombinant protein containing amino
acids 455‐552 of Loz1 forms a tight complex with Loz1‐reponse elements (LRE;
GNNGATC). In contrast, only a weak complex was formed when the LRE element was incubated with a recombinant Loz1 protein without the accessory domain (Loz1 427‐552)
[96]. These observations imply that amino acids 427‐455 are required for Loz1 to bind tightly to the LRE. Histidine 439 and 444 are both within this potentially important
region. To determine this hypothesis, further understandings of the mechanisms of Loz1
in adh4 gene repression are needed.
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CHAPTER 5
Understanding the zinc‐regulated mammalian cell proliferation
5.1. Introduction
Due to its essential but potentially toxic nature, intracellular zinc levels are tightly controlled in all biological creatures by zinc homeostasis mechanisms. Although it has been several decades since it was first discovered that zinc deficiency in humans results in severe health problem [98], the only known eukaryotic protein that senses cellular zinc deficiency is Zap1 from S. cerevisiae [33]. Although homologs of Zap1 can be found in other fungal species, there is no homolog of Zap1 in humans.
Based on the current knowledge of the molecular structure of Zap1, I hypothesized that intracellular zinc levels may modulate the activity of human proteins that contain related domains to those found in Zap1. By examining sequence homology
of human proteins, HiNF‐P protein was identified. This protein contains a domain that shares significant similarity to a key regulatory zinc finger pair in Zap1.
HiNF‐P is a transcriptional activator of histone H4 gene expression [177]. It directly binds to a specific recognition motif that is located within the promoter region 137
of 14 distinct genes that encode histone H4. Over‐expression of HiNF‐P gene increases
H4 promoter activity in a manner that is dependent on the conserved HiNF‐P
recognition motif in H4 gene promoter [178]. HiNF‐P, thus, plays critical roles in the cell
cycle‐dependent H4 expression.
This chapter tests the hypothesis that the activity of HiNF‐P is controlled by
cellular zinc availability. Specifically, the effects of zinc on the expression of HiNF‐P
target genes are investigated using 2 different methods that lead to zinc deficiency. The
zinc‐regulated activity of HiNF‐P is also examined.
5.2. Materials and methods
5.2.1. Cell culture
COS‐7, HEK293T and HepG2 cell lines were cultured in regular 10% FBS‐
supplemented DMEM (Gibco Life Technologies, Grand Island, NY) in incubator with 10%
CO2 at 37°C; this medium is expected to contain about 3.5 µM of zinc [101]. In order to
create zinc‐deficient or ‐excessive condition, the cells were treated with 5 µM of TPEN
(SigmaAldrich), a membrane‐permeable zinc chelator, or 100 µM of ZnSO4, respectively,
unless concentrations are indicated.
To make zinc‐deficient cell culture medium, 5 g of Chelex‐100 ion exchange resin
(BioRad) was added to 100 mL FBS, and stirred overnight at 4 . The FBS supernatant
138 was filter‐sterilized into polyethylene centrifuge tubes after the removal of resin. 10% of the chelexed FBS was then added to DMEM.
5.2.2. MTT assay
Cell proliferation was examined by colormetric assay measuring conversion of tetrazolium dye to formazan producing purple color. Cells were seeded in 96‐well plate and treated with 2, 5, 10, and 20 µM of TPEN for several time points. After incubation with the tetrazolium dye (0.5 mg/ml) for 4 h, MTT conversion, which correlates with the number of living cells, was determined in 96‐well plate reader.
5.2.3. Reverse Transcription PCR (RT‐PCR) and quantitative real‐time PCR (qRT‐PCR).
Histone H4, MT‐1 and GAPDH mRNA levels were determined by RT‐PCR
(BioRad)or qRT‐PCR (Applied Biosystems, Foster City, CA). RNA was extracted from each group of cells using TriZol and reverse transcribed to cDNA by M‐MLV reverse transcriptase (Invitrogen). The synthesized cDNAs were amplified by Taq DNA polymerase (New England Bio Labs) in RT‐PCR. For the qRT‐PCR, the cDNAs were amplified by SYBR green master mix with specific primers (Table C in Appendix) for each gene and the accurate result was confirmed via dissociation curve analysis. Expressions of H4 were presented with fold of Control which is normalized to GAPDH CT value.
5.2.4. Western blot
Western blot was used to measure H4 protein expression in the cells. Whole cell protein was extracted with RIPA lysis buffer (20 mM Trizma base, 1% Triton‐X100, 50
139 mM NaCl, 250 mM sucrose, 50 mM NaF, 5 mM Na4P2O7 ∙ 10H2O and protease inhibitor cocktail tablets) and then subjected to SDS‐polyacrylamide gel electrophoresis after protein concentration is determined. Separated proteins were transferred to PVDF membranes and incubated with specific antibodies for the H4 and actin in Odyssey
Blocking buffer. The protein expressions were visualized using the corresponding fluorescent secondary antibody under Odyssey imaging system.
5.2.5. siRNA Transfection
To test the significant role of HiNF‐P gene in zinc‐regulated H4 gene expression,
HiNF‐P expression was knocked down by siRNA transfection. Cells were seeded in 24‐ well plate with 0.5 x 105 / well 1 d before transfection. HiNF‐P specific or control scrambled siRNA (Life Technologies) was introduced using an X‐treme 9 transfection reagent (Roche, Indianapolis, IN) for 48 h. The efficiency of the respective gene suppression was determined by qRT‐PCR method.
5.2.6. Plasmid construction
The CMV promoter‐driven plasmid containing the full length of HiNF‐P fused with FLAG (DYKDDDDK) tag has been obtained from Dr. Masayuki Sekimata at
Fukushima Medical University School of Medicine. A firefly luciferase reporter vector inserted with 5 copies of core HiNF‐P binding sequences (pGL3‐Hpolβ) was generated using pGL3‐polβ construct which includes a human DNA polymerase β promoter. As a control reporter vector, a renilla luciferase vector (pRL‐TK) was obtained.
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5.2.7. Dual luciferase reporter assay
The transcriptional activities of HiNF‐P in different levels of zinc were measured
using dual luciferase reporter assay (Promega, Madison, WI). Cells were co‐transfected
with the HiNF‐P expression vectors, the pGL3‐Hpolβ reporter vector, and the pRL‐TK
control reporter vector using the X‐treme 9 transfection reagent. After 24 h incubation,
the cells were exposed to 1, 2, or 5 µM of TPEN or ZnSO4 and the luciferase activity was
determined by luminometer. The data is presented by ratio of firefly luciferase versus
renilla luciferase activity.
5.3. Results
5.3.1. HiNF‐P contains a ZF pair with similar amino acid residues ZF pair from Zap1
HiNF‐P is the only mammalian protein that contains 2 ZF pairs [179]. A ZF pair is
a structural motif that contains 2 adjacent zinc fingers. Proteins that contain zinc finger
pairs include Gli1 and Zap1 [138, 179‐181]. The unique ZF pair structure depends on the
existence of tryptophan residues between the 2 conserved cysteines of the tandem C2H2 type zinc fingers (CWCH2). In a recent database analysis, 587 genes containing the
CWCH2 were categorized into 11 classes of genes based on sequence similarity, domain
structure, and functional similarity [179]. Of the 11 gene classes, only Zap1 and HiNF‐P
classes have 2 individual ZF pairs per protein. All of the remaining classes have 1 ZF pair
per protein.
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Consistent with the importance of the ZF1/2 and ZF3/4 in Zap1 function in S. cerevisiae, the database analysis revealed that ZF1/2 and ZF3/4 are conserved in the
Zap1 orthologs. Using the sequence conservation analysis for the ZF pair domains from
the 11 gene classes [179], it is possible to compare the sequence of the regulatory ZF1/2
pair from Zap1 with other ZF classes. I hypothesized that another regulatory ZF pair
would share higher sequence conservation with Zap1 ZF1/2.
When all of the zinc finger classes are compared to the Zap1 ZF1/2 class, the
HiNF‐P ZF1/2 class has a number of similarities to the Zap1 ZF1/2 pair at the amino acid
level. These similarities include substitution of a conserved phenylalanine in ZF1. In most classical C2H2‐type zinc fingers, a conserved phenylalanine plays a central role in the
formation of the hydrophobic core of the zinc finger domain. An unusual feature of the
Zap1 and HiNF‐P ZF1/2 pairs is that this phenylalanine is absent (Fig. 5.1C). A cysteine residue, instead, is found in this position.
ZF2 is critical to the zinc‐dependent regulation of ZF1/2 from Zap1 [139]. In ZF2,
2 unique conserved amino acids are exclusively found in both HiNF‐P and Zap1. The first is a phenylalanine (F10) that is located at the start of the second β‐sheet in the ZF2. The high conservation of F10 in both HiNF‐P and Zap1 gene orthologs, suggests that this
residue may play a critical role in the ZF2 functions. Another conserved residue is
aspartate (D14), which is located immediately before the beginning of the α helix. Given
that substitution of an α‐helix motif of ZF4 with that of ZF2 created zinc lability to ZF3/4
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in Zap1 [139], this residue might be important for the zinc regulation of ZF2. A number
of acidic amino acids are also conserved in both HiNF‐P ZF1/2 and Zap1 ZF1/2.
Previous studies demonstrated that the conformation of the ZF1/2 pair structure is dependent on intracellular zinc levels, and this conformation change is critical for the regulation of AD2 by zinc [37, 38]. Based on the similarities of conserved amino acids in orthologs of Zap1 and HiNF‐P, ZF1/2 domain was hypothesized to confer the metal‐ regulatory activation of HiNF‐P transcription factor.
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Figure 5.1. Schematic structures and amino acid comparisons of ZF1/2 pair in Zap1 and HiNF‐P A. Olive cylinders and purple circle represent zinc finger motifs and zinc ions, respectively. In the CWCH2 ZF pair, the interaction between 2 zinc fingers is mediated by zinc ions and hydrophobic amino acids including well‐conserved tryptophan residue. The tryptophan (W) is shown in red. B. All ZFs are numbered and are shown as cylinders. Zap1 contains 2 ZF fingers, ZF1/2 and ZF3/4, which are indicated as linked‐dark blue cylinders. The ZF1/2 and ZF3/4 in HiNF‐P are represented as dark red cylinders and the other ZFs are shown in light red. C. Amino acid sequences of ZF1/2 in Zap1 and HiNF‐P
are shown and compared with the conserved sequences of the classic C2H2 ZF. The red
amino acid residues indicate the conserved sequences in all CWCH2 type ZFs. The blue amino acids are residues that are substituted or conserved in the ZF1/2 pairs in both Zap1 and HiNF‐P orthologs.
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Fig. 5.1
A
B
C
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5.3.2. Cell viability upon TPEN‐induced zinc deficiency
The high level of sequence homology between ZF1/2 domains from HiNF‐P and
Zap1 lead to hypothesize that HiNF‐P may be regulated by zinc availability. To test this
hypothesis, a number of mammalian cell lines were grown under various zinc conditions
(see below) and HiNF‐P target genes were measured by RT‐PCR and qRT‐PCR. One of
known targets of HiNF‐P is histone H4 gene [177]. The histone gene expression,
however, could be altered by various cellular stresses through another mechanism (e.g.
apoptosis). The zinc chelator TPEN used in this study is known to induce strong zinc‐
deficiency in various cells, while it also causes the apoptotic cell death in the use of high
concentrations [97]. Therefore, to investigate whether zinc regulates the HiNF‐P activity
and H4 gene expression using TPEN, it was required to know the optimal levels of TPEN,
which could not induce cell death.
To establish conditions for examining gene expression in response to zinc
deficiency, cell viabilities upon TPEN treatment were measured in 3 different cell lines
(Fig. 2). The human embryonic kidney (HEK293T), monkey kidney (COS‐7), and human
liver cell lines (HepG2) were selected for HiNF‐P protein is highly expressed in kidney
and liver tissues [182‐184]. Compared to COS‐7 and HepG2 cell lines, HEK293T cell is
very sensitive to TPEN‐induced zinc deficiency. The viability of the cell decreased with 2
µM TPEN for 5 h incubation. 20 µM of TPEN treatment for 5 h almost completely
inhibited cell survival in the cell. Due to the significant level of cell death in HEK293T cells on exposure to TPEN, the HEK293T cell line was excluded from further studies. On
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the other hand, both COS‐7 and HepG2 cell lines were resistant to 2, 5, and 10 µM of
TPEN for 5 h. Longer exposure of TPEN (10 h) significantly reduced cell viability in each cell line. These results indicate that zinc deficiency reduces cell proliferation in a dependent manner of the zinc chelator, TPEN. An extended exposure to severe zinc
deficiency induces cell death, so further studies were performed using TPEN within the
ranges of time and concentration where the cells can survive.
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Figure 5.2. Cell viabilities in different zinc levels A. COS‐7, HEK293T and HepG2 cells were treated with 2, 5, 10, and 20 µM of TPEN for 5 h. The cell viability was measured using MTT assay and calculated as % of OD of the TPEN‐treated group against the control group in each cell line. B. A similar experiment was performed to that described in Panel A with the exception that the incubation length was 10 h. Results are mean ± SD. * indicates p < 0.05 compared with control.
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5.3.3. Histone H4 mRNA expression in TPEN‐induced zinc deficient cells
To investigate whether zinc affects HiNF‐P activity in vivo, both COS‐7 and HepG2
cell lines were grown over a range of zinc levels using TPEN (Fig. 5.3). In both cells, the
expression of H4 mRNA genes was down‐regulated by 5 µM of TPEN (Fig. 5.3A and B).
Metallothionein 1 (MT1) mRNA levels were also decreased by TPEN and increased by
ZnSO4, validating that intracellular zinc status was changed by TPEN or ZnSO4 treatments. In Fig. 5.3C, zinc supplementation into the TPEN‐added cells rescued H4 gene expression. Thus, the effect of TPEN on H4 expressions could result from lack of its ligand, potentially zinc, in the cells. These results are consistent with the hypothesis that transcriptional activity of HiNF‐P may be regulated by zinc availability.
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Figure 5.3. Histone H4 and MT1 mRNA expressions in different zinc levels A. Histone H4 gene expression was examined in COS‐7 cell in zinc‐deplete (TPEN) or ‐ replete condition (ZnSO4). The cells were collected after 5 h incubation of each condition and their RNAs were extracted using Trizol. cDNAs were synthesized using 100 ng of total RNAs and M‐MLV reverse transcriptase. H4, Metallothionein 1 (MT1), and GAPDH cDNAs were amplified with specific primers for each gene. MT1 expression showed zinc responsiveness of cells while GAPDH was used for loading control. B. Zinc‐regulated H4 gene expression was detected in HepG2 cell line. TPEN or ZnSO4 was treated for 8 h in the culture medium. C. Zinc supplementation rescues H4 expression in TPEN‐treated cells. COS‐7 cells were incubated in the absence or presence of 5 µM TPEN and indicated concentration of ZnSO4 for 5 h.
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Fig. 5.3
A B
C
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5.3.4. Regulation of H4 mRNA expression by HiNF‐P
Histone gene modification can occur at the transcriptional and post‐ transcriptional level [160]. To determine if the effects of zinc on H4 expression were mediated by HiNF‐P transcription factor, specific siRNAs against HiNF‐P gene were introduced into cells, and histone H4 gene expression examined using qRT‐PCR. Fig. 5.4A
revealed that the HiNF‐P siRNA successfully decreases over 60% of HiNF‐P gene
expression. The siRNA also significantly reduces H4 mRNA expression, consistent with previous studies [102, 185]. However, this decrease was not observed in cells not treated with TPEN (Fig. 5.4B). These results support published results that HiNF‐P is
critical for H4 gene expression.
The data in Fig. 5.4B also revealed the essential role of HiNF‐P in the zinc‐ regulated H4 gene expression. In the cells transfected with control siRNA, H4 mRNA
expression was reduced about 70% by TPEN, which confirms the impact of zinc
depletion in H4 expression using real‐time PCR. Comparing H4 levels in HiNF‐P siRNA‐
transfected cells that were in normal or low zinc medium, the level of gene expression
was similar when cells were treated with siRNAs or TPEN. The HiNF‐P siRNA‐transfected
cells maintain the similar levels of H4 expression in both zinc‐deficient and sufficient
conditions. The findings suggest that HiNF‐P may be required for the down‐regulation of
H4 expression in response to zinc deficiency. Based on these observations as well as the
finding of constant HiNF‐P levels upon TPEN, zinc depletion may modify transcriptional
activity of HiNF‐P.
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A
B
Figure 5.4. H4 and HiNF‐P genes expressions in HiNF‐P deficient cells HiNF‐P specific siRNA (HiNFP siR) or nonspecific scrambled siRNA (Con siR) were transfected into COS‐7 cell for 48h. Cells were treated with (TPEN) or without (Con) 5 µM of TPEN for 5 h. cDNA of each group was synthesized by above methods and was used as template for qRT‐PCR using SYBR green. CT values for HiNF‐P, H4, and GAPDH were obtained and the result was presented as normalized ratio of each gene versus the GAPDH CT value. A. HiNF‐P siRNA suppresses the gene expression of HiNF‐P mRNA and
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TPEN does not change its expression. B. H4 gene expression is down‐regulated by HiNF‐ P siRNA transfection or TPEN treatment. (*p<0.05 vs. Control siRNA Control group) 5.3.5. Critical role of the ZF pair in transcriptional activity and zinc responsiveness of
HiNF‐P
The role of ZF pairs in transcriptional activity in HiNF‐P has not been investigated
[96]. To examine the function of the ZF1/2 pair, I generated plasmids expressing wild‐ type HiNF‐P (pHiNF‐P) or HiNF‐P mutant (pHiNF‐P ZF1mut) which were disrupted in a critical zinc binding residue (Fig. 5.5). To determine that the plasmids were functional, they were co‐transfected with luciferase reporter plasmids that contained a minimal
promoter with multiple HiNF‐P binding sites (pGL3‐polβ H5). In these co‐transfected
cells, luciferase activity increased indicating that this reporter system was functional. In
the presence of pHiNF‐P ZF1mut, cells showed lower luciferase activity, which suggests
that the ZF1/2 pair may play a critical role in HiNF‐P transcriptional activity of its target
genes.
Furthermore, the cells transfected with the wild‐type HiNF‐P or HiNF‐P mutant
vector were exposed to TPEN 5 µM for 5 h. In agreement with the anticipated role of
HiNF‐P in zinc‐regulated H4 gene transcription, TPEN‐induced zinc deficiency decreases
the transcriptional activity of wild type HiNF‐P. The ZF1/2 mutant, however, fails to
respond to zinc depletion in that its transcriptional activity is not significantly changed
by TPEN. As a result, the ZF1/2 may be the domain that is responsible for transcription
activity and zinc‐responsiveness of HiNF‐P.
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A
B
Figure 5.5. Importance of ZF1/2 for transcriptional activity and zinc responsiveness of HiNF‐P A. Luciferase reporter and HiNF‐P expression vectors are shown. A cross on the ZF indicates introduced mutations that disrupt zinc binding in ZF1. B. Dual luciferase assays were performed after transfection of the reporter and expression vectors for 48 h to COS‐7 cell. Data was displayed with ratios of luminescence values of firefly to renilla. Results are mean ± SD. * indicates p < 0.05 compared with control.
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5.3.6. Cell proliferation in zinc‐deficient medium with chelated‐FBS
TPEN is a membrane permeable zinc chelator. As a consequence, TPEN
treatment causes a direct impact on both intra‐ and extracellular levels of zinc [186]. As
strong zinc depletion by TPEN also induces apoptosis via complex cell signaling in
hepatocytes [187], I used an alternative method of generating zinc deficiency. This method depleted zinc from the medium by using chelating resin‐treated FBS [102].
To compare the growth rates in the medium containing different zinc levels,
HepG2 cells were grown in the zinc‐deficient medium (ZD) in which the Chelex‐treated
FBS was added to DMEM, or in zinc‐normal medium (ZN) supplemented with 4 µM of
ZnCl2, which contained normal zinc level in FBS. In Fig. 5.6, the cells grown in ZD for 3 d
showed the 80% of cell viability compared to that in cells in ZN. This growth defect is
similar to what was seen on the cells that was treated with 5 µM of TPEN for 10 h. The
finding suggests that, without the side effects of TPEN, the cells grown in zinc‐limited medium are impaired in proliferation.
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Figure 5.6. Cell viability in zinc‐normal (ZN) or ‐deficient (ZD) medium
HepG2 cells were grown in ZN or ZD for 3 d, and MTT conversion was measured in both
groups. Percentage of cell viability in ZD was calculated relative to the viability of cells in
ZN. Results are mean ± SD. * indicates p < 0.05 compared with ZN.
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5.3.7. H4 mRNA level in cells grown in medium containing different zinc concentrations
To determine whether growth in ZD resulted in a decrease in H4 expression, RT‐
PCR was performed with RNA extracts from the cells grown in ZD or ZN. In Fig. 5.7, MT1 expression showed robust regulation depending on zinc levels. H4 mRNA level, however, were only slightly down‐regulated in ZD in both 1.5 and 3 d incubations. This
extent of regulation was not as prominent as in TPEN‐treated cells (see Fig. 5.3B).
ATM and CKS2 are newly identified HiNF‐P target genes, which play a role in the
DNA damage response [188]. The promoter regions of both genes contain DNA binding
elements for HiNF‐P, suggesting that HiNF‐P directly controls their expression. It was
thus expected that the expression of both genes would be altered by zinc levels if HiNF‐
P activity was regulated by zinc. When RT‐PCR was used to examine ATM and CKS2
levels, their expression was not significantly regulated in ZD (Fig. 5.7). These results
suggest that HiNF‐P may not be modulated by zinc, or at least, the ZD medium.
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Figure 5.7. mRNA levels of HiNF‐P target genes and MT1 in HepG2 cells
The cells were grown in ZD or ZN for 1.5 or 3 d. Each gene level was examined using RT‐
PCR with specific primers and showed on agarose gel.
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5.4. Discussion
A homolog of ZAP1 is not present in humans despite its essential roles for zinc homeostasis in S. cerevisiae. Due to sequence similarities between zinc finger domain structures between HiNF‐P and Zap1, this chapter investigated whether HiNF‐P activity may also be regulated by zinc in humans.
Reviewing the amino acid sequences of ZF1/2 pair domain from Zap1 and HiNF‐P orthologs, it was hypothesized that the unique zinc finger pair domain is a zinc‐
regulatory region in HiNF‐P. According to a previous report, the N‐terminal region encompassing ZF1/2 and ZF3/4 in HiNF‐P is necessary for DNA binding and transcription
activation [178]. Masayuki and Yoshimi [189] suggested a DNA binding role for each zinc
finger in the N‐terminus, showing that zinc is required for HiNF‐P to bind DNA in vitro.
Later studies revealed the presence of additional zinc fingers in HiNF‐P, including zinc
fingers belonging to this ZF1/2 pair [177, 178, 190]. The role of the ZF1/2 pair domain in the protein was not determined in vivo.
To investigate the effect of zinc deficiency on cells, this study introduced 2 different methods to induce zinc depletion in cells: TPEN treatment and usage of zinc‐ deficient medium containing chelated FBS. When TPEN was added to the regular cell
culture medium that contains 10% FBS as a significant source of zinc, histone H4 mRNA
level was clearly reduced. In the case of HepG2 cells, band intensity from PCR and real‐
time PCR data displayed about 4.5 times higher H4 level in control group than TPEN‐
treated group.
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This effect of TPEN on H4 mRNA expression may be directly or indirectly
mediated through HiNF‐P inactivation considering critical roles of HiNF‐P in H4 gene
transcription. The real‐time PCR data showing that HiNF‐P knockdown cells did not
respond to TPEN, indicates that HiNF‐P is required in TPEN‐regulated H4 gene
expression. HiNF‐P also showed less transcriptional activity upon TPEN in luciferase assay. To exclude the chance that TPEN interferes other transcription machinery, the reporter gene plasmid pGL3‐polβ H5 was introduced. It contains a luciferase gene driven by a human DNA polymerase promoter proximal to 5 copies of the HiNF‐P binding
elements. As the reporter plasmid without the HiNF‐P binding site (pGL3‐polβ) did not
respond to TPEN, it is concluded that TPEN‐induced HiNF‐P inactivation is the only factor that changed luciferase activity in the experiment. TPEN treatment, hence, inactivates
HiNF‐P resulting in low mRNA expression of H4 gene.
On the other hand, cells grown in ZD containing chelated‐FBS showed less
decrease in H4 mRNA expression, compared to TPEN‐treated cells. Culturing cells in ZD
is regarded as a method to render mild zinc deficiency because there was no artificial
competitive agent for zinc in the cells. One of the reasons of less effects of ZD on H4 expression might also be that heterogeneous cell population experiences different level of zinc starvation stress. HiNF‐P activity and H4 mRNA level are tightly regulated according to cell cycle stage [178]. The basal and/or regulated expression level of H4 gene in different phases in the cell cycle may partly nullify the effect of ZD. In these reasons, further studies may need to use synchronized cells to examine H4 expressions.
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A common method to synchronize cell cycle is serum starvation [191]. For serum is crucial source of zinc in the usual cell culture medium, the method for synchronization should be attentively chosen.
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CHAPTER 6
Concluding remarks
Dietary zinc deficiency is a global health issue. Among the deficiencies of vitamins and minerals in children under 5 years of age, zinc deficiency was found to be the second‐largest factor responsible for death [192]. Other health problems known to be associated with zinc deficiency, including stunting, severe wasting, and low‐birth weight, have been estimated to constitute 2.2 million deaths worldwide [192].
Consistent with zinc deficiency having deleterious effects on growth and development, many studies have suggested a correlation between increased dietary zinc levels and favorable outcomes, especially in fetal development, birth weight, growth rate, and childhood disease treatment [193, 194].
Although zinc is vital for life, it can be toxic or inhibitory to growth when in excess. While the precise mechanism of zinc toxicity is not yet known, it may in part result from competition of zinc with other metal‐binding sites in enzymes and transport proteins [195‐197]. Due to the essential but potentially toxic nature of this nutrient, intracellular zinc levels are tightly controlled in all biological creatures.
163
Key factors in maintaining zinc homeostasis in eukaryotic cells include zinc
transporters, zinc buffering molecules, and zinc‐regulatory transcription factors. The
levels and activities of these factors are precisely regulated to preserve optimal intracellular zinc levels. To extend the knowledge of zinc homeostasis in eukaryotic cells, this dissertation investigated the role of specific zinc transporters, potential zinc buffering peptides, and zinc‐regulatory factors using Schizosaccharomyces pombe as a
model organism.
Most of our current knowledge about zinc‐responsive transcription factors in
eukaryotic cells has been obtained from studies with MTF‐1 or Zap1. S. pombe creates a
new system to explore zinc homeostasis as it does not express a homolog of MTF‐1 or
Zap1. Instead the fission yeast uses a different factor, Loz1, which transcriptionally
controls the expression of a zinc buffering metallothionein and zinc uptake transporter.
Using the many advantages of a yeast model system, this dissertation examined the role
of zinc homeostasis‐related genes, including zinc transporters and Loz1.
In chapters 2 and 3, genetically encoded zinc‐responsive FRET‐based sensors were utilized to measure the changes in the labile pool of cytosolic zinc. The results obtained in these chapters demonstrate that Zhf1, a zinc transporter in endoplasmic
reticulum membrane, and both Zrg17 and Cis4, which reside in Golgi membrane, play a
role in maintaining cytosolic labile zinc. I also show that although mutants lacking the
loz1 gene have increased expression of zrt1, which in turn leads to the
hyperaccumulation of zinc when zinc is in excess, loss of Loz1 function also leads to cells
164
having an increased tolerance to zinc. In addition, the findings of the unexpected role of
Loz1 in zinc homeostasis reveal that phytochelatins, small peptides that have a well‐
known role in the detoxification of toxic heavy metals, have an additional zinc buffering
role.
Chapter 4 investigated the molecular mechanisms by which Loz1 is zinc‐
regulated. Using chimeric proteins containing Loz1 and a related transcription factor
MtfA, I found that the accessory domain adjacent to a double zinc finger is necessary for zinc‐responsive gene regulation of the transcription factor. In chapter 5, the study examined whether HiNF‐P is controlled by zinc in vivo through its zinc finger pair domain. The study sought to find a novel zinc‐responsive transcription factor in mammalian cells, extending the current knowledge of the molecular structure of Zap1.
Overall, the studies in this dissertation shed light on the role of genes encoding
zinc transporters, zinc buffering molecules, and zinc‐responsive transcription factors in
eukaryotic cells. The knowledge of these key factors in zinc homeostasis at the cellular
level lay a cornerstone for understanding zinc homeostasis mechanisms in higher
eukaryotes. These studies using microorganisms will help to bridge the gaps of
knowledge in zinc homeostasis and to translate new knowledge from the basic sciences
into ways of addressing zinc‐associated public health issues in the future.
165
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APPENDIX. A. Additional Tables
Table A.1. S. pombe strains used in this study
Strain Genotype Reference
JW81 h‐ ade6‐M210 leu1‐32 ura4‐D18 1
SPAC22H10.13Δ h+ ade6‐M? leu1‐32 ura4‐D18 zym1Δ::kanMX6 Bioneer
SPAC25B8.19cΔ h+ ade6‐M210 leu1‐32 ura4‐D18 loz1Δ::kanMX4 Bioneer
SPAC25B8.19cΔ h+ ade6‐M? leu1‐32 ura4‐D18 zhf1Δ::kanMX6 Bioneer
SPAC3H1.10Δ h+ ade6‐M? leu1‐32 ura4‐D18 pcs2Δ::kanMX6 Bioneer
h‐ ade6‐M210 leu1‐32 ura4‐D18 loz1Δ::KanMX6 pZHF1‐loz1::leu1+ 4
h‐ ade6‐M210 leu1‐32 ura4‐D18 loz1Δ::KanMX6 pZHF1‐AL::leu1+ 4
ABY46 h‐ ade6‐M210 leu1‐32 ura4‐D18 pJK148::leu1+ 2
ABY87 h‐ ade6‐M210 leu1‐32 ura4‐D18 zrt1Δ::kanMX6 3
ABY385 h+ ade6‐M210 leu1‐32 ura4‐D18 loz1Δ::kanMX4 pJK148::leu1+ 3
This study; ABY577 h‐ ade6‐M210 leu1‐32 ura4‐D18 JK pgk1‐zrt1::leu1+ generated by SYC This study; ABY578 h‐ ade6‐M210 leu1‐32 ura4‐D18 zrt1Δ::kanMX6 JK pgk1‐zrt1::leu1+ generated by AJB ABY651 h‐ ade6‐M210 leu1‐32 ura4‐D18 loz1Δ::KanMX6 pZHF1‐mtfA::leu1+ 4
ABY653 h‐ ade6‐M210 leu1‐32 ura4‐D18 loz1Δ::kanMX6 ploz1::leu1+ 3
ABY751 h‐ ade6‐M210 leu1‐32 ura4‐D18 loz1Δ::KanMX6 pZHF1‐ALA::leu1+ 4 This study; h+ ade6‐M210 leu1‐32 ura4‐D18 loz1Δ::kanMX4 pZHF1‐loz1(H432A)‐ ABY771 generated by GFP::leu1+ SYC Continued
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Table A.1 continued
This study; h+ ade6‐M210 leu1‐32 ura4‐D18 loz1Δ::kanMX4 pZHF1‐loz1(H439A)‐ ABY772 generated by GFP::leu1+ SYC This study; h+ ade6‐M210 leu1‐32 ura4‐D18 loz1Δ::kanMX4 pZHF1‐loz1(H444A)‐ ABY773 generated by GFP::leu1+ SYC This study; h+ ade6‐M210 leu1‐32 ura4‐D18 loz1Δ::kanMX4 pZHF1‐loz1(H454A)‐ ABY774 generated by GFP::leu1+ SYC This study; ABY795 h‐ ade6‐M210 leu1‐32 ura4‐D18 pZapCY1::leu1+ generated by SYC This study; ABY797 h‐ ade6‐M210 leu1‐32 ura4‐D18 pZapCY2::leu1+ generated by SYC This study; ABY814 h+ ade6‐M210 leu1‐32 ura4‐D18 loz1Δ::kanMX6 pZapCY2::leu1+ generated by SYC This study; ABY824 h+ ade6‐M? leu1‐32 ura4‐D18 zhf1Δ::kanMX6 pZapCY1::leu1+ generated by SYC This study; ABY825 h+ ade6‐M? leu1‐32 ura4‐D18 zhf1Δ::kanMX6 pZapCY2::leu1+ generated by SYC This study; ABY829 h‐ ade6‐M210 leu1‐32 ura4‐D18 zrt1Δ::kanMX6 pZapCY2::leu1+ generated by SYC This study; ABY851 h? ade6‐M? leu1‐32 ura4‐D18 loz1Δ::KanMX6 zrt1Δ::KanMX6 pZapCY1::leu1+ generated by SYC This study; ABY853 h? ade6‐M? leu1‐32 ura4‐D18 loz1Δ::KanMX6 zrt1Δ::KanMX6 pZapCY2::leu1+ generated by SYC This study; ABY859 h+ ade6‐M? leu1‐32 ura4‐D18 zrg17Δ::kanMX6 pZapCY1::leu1+ generated by SYC This study; ABY860 h+ ade6‐M? leu1‐32 ura4‐D18 zrg17Δ::kanMX6 pZapCY2::leu1+ generated by SYC This study; h‐ ade6‐M216 leu1‐32 ura4‐D18 loz1Δ::KanMX6 zym1Δ::KanMX6 ABY869 generated by pZapCY2::leu1+ SYC This study; h‐ ade6‐M216 leu1‐32 ura4‐D18 loz1Δ::KanMX6 zym1Δ::KanMX6 ABY870 generated by pZapCY1::leu1+ SYC This study; ABY929 h‐ ade6‐M210 leu1‐32 ura4‐D18 JK pZapCY2::leu1+ pTN‐ZRT1‐lacZ::ade6+ generated by AJB Continued
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Table A.1 continued
This study; ABY930 h‐ ade6‐M210 leu1‐32 ura4‐D18 loz1Δ::kanMX6 pTN::Ade+ pZapCY2::leu1+ generated by SYC This study; h‐ ade6‐M210 leu1‐32 ura4‐D18 loz1Δ::kanMX6 pTN pgk1‐zym1::Ade+ ABY931 generated by pZapCY2::leu1+ SYC This study; h‐ ade6‐M210 leu1‐32 ura4‐D18 loz1Δ::kanMX6 pTN pgk1‐MT1::Ade+ ABY932 generated by pZapCY2::leu1+ SYC This study; ABY942 h‐ ade6‐M210 leu1‐32 ura4‐D18 JK pZapCY1::leu1+ pTN‐ZRT1‐lacZ::ade6+ generated by AJB This study; ABY943 h? ade6‐M210 leu1‐32 ura4‐D18 loz1Δ::kanMX6 pZapCY1::leu1+ generated by SYC This study; ABY949 h? ade6‐M? leu1‐32 ura4‐D18 SPAP8A3.03Δ::kanMX6 pZapCY1::leu1+ generated by SYC This study; ABY950 h? ade6‐M? leu1‐32 ura4‐D18 cis4Δ::kanMX6 pZapCY1::leu1+ generated by SYC This study; ABY951 h? ade6‐M? leu1‐32 ura4‐D18 SPAP8A3.03Δ::kanMX6 pZapCY2::leu1+ generated by SYC This study; ABY952 h? ade6‐M? leu1‐32 ura4‐D18 cis4Δ::kanMX6 pZapCY2::leu1+ generated by SYC This study; ABY960 h‐ ade6‐M210 leu1‐32 ura4‐D18 JK148::leu1+ pTN‐lacZ::ade6+ generated by AJB This study; ABY961 h‐ ade6‐M210 leu1‐32 ura4‐D18 JK148::leu1+ pTN‐zrt1‐lacZ::ade6+ generated by AJB This study; ABY969 h? ade6‐M? leu1‐32 ura4‐D18 loz1Δ::kanMX6 pcs2Δ::kanMX6 generated by SYC This study; ABY970 h? ade6‐M? leu1‐32 ura4‐D18 loz1Δ::KanMX6 pcs2Δ::KanMX6 pZapCY2::leu1+ generated by SYC This study; ABY971 h? ade6‐M? leu1‐32 ura4‐D18 pcs2Δ::KanMX6 pZapCY2::leu1+ generated by SYC This study; ABY972 h? ade6‐M? leu1‐32 ura4‐D18 pcs2Δ::KanMX6 pZapCY1::leu1+ generated by SYC This study; ABY973 h? ade6‐M? leu1‐32 ura4‐D18 loz1Δ::KanMX6 pcs2Δ::KanMX6 pZapCY1::leu1+ generated by SYC Continued
183
Table A.1 continued This study; ABY974 h? ade6‐M210 leu1‐32 ura4‐D18 zrt1Δ::kanMX6 pZapCY1::leu1+ generated by SYC
1. Wu JQ, Kuhn JR, Kovar DR, Pollard TD (2003) Spatial and temporal pathway for assembly and constriction of the contractile ring in fission yeast cytokinesis. Dev Cell 5(5):723–734. 2. Ehrensberger KM, et al. (2013) Zinc‐dependent regulation of the Adh1 antisense transcript in fission yeast. J Biol Chem 288(2):759–769. 3. Corkins ME, et al. (2013) Zinc finger protein Loz1 is required for zinc‐responsive regulation of gene expression in fission yeast. PNAS 110(38):15371‐15376 4. Ehrensberger KM, et al. (2014) The double zinc finger domain and adjacent accessory domain from the transcription factor Loss of zinc sensing 1 (Loz1) are necessary for DNA binding zinc sensing. J Biol Chem 289(26):18087‐18096
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Table A.2. Plasmids used in this study
AJB Database Function Plasmid Name Reference No.
Published plasmid pJK148 12 1
backbones pTN215 280 2
pZapCY1 454 This study FRET sensors pZapCY2 455 This study
pTN‐lacZ 465 2 LacZ constructs pTN‐zrt1‐lacZ 470 2
pLoz1 247 3
Integrating expression JK pgk1‐zrt1 274 This study
plasmids TN pgk1‐zym1 473 This study
TN pgk1‐MT1 472 This study
pZHF1‐loz1‐GFP 287 2
pZHF1‐AN8741‐GFP 308 2
pZHF1‐AN8741/loz1 C‐terminus‐ Loz1 chimera plasmids 416 2 GFP
pZHF1‐AN8741/loz1 accessory 414 2 region‐GFP
pZHF1‐loz1 H444A 422 2
pZHF1‐loz1 H432A 433 2 Loz1 mutagenesis plasmids pZHF1‐loz1 H439A 434 2
pZHF1‐loz1 H454A 432 2
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1. Keeney, J.B. and Boeke, J.D (1994) Efficient targeted integration at leu1‐32 and ura4‐294 in Schizosaccharomyces pombe. Genetics 136, 849‐856. 2. Ehrensberger KM, et al. (2014) The double zinc finger domain and adjacent accessory domain from the transcription factor Loss of zinc sensing 1 (Loz1) are necessary for DNA binding zinc sensing. J Biol Chem 289(26):18087‐18096 3. Corkins ME, et al. (2013) Zinc finger protein Loz1 is required for zinc‐responsive regulation of gene expression in fission yeast. PNAS 110(38):15371‐15376
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Table A.3. Primers used in this study
Functions Name Sequences
Adh4F 5’‐GACGCTTCAGAATATTCCAATTC‐3’
Adh4RT7 5’‐TAATACGACTCACTATAGGGAGCTCGCTTAACGAACACTTATCG‐3’
Pgk1F 5’‐GGAGGATGCTCTTTCCCACGTC‐3’
Pgk1RT7 5’‐TAATACGACTCACTATAGGGAGCAAATTCACGAATGTTAGAGAAC‐3’ RNA probes Zrt1F 5’‐GGCAGCTGGTTTAGGTGTTCGTG‐3’
Zrt1RT7 5’‐TAATACGACTCACTATAGGGAGCACCAAATCAAGCCCATTTACC‐3’
Zym1F 5’‐CGATGGAACACACTACCCAATGTAAG‐3’
Zym1RT7 5’‐TAATACGACTCACTATAGGGAGCGATGATGCCCTTACTCTACGC‐3’
Loz1 H432A F 5'‐TCAAATTATTCTFATCATGCCAACAACGATAAGAGAGCTCATG‐3'
Loz1 H432A R 5'‐CATGAGCTCTCTTATCGTTGTTGGCATGATCAGAATAATTTGA‐3'
Loz1 H439A F 5'‐CAACGATAAGAGAGCTGCTGTATCGCGGCGACATTCAAC‐3'
Loz1 H439A R 5'‐GTTGAATGTCGCCGCGATACAGCAGCTCTCTTATCGTTG‐3' Site‐directed mutagenesis Loz1H444AF 5'‐CATGTATCGCGGCGAGCTTCAACTTCACGC‐3'
Loz1H444AR 5'‐GCGTGAAGTTGAAGCTCGCCGCGATACATG‐3'
Loz1H454AF 5'‐CAAAATTGCACAATCCTGTACCGGTTCTTCGTC‐3'
Loz1H454AR 5'‐GACGAAGAACCGGTACAGGATTGTGCAATTTTG‐3'
Zap1CY1BamH1 5'‐ATGAAGGATCCTTACTTGTATAGCTCGTCCATG‐3'
Zap1CY1EcoR1 5'‐GACCGAATTCATGGTGAGCAAGGGCGAGGAG‐3'
Zrt1promEagI 5'‐GATCTGCGGCCGGATCATGGATTGTTCCATGTG‐3'
Plasmid Zrt1promBamHI 5'‐ACTGGGATCCATTTTAATGTACGAGAAAGATAC‐3'
ZYM1EcoR1 5'‐TCAACGAATTCATGGAACACACTACCCAATG‐3'
ZYM1BamH1 5'‐TCAACGGATCCTTACTTCGAAGCACATTTGC‐3'
MT1EcoR1 5'‐TCAACGAATTCATGGACCCCAACTGCTCCTG‐3' Continued
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Table A.3 continued
MT1BamH1 5'‐TCAACGGATCCTCAGGCACAGCAGCTGCACTTCTC‐3'
NidAN8741EcoRIF1 5'‐GTCTGAGAATTCATGGATCTCGCCAACCTCATC‐3'
NidAN8741R1 5'‐GGAGAGCTGAGCCGTTGGCGCTTTGGAGCTTGTGTCGAG‐3'
NidAN8741IrF2 5'‐CTCGACACAAGCTCCAAAGCGCCAACGGCTCAGCTCTCC‐3'
NidAN8741BamHIr2 5'‐CCCGGGGATCCGCACCATCGCGACAGCCCTCCC‐3'
H4 F 5’‐AGCTGTCTATCGGGCTCCAG‐3’
H4 R 5’‐GGCCAGAGATCCGCTTAACGC‐3’
MT1 F 5’‐ GCACCTCCTGCAAGAAAAGCT‐3’
MT1 R 5’‐ GCAGCCTTGGGCACACTT‐3’
HiNF‐P F 5’‐GAGGAGGATGACCCACTTGA‐3’
HiNF‐P R 5’‐TCAGCTTGGTGTGGTAGCAG‐3’ RT‐PCR GAPDH F 5'‐GAGAAGTATGACAACAGCCTCAAG‐3'
GAPDH R 5'‐GGCAGGTCAGGTCCACCACTG‐3'
ATM F 5’‐CTGTGGTGGAGGGAAGATGT‐3’
ATM R 5’‐TGTTGATGAGGGGATTGCTGT‐3’
CKS2 F 5’‐ACCGGCATGTTATGTTACCC‐3’
CKS2 R 5’‐TGTGGTTCTGGCTCATGAAT‐3’
188