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THE DUAL NATURE OF REACTIVE OXYGEN SPECIES: REGULATION OF PH HOMEOSTASIS AND SURVIVAL IN SACCHAROMYCES CEREVISIAE BY ROS DAMAGE
AND SIGNALING
by James Allen Baron
A dissertation submitted to Johns Hopkins University in conformity with the requirements for the degree of Doctor of Philosophy
Baltimore, Maryland October, 2013
© 2013 J. Allen Baron All Rights Reserved
ii
ABSTRACT
pH homeostasis is intimately linked with the metabolic state of cells. The baker’s
yeast, Saccharomyces cerevisiae, is an excellent model of this as it readily switches from
fermentation to respiration contingent on carbon source availability. When grown in
abundant glucose, yeast obtain energy by fermentation and maintain a stable intracellular
pH over a variety of environmental pH’s by activating energetically expensive H+-
ATPases. When glucose is limited or absent, yeast switch to respiration, decrease the
activity of H+-ATPases to conserve energy, and pH homeostasis becomes highly
dependent on environmental pH and buffering by cellular metabolites. In this thesis, the
regulation of pH homeostasis by reactive oxygen species (ROS) is examined at these two
extremes of glucose abundance and glucose starvation using a variety of genetic and
biochemical techniques.
During long-term growth in the absence of glucose, yeast alternately alkalinize
and acidify their environment which affects growth and survival. We demonstrate that
mitochondrial ROS initiate the alkali-to-acid shift by inactivating the Fe-S cluster
enzymes aconitase and succinate dehydrogenase of the tricarboxylic acid (TCA) cycle
and that this shift is accelerated by deletion of the mitochondrial superoxide dismutase,
SOD2. Inhibition of the TCA cycle enzymes alters metabolite flux through the cycle and
leads to the buildup and secretion of the upstream metabolite acetic acid, which is
generated by the aldehyde dehydrogenase Ald4p. Acetic acid secretion acidifies the
environment without activating the plasma membrane H+-ATPase, Pma1p, and promotes
the growth of yeast under these conditions by providing a new carbon source for survival.
iii
This work demonstrates that inhibition of Fe-S cluster enzymes by ROS can be beneficial
under conditions of glucose starvation.
In abundant glucose the principally cytosolic superoxide dismutase, Sod1p, is
required for maximal activation of Pma1p, although the mechanism was not fully
understood. In this work we discovered that deletion of SOD1 in combination with a
specific mutation in the C-terminal autoinhibitory region of Pma1p, Pma1-T912D, is
lethal to yeast cells. This lethality can be rescued by treatment with Mn-based
antioxidants and by hypoxia. Furthermore, we identified spontaneous second-site
mutations in PMA1 that reverse the aerobic lethality by activating the H+-ATPase. Thus,
lethality results from profound inhibition of essential Pma1p activity. Together these
results support a model in which Sod1p helps protect Pma1p from oxidative damage,
particularly in cases where auto-inhibition of Pma1p is disrupted, as with Pma1-T912D.
Thesis Advisor: Dr. Valeria Culotta
Thesis Readers: Dr. Michael Trush
Dr. Rajini Rao
Dr. Jiou Wang
iv
ACKNOWLEDGEMENTS
The first line of a song released in 1965 by the Lettermen speaks an important
truth “No man is an island, no man stands alone.” This is especially true in the life
sciences where expertise, reagents, and ideas are constantly shared. With this in mind, I
acknowledge those who contributed during the course of my PhD studies to this thesis.
Apart from myself, Dr. Valeria Culotta has done the most to ensure both the
success of this work and of my PhD career. She provided a place to do research and a
project to work on when I was required to find a new lab in my 4th year of study. Her
expertise established the basis of my knowledge about reactive oxygen species and
metals in biology. Her advice and guidance in both writing and presenting has been
essential in developing this thesis and improving my technical communication skills.
Janice Chen worked with me as an undergraduate from 2011 - 2013 and
contributed to this work by producing strains and plasmids, measuring intracellular
metabolites, performing growth tests and investigating a number of side projects. Of
particular note, Janice helped sort out the role of acetate and succinate in the acid burst.
To acknowledge others who contributed to the experiments found in this thesis:
Kaitlin Laws initiated the studies of Chapter 2 as a rotation student and some of her data
is included in her thesis; David Corrigan did early work that contributed to Chapter 3;
Dr. Aaron Haeusler and Sabin Mulepati helped me use a french press and microfluidizer,
respectively; Paul Miller helped me freeze samples for TCA metabolite analysis; Dr.
Yana Sandlers, assistant director of the Biochemical Genetics Laboratory at KKI,
performed the stable-isotope dilution to measure TCA metabolites; Caelin Potts provided
sage advice about indirect immunofluorescence and use of the microscope; Dr. Yong-
v
Qiang Zhang provided protocols for crude membrane isolation and Pma1p activity
assays; Dr. Brian Learn provided technical expertise and access to an ultracentrifuge
rotor required for crude membrane isolations; various members of the Culotta lab
including Christine Vasquez, Dr. Julie Gleason, Chynna Broxton, Cissy Li, Dr. Daphne
Aguirre, and Dr. Jeffry Leitch inoculated/collected cells and provided reagents, among
other things; Margaret Dayhoff-Brannigan provided access to a dissecting scope which
helped me to sort out which strains develop rho- mutations; lastly, Brendan Cormack
provided GFP which will be fused to Pma1p for future experiments.
Dr. Amit Reddi deserves special recognition. He was examining the regulation of
Yck1p by Sod1p while I was doing the experiments in Chapter 3. This proved to be very
advantageous for comparison and discussion of results. He also introduced me to protein
activity assays and pH calculations using pH indicator dyes (Chapter 2). His influence
and professional expertise has proved invaluable.
To acknowledge those whose support was essential to the work of my PhD and
the production of this thesis: Rob Jensen, Rajini Rao, and Jiou Wang served as my thesis
advisory committee; Sharon Warner and Geraldine Graziano guided my academic
pursuits and helped with arranging the thesis defense and committee; Michael Matunis
provided guidance when switching labs; Gabriel Brandt provided assistance with
topology maps; Martha Baron helped with organization and formatting of this thesis; and
Marsha Baron cared for our son so I could devote the time needed to writing.
Drs. Eric Grote and Valerie Olmo worked closely with me in my early PhD career
examining the role of calcium in cell-cell fusion. Their contributions were great and
cannot be properly enumerated here.
vi
TABLE OF CONTENTS
Abstract ……………………………………………………………………………... ii
Acknowledgements …………………………………………………………………. iv
Table of Contents …………………………………………………………………… vi
List of Tables ……………………………………………………………………….. viii
List of Figures ………………………………………………………………………. ix
List of Abbreviations ……………………………………………………………….. xi
Chapter 1 – Introduction
Introduction to Reactive Oxygen Species …………………………………... 2
Sources of ROS ……………………………………………………………... 5
Targets of ROS ………………………………………………………........... 10
Antioxidant Enzymes ……………………………………………………..... 12
ROS in Signaling …………………………………………………………… 16
pH Homeostasis in the Baker’s Yeast S. cerevisiae …………………........... 18
The Research in This Thesis ………………………………………….......... 24
References …………………………………………………………….......... 30
Chapter 2 – Superoxide Triggers an Acid Burst in Saccharomyces cerevisiae to
Condition the Environment of Glucose-starved Cells
Introduction ………………………………………………………………… 61
Materials and Methods ……………………………………………………... 63
Results ……………………………………………………………………… 68
Discussion …………………………………………………………….......... 76
vii
References …………………………………………………………….......... 96
Chapter 3 – Oxidative Stress and Lethality in S. cerevisiae with Mutations in SOD1
and the Proton ATPase PMA1
Introduction ………………………………………………………………… 108
Materials and Methods ……………………………………………………... 110
Results ……………………………………………………………………… 114
Discussion …………………………………………………………….......... 119
References …………………………………………………………….......... 136
Chapter 4 - Review and Future Directions
Review of Thesis Research ………………………………………………. 148
Future Directions …………………………………………………………… 152
References …………………………………………………………….......... 156
Curriculum Vitae …………………………………………………………………… 159
viii
LIST OF TABLES
TABLE 2-1. Effects of TCA and ETC mutants on extracellular pH ........................ 79
TABLE 2-2. Contribution of acetic acid to pH of growth medium ……………….. 81
TABLE 3-1. Effect of Pma1 mutations on activity and C-terminal conformation ... 122
TABLE 3-2. BY4741 strains used in this study …………………………………... 123
TABLE 3-3. NY13 strains used in this study ……………………………………... 124
TABLE 3-4. Plasmids used in this study ………………………………………….. 125
ix
LIST OF FIGURES
FIGURE 1-1. Topology model of the plasma membrane H+-ATPase, Pma1p ……. 26
FIGURE 1-2. Diagram of extracellular pH changes and ammonia signaling by yeast
during long-term growth in the absence of glucose ………………………………… 28
FIGURE 2-1. Yeast cells alkalize their environment under respiratory conditions .. 82
FIGURE 2-2. Acid production following the alkaline phase of yeast cell growth is
accelerated by sod2∆ mutations and eliminated by ald4∆ mutations ……………… 84
FIGURE 2-3. Role of mitochondrial SOD2 and Mn-antioxidants in preventing the acid
burst of respiratory cells ……………………………………………………………. 85
FIGURE 2-4. The redox cycler paraquat and sdh∆ induce a rapid acid burst under
respiratory conditions and are not additive ………………………………………… 86
FIGURE 2-5. Activities of mitochondrial Fe-S cluster enzymes in acid producing cells
……………………………………………………………………………………… 88
FIGURE 2-6. Acid production with low glucose does not correlate with activation of
Pma1p, the proton ATPase ………………………………………………………… 90
FIGURE 2-7. Cells produce high levels of extracellular acetate during the acid burst
……………………………………………………………………………………… 91
FIGURE 2-8. The mitochondrial Ald4p aldehyde dehydrogenase is required for the acid
burst ………………………………………………………………………………… 93
FIGURE 2-9. Medium conditioned during the acid burst can promote yeast growth
……………………………………………………………………………………... 94
x
FIGURE 3-1. Effect of sod1∆ on Pma1 localization and growth of Pma1 mutants with
altered glucose regulation ………………………………………………………….. 126
FIGURE 3-2. Mutations in Pma1 stalk segment 5 that alter glucose regulation are not
synthetically lethal with sod1∆ …………………………………………………….. 128
FIGURE 3-3. T912D sod1∆ cells develop suppressors with secondary mutations in
PMA1 ………………………………………………………………………………. 129
FIGURE 3-4. The PMA1-A906G mutation increases Pma1p ATPase activity but does
not alter Pma1p localization or protein level ………………………………………. 131
FIGURE 3-5. T912D sod1∆ synthetic lethality is due to oxidative stress and not Yck1p
misregulation ………………………………………………………………………. 132
FIGURE 3-6. Pma1-T912D mutant cells are not sensitive to environmental oxidants or
hyperoxia …………………………………………………………………………... 134
FIGURE 4-1. Models depicting ROS regulation of pH homeostasis at the metabolic
extremes of glucose abundance and glucose starvation ……………………………. 150
xi
LIST OF ABBREVIATIONS
A absorbance
ACO1 aconitase
AHP1 thioredoxin peroxidase
AKR1 palmitoyl transferase, palmitoylates Yck1p
Ala alanine
ALD aldehyde dehydrogenase
ALD2 minor cytosolic ALD, stress-induced
ALD3 minor cytosolic ALD, stress-induced
ALD4 major mitochondrial ALD, glucose-repressed
ALD5 minor mitochondrial ALD, constitutively expressed
ALD6 major cytosolic ALD, constitutively expressed
Asp asparagine
ATO1 putative acetate importer/ammonia exporter
ATO2 putative acetate importer/ammonia exporter
ATO3 putative ammonia exporter
ATP adenosine triphosphate
BCG bromocresol green
BCP bromocresol purple
BSA bovine serum albumin
BSD2 heavy metal homeostasis protein, deletion suppresses sod1Δ
phenotypes
CaCl2 calcium chloride
xii
CCS copper chaperone for SOD1
CIT1 citrate synthase
CO2 carbon dioxide
CoA coenzyme A
CTT1 cytosolic catalase
Cu copper
Cys cysteine
DABCO 1,4-diazabicyclo[2.2.2]octane
DAPI 4’,6-diamidino-2-phenylindole
DCIP 2,6-Dichloroindophenol
DNA deoxyribonucleic acid
DTT dithiothreitol
DUOX dual oxidase
E redox potential
EDTA ethylenediaminetetraacetic acid
EGF epidermal growth factor
EGTA ethylene glycol-bis(2-aminoethylether)-N,N,N′,N′-tetraacetic
acid
ETC electron transport chain
FAD flavin adenine dinucleotide
Fe iron
Fe-S iron-sulfur
FGF-2 fibroblast growth factor-2
xiii
FUM1 fumarase
GC-MS gas chromatography-mass spectrometry
GLR1 glutathione reductase
GPX1 glutathione peroxidase
H+ proton
H2O2 hydrogen peroxide
H2SO4 sulfuric acid
HCl hydrochloric acid
HO• hydroxyl radical
HO2• hydroperoxyl radical
HOCl hypochlorous acid
hrs hours
IDH1 isocitrate dehydrogenase
IMS intermembrane space
KCN potassium cyanide
KGD1 α-ketoglutarate dehydrogenase
Km Michaelis constant
KNO3 potassium nitrate
KPhos potassium phosphate
Leu leucine
LEU1 isopropylmalate dehydratase
LSC1 succinyl-CoA ligase
Lys lysine
xiv
LYS4 homoaconitase
MDH1 malate dehydrogenase
MES 2-(N-morpholino)ethanesulfonic acid
Met methionine
MgSO4 magnesium sulfate
mins minutes
mito mitochondrial
Mn manganese
MnCl2 manganese chloride
MOPS 3-(N-morpholino)propanesulfonic acid
NAD+ (NADH) nicotinamide adenine dinucleotide
NADP+ (NADPH) nicotinamide adenine dinucleotide phosphate
NaN3 sodium azide
NHA1 yeast plasma membrane Na+/H+ antiporter
NHE1 mammalian Na+/H+ exchanger
NHX1 yeast endocytic Na+(K+)/H+ exchanger
Ni nickel
NOX NADPH oxidase
Nox2 phagocytic NOX enzyme, also referred to as gp91phox
O2 molecular oxygen
O2• −
superoxide
OD600 optical density, 600 nm
PBS phosphate-buffered saline
xv
PCD programmed cell death
PCR polymerase chain reaction
PDGF platelet-derived growth factor
pHex extracellular pH
pHi intracellular pH
pKa acid dissociation constant
PMA1 major plasma membrane proton ATPase
PMA2 minor plasma membrane proton ATPase
PMR1 golgi Ca2+/Mn2+ P-type ATPase, deletion suppresses sod1Δ
phenotypes
PMSF phenylmethylsulfonyl fluoride
PQ paraquat
psi pounds per square inch
PTK2 yeast Ser/Thr kinase, phosphorylates Pma1p
PTP protein tyrosine phosphatase
RHO+ WT mitochondrial DNA
rho- mitochondrial DNA mutant
ROS reactive oxygen species
rpm revolutions per minute
S. cerevisiae Saccharomyces cerevisiae
SC synthetic complete
SD standard deviation
SDH succinate dehydrogenase
xvi
SDS sodium dodecyl sulfate
SERCA sarcoplasmic/endoplasmic reticulum Ca2+-ATPase
SOD superoxide dismutase
SOD1 cytosolic Cu/Zn-SOD
SOD2 mitochondrial Mn-SOD
-SOH sulfenic acid
-SO2H sulfinic acid
-SO3H sulfonic acid
SOK2 yeast negative transcriptional regulator of pseudohyphal
differentiation
TCA tricarboxylic acid
TNF tumor necrosis factor
TNFR tumor necrosis factor receptor
TRR2 mitochondrial thioredoxin reductase
TSA1 thioredoxin peroxidase
TUF1/RAP1 positive transcriptional regulator of Pma1p
URA3 orotidine-5'-phosphate (OMP) decarboxylase
V-ATPase vacuolar proton ATPase
VEGF vascular endothelial growth factor
WT wild-type
YCK1 yeast casein kinase I homologue
YNO1 yeast NADPH oxidase
YP yeast extract, peptone
xvii
YPD yeast extract, peptone, 2% dextrose
YPDE YPD + 15 mg/L ergosterol and 0.5% Tween 80
Zn zinc
1
CHAPTER 1
Introduction
2
1. Introduction to Reactive Oxygen Species
1.1. Definition of Reactive Oxygen Species
Reactive oxygen species (ROS) comprise a set of molecules/compounds derived
from molecular oxygen (O2) that have increased reactivity relative to O2 in its ground
state. This includes the three species derived from the consecutive univalent reduction of
O2 prior to the formation of water: superoxide (O2• −), hydrogen peroxide (H2O2), and
hydroxyl radical (HO•). These ROS are the focus of this thesis.
ROS also include O2 molecules in which the normally unpaired electrons of parallel spin
have spin-flipped (i.e. have antiparallel spin) referred to as singlet oxygen.
1.2. The Dual Nature of ROS
For more than 40 years, it has been clear that ROS are significant damaging
agents in cells [1]. Discovery of the wide array of cellular antioxidant defenses in nearly
all aerobic organisms, as well as, a number of strict anaerobes [2] and the production of
ROS by cells of the immune system to eliminate invading pathogens [3] underscores their
destructive nature. More recently, a second role for ROS in cell signaling has been
identified. This much younger field of study began with the discovery of enzymes in
cells outside of the immune system that purposely produce ROS in response to specific
stimuli and the pathogenic states that arise when ROS production is diminished or
eliminated [4]. Although many roles for ROS in signaling have been uncovered, we still
know very little. This dual nature of ROS as both damaging and signaling agents
requires cells to exquisitely balance ROS generation and removal to ensure their required
O2
• − O2 H2O2 HO
• H2O
e-
e-
2H+
e
-
e-
2H+
3
role in signaling is achieved without causing significant damage to the cell. What
remains unclear is the line that divides this dual nature.
1.3. ROS in Human Health and Disease
Numerous human diseases have been linked to altered ROS status, including
cancer, diabetes, cardiovascular disease, neurologic disorders, and Down Syndrome to
name a few. The exact contribution of ROS to these diseases is not clearly understood
but there is plenty of evidence that suggests ROS play both damaging and signaling roles.
In the development of cancer, for example, high levels of ROS can be a driving factor by
causing damage to DNA [5,6]. At the same time, increased levels of ROS may activate
growth signaling pathways and enhance the production of factors that promote metastasis
[7]. Another interesting example is insulin signaling and the development of insulin
resistance, a factor contributing to both type 2 diabetes and obesity. In this case, ROS
appears to be required for normal insulin signaling, as well as, to contribute to insulin
resistance [8]. Normally, insulin stimulates a cascade of events that produce ROS, which
then amplify the cellular response to insulin. However, elevated levels of ROS may
contribute to insulin resistance by causing mitochondrial damage and dysfunction or by
activating signaling pathways that antagonize the cellular response to insulin.
1.4. Basic Properties of ROS
Although by definition ROS are all more reactive than O2, their properties are not
exactly the same. While some conclusions can be drawn about the reactivity of
superoxide, hydrogen peroxide, and hydroxyl radical with biomolecules (addressed
generally in Section 3), the direct targets of each may be difficult to ascertain because
these three ROS are readily interconverted in the cellular environment. To establish a
4
foundation the basic properties of superoxide, hydrogen peroxide, and hydroxyl radical
will be introduced.
1.4.1. Superoxide
Superoxide (O2• −) is essentially molecular oxygen with an extra electron. As
such, it is a free radical and has a negative charge. With a single-electron redox potential
of E = +0.89 V it is a much stronger oxidant than triplet oxygen (E = -0.16 V) [9].
Because of its unpaired electron, it primarily participates in single-electron transfer
reactions which limits its direct reactivity with biomolecules. Its anionic nature also
limits its reactivity with negatively charged biomolecules and can prevent it from
crossing cellular membranes [10]. The protonated form of superoxide, hydroperoxyl
radical (HO2•), is not limited in this manner but, with a pKa ≈ 4.88, less than 1% of
superoxide is protonated in cells at neutral pH [11-13].
An important property of superoxide is that it can act as both a reductant and an
oxidant. This permits the disproportionation of superoxide to hydrogen peroxide and O2.
The rate of this reaction is dependent on pH due to charge repulsion. At neutral pH the
second-order rate constant is k ≈ 105 [13-15].
1.4.2. Hydrogen Peroxide
Hydrogen peroxide (H2O2), unlike the other ROS described here, is not a free
radical. It is not as strong of a single-electron oxidant as superoxide (E = +0.38 V) but is
a very strong two-electron oxidant (E = +1.349 V) [9]. It is may also react with
biomolecules by electrophilic attack.
O2
• − O2 H2O2 O2
• −
2H+
+ +
5
With a pKa ≈ 11.6, hydrogen peroxide is primarily neutral in cells [16]. It is also
the most stable of the ROS, with a half-life measured in seconds to minutes [17,18], and
is able to cross biological membranes.
1.4.3. Hydroxyl Radical
Hydroxyl radical (HO•) is the strongest oxidant of the ROS described here (E =
+2.32 V) [9]. Furthermore, its reactivity is not limited by charge repulsion or kinetic
restraints. It is, therefore, very unstable and reactive. Many of the effects attributed to
ROS are likely to be caused by hydroxyl radical, as it is readily generated from hydrogen
peroxide and redox-active metals by Fenton & Fenton-like chemistry (see Section 2.3).
2. Sources of ROS
ROS are produced in cells by enzymes, redox-active metals, and by a variety of
environmental/exogenous sources. Three significant sources of cellular ROS have been
implicated in this thesis: the mitochondria, primarily as a by-product of the electron
transport chain (ETC) [19], the NADPH oxidases, and metal-catalyzed ROS.
2.1. Mitochondrial ETC
The ETC is widely believed to be a large producer of ROS in cells. It is
comprised of four redox-active, multi-subunit complexes that collectively couple electron
transfer to H+ transport across the inner mitochondrial membrane to generate an
electrochemical gradient. The final step in the ETC is a 2-electron transfer to the ultimate
electron acceptor O2 along with protons to generate water [20,21]. All other steps in the
ETC catalyze single-electron transfers [21]. For this reason, the primary ROS produced
by the ETC is superoxide.
6
Examining the redox potentials of all reactions in the ETC and comparing them
with that of O2 permits identification of the complexes with the potential for superoxide
production. The redox potential of O2 is estimated to be E = -130 mV [21,22]. Only
complexes I (E = -20 to -380 mV) and III (E = -400 to 700 mV) have redox potentials
lower than that of O2 and are able to reduce it [21]. Complexes II (E = -90 to 140 mV)
and IV (E = 210 to 815 mV) do not [21]. A large body of experimental evidence
validates complex I and III as ROS producers [23-32]. The baker’s yeast,
Saccharomyces cerevisiae, used in these studies does not have a true, multisubunit
complex I [33]. Therefore, complex III is likely to be the major ROS producer of the
ETC in yeast.
It is worth noting that the ETC, more particularly complex III, can produce
superoxide on both sides of the inner mitochondrial membrane [23-27]. Superoxide
produced in the intermembrane space (IMS) may also escape to the cytoplasm, with
voltage dependent anion channels implicated in its transport [34]. A small fraction could
also exit mitochondria in the protonated hydroperoxyl radical form. Therefore, the ROS
produced by the ETC are not strictly limited to affects in the mitochondria but can also
affect the remainder of the cell.
A variety of factors influence the quantity of ROS produced by the ETC. While
some of these are exogenous in origin (e.g. inhibitors such as rotenone or antimycin A)
[24,28,29,32,35], others are genetic (e.g. mutations in complex II) [36-39] or metabolic in
nature (e.g. NADH/NAD+ ratio, carbon source utilization) [28,30-32,35]. We examine
the effects of mitochondrial ROS under specific metabolic conditions in Chapter 2.
2.2. NADPH Oxidases
7
The NADPH oxidases (NOX) are a family of multi-subunit enzymes that utilize
electrons from NADPH to produce superoxide. At their core, the catalytic subunit of all
NOX enzymes has six membrane-spanning domains, two hemes, and conserved
NADPH- and FAD-binding sites [40,41]. Some members have additional
transmembrane domains and/or calcium-binding EF-hand motifs [40]. The dual oxidases
(DUOX) also have a peroxidase-like domain and hydrogen peroxide appears to be the
primary ROS they produce [40].
NOX enzymes have solidified our acceptance of the dual nature of ROS. They
create ROS as damaging agents during innate immunity and in non-immune cells can
regulate a whole host of signaling pathways [40,42]. A NOX enzyme was also recently
identified in yeast with potential implications for this thesis (see Sections 2.2.2 and 5.3;
also Chapter 3) [43].
2.2.1. Nox2 - ROS Produced During the Mammalian Immune Response
The first NOX enzyme discovered, Nox2 (also referred to as gp91phox), was
identified in phagocytic cells where it generates the ROS responsible for destroying
invading pathogens, referred to as the “respiratory burst”. Individuals with defects in
Nox2 subunits develop chronic granulomatous disease and exhibit an inability to
eliminate certain microbial and fungal infections [41,44]. The ROS produced by Nox2
can be converted to other damaging species, such as the hypochlorous acid (HOCl; i.e.
the active agent in bleach) formed by myeloperoxidase [45-47].
2.2.2. Nox Enzymes in Non-immune Cells and Yeast
Some time after discovery of Nox2, other mammalian NOX enzymes were
identified. These enzymes are found in tissues throughout the body, including the
8
vascular smooth muscle, kidneys, gastrointestinal tract, inner ear, thyroid, testis, and
uterus [40,48]. While some may be involved in immunity, these enzymes are primarily
involved in cellular signaling pathways (see Section 5.1 for an example) [40,48].
The yeast NADPH oxidase, Yno1p (also known as Aim14p), was identified by
Rinnerthaler and colleagues only recently [43]. Yno1p, formerly, was believed to be a
ferric reductase based on homology but it has no ferric reductase activity and is, in fact, a
bona fide NOX enzyme [43,49]. Though little is known about its cellular role, Yno1p
has been implicated in regulating actin filaments, cell death, and glucose and oxygen
signaling [43,50]. Its role in glucose and oxygen sensing will be addressed in Section
5.3.
2.3. Metal-catalyzed ROS
Transition metals, particularly Fe and Cu, are both essential and toxic to
eukaryotic organisms. Because they are able to redox cycle they are important producers
of cellular ROS, catalyzing the conversion of hydrogen peroxide to the significantly more
reactive hydroxyl radical by Fenton and Fenton-like chemistry [51-53]:
To prevent the formation of hydroxyl radical by these metals their availability is strictly
regulated in cells. However, as will be discussed in Section 3.1, ROS themselves can
increase bioreactive levels of Fe.
2.4. Other Enzymatic Sources
Many other enzymes in cells are known to generate ROS. These enzymes are
located throughout mitochondria, peroxisomes, the endoplasmic reticulum, and the
cytosol, and include but are certainly not limited to cytochrome p450s, various oxidases,
H2O2 + Fe2+
/Cu+
HO• + HO
− + Fe
3+/Cu
2+
9
glycerol 3-phosphate dehydrogenase, pyruvate and α-ketoglutarate dehydrogenases, and
the electron-transfer flavoprotein of fatty acid beta-oxidation [29,35,54-59]. These will
not be reviewed in detail.
2.5. Environmental/Exogenous Sources
Environmental stimuli that produce ROS include ionizing radiation [60],
ultraviolet light [61], and exogenous toxicants [62,63]. These environmental sources are
pervasive and the ROS they produce are not always neutralized by cells. Two exogenous
ROS producers that are particularly relevant to this thesis include paraquat and
menadione.
Paraquat is a widely used herbicide, killing plants by accepting electrons from
photosynthetic machinery and subsequently transferring them to O2, effectively
regenerating the compound and producing superoxide [64]. In non-photosynthetic
organisms, the cellular reductant NADPH and the ETC can reduce paraquat to incite
superoxide production [63,65-73]. For this reason, mitochondrial damage due to
superoxide production appears to be the primary cause of paraquat toxicity but depletion
of cellular NADPH may also be important [65,67-69,74,75].
Menadione is a synthetic compound that has been used as a supplement based on
its vitamin K activity but is currently banned in the United States due to concerns about
toxicity. Like paraquat, menadione is a redox cycling compound that generates
superoxide [63,76-78]. However, it may also induce toxicity by forming glutathione
conjugates leading to diminished glutathione levels in cells [79-81]. For this reason,
menadione and paraquat may have different effects on yeast cells [82].
10
3. Targets of ROS
ROS can react with essentially all classes of biomolecules: DNA, lipids,
carbohydrates, and proteins. The consequences of ROS production can therefore, be
extremely detrimental to life. Since the work of this thesis focuses on the effects of ROS
on proteins, this section will be restricted to protein modifications that result from ROS.
These modifications include damage to Fe-S cluster cofactors, cysteine oxidation,
methionine sulfoxide formation, and protein carbonylation.
3.1. Damage to Fe-S Clusters
Fe-S clusters are redox-active cofactors composed of two to four Fe atoms
bridged by inorganic sulfur and are usually ligated to proteins by cysteine residues. The
reaction of superoxide with Fe-S clusters occurs with a second order rate constant ≈ 106-
107 M-1 s-1 [83-86], faster than the spontaneous disproportionation rate of superoxide (≈
105 M-1 s-1). Fe-S clusters are widely regarded as the primary targets of superoxide but
they are also inactivated at appreciable rates by hydrogen peroxide (≈ 103 M-1 s-1) [83-
85,87].
A classic Fe-S cluster protein inactivated by ROS is aconitase. Aconitase (Aco1p
in yeast) functions in the tricarboxylic acid (TCA) cycle and catalyzes the reversible
isomerization of citrate to isocitrate, forming cis-aconitate as an intermediate. It is one of
a class of [4Fe-4S] enzymes especially susceptible to ROS inactivation because one of
the Fe atoms is solvent exposed [83-85,87]. ROS oxidize this Fe atom resulting in cluster
destabilization and subsequent release of Fe2+ [86].
O2
• − + 2H
+ [4Fe – 4S]
2+ [4Fe – 4S]
3+ H2O2 + +
[4Fe – 4S] 3+
[3Fe – 4S] 1+
+ Fe2+
11
Though Fe-S cluster oxidation results in protein inactivation, it is reversible,
largely because Fe-S clusters can be repaired and re-assembled [88-91]. However,
release of the reduced Fe from the cluster may catalyze the formation of hydroxyl radical
by Fenton chemistry and result in significant secondary, irreversible damage to
surrounding biomolecules [92-94].
Fe-S cluster enzymes susceptible to ROS inactivation exist in both the
mitochondrial matrix and the cytoplasm and are involved in biosynthesis of various
amino acids (including Leu, Met, Lys), the TCA cycle (aconitase, succinate
dehydrogenase) and the ETC, among others [95-99]. We revisit superoxide inactivation
of the mitochondrial Fe-S cluster proteins, aconitase and succinate dehydrogenase (SDH),
in Chapter 2 of this thesis and show surprisingly, that their inactivation can actually be
beneficial in glucose starvation conditions.
3.2. Other ROS Targets
Cysteine oxidation, methionine sulfoxide formation, and carbonyl formation are
more general markers of oxidative damage that affect a large number of proteins
[11,16,100].
Protein cysteine residues can be oxidized by ROS to sulfenic acid (-SOH).
Frequently, the sulfenic acid derivative reacts with another cysteine to form a disulfide
bond [16]. These cysteine modifications are reversible and evidence is mounting that
they are important in signaling. A very important class of targets are the protein tyrosine
phosphatases, which have a catalytic cysteine that is inactivated by hydrogen peroxide
(see Section 5.1) [101]. Successive oxidation of sulfenic acid by ROS generates sulfinic
12
(-SO2H) and sulfonic acid (-SO3H) residues that are generally considered irreversible
[102,103].
Methionine residues in proteins are oxidized by ROS to methionine sulfoxide
[104,105]. This modification is enzymatically reversible by NADPH-requiring
methionine sulfoxide reductases [106,107] and may both activate and inactivate enzymes
[104], indicating it may not strictly be damaging but may also play a role in signaling.
Protein carbonylation is essentially non-specific damage to proteins [11,108].
Carbonyls are formed by ROS oxidation of arginine, lysine, proline, cysteine, threonine,
leucine, and histidine side-chains [108]. Carbonyls are irreversible and are often used as
biomarkers for oxidative damage [108,109].
4. Antioxidant Enzymes
The antioxidant defenses of cells are designed to prevent the accumulation of
ROS species to damaging levels and to repair damage when it occurs. They must also
carefully balance the level of ROS to allow signaling. The antioxidant enzymes used by
cells include superoxide dismutases, catalases, peroxidases, and the general reducing
factors glutathione and thioredoxin.
4.1. Superoxide Dismutases
Superoxide dismutases (SODs) are the first line of defense. They catalyze the
disproportionation of superoxide to hydrogen peroxide and O2 using Cu, Fe, Mn, or Ni as
a catalytic metal cofactor [110,111].
O2
• − + Cu
2+/Mn
3+ Cu
+/Mn
2+ + O2
O2
• − + Cu
+/Mn
2+ + 2H
+ Cu
2+/Mn
3+ + H2O2
13
Although superoxide spontaneously disproportionates at a fast rate (105 M-1 s-1 see
Section 1.4.1), SOD-catalyzed disproportionation occurs with rate constants in the 109 M-
1 s-1 range [15,112,113]. Based on homology, there are three major types of SODs
frequently referred to by the metal cofactors they use: Cu/Zn-SOD, Fe- or Mn-SOD, and
Ni-SOD [110]. Most eukaryotes have at least one largely cytosolic Cu/Zn-SOD and one
mitochondrial matrix Mn-SOD.
4.1.1. Cu/Zn-SOD
The largely cytosolic Cu/Zn-SOD (known in many organisms as SOD1) is a
homodimer where each subunit contains a Zn that serves as structural support, a catalytic
Cu, and a conserved and essential disulfide bond [110,111]. Free Cu levels in cells are
generally extremely low [114]. For this reason, most eukaryotes have a dedicated
chaperone known as CCS that delivers Cu and catalyzes disulfide bond formation in an
oxygen-dependent manner [114-119]. In baker’s yeast, this copper chaperone is
absolutely essential for activation of the Cu/Zn-SOD, known as Sod1p [120].
Cu/Zn-SOD is found primarily in the cytoplasm and mitochondrial IMS, though it
has been found in other locations throughout the cell [121-123]. In studies that have been
done in baker’s yeast and in mammals, the import of Cu/Zn-SOD1 into the IMS is
dependent on its interaction with CCS and a disulfide relay system [121,124-127]. IMS
Cu/Zn-SOD1 is important for scavenging superoxide released by the ETC on the outside
of the inner membrane [124]. Outside of the mitochondria, Cu/Zn-SOD protects
cytosolic proteins from oxidation and can also operate in cell signaling through NOX-
dependent pathways (see Section 5.1 and 5.3).
14
The importance of Cu/Zn-SOD is underscored by its abundance (≈90% of the
total SOD activity [128,129]) and the phenotypes that develop in sod1 null mice and
yeast. Sod1-/- mice develop normally and survive into adulthood but exhibit, among
other things, reduced lifespan, increased incidence of liver cancer, motor neuropathy, loss
of muscle mass, and reduced female fertility [130-136].
sod1Δ null mutants of baker’s yeast exhibit a plethora of phenotypes resulting
from superoxide damage. The activities of the Fe-S cluster proteins aconitase (Aco1p),
homoaconitase (Lys4p), and isopropylmalate dehydratase (Leu1p) are all reduced, and
the Lys4p and Leu1p defects result in aerobic auxotrophies for lysine and leucine, or a
dependency on these amino acids for aerobic growth [137]. Interestingly, Lys4p and
Aco1p are both mitochondrial matrix proteins which suggests Sod1p somehow provides
cross-compartment protection against superoxide. sod1Δ null yeast also exhibit a
dependence on exogenous methionine for aerobic growth, a defect resulting from reduced
levels of NADPH needed for methionine biosynthesis [99]. The mutants are additionally
sensitive to environmental oxidants including hyperoxia and the redox-cycling compound
paraquat [99,128,138], have defects in vacuolar morphology [139], and exhibit a high
mutation rate [140].
Deletion of SOD1 also has effects on signaling in yeast. For example, Sod1p is
required for stabilization of the Ca2+-activated phosphatase calcineurin [141]. More
recently, Sod1p has been shown to play a role in regulating the switch between
respiration and fermentation [50,140]. In this case, Sod1p stabilizes a key regulator of
glucose repression, the yeast casein kinase I homologue Yck1p, in oxygen- and glucose-
15
dependent manners. Details on this signaling pathway involving Sod1p and Yck1p are
presented below in Section 5.3.
4.1.2. Mn-SOD
The mitochondrial Mn-SOD (also known SOD2 in many eukaryotes) is a
homotetramer with a single Mn per subunit that localizes to the mitochondrial matrix
where it plays an important role in detoxifying superoxide produced within the
mitochondrial matrix [142,143]. Sod2-/- mice are neonatal lethal due to dilated
cardiomyopathy and possible central nervous system defects [144,145]. At the molecular
level these mice exhibit defects in the TCA cycle and ETC enzymes aconitase, SDH, and
complex I, as well as, extensive DNA damage [96].
Unlike Sod2-/- mice, the phenotypes of sod2Δ null yeast appear relatively minor.
SOD2 deletion in the baker’s yeast S. cerevisiae causes increased sensitivity to hyperoxia
and paraquat, reduced chronological lifespan, aconitase deficiency, and a hypermutation
rate but has no apparent effect on growth rate [98,146]. Interestingly, during long-term
periods of glucose starvation, sod2Δ null yeast are unable to enter a quiescent state
characterized by alkalinization of the extracellular environment [147]. This phenotype is
described in more detail in Section 6.4.2 and forms the basis of work in Chapter 2 of this
thesis.
4.2. Catalases, Peroxidases, Glutathione and Thioredoxin
Catalases and peroxidases are similar in that they both catalyze the decomposition
of hydrogen peroxide [103]. However, they differ in that catalases use heme to strictly
catalyze hydrogen peroxide decomposition to water and oxygen, whereas the peroxidase
enzymes use various cofactors to detoxify hydrogen peroxide or alkyl peroxides and to
16
produce a variety of other compounds [148]. Two general classes of peroxidases that
function in ROS scavenging include the glutathione and thioredoxin peroxidases, which
use glutathione and thioredoxin as cofactors, respectively [103,149]. These two cofactors
are also the primary regulators of the reducing environment of cells.
5. ROS in Signaling
The roles identified for ROS in signaling are increasing rapidly. For the sake of
brevity we will focus on three examples involving superoxide dismutase enzymes. The
first two demonstrate how regulation of the cytoplasmic Cu/Zn-SOD1 and mitochondrial
matrix Mn-SOD2 proteins can influence ROS signaling, respectively. The third is a
specific signaling pathway involving Sod1p in yeast that is directly pertinent to Chapter 3
of this thesis.
5.1. SOD1 Regulation of Growth Factor Signaling
Stimulation of growth factor tyrosine kinase receptors has been broadly shown to
induce ROS production by NOX enzymes [4,100]. The downstream result of this ROS
production is the reversible inactivation of protein tyrosine phosphatases (PTPs) which
amplifies the phosphorylation signals induced during growth factor signaling [101].
Juarez and colleagues demonstrated in mammalian cell culture systems that the NOX-
produced superoxide during growth factor signaling is converted to hydrogen peroxide by
SOD1 [150]. This SOD1-produced peroxide can then inactivate the PTPs for a variety of
tyrosine kinase receptors including those for EGF, PDGF, FGF-2 and VEGF.
17
5.2. Mitochondrial ROS Stimulation by TNF-α, an Apoptotic Signal
Tumor necrosis factor (TNF)-α is an initiating factor of programmed cell death
(PCD). It binds to the TNF receptor which results in a wide variety of downstream
signaling, eventually resulting in cell death [151]. One of the pathways activated by
TNFR stimulates ROS production by the ETC which in turn activates the c-Jun NH2-
terminal kinase initiating a positive feedback loop that enhances cell death [151,152]. In
support of the role of mitochondrial ROS in PCD, NF-kB mediated activation of the
SOD2 gene was shown to inhibit PCD [151]. The exact mechanism by which
mitochondrial ROS contribute to PCD has not been completely elucidated. In Chapter 2
of this thesis, we examine the effects of mitochondrial ROS and SOD2 in yeast under
conditions of glucose starvation and demonstrate that ROS have a beneficial effect.
5.3. ROS Regulation of Yck1p in Yeast
In the budding yeast S. cerevisiae, fermentation is preferred over respiration even
with abundant oxygen, similar to the Warburg effect of cancers and other rapidly dividing
cells. This repression of respiration is mediated in part by a signaling casein kinase at the
cell surface, Yck1p [153]. Recent work in our lab has shown that Yck1p-control of
glucose repression requires active Sod1p enzyme [50]. sod1∆ mutants respire with high
glucose and this correlates with loss of the Yck1p polypeptide. Examination of the
underlying mechanism demonstrated that Sod1p stabilizes Yck1p in an oxygen- and
glucose-dependent manner. Sod1p binds directly to Yck1p and the peroxide produced by
Sod1p protects Yck1p from degradation. Oxygen and glucose fermentation conditions
are needed to provide the superoxide substrate for Sod1p, which appears to be derived
from the aforementioned Yno1p NADPH oxidase (see Section 2.2.2) as opposed to the
18
mitochondria. When Yck1p is stabilized by Sod1p and peroxide, the casein kinase
triggers downstream signaling of many events favoring fermentation over respiration,
including activation of the plasma membrane H+-ATPase, Pma1p. In fact, sod1∆ mutants
show a decrease in Pma1p activity with high glucose, consistent with the requirement for
Sod1p in stabilizing Yck1p. However, other possibilities for Sod1p control of Pma1p
could not be excluded including those involving oxidative stress protection. The
mechanism by which Sod1p maintains essential Pma1p activity is examined in more
detail in Chapter 3 of this thesis.
6. pH Homeostasis in the Baker’s Yeast S. cerevisiae
Regulating pH is essential for life. To survive and function, cells must maintain
their internal pH (pHi) within strict tolerances to ensure the proper functioning of
enzymes and maintain electrochemical gradients required for the uptake of nutrients and
energy generation. In general, this is accomplished by the combined action of buffering
and proton/metabolite transport. An excellent conceptual review of pH regulation was
presented by Walter Boron [154].
6.1. Metabolism and pH in Baker’s Yeast
The baker’s yeast Saccharomyces cerevisiae provides an attractive model system
to study pH homeostasis as it relates to metabolism. When glucose is abundant, the
organism prefers to ferment rather than respire, and fermentation results in acidification
of the extracellular environment. Conversely, when glucose has been depleted or cells
grow with non-fermentable carbon sources, the organism respires and if anything, the pH
of the extracellular environment (pHex) is elevated [155-157]. The effects of carbon
19
source on pH homeostasis in yeast are a major topic of Chapters 2 and 3 of this thesis.
Thus, this section of the introduction focuses on mechanisms of pH control in baker’s
yeast.
S. cerevisiae relies on proton transport for both pHi regulation and the generation
of electrochemical gradients that drive secondary transport. Because of this, yeast cells
grow optimally in environments with low extracellular pH (pHex 3.5-6.0), though they are
able to grow in nutrient-rich media ranging from pHex 2.5-8.5 [158,159]. Since S.
cerevisiae has a strong preference for fermentation as its primary mode of energy
production, copious amounts of intracellular acid equivalents, including organic acids and
H+ are continuously generated [160]. Despite this acid production and the constant influx
of H+ driving nutrient uptake, yeast are able to maintain steady pHi ≈ 7.2 under
fermentative conditions independent from pHex between 3.0 and 7.5 [161], indicating a
significant pHi regulatory capacity. The primary machinery yeast employ to prevent pHi
acidification includes the plasma membrane H+-ATPase, Pma1p, and the vacuolar H+-
ATPase (V-ATPase). Yeast also export organic acids such as acetate generated from
fermentative metabolism to prevent their buildup in the cell [162-164]. The combined
action of Pma1p and organic acid secretion acidifies the extracellular environment, which
maintains a pH ≈ 4-5, an ideal range for yeast growth.
6.2. The Major pH Regulator of Yeast, Pma1p
Plasma membrane H+-ATPases, found throughout the fungal and plant kingdoms,
pump protons out of the cell and, therefore, serve as major regulators of pHi [165]. They
also generate an electrochemical H+ gradient that drives secondary nutrient uptake [166].
Yeast have two plasma membrane H+-ATPases designated Pma1p and Pma2p. Due to
20
low expression, Pma2p is not a significant regulator of pHi [167]. Pma1p, on the other
hand, is one of the most abundant proteins in the plasma membrane [168]. Its activity
correlates with cellular growth rate and the transporter is essential for life [169,170].
Pma1p, like all plasma membrane H+-ATPases, is a P-type ATPase. This family
includes the well-known sarcoplasmic/endoplasmic reticulum Ca2+-ATPase (SERCA)
and the plasma membrane Na+/K+-ATPase. Thus, understanding Pma1p regulation and
function may shed light on other important members of this family.
Structurally Pma1p is an integral membrane protein comprised of 10 α-helical
transmembrane segments with both the N- and C-termini located in the cytoplasm (see
Fig. 1-1). The large central portion of the protein between membrane segments 4 and 5 is
located in the cytoplasm and constitutes the ATPase domain. The C-terminus of the
protein contains an α-helix that acts as an inhibitory domain by binding to the ATPase
domain, presumably through interactions involving the extended α-helix of membrane
segment 5, referred to as stalk segment 5 [171-174]. Interactions between the C-terminal
inhibitory domain and the ATPase domain are particularly important under glucose
control as described below in Section 6.4.1. The function of the cytosolic N-terminus of
Pma1p is largely unknown.
Pma1p is not only essential, it is also a major consumer of cellular ATP [175].
Therefore, this proton ATPase must fall under tight control. The PMA1 gene is
transcriptionally regulated by carbon source through the TUF1/RAP1 transcription factor
[170,176]. Post-translationally, Pma1p can be regulated by phosphorylation,
ubiquitination and protein degradation, calcium signaling, glucose sensing, and glycolysis
[50,177-182]. Pma1p activity can also be controlled at the level of trafficking through
21
the secretory pathway. For example, deletion of essential subunits from the V-ATPase
results in mis-localization of Pma1p to the vacuolar membrane [183]. Numerous
conditions have been shown to enhance Pma1p activity which are at best poorly
understood. These include intracellular acidification, nitrogen starvation, weak acid
stress, and ethanol stress [184-187]. By comparison, the response of Pma1p to glucose
availability has been studied extensively and will be reviewed in Sections 6.4.1 below.
6.3. Other Transport Systems for pH Homeostasis
Along with Pma1p, a number of other ATPases contribute to cellular pH in
baker’s yeast. Foremost among these is the V-ATPase. The V-ATPase indirectly
regulates pHi by affecting Pma1p localization, as mentioned previously [183]. More
importantly, the V-ATPase directly regulates pHi by pumping H+ from the cytosol into
the vacuole where acidification is critical for vacuolar function and for providing an
electrochemical gradient for vacuolar membrane transport [160,183]. The Na+(K+)/H+
exchangers, Nhx1p (in the endosomal pathway) and Nha1p (on the plasma membrane)
are also important in pHi regulation. Deletion of NHX1 causes intracellular acidification
and defects in vesicle trafficking out of endosomal compartments [188]. Deletion of
NHA1, on the other hand, results in intracellular alkalinization [188,189].
6.4. Effect of Carbon Source on Regulation of pH Homeostasis in Yeast
As mentioned earlier, S. cerevisiae prefers fermentation over respiration to obtain
energy. Because of this, the presence of the fermentable carbon source glucose
significantly effects yeast biology, including pH homeostasis.
22
6.4.1. Glucose Control of pH
When added to media, glucose induces a rapid decrease in pHi followed by an
increase in pHi to a resting pH of 7.2 [1]. The initial decrease is attributed to acid
production during glycolysis, while the subsequent increase in pHi is attributed to
activation of the proton ATPases, Pma1p and the V-ATPase. Glucose enhances the H+-
pumped out of the cytosol and into the vacuole by increasing the number of V-ATPase
complexes assembled on the vacuolar membrane [190,191]. At the same time, glucose
increases the Vmax of ATP hydrolysis and Km for ATP of the cell surface proton ATPase
Pma1p, in part by inducing phosphorylation of its C-terminal autoinhibitory domain at
residues S899, S911, and T912 (Fig. 1-1) [192-194]. S899 is phosphorylated by the
kinase PTK2 and phosphorylation at this residue appears to regulate the decrease in Km
[194,195]. Successive phosphorylation of T912 followed by S911 is responsible for the
Vmax increase [192,194]. The kinases responsible for phosphorylating these residues have
not yet been identified. Together the increased activity of Pma1p and the V-ATPase are
able to maintain pHi in this state until glucose is consumed.
As the fermentation of glucose progresses numerous organic acids are generated.
The exact acids and their quantity can vary with growth conditions but the primary
organic acids secreted are acetic and succinic acid [164,196-198]. Carbonic acid may
also accumulate as the CO2 released absorbs protons in the acidic environment. Along
with the H+ pumped out by Pma1p these organic acids lower and maintain pHex ≈ 4-5, an
optimal condition for yeast growth [164]. We examine the secretion and contribution of
both acetic and succinic acids to pHex in Chapter 2 of this thesis.
23
6.4.2. Effects of Glucose Starvation on pH
When cells become starved for glucose or are grown in non-fermentable carbon
sources that promote respiration (e.g., glycerol or ethanol), pHi and pHex are dramatically
altered. The most significant difference is that yeast cannot maintain a stable pHi
independent of pHex [158]. In fact, pHi under these conditions slowly equilibrates with
pHex [158,161]. The primary cause for this is the reduced level of H+-pumping by Pma1p
and V-ATPase activity, essentially making the buffering within cells a primary factor in
regulating pHi. This buffering comprises the sum total of the buffering capacity of all the
weak acid/base pairs in a cell, including proteins, peptides, organic and inorganic
metabolites.
When extracellular glucose has largely been consumed following a long period of
fermentation, cells shift to a period of growth dependent on respiration (known as the
diauxic shift) [199]. The high levels of ethanol and organic acids such as acetate that
filled the extracellular environment during fermentation begin to enter the cell [199-201].
The uptake of acetic acid into the cell is believed to be toxic [186,196,202,203].
However, in studies reported in Chapter 2, we provide evidence that acetate can actually
be beneficial with prolonged periods of nutrient starvation.
On the other hand, if cells are continuously cultured in non-fermentable carbon
sources or in media containing very low glucose, the environment is not acidified. Instead
cells alkalinize their environment. This occurs because cells consume low levels of
organic acids in the environment (such as acetate) as a carbon source while also
producing weak bases such as ammonia and bicarbonate [155,157,204]. Media
alkalinization slows growth but prevents pHi from dropping too low [157,158,161].
24
Interestingly, S. cerevisiae colonies starved for glucose over long periods
alternately acidify and alkalinize their environment (Fig. 1-2A) [204,205]. The acidic
phases are thought to correspond to cell growth and the alkaline to a resting phase. In
depth studies by Palkova and colleagues have demonstrated that the alkaline phase of
growth coincides with release of volatile ammonia which then serves as a signal to
neighboring colonies to synchronize growth phase (Fig. 1-2B) [204-207]. As
neighboring colonies synchronize, the ammonia concentrations between them
preferentially increase over that on their periphery. This predominantly limits growth at
the colony interface preventing colonies from growing into one another and has been
proposed as a strategy yeast use to prevent colony competition for nutrients.
Palkova and co-workers have investigated the mechanism of this alternating
acidification and alkalinization using a yeast genetic approach. They identified a number
of genes which when deleted appear to prevent or limit entry into the alkaline phase,
including the putative ammonia and/or acetate transporters, ATO1, ATO2, and ATO3; the
transcriptional regulator SOK2, which represses pseudohyphal growth; the cytosolic
catalase, CTT1, and of great interest to us, the mitochondrial Mn-SOD, SOD2
[147,205,208]. The mechanism by which the mitochondrial matrix Sod2p controlled
extracellular pH was not known, and is the focus of work presented in Chapter 2.
7. The Research in This Thesis
In this thesis, we examine the role of ROS in regulating pH homeostasis of the
baker’s yeast, S. cerevisiae, at two metabolic extremes: that of glucose abundance, when
25
cells prefer to ferment, and that of glucose starvation, when cells obtain energy by
respiration.
The work in Chapter 2 addresses ROS regulation of pH under glucose starvation.
Specifically, it explores the mechanism by which yeast cells grown over long periods in
the absence of glucose shift from alkalinizing their surrounding environment to
producing acid and addresses why deletion of the mitochondrial Mn-SOD, SOD2, affects
this transition. We demonstrate that media acidification is initiated by ROS in the
mitochondrial matrix, which can be further enhanced by deletion of SOD2. These ROS
inactivate the Fe-S cluster enzymes, aconitase and SDH, which inhibits metabolite flow
through the TCA cycle leading to a significant build up and secretion of acetic acid by the
mitochondrial aldehyde dehydrogenase, Ald4p. This not only acidifies the extracellular
environment but provides a new carbon source for neighboring yeast cells that can
enhance growth under these conditions.
The work in Chapter 3 examines the role of cytosolic Sod1p in ensuring
activation of the plasma membrane H+-ATPase, Pma1p, during growth in abundant
glucose. During the course of these studies, we uncovered a surprising genetic
interaction between PMA1 and SOD1. Namely, a specific mutation in the C-terminal
autoinhibitory domain of Pma1p, Pma1-T912D, is lethal in sod1∆ cells, but not in SOD1+
cells. By exploring the mechanism underlying this “synthetic lethality” [209,210] we
provide evidence that Sod1p guarantees maximal activation of Pma1p by preventing
oxidative damage. This is likely in addition to its role in signaling via stabilizing the
Pma1p-regulatory kinase, Yck1p, as described previously [50].
26
FIGURE 1-1. Topology model of the plasma membrane H+-ATPase, Pma1p.
A topology map of Pma1p based on the previously published map of Lecchi, et al. [174]
with zigzag lines representing α-helices. The N- and C-termini are indicated with an “N”
and “C,” respectively and the 10 transmembrane α-helices are numbered from M1 to
M10. The ATPase domain of Pma1p is indicated by the grey dashed box with the
approximate binding site for ATP designated. The auto-inhibitory C-terminal α-helix
binds to the ATPase domain in the absence of glucose, presumably through interactions
involving the extended α-helix of membrane segment 5 (indicated by grey arrow) [171-
174]. This inhibition is relieved by glucose-induced phosphorylation of residues S899,
S911, and T912 found in the auto-inhibitory α-helix. The approximate location of these
residues is specified.
27
28
FIGURE 1-2. Diagram of extracellular pH changes and ammonia signaling by yeast
during long-term growth in the absence of glucose.
(A) Over a period of days to weeks on nonfermentable (i.e. respiratory) carbon sources,
yeast cells alternately alkalinize (purple color) and acidify (yellow color) their
extracellular environment [204]. The alkalinization slows cell growth while acidification
enhances growth. (B) The alkaline period is marked by production of volatile ammonia
(purple arrows). If colonies are not synchronized the ammonia produced by one causes
the neighboring colonies to enter the alkaline phase as well. Once synchronized, volatile
ammonia accumulates with the highest concentration between colonies (purple cloud).
This preferentially limits growth at the colony interface over that on the periphery
preventing colony competition for nutrients. (C) The mechanism by which cells enter the
acid phase and the effect this phase has on cells was previously unknown (indicated by
question marks). This is the subject of Chapter 2 of this thesis.
29
30
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60
CHAPTER 2
Superoxide Triggers an Acid Burst in Saccharomyces
cerevisiae to Condition the Environment of
Glucose-starved Cells
This research was originally published in the Journal of Biological Chemistry.
J.A. Baron, K.M. Laws, J.S. Chen, V.C. Culotta. Superoxide triggers an acid burst in
Saccharomyces cerevisiae to condition the environment of glucose-starved cells. J. Biol.
Chem. 2013; 288: 4557-4566. © the American Society for Biochemistry and Molecular
Biology.
61
1. Introduction
Cells exist and thrive in environments that are both supportive of life and
potentially hostile. The environment is in a constant state of change and the onset of
stress conditions, such as nutrient deprivation and exposure to toxins, require appropriate
adaptation strategies. Cells have evolved a variety of mechanisms to appropriately sense
and respond to variations in the environment through induction of stress pathways and
adjustments in metabolism. Yet in some cases, cells respond by modifying the
microenvironment itself to enhance its suitability for their fitness and survival. The
pathogen Plasmodium falciparum, for example, is known to secrete a variety of proteins
that enhance the permeability of the red blood cell membrane to increase the availability
of nutrients [1]. Cancer cells are particularly adept at altering microenvironments. During
the process of invasion, these cells secrete a variety of metalloproteases [2] and also
acidify their surrounding environment to inhibit the toxic activity of immune cells and
chemotherapies, and promote migration to more nutrient-rich environments [3,4]. To
make nutrients more accessible, the bakers' yeast Saccharomyces cerevisiae secretes
phospholipases [5], a variety of enzymes that break down sugars [6], and phosphatases
[7].
To S. cerevisiae, the pH of the environment is critical for regulating growth and
survival [8-10]. Cells growing on the preferred carbon source glucose acidify the
surrounding media by activating the proton ATPase, Pma1p [11], and secreting organic
acids [12,13]. This acidic environment (≈ pH 4-5) helps to maintain the electrochemical
gradient across the plasma membrane and drive uptake of many essential nutrients [14].
On the other hand, when yeast cells grow on the less preferred non-fermentable carbon
62
sources or are starved for glucose, they alkalize their surrounding environment, which
slows growth but enhances survival under stress [8,15,16]. This period of alkalization
correlates with increases in volatile ammonia that serves as a signaling molecule to
synchronize colony growth [15-18]. Interestingly, following prolonged growth on the
non-fermentable carbon source glycerol, yeast cells can switch from this alkalization
phase to producing acid [17,19,20]. The mechanism or rationale for this sudden “acid
burst” has been unknown. In previous studies completed by Palkova's group [17,19,20],
the production of ammonia during the alkalization phase was reportedly defective in S.
cerevisiae cells lacking the mitochondrial matrix manganese-containing superoxide
dismutase, SOD2 (or yeast Sod2p). Sod2p was suggested to affect ammonia signaling,
but through an unknown pathway [19,21].
Superoxide dismutases (SODs) represent a family of metalloenzymes responsible
for detoxifying superoxide radicals generated as a by-product of aerobic metabolism.
Most eukaryotes, including S. cerevisiae, contain both a Mn-SOD2 in the mitochondrial
matrix as well as a Cu/Zn-containing SOD14 that is largely cytosolic but also found in
the mitochondrial intermembrane space [22,23]. The presence of SODs on both sides of
the mitochondrial inner membrane collectively protect against superoxide generated by
the electron transport chain. The importance of SOD2 in guarding against mitochondrial
oxidative damage has been underscored in various model organisms for SOD2
deficiency. SOD2-/- mice are neonatal lethal and early death in this case is thought to
arise from severe oxidative damage to the mitochondrial respiratory chain [24,25]. Loss
of SOD2 also induces early mortality in Drosophila adults, associated with damage to the
mitochondrial respiratory chain and TCA (tricarboxylic acid) cycle enzymes [26]. In the
63
bakers' yeast S. cerevisiae, cells lacking Sod2p are viable but are highly sensitive to
hyperoxia and redox cycling agents [27-29], defects ascribed to increased superoxide
damage to mitochondrial components. However, the means by which loss of yeast
mitochondrial matrix Sod2p can control extracellular pH as described earlier [19] is not
clear, particularly because the superoxide substrate for Sod2p should not cross the
mitochondrial membranes in significant quantities [30-33].
Here, we describe a new connection between mitochondrial superoxide and the
control of environmental pH. Specifically we find that the acid burst produced by cells
during long-term respiratory conditions can be induced by mitochondrial superoxide.
With either mutation in yeast SOD2 or by treatment with the redox cycler paraquat, the
acid burst is accelerated in yeast cells starved for glucose. Moreover, the acid burst is
eliminated through Mn-antioxidants that act as SOD mimics and remove intracellular
superoxide. We provide evidence that superoxide damage to Fe-S enzymes in the TCA
cycle results in massive production of acetate involving the mitochondrial aldehyde
dehydrogenase Ald4p. The concomitant acetate burst during nutrient starvation provides
a new carbon source to enhance cell growth during long-term nutrient deprivation.
2. Materials and Methods
2.1. Yeast Strains
The yeast strains in this study were all derived from the parent strain BY4741
(MATa, his3Δ1, leu2Δ0, LYS2, met15Δ0, ura3Δ0). Strains with single gene deletions
were obtained from the MATa haploid deletion library (Open Biosystems) and verified by
sequencing in earlier studies [27] or in the current studies: sod2Δ, sod1Δ, sdh2Δ, sdh4Δ,
64
cyt1Δ, cox7Δ, cit1Δ, idh1Δ, kgd1Δ, lsc1Δ, fum1Δ, mdh1Δ, ald2Δ, ald3Δ, ald4Δ, ald5Δ,
ald6Δ, pmr1Δ. Strains LJ111 (aco1::LEU2) (gift of L. Jensen), CO217 (coq1::LEU2)
[34], and LJ109 (rho−) [35] were described previously. Strains JAB075
(ald4::kanMX, sod2::URA3) and AR148 (pmr1::kanMX, sod2::URA3; gift of A. Reddi)
were created by transformation of the aforementioned ald4Δ andpmr1Δ strains,
respectively, with the sod2Δ::URA3 plasmid, pGSOD2 as described [36]. Strain JAB069
(ald4::kanMX, sdh4::HIS3) was created by transformation ofald4Δ with a PCR-
based sdh4::HIS3 deletion cassette generated by amplifyingHIS3 from pRS403 [37] using
as primers: forward primer, 5′-
ACGCTTTCGACTTTCTTCCTACGCGCTTTATAATAGCTATGGCGGCATCAGAG
CAGATTG-3′; reverse primer, 5′-
GTTACATGACCGAACAAATGATTCGTGGTGATTTATCTACGTTTACAATTTCC
TGATGCG-3′. Transformations were performed by the standard lithium acetate
procedure [38].
2.2. Culture Conditions and pH Measurements
To examine the effects of yeast colony growth on extracellular pH, solid growth
medium containing 3% glycerol (or where indicated, 2% glucose), 1% yeast extract,
0.01% bromocresol purple (BCP; Sigma, B5880), and 2% bacto-agar was prepared
precisely as described by Palkova and co-workers [15], except supplemental CaCl2 was
generally omitted. pH was typically adjusted to 5.75 with HCl prior to autoclaving but
could range from 4.35 (enhance detection of media alkalization) to 6.5 (enhance detection
of media acidification) without altering the cell alkaline and acid phases. When needed,
the specified concentrations of paraquat (MP Biomedicals) or MnCl2 were added
65
immediately before pouring. In all cases, extracellular pH was monitored using “giant
colony” growth as prescribed by Palkova et al. [15] where 2 × 105 cells in 10 μl were
spotted onto plates and incubated at 30 °C. Images were taken with a Sony Cybershot
DSC-F828 on the days indicated. Cell viability measurements were obtained by removing
cells at the designated times and measuring colony-forming units on YPD (1% yeast
extract, 2% peptone, 2% glucose) averaged over 3 giant colonies as described (20).
Results were normalized to WT at day 4. WT and sod2Δ strains retain full viability
through day 8 and at day 11 exhibited 92 (S.D. ± 8) and 89% (S.D. ± 11) viability,
respectively.
For monitoring effects of yeast growth under low glucose conditions, cells were
inoculated at an initial A600 = 0.05 and grown shaking at 220 rpm, at 30 °C for 24 hrs in a
liquid media consisting of 1% yeast extract and 0.2% glucose, adjusted to pH 5.5 with
HCl. To determine the extracellular pH of these cultures, a colorimetric assay based on
the pH-sensitive dyes BCP and bromocresol green (BCG; Sigma, 114359) was
developed. Briefly, cells were removed by centrifugation at 17,000 × g for 1 min and the
media was collected (90 μl each) in triplicate and applied to a 96-well plate. Water
(blank), 0.1% BCP, or 0.1% BCG (10 μl) were added and absorbance was measured
at A430, A589, A514, and A616 in a Synergy HT Multi-Mode Microplate Reader (BioTek). pH
was calculated by subtracting the absorbance of the blank at each wavelength and using
the following empirically derived equations:
For BCP (pH ≥ 5.3),
pH = 5.852 + 1.081 × LOG �𝐴589𝐴430
�
66
For BCG (pH < 5.3),
pH = 4.590 − LOG�5.963
�𝐴616𝐴514� − 0.045
− 1�
Plates were imaged on an HP Scanjet 8200 scanner. For conditioned media tests,
cultures from low glucose conditions as described earlier were filter-sterilized (Millipore,
Millex-GV, 0.22 μm) to remove cells and the conditioned medium collected was then
used for inoculation of WT cells at A600 = 0.1 followed by aerobic growth for the
indicated time points.
2.3. Metabolite Measurements
To measure glucose, acetate, and succinate concentrations in media, conditioned
medium was collected from cells grown in low glucose as described earlier and cells
were removed by centrifugation. Measurements were performed with a Synergy HT
Multi-Mode Microplate Reader using the QuantiChrom Glucose Assay Kit (BioAssay
Systems), the Acetic Acid kit (R-Biopharm), and the Succinic Acid kit (R-Biopharm),
respectively, according to the manufacturers' specifications. To adapt the Acetic Acid and
Succinic Acid kits to a plate reader format, one-tenth the recommended volumes were
used and each reading was path length corrected using the standard correction of the plate
reader: (A977 −A900)/0.18.
Intracellular acetate measurements were performed in the same manner as those
for media, except cells were first lysed by glass bead homogenization in 20 mM
potassium phosphate, 1.2 M sorbitol, 1.0 mM PMSF, pH 7.4, and clarified by
centrifugation at 17,000 × g for 10 mins. After determining the total protein by Bradford
assay, lysates were heated 15 mins at 85 °C and the supernatant was collected for analysis
67
after a second centrifugation. Intracellular acetate concentrations were normalized to total
protein.
To measure intracellular metabolites of the TCA cycle, cells grown as described
earlier in low glucose media were harvested at A600 = 1-1.2 and washed in PBS. Cells
were resuspended in PBS at a ratio of 5 μl of PBS/A600 cells and lysed by glass bead
homogenization as described [39]. Lysates were clarified by centrifugation at 17,000
× g for 10 mins and total protein was determined by Bradford assay. Duplicate samples
were analyzed by stable-isotope dilution GC-MS by the Kennedy Krieger Institute
Biochemical Genetics Laboratory.
2.4. Enzymatic Assays
For measurements of aconitase and succinate dehydrogenase (SDH) activity, cells
were collected from low glucose liquid media (10 ml) after 24 hrs growth by
centrifugation or scraped from glycerol plates containing BCP (2-9 × 108 cells collected
from strains plated as in Fig. 2-1A) on the days indicated and stored at -20 °C. Cells were
then resuspended in lysis buffer consisting of 20 mM potassium phosphate,
1.2 M sorbitol, 1.0 mM PMSF, 0.1% Triton X-100, pH 7.4, at a ratio of 10 μl of buffer/2
× 107 cells and lysed by glass bead homogenization. Protein concentrations were
determined by Bradford assay and enzymatic assay measurements were performed with
10-50 μg of lysate protein per reaction. Aconitase activity was measured as described
[39] in 250 μl of total reaction volume and calculated as the nanomoles of cis-aconitate
consumed per mg of protein per min using the Beer-Lambert law and an extinction
coefficient of 4.88 mM−1 cm−1. SDH activity was measured with the procedure of
Ackrell et al. [40], adapted to a 200-μl 96-well plate format. Briefly, SDH in whole cell
68
lysates was activated at 30 °C for 10 mins in the reaction mixture without electron
acceptors/dyes (178 μl volume; final concentration after addition of dyes = 20
mM potassium phosphate, 20 mM succinate or malonate, 1 mM KCN, pH 7.4). After 10
mins, pre-warmed dyes (22 μl volume; final concentration in complete reaction mix = 1
mM phenazine methosulfate (Sigma), 0.05 mM 2,6-dichlorindophenol (Sigma)) were
added to initiate the reaction. SDH activity was calculated as the malonate-sensitive
nanomoles of 2,6-dichlorindophenol reduced per mg of protein per minute using an
extinction coefficient of 21 mM−1 cm−1.
For Pma1p ATPase activity, cells were grown in 1% yeast extract with 2 or 0.2%
glucose, pH 5.5 to an A600 = 0.75-0.9. Plasma membranes were isolated [41] and Pma1p
ATPase activity was measured by assaying orthovanadate-sensitive release of inorganic
phosphate precisely as described by Monk et al. [42] and normalized to the activity of
WT cells grown in 0.2% glucose.
3. Results
3.1. A Role for Superoxide and SOD2 in Controlling the Acid Burst of Respiring Cells
Extracellular pH in yeast growth medium is readily monitored by pH indicators
such as BCP and BCG. When grown in high glucose, fermenting yeast cells rapidly
acidify the medium (Fig. 2-1A top), but when grown on non-fermentable carbon sources,
the respiring yeast cells initially elevate the extracellular pH (Fig. 2-1A,
middle and bottom). As prescribed by Čáp et al. [19], this alkalization is observed on
glycerol medium supplemented with 30 mM calcium (Fig. 2-1A, middle), yet is also seen
without added calcium (Fig. 2-1A, bottom); hence all subsequent studies were conducted
69
in the absence of calcium supplements. Alkalinization of the growth medium is not
unique to non-fermentable carbon sources, but is also observed in liquid cultures with
low glucose, conditions where cells switch from fermentation to respiration [8] (Fig. 2-
1B). Cells grown in 0.2% glucose rapidly deplete their glucose (Fig. 2-1C) and the pH of
the extracellular medium rises from 5.5 to ≥7.0 within 24 hrs (Fig. 2-1B).
Following long-term growth on glycerol containing medium, S. cerevisiae cells
can enter an acid phase as previously described, which we refer to herein as the acid burst
(Fig. 2-2) [17,19,20]. We observe that the initiation of the acid burst is substantially
accelerated in strains lacking Mn-SOD2, the mitochondrial matrix manganese-containing
SOD. The sod2Δ null cells show no defect in the initial alkalization of the medium (Fig.
2-2, Fig. 2-3A); however their onset of entry into the acid phase is accelerated compared
with WT strains (Fig. 2-2). This sod2Δ impact on extracellular pH appears specific to
respiratory conditions, as sod2Δ cells are indistinguishable from WT cells in acid
production under fermentative conditions with high glucose (Fig. 2-3A, right). Moreover,
the early acid phase of sod2Δ cells under respiratory conditions is not due to accelerated
death of these cells, because our viability tests (see Materials and Methods) estimate that
WT andsod2Δ both retain ≈ 90% viability after 11 days on glycerol.
The premature acid burst appears specific to sod2Δ strains and was not observed
in other yeast mutants we tested that control oxidative stress resistance. For example,
deletion of genes required for glutathione and thioredoxin reduction in the mitochondria
(GLR1 [27] and TRR2 [43]) did not show the same accelerated acid burst of sod2Δ cells,
nor did deletion of the thioredoxin and glutathione peroxidases AHP1 [44], TSA1 [45],
and GPX1 [46] (data not shown). We also observed that loss of SOD1, encoding the
70
cytosolic and mitochondrial intermembrane space Cu/Zn-SOD, does not induce a
premature acid burst (Fig. 2-3A). The effects on extracellular pH appear specific to loss
of SOD activity on the matrix side of the mitochondrial inner membrane.
In addition to SOD enzymes, yeast cells have the capacity to remove superoxide
through the action of small non-proteinaceous complexes of manganese such as Mn-
phosphate. These redox active compounds of manganese, so called Mn-antioxidants, are
effective SOD mimics in vitro and in yeast cells in vivo [47-53]. Mn-antioxidants are
elevated when yeast cells accumulate high manganese through either supplementation of
manganese salts to the growth medium or by mutations in the pmr1Δ Golgi pump for
manganese [49-51]. We observed that in both cases, the Mn-antioxidants prevented the
premature acid burst of sod2Δ strains (Fig. 2-3, B and C left). Moreover, boosting levels
of Mn-antioxidants in WT cells completely prevented cells from exiting the alkaline
phase and the acid burst was eliminated (Fig. 2-3C, right). These studies together with
results from sod2Δ cells strongly indicate that the transition to acid phase following long-
term respiratory conditions is triggered by superoxide.
Redox cycling agents such as paraquat can induce production of cellular
superoxide, including mitochondrial matrix superoxide [54-56]. As seen in Fig. 2-4A,
paraquat treatment dramatically accelerated the transition to the acid burst in glycerol and
within 24 hrs, both WT and sod2Δ cells treated with paraquat were producing copious
levels of acid. Much less paraquat was required in the case of sod2Δ cells (Fig. 2-4A),
consistent with the anticipated high superoxide in these mutants that cannot catalytically
remove mitochondrial matrix superoxide. In addition to effects on glycerol plates,
paraquat treatment also caused cells grown in low glucose to produce acid rather than
71
base (Fig. 2-4B). The paraquat-induced burst of acid was not due to cell death; in fact
total cell growth over 24 hrs was slowed by only 25-30% with doses of paraquat that
induced maximal acidification (Fig. 2-4C), which translates to an overall reduction in
growth rate of only ≈ 15%. These studies support the notion that acid production during
respiratory conditions can be induced by superoxide, specifically the mitochondrial
superoxide relevant to Sod2p.
3.2. Fe-S Cluster Enzymes of the TCA Cycle as Targets of Superoxide during the Acid
Burst
Superoxide can disrupt Fe-S clusters [57,58], and we tested whether the
mitochondrial Fe-S proteins SDH and aconitase were inhibited in cells producing acid. In
WT cells, both aconitase and SDH are affected by paraquat. Aconitase is inhibited even
by low (50 μM) levels of paraquat when cells are not producing acid, but maximal levels
of SDH inhibition requires very high (800 μM) paraquat, conditions where WT cells
produce acid (Fig. 2-5A). In sod2Δ cells, aconitase is constitutively low without paraquat
consistent with previous studies [58,59], and the main effect of 50 μM paraquat is
inhibition of SDH activity (Fig. 2-5A). Hence with WT and sod2Δ cells, acid production
is associated with losses in both aconitase and SDH. The same trends were apparent with
cells grown for extended periods on glycerol without paraquat: WT cells exhibited
reductions in both aconitase and SDH, whereas sod2Δ cells are constitutively low in
aconitase and lose SDH activity over time (Fig. 2-5B).
We tested whether the degree of SDH and aconitase inhibition observed in vitro
reflected disruptions in the TCA cycle, in vivo. TCA cycle metabolites were measured
from cells grown in low glucose and treated with paraquat. As controls, mutants of SDH
72
and aconitase were used. Yeast sdh4Δ mutants have high succinate but low citrate,
whereas aco1Δ mutants lacking aconitase have high citrate and low succinate (Fig. 2-
5C). WT cells treated with 800 μM paraquat exhibited the elevation in succinate typical
of sdh mutants, but did not lose citrate (Fig. 2-5C), consistent with the double inhibition
of SDH and aconitase (as in Fig. 2-5A). In sod2Δ null cells treated with 50 μM paraquat,
citrate was elevated consistent with losses in aconitase, but there was no loss of succinate
(Fig. 2-5C), again consistent with TCA blockages in both SDH and aconitase.
The correlation between the acid burst and TCA cycle inhibition prompted us to
test whether mutations in TCA cycle genes were by themselves able to induce an acid
burst. As seen in Fig. 2-4D, the aco1Δ and sdh2Δ mutants lacking aconitase and SDH
showed acid production in low glucose, analogous to what was seen with paraquat
treatment. Moreover, there was no additive effect of paraquat, indicating that paraquat
and loss of the TCA cycle operate in the same pathway to induce acid. Acid production is
not specific to disruptions at aconitase and SDH, as mutations in citrate synthase (cit1Δ),
α-ketoglutarate dehydrogenase (kgd1Δ), and fumarase (fum1Δ) all resulted in acid
production (Table 2-1). Disruptions in 5 of 8 steps in the TCA cycle mimicked the effects
of paraquat and induced acid production in low glucose. Only deletions in isocitrate
dehydrogenase (idh1Δ), succinyl-CoA ligase (lsc1Δ), and malate dehydrogenase
(mdh1Δ) resulted in little to no acid production (Table 2-1).
Because the TCA cycle is required for electron transport chain functioning, we
addressed whether blockages in the electron transport chain also lead to the acid burst. As
seen in Table 2-1, mutations that inhibit the electron transport chain, other than SDH loss,
including deletions affecting ubiquinone, complex III, or complex IV, and rho− mutations
73
lacking mitochondrial DNA, did not rapidly acidify the medium. Thus, it is not a general
defect in mitochondrial function that leads to acid production in low glucose, but specific
disruptions in the TCA cycle.
3.3. The Acid Burst Results from Acetate Production by the Aldehyde Dehydrogenase
Ald4p
What is the source of the acid burst with low glucose? In the case of high glucose,
yeast acidification of the environment involves activation of a cell surface proton ATPase
Pma1p, as well as cellular export of organic acids [11-13]. As seen in Fig. 2-6A, cells
grown in low glucose have extremely low levels of Pma1p activity compared with cells
grown in high glucose. However, Pma1p activity remained low in the TCA cycle mutants
that produce acid (Fig. 2-6, B and C). We therefore examined effects of organic acids.
Previous studies have shown that Fe-S cluster assembly or delivery defects result in
extracellular acidification, marked by secretion of various organic acid mixtures [60-62]
and, in yeast, sdh3 and sdh4 mutants secrete succinate [62]. However, sdh mutants grown
under glucose starvation conditions exhibit only small increases in extracellular succinate
(Fig. 2-6D). The same is true with WT cells treated with paraquat (Fig. 2-6D). These
small increases in extracellular succinate are not likely to account for the large changes in
extracellular pH, and other acid producing cells (e.g. cit1Δ, aco1Δ, kdg1Δ, and fum1Δ)
do not elevate extracellular succinate (Fig. 2-6D). Another organic acid more highly
elevated in sdh mutants is acetate [63], which feeds into the TCA cycle via acetyl-CoA.
WT yeast cells grown in high glucose produce abundant acetate [13] but with glucose-
starved WT cells, extracellular acetate is virtually undetectable (Fig. 2-7A). Remarkably,
when glucose-starved WT cells are treated with paraquat, extracellular acetate levels rise
74
dramatically to levels comparable with that seen with high glucose (Fig. 2-7, A and B).
The high expulsion of acetate into the growth medium was also seen in sod2Δ cells
treated with low paraquat (Fig. 2-7B) and with all the acid producing TCA cycle mutants
(sdh2Δ, cit1Δ, aco1Δ, kgd1Δ, and fum1Δ) (Fig. 2-7D). In every case where cells make
acid with low glucose, there is ≥100-fold higher levels of extracellular acetate in the
growth medium than is seen with control cells not producing acid (Fig. 2-7, A, B, and D,
and Table 2-2). Intracellular acetate is also elevated, albeit not to the same degree as
extracellular acetate (Fig. 2-7, C and E). The high level of extracellular acetate can easily
account for the acid burst observed, as spiking the growth medium with levels of acetic
acid produced in paraquat-treated cells (0.276 μg/μl) was sufficient to lower the pH to
roughly the same degree (Table 2-2).
The acetate that normally enters the TCA cycle can be derived from the cytosolic
and/or mitochondrial pyruvate dehydrogenase bypass pathways or from the hydrolysis of
acetyl-CoA. In the pyruvate dehydrogenase bypass pathways, pyruvate is converted to
acetaldehyde, which in turn, is converted to acetate via the action of aldehyde
dehydrogenase (ALD) enzymes [64]. S. cerevisiae expresses 5 aldehyde dehydrogenase
isoforms [65] and we tested if mutants in any of these would prevent the acid burst. Of
these, deletion of the major mitochondrial isoform Ald4p most significantly inhibited
paraquat-induced acidification of the growth medium with low glucose (Fig. 2-8A). Loss
of Ald4p also attenuated the constitutive acidification of low glucose medium by sdh4Δ
mutants (Fig. 2-8B). Consistent with these reversals of acidification, we observed
corresponding decreases in acetate production when ald4Δ is disrupted in paraquat-
treated cells and in sdh4Δ mutants grown with low glucose (Fig. 2-7F). The effect
75
of ald4Δ mutations was also examined in cells grown under long-term glycerol
conditions. As seen in Fig. 2-8C, ald4Δ mutations prevented the early acid burst of sod2Δ
cells on glycerol. Moreover, loss of ALD4 in the background of SOD2+ cells completely
prevented the acid burst with long-term respiratory conditions, and the extracellular pH
continued to rise over time with ald4Δ mutants (Fig. 2-2). Under all cases studied, the
acid burst produced by cells under glucose starvation is dependent on mitochondrial
pyruvate dehydrogenase bypass enzyme, Ald4p.
3.4. Conditioning the Growth Medium by the Acid Burst
What is the significance of the Acid Burst? As one possibility, cells might
produce copious amounts of acetate to provide a carbon source for neighboring cells
under conditions of long-term carbon source starvation. To determine whether acetate
production and/or media acidification could enhance growth of yeast cells, we collected
“conditioned” growth media that was conditioned by cells grown 24 hrs in low glucose.
Low acetate/high pH medium was obtained from cultures of WT and ald4Δ cells, and
high acetate/low pH medium was obtained from cultures of sdh4Δ cells and from sod2Δ
cells treated with 50 μM paraquat, a low dose that is non-toxic to WT cells (Fig. 2-4C).
After removal of cells by filtration, these various conditioned media were used to
inoculate WT cells and growth was monitored over 10 hrs. As seen in Fig. 2-9A, the high
acetate/low pH media of sdh4Δ and paraquat-treated sod2Δ cells supported strong growth
of WT cells, whereas the low acetate/high pH media from WT cells or from ald4Δ
mutants was poorly effective at supporting growth. To determine whether acetate alone
accounts for these differences, we supplemented the low acetate medium from WT
and ald4Δ cells with levels of glacial acetic acid (0.20-0.22 μg/μl) equivalent to that in
76
cultures of sdh4Δ and paraquat-treated sod2Δ cells. This supplementation of acetic acid
was sufficient to eliminate differences in the various conditioned media, and all four
media types supported growth of WT cells (Fig. 2-9B). Thus, the production of acetate
can enhance growth of yeast cells under glucose starvation. This event triggered by
superoxide disruptions in the TCA cycle may enable the organism to withstand long-term
periods of starvation.
4. Discussion
Herein, we describe a mechanism by which the S. cerevisiae yeast modifies its
extracellular environment during prolonged nutrient deprivation. Under long-term
colonization on respiratory carbon sources, yeast cells shift from alkalization to
producing acid and we demonstrate here that this acid burst is dependent on the acetate-
producing Ald4p aldehyde dehydrogenase. Media conditioned by Ald4-acetate is indeed
able to promote yeast cell growth under glucose starvation conditions. Our findings are
consistent with a model in which the acid burst is produced in response to disruption of
the TCA cycle by superoxide (Fig. 2-9C).
Perhaps the most intriguing finding of this work is the apparent role of
mitochondrial matrix superoxide in conditioning the extracellular environment.
Mitochondrial superoxide is not likely to cross mitochondrial membranes in significant
quantities [30-33] and yet it can influence extracellular pH by modulating organic acid
production. In support of this effect of mitochondrial superoxide, we observed that the
acid burst under long-term glycerol conditions is accelerated by sod2Δ mutations lacking
mitochondrial matrix Mn-SOD, but not by sod1Δ mutations affecting the largely
77
cytosolic Cu/Zn-SOD1. Previously, the loss of Sod2p was shown to reduce volatile
ammonia production and prevent transition to the alkali phase under long-term glycerol
conditions but the mechanism was unknown [19]. This may be explained by the early
acid burst of sod2Δ cells which by lowering pH would trap a significant portion of
ammonia in the non-volatile ammonium form (pKa ≈ 9.25). Alternatively, Sod2p may
have separable roles in regulating extracellular pH through ammonia and acetate
production.
In further support of the role of superoxide in triggering the acid burst, we
observe that acid production under both glycerol and low glucose conditions is
dramatically accelerated when WT cells are treated with the redox cycler paraquat;
conversely, the acid burst with long-term glycerol conditions is eliminated by Mn-
antioxidants, chemical mimics of SOD enzymes that can remove superoxide in yeast [47-
53]. Superoxide, more specifically mitochondrial superoxide, represents an ideal
candidate to initiate the acid burst. Key Fe-S cluster enzymes in the TCA cycle (i.e. SDH
and aconitase) are sensitive to superoxide inactivation [24-26,57,58], and we show here
that disruption in the TCA cycle can cause build-up of acetate derived from
mitochondrial Ald4p. Although multiple reports have established a role for ROS in cell
signaling and regulation [66-69], the inactivation of TCA cycle enzymes by
mitochondrial superoxide was up until now considered a detrimental outcome of
oxidative damage. This is, perhaps, the first example where superoxide inactivation of
mitochondrial enzymes can be beneficial, particularly under the stress conditions of
nutrient deprivation.
78
Superoxide is a natural by-product of respiration, yet in WT cells the acid burst
only occurs after prolonged incubations under respiratory conditions (as in Fig. 2-2). The
products of superoxide damage appear to accumulate slowly in these cells, as we observe
a gradual loss in SDH activity over prolonged respiratory growth (Fig. 2-5B). In sod2Δ
cells the combined effects of this slow loss in SDH together with the already low
aconitase of these cells can explain the acceleration in the acid burst observed. Once SDH
and/or aconitase activity have been sufficiently inhibited to disrupt TCA cycle
functioning, the Ald4p acid burst occurs. In the simplest model, the blockage in the TCA
cycle itself prevents mitochondrial consumption of Ald4-acetate, and the accumulated
metabolite is expelled from the cell. Ald4 production of acetate might also be enhanced
in a futile attempt to correct the TCA cycle defect. It is noteworthy that blockages in 5 of
8 steps in the TCA cycle resulted in an acetate burst, including losses in SDH and
aconitase, as well as citrate dehydrogenase, α-ketoglutarate dehydrogenase, and
fumarase. It is possible that mutants for the remaining three steps (idh1Δ, mdh1Δ,
and lsc1Δ [70,71]) do not yield the same disruption in TCA cycle functioning.
Although high acetate has previously been associated with increased
chronological aging in yeast [8,9], these previous studies were conducted with fermenting
yeast cells with abundant glucose. In contrast, our studies with respiring yeast (as in Fig.
2-9) indicate that high acetate can be beneficial in environments with scarce nutrients.
Overall, these studies provide the first line of evidence for mitochondrial superoxide and
the TCA cycle in conditioning the extracellular environment under glucose starvation
conditions.
79
TABLE 2-1. Effects of TCA and ETC mutants on extracellular pH.
pH was measured as described in Materials and Methods after the indicated strains were
grown 24 hrs in 0.2% glucose growth medium. The average pH of triplicate samples is
denoted with standard deviation in parentheses. For WT + PQ, WT cells were grown in
media containing 800 μM paraquat. TCA = tricarboxylic acid cycle, ETC = electron
transport chain, mito = mitochondrial
ET
C m
utan
ts
T
CA
mut
ants
Sa
mpl
e D
escr
iptio
n pH
Sam
ple
Des
crip
tion
pH
No
cells
5.43
(± 0
.00)
No
cells
5.53
(± 0
.00)
W
T
6.90
(± 0
.03)
WT
7.
17 (±
0.0
5)
WT
+ PQ
4.67
(± 0
.03)
WT
+ PQ
4.59
(± 0
.03)
sd
h2∆
Com
plex
II
4.47
(± 0
.02)
sdh2∆
Succ
inat
e de
hydr
ogen
ase
4.54
(± 0
.05)
sd
h4∆
4.47
(± 0
.02)
sdh4∆
4.55
(± 0
.04)
co
q1∆
Coe
nzym
e Q
5.
19 (±
0.0
1)
ci
t1∆
Citr
ate
synt
hase
4.
54 (±
0.0
2)
cyt1∆
Com
plex
III
5.21
(± 0
.03)
aco1∆
Aco
nita
se
4.64
(± 0
.02)
co
x7∆
Com
plex
IV
5.20
(± 0
.03)
idh1∆
Isoc
itrat
e de
hydr
ogen
ase
5.46
(± 0
.10)
rh
o−
No
mito
DN
A
5.17
(± 0
.03)
kgd1∆
α-ke
togl
utar
ate
dehy
drog
enas
e 4.
72 (±
0.0
1)
lsc1∆
Succ
inyl
-CoA
liga
se
6.42
(± 0
.08)
fu
m1∆
Fu
mar
ase
4.71
(± 0
.01)
m
dh1∆
M
alat
e de
hydr
ogen
ase
7.14
(± 0
.07)
80
81
TABLE 2-2. Contribution of acetic acid to pH of growth medium.
Acetic acid and pH levels were measured as described in Materials and Methods in 0.1%
glucose growth medium (starting pH 5.55) that was cultured where indicated (WT cells)
for 24 hrs with WT yeast grown ± paraquat. Supplementation of medium with 0.25 µg/µl
acetic acid is sufficient to drop pH to levels seen in medium from paraquat treated WT
cells.
Sample Acetate (μg/μl) pH
WT cells undetectable 6.46 WT cells + 800 μM PQ 0.276 4.87 No cells 0.026 5.55 No cells + 0.25 μg/μl
glacial acetic acid 0.278 4.93
82
FIGURE 2-1. Yeast cells alkalize their environment under respiratory conditions.
A) “Giant colonies” [15] of WT BY4741 yeast cells were obtained by spotting 2x105
cells onto the indicated growth medium containing the pH indicator BCP and as carbon
source either 2% glucose or 3% glycerol. Where indicated, medium was supplemented
with 30 mM CaCl2. Cells were photographed on the second day of growth. (B,C) WT
yeast cells were inoculated at OD600 = 0.05 in a liquid media containing 0.2% glucose
and following specific time points, cells were removed and extracellular medium
analyzed for either glucose or pH. These data were provided by K.M. Laws. (B) Media
pH was determined following 24 hrs of growth using BCP and BCG indicator dyes.
Shown is a photograph of media color change with the pH indicators and the
corresponding pH value calculated as outlined in Materials and Methods. Numerical
results represent the averages of triplicate analyses where SD ≤0.03. “Cells: -” = growth
media alone. (C) Extracellular glucose was determined at the indicated time points as
described in Materials and Methods. Results represent the averages of triplicate samples
where error bars denote SD. Illustrated in 2-1A bottom and 2-1B right are the gradient of
color changes with BCP and BCG as pH ranges from acidic (BCP yellow; BCG green) to
basic (BCP purple; BCG blue).
83
84
FIGURE 2-2. Acid production following the alkaline phase of yeast cell growth is
accelerated by sod2∆ mutations and eliminated by ald4∆ mutations.
The indicated yeast strains were analyzed for acid and base production following the
designated times of incubation on glycerol containing plates (no calcium) as described in
Fig. 2-1A.
85
FIGURE 2-3. Role of mitochondrial SOD2 and Mn-antioxidants in preventing the
acid burst of respiratory cells.
Giant colonies of the indicated strains were plated under either respiratory, 3% glycerol
(A left, B-C), or fermenting, 2% glucose (A right), conditions as in Fig. 2-1A and
photographed on the designated days of incubation (part A, B) or on day 11 (sod2∆) or
day 16 (WT) (part C). (C) Where indicated, plates were supplemented with 0.5 mM
MnCl2, a concentration that is non-toxic but sufficient to maximize intracellular levels of
Mn-antioxidants that remove superoxide [47-53]. Data in parts A and B were provided
by K.M. Laws.
86
FIGURE 2-4. The redox cycler paraquat and sdh∆ induce a rapid acid burst under
respiratory conditions and are not additive.
The indicated yeast strains were grown either on glycerol containing medium for 2 days
(A), or in low glucose liquid medium for 24 hrs (B, C, D) as in Fig. 2-1. Where
indicated, the growth medium was supplemented with the designated concentrations of
paraquat (PQ). (B, C, open triangles and D) pH was calculated in triplicate as in Fig. 2-
1B where values in parentheses (B,D) denote SD. (C) Cell growth (black circles) was
measured turbidimetrically at an optical density of 600 nm (OD600) and plotted as a
function of control cell growth with no paraquat.
87
88
FIGURE 2-5. Activities of mitochondrial Fe-S cluster enzymes in acid producing
cells.
The designated yeast strains were grown for 24 hrs in 0.2% glucose medium that was
supplemented with either 800 µM (WT) or 50 µM (sod2∆) paraquat. (A) Extracellular
pH shown in left column was measured in triplicate samples as in Fig. 2-1B, and whole
cell lysates were prepared and assayed for aconitase and SDH activity as described in
Materials and Methods. Units are represented as either nmol cis-aconitate converted /
mg protein /min (aconitase), or nmol DCIP reduced / mg protein / min (SDH); values
represent the average of triplicate samples; error bars = SD. (B) The indicated
intracellular metabolites were measured by gas chromatography mass spectrometry as
described in Materials and Methods. Values represent averages of duplicate samples;
error bars = SD.
89
90
FIGURE 2-6. Acid production with low glucose does not correlate with activation of
Pma1p, the proton ATPase.
The indicated yeast strains were grown in either 2% (A) or 0.2% (A-C) glucose to mid
log phase, and Pma1p activity was measured in membranes by orthovanadate-sensitive
ATPase activity as described in Materials and Methods. Values are shown normalized to
WT cells in 0.2% glucose, and represent the average of duplicate measurements; error
bars = SD. (C) “Base” and “Acid” reflects the corresponding effect of the yeast strain on
extracellular medium in 0.2% glucose.
91
FIGURE 2-7. Cells produce high levels of extracellular acetate during the acid
burst.
Extracellular acetate/acetic acid was measured in the growth medium of WT cells (A) or
the indicated strains grown in either 2% (A) or 0.2% (A-C) glucose for 24 hrs as
described in Materials and Methods. Asterisks donate the particular conditions and
strains in which cells acidify the growth medium. “Media” = levels of acetate in the
starting growth medium with no cells. “+PQ” = either 50 µM (sod2∆ cells) or 800 µM
(all other strains) paraquat added during growth. Results represent the averages of
triplicate samples; error bars = SD.
92
93
FIGURE 2-8. The mitochondrial Ald4p aldehyde dehydrogenase is required for the
acid burst.
The indicated strains were grown either in low glucose liquid medium (A and B) or on
glycerol containing plates (C) and pH monitored as in Fig. 2-1. (A, B) pH values
represent the results of duplicate (A) or triplicate (B) samples where SD ≤0.12. +PQ =
cultures supplemented with 800 µM paraquat.
94
FIGURE 2-9. Medium conditioned during the acid burst can promote yeast growth.
Growth medium was collected from the indicated cultures following 24 hrs incubation in
0.2% glucose medium. This cell-free “conditioned medium” was used to inoculate WT
cultures of yeast as described in Materials and Methods and growth monitored
turbidimetrically at 600 nm. Results represent the averages of triplicates cultures; error
bars = SD. In some cases, the error was too small to be seen by error bars. (A)
Conditioned medium was used as isolated from cells. (B) The levels of acetic
acid/acetate in all four types of conditioned medium were normalized by supplementing
0.20-0.22 μg/μl glacial acetic acid to the medium conditioned by WT and ald4∆ cells.
(C) A model for how mitochondrial superoxide produced during long-term respiratory
conditions can lead to the acid burst and conditioning of the extracellular environment.
Superoxide can damage the Fe-S enzymes aconitase and SDH in the mitochondrial
matrix and the concomitant loss of TCA cycle activity induces massive production of
acetate from the mitochondrial Ald4p aldehyde dehydrogenase. The high levels of
acetate expelled into the growth medium conditions the extracellular environment to
promote growth of neighboring yeast cells.
95
96
5. Reference List
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107
CHAPTER 3
Oxidative Stress and Lethality in S. cerevisiae with
Mutations in SOD1 and the Proton ATPase PMA1
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1. Introduction
The plasma membrane H+-ATPase is found in species throughout the fungal and
plant kingdoms and regulates intracellular pH and the membrane electrochemical
potential required for secondary nutrient uptake [1]. In the yeast Saccharomyces
cerevisiae, the major plasma membrane H+-ATPase, Pma1p, is essential for life and its
activity correlates directly with growth [2,3]. Furthermore, Pma1p activity significantly
affects viability under a variety of stress conditions [4], making it a key regulator of cell
survival and growth.
Pma1p itself is regulated by a variety of conditions with the most notable and well
studied being glucose availability [5-8]. In response to glucose, Pma1p is both
transcriptionally up-regulated [3,9] and post-translationally activated [8]. However,
given the high expression of Pma1p even in the absence of glucose [3] and its long half-
life [10,11], post-translational regulation plays a more significant role in modulating its
activity. This regulation depends primarily on an alpha-helix in the C-terminus of the
protein which binds to the ATPase domain and inhibits Pma1p activity in the absence of
glucose [12-14]. The addition of glucose stimulates phosphorylation of three residues
(Ser-899, Ser-911, Thr-912) in the C-terminal inhibitory domain triggering its release
from the ATPase domain coincident with an increase in the Vmax for ATP hydrolysis and
decrease in the Km for ATP [14-17]. Analysis of mutations at these residues, either to
Ala to prevent, or Asp to mimic phosphorylation, has suggested a unique role for each
phosphorylation site in glucose-induced Pma1p activation (see Table 3-1).
Recently our lab demonstrated that glucose activation of S. cerevisiae Pma1p is
inhibited by mutations in SOD1 encoding the largely cytoplasmic Cu/Zn superoxide
109
dismutase [18]. The mechanism was not fully understood. Sod1p catalyzes the
disproportionation of superoxide to hydrogen peroxide and molecular oxygen and is thus
best known for its role in cellular antioxidant defense [19-22]. However, Sod1p can also
function in cell signaling independent of its role in oxidative stress protection. For
example, in S. cerevisiae the peroxide produced by Sod1p can stabilize the casein kinase
I homologue needed for glucose signaling, Yck1p [18]. Pma1p is one known target of
Yck1p [18,23,24], suggesting that the aforementioned effect of sod1∆ on Pma1p may
involve loss of Yck1p. However, we could not exclude the possible contribution of
oxidative stress protection in Sod1p control of Pma1p.
The connection between Sod1p and Pma1p was explored further in the current
study using mutants of Pma1p known to disrupt glucose control and/or ATPase activity.
These Pma1p mutations targeted the phosphorylation sites at the C-terminus, as well as,
potential interacting residues in the ATPase domain. Out of 14 Pma1p alleles examined,
one variant exhibited a striking “synthetic” interaction [25,26] with sod1∆. Specifically,
the T912D substitution in the C-terminal inhibitory domain of Pma1p was synthetically
lethal with the sod1∆ null mutation, i.e. the double mutant was not viable in air. This
synthetic lethality was reversed by second-site suppressor mutations in Pma1p that
hyperactivate the ATPase, indicating the lethality results from profound loss in activity of
the essential Pma1p proton ATPase. The synthetic lethality was also reversed under
anaerobic conditions and by antioxidant treatments that reverse oxidative damage of
sod1∆ strains but do not rescue the sod1∆ loss of Yck1p. Together, our data support a
model in which Sod1p maintains Pma1p activity through both oxidative stress-dependent
and -independent (Yck1p signaling) pathways.
110
2. Materials and Methods
2.1. Yeast Growth, Yeast Strains and Plasmids
S. cerevisiae yeast strains were maintained at 30°C either in an enriched YPD
(yeast extract, peptone, 2% dextrose) or a minimal synthetic complete (SC) medium
devoid of lysine where indicated. Anaerobic growth was carried out using the GasPak
EZ Anaerobe Container System (Becton Dickinson) in medium supplemented with 15
mg/L ergosterol and 0.5% Tween 80 (YPDE). For plate-based growth tests, 104, 103, and
102 cells were spotted onto the designated medium and growth proceeded at 30°C for
three to five days. For liquid growth assays, 106 cells/ml were inoculated into YPD with
or without the concentrations of drug indicated (Fig. 3-6A and B) and grown shaking at
30°C for 18 hrs (paraquat) or 15 hrs (menadione) after which cell growth was determined
turbidimetrically at 600 nm using a spectrophotometer.
The S. cerevisiae strains used in this study are either of the BY4741 (MATa
his3∆1 leu2∆0 met15∆0 ura3∆0) or NY13 (MATa ura3–52) backgrounds and can be
found in Tables 3-2 and 3-3. Pma1p mutants in the NY13 background were generous
gifts of Ken Allen and Carolyn Slayman and where indicated, sod1∆ mutations were
introduced using the sod1::KanMX4 plasmid, pJAB002, as described [27]. The Pma1-
T912D sod1∆ strain, JAB020, is only viable under anaerobic conditions, but aerobically
viable suppressors with intragenic mutations in PMA1 spontaneously appeared during
yeast transformation procedures (e.g., A906G, strain JAB142) or could be isolated at a
frequency of 105 by plating 107 cells for two days in air on YPD media (e.g., T540A,
strain JAB124; L586V, strain JAB125 and F666L, strain JAB126). Site specific single or
double mutations in PMA1 were introduced in BY4741 or the isogenic sod∆1::KanMX or
111
akr1∆::KanMX (Open Biosystems) strains by gene replacement using derivatives of the
URA3 vector pGW201 vector expressing specific alleles of PMA1. To create the Pma1-
T912D/A906G sod1∆ strain, JAB140, the sod1::LEU2 plasmid, pKS1, was introduced as
described previously [28] into strain JC20. It is noteworthy that in the BY4741
background, the single Pma1-T912A (JC02) and Pma1-T912D (JC03) strains developed
rho- mutations at a very high frequency, but not in the background of the isogenic sod1∆
or akr1∆ strains. Tests for rho- mutations were conducted by plating cells on YP
medium containing 3% glycerol rather than 2% glucose and incubating at 5% oxygen that
is sufficient to drive mitochondrial respiration but is not lethal to the oxygen-sensitive
T912D sod1∆ strain.
The plasmids used in this study are outlined in Table 3-4. The vectors for
generating site-specific substitutions in PMA1 were all from the phagemid pGW201 and
contained PMA1 mutations introduced by Quikchange site-directed mutagenesis
(Stratagene). pGW201 [29] contains a PMA1 cassette harboring URA3 at the 3′ non-
coding end that is liberated by HindIII digestion. After transformation into yeast,
successful introduction of the desired pma1 allele was verified by sequencing of the C-
terminal PMA1 coding region. The sod1:LEU2 and sod1::KanMX vectors were as
previously described [27,28].
2.2. Biochemical and Microscopy Analyses
For western blots of Pma1p, cells were grown at 30°C in YPD to mid-log phase.
Cells were harvested, washed in water, and resuspended in buffer comprised of 20 mM
potassium phosphate, 1.2 M sorbitol, 1% Triton X-100, 1.0 mM PMSF, 1:100 protease
inhibitor cocktail (P8340, Sigma), pH 7.4. Cells were lysed in the presence of zirconium
112
oxide beads by vortexing twice for 2 mins at 50 Hz in a TissueLyzer LT (Qiagen).
Lysates were clarified by centrifugation and 20 μg total protein was then prepared and
loaded onto NuPAGE Novex 4-12% Bis-Tris Gels (Life Technologies) according to the
manufacturer’s instructions, except lysates were incubated at room temperature for 15
mins. Gel electrophoresis was carried out in MOPS buffer according to the
manufacturer’s instructions and then transferred to a nitrocellulose membrane with the
iBlot Gel Transfer Device (Life Technologies) using the standard settings. Membranes
were probed with anti-Pma1p primary (1:5000, Abcam, ab4645) and Alexa Fluor 680
Donkey Anti-Mouse IgG secondary (1:5,000, Life Technologies, A10038).
For Pma1p activity assays, a culture of 4 x 106 cells was seeded into 1L YPD and
grown to mid-log phase at 30°C. Cells were washed once with water and pellets stored at
-80°C until ready for membrane isolation. Plasma membranes were isolated essentially
as described [18,30,31]. Briefly, pellets were resuspended in 25 ml ice cold
homogenization buffer (50 mM Tris-HCl, pH 7.5, 0.3 M sucrose, 5 mM EDTA, 1 mM
EGTA, 5 mg/ml BSA, 2 mM DTT, 1 mM PMSF, 1:100 protease inhibitor cocktail) and
passed twice through a French press at ≈ 20,000 psi for lysis. Lysates were clarified by
centrifugation at 3,500×g for 5 mins and then at 14,000×g for 10 mins. Membranes were
then pelleted by centrifugation at 200,000×g, washed once by dounce homogenization in
a wash buffer containing 10 mM Tris-HCl, pH 7.0, 1 mM EGTA, 10% glycerol, 1 mM
PMSF, 1:100 protease inhibitor cocktail and harvested again by centrifugation as above.
Membranes were resuspended in 1 ml wash buffer by dounce homogenization and stored
at -80°C. After thawing, protein concentrations were estimated using the Bradford assay
and Pma1p activity was determined by the orthovanadate-sensitive phosphate release
113
from ATP following published protocols [18,31,32]. Basically, 10-50 μg purified
membrane protein in 20 μl membrane wash buffer was added to 120 μl Pma1p activity
buffer (50 mM MES-Tris, pH 6.0, 15 mM MgSO4, 50 mM KNO3, 0.2 mM ammonium
molybdate, 5 mM NaN3, 3 mM ATP) with or without 100 μM orthovanadate and
incubated for 10 mins at room temperature in a 96-well plate. The reaction was stopped
by the addition of 140 μl of the developing reagent containing 1% SDS, 100 mM sodium
molybdate, 0.6 M H2SO4, and 0.8% ascorbate. The A620 was recorded within 5 mins after
addition of the developing reagent and activity was subsequently normalized to WT.
Indirect immunofluorescence was performed largely as described [33,34]. Briefly,
cells were grown in YPD to OD600 = 1.0 and then fixed with 4.4% formaldehyde added
directly to the shaking culture for 30 mins. The cells were then washed once with 100
mM potassium phosphate, pH 6.5 (KPhos buffer) and then fixed a second time overnight
in KPhos buffer with 4% formaldehyde. Cells were washed once in KPhos buffer and
spheroplasts were generated by incubation in 200 mM Tris-HCl, pH 8.0, 20 mM EDTA,
1% 2-mercaptoethanol for 10 mins, followed by a second incubation in 1.2 M sorbitol,
100 mM potassium phosphate, pH 6.5 with 1.5 mg/ml Zymolyase 20T for 30-60 mins.
Cells were washed once in 1.2 M sorbitol and permeabilized in 1.2 M sorbitol, 2% SDS
for 2 mins. Cells were then washed twice in 1.2 M sorbitol, allowed to adhere to poly-
lysine coated slides to form a monolayer, and then treated with primary (1:25 anti-Pma1p
mouse monoclonal [40B7]; Abcam, ab4645) and secondary antibodies (1:250 Goat Anti-
Mouse Alexa Fluor 488; Molecular Probes, A11001) in PBS with 1% BSA. After final
aspiration, mounting solution containing 2.5% DABCO, 100 mM Tris HCl, pH8.8, 50%
Glycerol, 0.2 μg/ml DAPI was added and slides sealed. Images were taken on a Zeiss
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Observer.Z1 fluorescence microscope with an Apotome VH optical sectioning grid
(Zeiss, Gena, Germany) under 100X magnification.
3. Results
3.1. Pma1p Mutants and Sod1p Deficiency
We previously reported that Pma1p activity is low in S. cerevisiae sod1∆ mutants
based on assays of isolated membranes [18]. One possible mechanism might involve
mis-sorting of Pma1p in the secretory pathway, as has been shown for yeast mutants
defective in vacuolar function [33]. Since S. cerevisiae sod1∆ cells have abnormal
vacuoles [35], we tested whether loss in Pma1p activity might reflect Pma1p mis-sorting.
By indirect immunofluorescence microscopy, Pma1p in WT cells is predominantly
localized to the cell-surface as seen in Fig. 3-1A. This precise pattern of Pma1p
localization was mirrored in sod1∆ cells. Furthermore, the intensity of Pma1p staining
was comparable in WT and sod1∆ cells (Fig. 3-1A), consistent with normal Pma1p
polypeptide levels in sod1∆ cells [18]. Thus, sod1∆ mutations do not alter Pma1p protein
levels or localization. Pma1p ATPase activity at the cell surface must be affected.
Pma1p activity at the cell surface is normally controlled by glucose and we sought
to determine the effects of sod1∆ mutations when such glucose control of Pma1p has
been altered. To this end, we utilized mutant alleles of Pma1p known to affect glucose
control and activity of the ATPase. These alleles target the C-terminal inhibitory and
interacting ATPase domains (Fig. 3-1B) and include Ala and phospho-mimic Asp
substitutions at the phosphorylation sites S899, S911, and T912, and Cys scanning
substitutions in stalk segment 5 that increase or decrease Pma1p activity as summarized
115
in Table 3-1 [36,37]. Strains expressing these various Pma1p alleles in combination with
sod1∆ were initially created under anaerobic conditions to minimize sod1∆ oxidative
damage. When tested for growth in atmospheric oxygen, we were surprised to find a
strong “synthetic” effect [25,26] when sod1∆ mutations were combined with a specific
allele of Pma1p. Specifically, the Pma1-T912D mutant was not viable when combined
with sod1∆ mutations (Figs. 3-1C and D). The results were reproducible in two distinct
strain backgrounds and the synthetic lethality was only seen with oxygen, as these same
strains grew well under anaerobic conditions (Figs. 3-1C and D). For the purposes of this
study, this synthetic lethal mutant will be referred to as T912D sod1∆.
The lethal effect of combining sod1∆ and Pma1p mutations appeared highly
specific to T912D sod1∆. As seen in Fig. 3-1C and Fig. 3-2, 13 out of 14 Pma1p mutants
in the C-terminal inhibitory and ATPase domain showed no evidence of a synthetic effect
with sod1∆, i.e., there was no obvious changes in cell growth of the double mutant
compared to the single Pma1p or sod1∆ mutant. Even Pma1-T912A showed no
inhibitory effect when combined with sod1∆; the lethality appeared unique to Pma1-
T912D (Fig. 3-1C). In the course of our studies, we noted that single T912D mutants of
Pma1p have a high tendency to accumulate rho- mutations, or losses of mitochondrial
DNA, particularly in the BY4741 strain background (see Materials and Methods).
However, T912A Pma1p mutants also have a high tendency to go rho- but are not lethal
with sod1∆, and more importantly, all of our T912D sod1∆ mutants are RHO+ (see
Materials and Methods). Thus the synthetic lethality of T912D sod1∆ cannot be
attributed to loss of mitochondrial DNA and other mechanisms must be involved.
3.2. Suppressors of the T912D sod1Δ Aerobic Lethality
116
To help understand the basis for the T912D sod1Δ lethality, we used a genetic
suppressor approach. After prolonged incubation in air, viable colonies of T912D sod1Δ
could be identified at a frequency of ≈105. When isolated and retested, those colonies
retained full aerobic growth (Fig. 3-3A, representative sample), indicating stable genetic
suppression of either Sod1- or Pma1-deficiency. To decipher between these two, we
tested for aerobic lysine auxotrophy, a hallmark of Sod1p deficiency [38]. Mutants of
sod1∆ cannot grow aerobically without lysine due to oxidative damage to the lysine
biosynthetic pathway, and previously characterized genetic suppressors of sod1∆ (e.g.,
pmr1∆, bsd2∆) correct this lysine defect [39-44]. However, none of the suppressors of
T912D sod1∆ could grow aerobically without lysine (Fig. 3-3B), suggesting suppression
reversed the deficiency of Pma1p, not Sod1p. To test for intragenic suppressors in
Pma1p, the entire PMA1 gene was sequenced in four representative suppressors and each
was found to have a distinct additional missense mutation within the PMA1 coding region
(Fig. 3-1B). F666L lies within the aforementioned Pma1p stalk region and in fact F666C
was previously shown by Miranda and colleagues to result in constitutive activation of
Pma1p [36]. The T540A, L586V mutations lie within the ATPase domain and A906G is
within the C-terminal inhibitory domain (Fig. 3-1B). These findings are highly
reminiscent of intragenic suppressors previously reported for the Pma1-S911A/T912A
double mutant. Like T912D sod1∆, the double Pma1-S911A/T912A is a synthetic lethal
mutant, and this lethality is rescued by mutations in the Pma1p ATPase domain (P536L,
A565T, G587N, G648S), the stalk domain (P669L, G670S) and the C-terminus (M907I)
[16,17]. M907I is in close juxtaposition to the A906G suppressor identified here and was
117
reported by Eraso and Portillo [17] to be a hyperactive Pma1p allele. We therefore chose
Pma1-A906G for further analysis.
To confirm that Pma1-A906G by itself suppresses T912D sod1∆, we introduced
this identical mutation into an independent strain background. As seen in Fig. 3-3C,
A906G was capable of restoring aerobic growth to the T912D sod1∆ strain in the
BY4741 background and as before (Fig. 3-3B), this reflects specific suppression of
Pma1p and not Sod1p deficiency, since the strain remains incapable of aerobic growth
without lysine (Fig. 3-3C). To explore how A906G suppresses T912D, we examined the
polypeptide levels, cellular localization and activity of Pma1-T912D with and without the
A906G suppressor. As seen in Fig. 3-4, the A906G mutation significantly enhances
Pma1p ATPase activity without changing Pma1p polypeptide levels or its localization at
the cell surface. In fact, activity with the Pma1-T912D/A906G double mutant is even
higher than that of WT Pma1p (Fig. 3-4C). A906G behaves as a hyperactive allele of
Pma1p, as has been reported for its close neighbor M907I [17]. Thus, the synthetic
lethality of T912D sod1∆ can be rescued by hyperactivation of Pma1p. These findings
strongly implicate profound loss of essential Pma1p activity as the cause of death in the
T912D sod1∆ strain.
3.3. Oxidative Damage, Not Loss of Yck1p Signaling Causes the Synthetic Lethality of
T912D sod1Δ
Our previous studies suggested that inhibition of Pma1p activity in sod1∆ cells
may reflect defective signaling by casein kinases. Sod1p stabilizes the yeast casein
kinase I homologue, Yck1p, which in turn regulates Pma1p [18,23,24]. Activity of
Yck1p at the plasma membrane can be abolished by deleting the palmitoyl transferase
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akr1∆ [45,46], and like sod1∆ mutations, akr1∆ mutants have low Pma1p activity [18].
We therefore predicted that akr1∆ mutations would phenocopy sod1∆ mutations and
show synthetic lethality with Pma1-T912D. However, in contrast to T912D sod1Δ, the
T912D akr1Δ double mutant was quite viable under all conditions tested (Fig. 3-5A).
This unexpected viability did not reflect rapid accumulation of genetic suppressors in this
strain. There were no second-site mutations in PMA1, as determined by complete
sequence analysis of the gene, and there was no obvious suppression of akr1∆, as the
large cell size and elongated bud morphology of akr1∆ remained intact (data not shown)
[46]. Since akr1∆ does not phenocopy sod1∆ when combined with Pma1-T912D, the
synthetic lethality of T912D sod1Δ may not be due to loss of Yck1p activation.
Aside from its role in regulating Yck1p, S. cerevisiae Sod1p is also important for
oxidative stress protection [47-49] and the oxidative damage of sod1∆ cells can be
reversed by growing cells under anaerobic conditions or by supplying high levels of non-
SOD antioxidants, such as reactive manganese complexes that can scavenge superoxide
[50-55]. Neither of these treatments will correct the Yck1p-defect of sod1∆ cells which
is independent of oxidative stress [18]. The synthetic lethality of T912D sod1∆ strains is
completely reversed by anaerobic conditions (Figs. 3-1C, 3-1D, and 3-3A), strongly
indicative of oxidative stress. The strain begins to grow when oxygen levels drop to 5%
(Fig. 3-5B). Furthermore, the synthetic lethality of T912D sod1 could be reversed by
treatment with high levels of manganese, which form non-proteinaceous antioxidants
(Fig. 3-5C) [41,50-55]. Together, these observations strongly indicate that oxidative
damage from sod1∆, rather than loss of Yck1p casein kinase signaling, contributes to the
synthetic lethality of T912D sod1∆. The combined effect of this damage together with
119
Pma1-T912D must sufficiently inhibit essential Pma1p activity to halt S. cerevisiae
growth.
4. Discussion
The cytosolic Cu/Zn-SOD enzyme can broadly impact cell metabolism and
growth. On one hand, Sod1p eliminates superoxide that can cause oxidative damage to
biomolecules, while on the other hand, Sod1p generation of peroxide can act in signaling
pathways independent of oxidative stress protection [18,56]. It was previously shown
that in S. cerevisiae, Sod1p can signal glucose repression through a mechanism involving
peroxide stabilization of Yck1p, and one downstream target of this signal is the proton
ATPase Pma1p [18]. We now provide evidence that Sod1-control of Pma1p extends
beyond Yck1p. Specifically, we find that through certain oxidative damage effects,
sod1∆ mutations are lethal to cells expressing Pma1p with a T912D mutation in the C-
terminal inhibitory domain. This synthetic lethality must be independent of Yck1p, as
Pma1-T912D is without consequence when Yck1p is inactivated in SOD1+ strains (e.g.,
by akr1∆ mutations). Furthermore, the T912D sod1∆ lethality is reversed by anti-oxidant
treatments that do not rescue the sod1∆ defect in Yck1p. We propose that Sod1p helps to
maintain Pma1p in the active state in two distinct ways: one dependent on Yck1p-
glucose signaling and the other via protection against oxidative damage.
Why are T912D sod1∆ mutants inviable in air? Since lethality is reversed by
hyper-activating mutations in Pma1p (e.g., A906G), death likely results from profound
inhibition of the essential proton ATPase Pma1p. As an alternative, lethality might
reflect severe oxidative damage to the cell from the combined effects of sod1∆ and
120
T912D PMA1 mutations. However, we have no evidence that Pma1-T912D mutations by
themselves cause oxidative damage. The single Pma1-T912D mutants are not more
sensitive to the environmental oxidant paraquat or to conditions of hyperoxia that are
lethal to sod1∆ strains (Fig. 3-6A,C). We did observe a slight sensitivity towards the
oxidant menadione compared to WT Pma1p strains, but this effect was orders of
magnitude diminished compared to the menadione sensitivity of sod1∆ strains (Fig. 3-
6B). We therefore favor a model in which profound inhibition of the essential Pma1p
ATPase is the cause of the aerobic lethality of T912D sod1∆ strains.
Of all the Pma1p mutants tested, only T912D showed synthetic lethality with
sod1∆; even T912A was without effect. In some studies, the T912D mutation has been
reported to inhibit the ATPase but the defect in Pma1p caused by T912A was more
severe [14,17] (Table 3-1). Thus the allele specificity for synthetic lethality cannot be
explained simply by T912D inhibition of the ATPase. Instead, effects on protein
conformation may be key. Based on trypsin digestion experiments by Lecchi and
colleagues [14], Pma1-T912D is predicted to adopt a more open conformation compared
to Pma1-WT or -T912A (summarized in Table 3-1). WT Pma1p has been reported to be
relatively resistant to oxidative damage compared to other P-type ATPases [57], but the
conformation of Pma1-T912D may render this transporter hypersensitive to such insults.
Pma1p has nine cysteine residues including five in the catalytic domain and modification
of these can inhibit the ATPase [58]. It will be of interest to study the effect of T912D
and sod1∆ mutations on the oxidation status of these cysteines as a potential target for
oxidative damage.
121
While direct oxidative damage of Pma1p is one possibility, it is also possible that
sod1∆-oxidative stress can affect Pma1p indirectly by damaging one of the Pma1p
regulatory proteins. Pma1p regulation is complex and numerous factors have been
implicated in its regulation, including those for ubiquitination and protein degradation
(RSP5, UBC4, proteasome) [59], glucose sensing (SNF3, GPA2) [60], phosphorylation
(Yck1p, Ptk2, Hxk1, Hxk2, Glk1, etc.) [18,23,24,61] and calcium signaling through
calcineurin [62]. In this regard, Sod1p has been shown to bind to and protect calcineurin
from inactivation [63]. Regardless of whether Sod1p is protecting Pma1p directly or
indirectly, these studies underscore the multiple ways this antioxidant enzyme can affect
growth and metabolism of aerobic cells.
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TABLE 3-1. Effect of Pma1 mutations on activity and C-terminal conformation
The effect of mutations used in this study on Pma1p activity and C-terminal
conformation was compiled from the literature cited. For Pma1p activity, “↑” indicates
increased activity, “↓” decreased activity, and “↓↓” severely decreased activity relative to
Pma1-WT in the same study. The C-terminal conformation of Pma1p mutants is based
on the ability of trypsin to cleave off the C-terminus of Pma1p, as performed by Lecchi
and colleagues in the presence of glucose [14]). Here, “closed” and “open” refer to
constitutive states unaffected by glucose availability, whereas “≈ WT” indicates the
mutations retain glucose-responsive conformational changes. N.D. = not determined
Pma1 Mutant Activity* C-terminal Conformation References
3CHT ≈ WT N.D. [36] V665C ↓ N.D. [36] A668C ↑ N.D. [36] D676C ↓ N.D. [36] L678C ↓ N.D. [36]
I684C ↑ ATPase, ↓ H+ transport N.D. [36]
H686C ↓ N.D. [36] R687C ↓ N.D. [36] Y689C ↑ N.D. [36] S899A ≈ WT or ↓ ≈ WT [14,17] S899D ≈ WT or ↑ ≈ WT [14,17] S911A ≈ WT N.D. [16] S911D N.D. N.D. - T912A ↓↓ closed [14,16,17,64] T912D ≈ WT or ↓ open [14,17]
* Primarily refers to ATPase activity but may also reflect H+-pumping activity where
measured. Unless otherwise noted, the identified mutation affected ATPase and H+-
pumping activity in a similar manner.
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TABLE 3-2. BY4741 strains used in this study
Strain Parent Strain Partial Genotype Plasmid Reference
BY4741 - MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 - [65] sod1Δ BY4741 sod1::kanMX - [66] akr1Δ BY4741 akr1::kanMX - [66] JC02 BY4741 PMA1-T912A (URA3), rho- pJC001 This study JC03 BY4741 PMA1-T912D (URA3), rho- pJC002 This study JC04 BY4741 PMA1-S911A (URA3) pJC003 This study JC05 BY4741 PMA1-S911D (URA3) pJC004 This study JC06 BY4741 PMA1-S899A (URA3) pJC005 This study JC07 BY4741 PMA1-S899D (URA3) pJC006 This study JC08 BY4741 PMA1-A906G (URA3) pJC007 This study JC09 sod1Δ sod1::kanMX, PMA1-T912A (URA3) pJC001 This study JC10 sod1Δ sod1::kanMX, PMA1-T912D (URA3) pJC002 This study JC11 sod1Δ sod1::kanMX, PMA1-S911A (URA3) pJC003 This study JC12 sod1Δ sod1::kanMX, PMA1-S911D (URA3) pJC004 This study JC13 sod1Δ sod1::kanMX, PMA1-S899A (URA3) pJC005 This study JC14 sod1Δ sod1::kanMX, PMA1-S899D (URA3) pJC006 This study JC15 sod1Δ s sod1::kanMX, PMA1-A906G (URA3) pJC007 This study JC20 BY4741 PMA1-T912D/A906G (URA3) pJAB008 This study JC61 BY4741 akr1::kanMX, PMA1-T912D (URA3) pJC002 This study JAB140 JC20 PMA1-T912D/A906G (URA3), sod1::LEU2 pKS1 This study
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TABLE 3-3. NY13 strains used in this study
Strain Parent Strain Partial Genotype Plasmid Reference PMA1-WT NY13 PMA1-WT (URA3) - [14] PMA1-T912D NY13 PMA1-T912D (URA3) - [14] PMA1-3CHT NY13 10His-PMA1-3C (lacks all Cys
residues except C376, C409, & C472) - [37]
PMA1-V665C NY13 PMA1-V665C (URA3) - [36] PMA1-A668C NY13 PMA1-A668C (URA3) - [36] PMA1-D676C NY13 PMA1-D676C (URA3) - [36] PMA1-L678C NY13 PMA1-L678C (URA3) - [36] PMA1-I684C NY13 PMA1-I684C (URA3) - [36] PMA1-H686C NY13 PMA1-H686C (URA3) - [36] PMA1-R687C NY13 PMA1-R687C (URA3) - [36] PMA1-Y689C NY13 PMA1-Y689C (URA3) - [36] JAB020-1 PMA1-T912D PMA1-T912D (URA3), sod1::kanMX pJAB002 This study JAB021 PMA1-WT PMA1-WT (URA3), sod1::kanMX pJAB002 This study JAB124 JAB020-1 PMA1-T912D/T540A (URA3),
sod1::kanMX - This study
JAB125 JAB020-1 PMA1-T912D/L586V (URA3), sod1::kanMX
- This study
JAB126 JAB020-1 PMA1-T912D/F666L (URA3), sod1::kanMX
- This study
JAB142 PMA1-T912D PMA1-T912D/A906G (URA3), sod1::kanMX
- This study
JC22 PMA1-3CHT PMA1-3CHT, sod1::kanMX pJAB002 This study JC23 PMA1-L678C PMA1-L678C, sod1::kanMX pJAB002 This study JC24 PMA1-Y689C PMA1-Y689C, sod1::kanMX pJAB002 This study JC25 PMA1-A668C PMA1-A668C, sod1::kanMX pJAB002 This study JC27 PMA1-D676C PMA1-D676C, sod1::kanMX pJAB002 This study JC28 PMA1-H686C PMA1-H686C, sod1::kanMX pJAB002 This study JC29 PMA1-I684C PMA1-I684C, sod1::kanMX pJAB002 This study JC30 PMA1-V665C PMA1-V665C, sod1::kanMX pJAB002 This study JC31 PMA1-R687C PMA1-R687C, sod1::kanMX pJAB002 This study
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TABLE 3-4. Plasmids used in this study
Plasmid Description Digest* Reference pGW201 Pma1-WT (URA3) HindIII [29] pJAB002 sod1::kanMX BamHI [27] pJAB008 Pma1-T912D/A906G (URA3) HindIII This Study pJC001 Pma1-T912A (URA3) HindIII This Study pJC002 Pma1-T912D (URA3) HindIII This Study pJC003 Pma1-S911A (URA3) HindIII This Study pJC004 Pma1-S911D (URA3) HindIII This Study pJC005 Pma1-S899A (URA3) HindIII This Study pJC006 Pma1-S899D (URA3) HindIII This Study pJC007 Pma1-A906G (URA3) HindIII This Study pKS1 sod1::LEU2 PstI [28]
*Digest indicates the restriction enzyme used to linearize the cassette prior to
transformation to ensure proper integration.
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FIGURE 3-1. Effect of sod1∆ on Pma1p localization and growth of Pma1p mutants
with altered glucose regulation
(A) Indirect immunofluorescence of endogenous Pma1p was performed on WT and
sod1∆ cells in the BY4741 background as described in Materials and Methods. (B)
Topology map of Pma1p based on a map previously published by Lecchi, et al. [14] with
circles representing individual amino acids. The N- and C-termini are designated by “N”
and “C”, respectively. Red circles indicate the location of mutants in stalk segment 5 and
the C-terminus that were tested for synthetic lethality with sod1∆ in parts C, D, and Fig.
3-2. The specific effects of these mutants on Pma1p activity can be found in Table 3-1.
Green circles and bold type denote the Pma1p residues in the central ATPase region and
C-terminus that were identified as secondary mutations by sequencing in strains with
suppressors of the T912D sod1∆ synthetic lethality (see Fig. 3-3). (C, D) Plate-based
growth tests were performed with 104, 103, and 102 cells spotted onto YPD (aerobic) or
YPDE (anaerobic) and incubated at 30°C for 5 days. Plates were maintained
anaerobically using the GasPak EZ Anaerobe Container System. (C) pma1 single and
pma1sod1∆ double mutants in the BY4741 strain background. (D) pma1 single and pma1
sod1∆ double mutants in the NY13 background.
127
128
FIGURE 3-2. Mutations in Pma1p stalk segment 5 that alter glucose regulation are
not synthetically lethal with sod1∆
sod1∆ was introduced into the Pma1p stalk segment 5 mutants indicated (NY13
background) by transformation of pJAB002 [27] and tested for growth aerobically and
anaerobically as described for Fig. 3-1C,D.
129
FIGURE 3-3. T912D sod1∆ cells develop suppressors with secondary mutations in
PMA1
Individual colonies of T912D sod1∆ in the NY13 strain background that grew aerobically
were isolated (see Materials and Methods) and along with appropriate controls were (A)
retested for their ability to grow aerobically as described for Fig. 3-1C,D or (B) tested to
determine if they suppress the sod1∆ lysine auxotrophy by spotting onto SC media
(+Lys) or SC media lacking Lys (-Lys) and incubating at 30°C for 3 days. Four
representative suppressors are shown. Each had a unique secondary mutation in the
Pma1p coding region as indicated. (C) In the BY4741 background, the WT PMA1 gene
was replaced with PMA1-A906G or PMA1-T912D/A906G in both WT and sod1∆ strains
as described in Materials and Methods and tested for lysine auxotrophy alongside
appropriate controls as described for part B.
130
131
FIGURE 3-4. The PMA1-A906G mutation increases Pma1p ATPase activity but
does not alter Pma1p localization or protein level
The strains indicated were grown at 30°C shaking in YPD to mid-log phase and tested by
(A) indirect immunofluorescence to determine Pma1p localization and (B) western blot
to determine total cellular Pma1p protein level using a monoclonal antibody against
Pma1p as described in Materials and Methods. (C) Pma1p activity was determined in
membranes prepared from cells grown in the same manner by measuring the
orthovanadate-sensitive ATPase activity as described in Materials and Methods. Values
shown represent the average of two biological replicates normalized to WT; error bars
indicate standard error. All strains in this figure are in the BY4741 background.
132
FIGURE 3-5. T912D sod1∆ synthetic lethality is due to oxidative stress and not
Yck1p misregulation
(A) Pma1-WT was replaced with PMA1-T912D in WT and akr1∆ null cells in the
BY4741 background and tested for aerobic and anaerobic growth as described in Fig. 3-
1C,D. (B) 104, 103, and 102 cells of each strain indicated (NY13 background) were
spotted onto YPDE (0% oxygen) or YPD (1-20% oxygen) and incubated in the oxygen
concentration specified for 3 days at 30°C. (C) Cells were spotted and grown
anaerobically on YPDE or aerobically on YPD with or without 1 mM MnCl2 as described
in Fig. 3-1C,D.
133
134
FIGURE 3-6. Pma1-T912D mutant cells are not sensitive to environmental oxidants
or hyperoxia
(A, B) 106 cells/ml were inoculated into liquid YPD containing the concentration of
paraquat (A) or menadione (B) indicated and shaken at 30°C for either 18 hrs (A) or 15
hrs (B). (B) Growth in menadione is graphed on a log scale. Error bars indicate standard
error of duplicate biological measurements. (C) 104, 103, and 102 cells of the strains
indicated (NY13 background) were plated onto YPD and incubated in air or at 100% O2
or onto YPDE and incubated anaerobically using the GasPak system, as described in
Materials and Methods, at 30°C for 2 days.
135
136
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CHAPTER 4
Review and Future Directions
148
1. Review of Thesis Research
Cellular pH homeostasis is regulated in a manner highly dependent on the
metabolic state of cells. This is particularly evident in yeast which readily shift from
respiratory to fermentative metabolism depending on carbon source availability. In the
presence of glucose, H+-ATPases rapidly extrude protons to tightly maintain pH
consuming significant amounts of cellular ATP in the process. In the absence of glucose,
H+-ATPase activities are diminished to conserve energy and environmental factors and
buffering become much more important.
Similarly, metabolism can regulate the ROS balance in cells. In fermenting yeast
growing with abundant glucose, the NADPH oxidase, Yno1p, along with other sources
produce a significant amount of ROS in the cytoplasm [1]. During respiration in the
absence of glucose, on the other hand, ROS in the mitochondrial matrix may be high.
The research in this thesis focuses on the role of ROS in regulating pH at these
two extremes of glucose abundance and glucose starvation. In abundant glucose, the
largely cytosolic superoxide dismutase, Sod1p, promotes activation of the major plasma
membrane H+-ATPase, Pma1p (see Fig. 4-1). Although earlier studies suggested Sod1p
works through a mechanism involving the Pma1p regulatory kinase, Yck1p [2], the
studies in this thesis indicate that the Sod1p-Pma1p connection extends beyond this
signaling pathway and involves oxidative stress protection as well (Chapter 3). While
Sod1p is important for pH homeostasis during glucose abundance, we find that the
mitochondrial matrix Sod2p and mitochondrial ROS become important during conditions
of glucose starvation. ROS in the mitochondrial matrix can actually be beneficial to yeast
during periods of long term glucose starvation. By damaging the TCA enzymes
149
aconitase and SDH (Fig. 4-1B; Chapter 2), an acid burst occurs that is marked by large
increases in Ald4p-derived acetate, providing a new carbon source for cell survival.
150
FIGURE 4-1. Models depicting ROS regulation of pH homeostasis at the metabolic
extremes of glucose abundance and glucose starvation.
(A) A model depicting Sod1p regulation of Pma1p during growth in abundant glucose
(Chapter 3). The plasma membrane H+-ATPase, Pma1p, is highly active in the presence
of glucose and is the primary protein responsible for ensuring pH homeostasis,
maintaining pHi ≈ 7.2 [3] while acidifying the surrounding media. Organic acids are also
secreted [4-6]. Sod1p acts through two pathways to ensure maximal Pma1p activation in
abundant glucose. Pathway 1 (top): Sod1p prevents degradation of the yeast casein
kinase, Yck1p, by producing H2O2 in a manner requiring oxygen and glucose. Yck1p
then activates Pma1p [2]. Pathway 2 (bottom): Sod1p scavenges ROS to prevent
oxidative damage, which inhibits Pma1p either directly or indirectly. (B) A model
depicting how mitochondrial ROS produced during glucose starvation can lead to the
acid burst and acidify the extracellular environment (Chapter 2). In the absence of
glucose, activity of the plasma membrane H+-ATPase, Pma1p, is low and yeast initially
alkalinize the surrounding media, presumably by producing ammonia (not shown) [7].
Later, yeast shift to a phase of acid production termed the acid burst. This period is
initiated by increased ROS in the mitochondrial matrix that inhibits the TCA cycle
enzymes aconitase (Aco1p) and SDH. TCA cycle inhibition prevents Ald4p-produced
acetate from entering the cycle (as acetyl-CoA) which subsequently builds up and is
secreted into the media. Acetate secretion alone causes acidification of the media, as
Pma1p activity remains low, and promotes yeast growth during glucose starvation.
151
152
2. Future Directions
2.1. Examination of the acid burst during glucose starvation (Chapter 2)
Under glucose starvation conditions, ROS stimulate the acid burst to cause media
acidification and enhance growth (Fig. 2-9C). This occurs quickly in sod2Δ null cells but
is also seen in WT yeast, where it occurs more gradually (Fig. 2-2). Čáp and colleagues
[8] previously examined ROS levels in the mitochondrial matrix during long-term
glucose starvation and demonstrated that matrix ROS increase in a subset of WT cells
and in a greater number of sod2Δ null cells after entrance into the alkali phase, which
agrees well with our proposal that ROS stimulate the shift from alkali-to-acid. One
important question that remains unanswered is why ROS increase in the mitochondrial
matrix. Is this through loss of antioxidant activities or through an increase in production
of mitochondrial ROS? Though Čáp and colleagues examined the activity of cytoplasmic
Sod1p and the catalase Ctt1p they did not assess the all important mitochondrial matrix
Sod2p [8]. As decreased activity of Sod2p is the most likely explanation for the
increased ROS, further work could evaluate the activity and protein levels of Sod2p
during the transition from alkaline to acidic phases. Enhanced ROS production might
also be the culprit and could be examined by comparing ROS production in mitochondria
or mitochondrial particles isolated at various times during growth using available probes
such as MitoSOX for superoxide [8] and Amplex Red for hydrogen peroxide [9,10].
The major mitochondrial aldehyde dehydrogenase, Ald4p, is required for
production of the acetate secreted during the acid burst (Fig. 2-7F and 2-8). In the
simplest model we proposed, Ald4p produces normal levels of acetate which build up due
to ROS-disruption of the TCA cycle and are eventually secreted. Another possibility is
153
that Ald4p-acetate production is enhanced concomitantly with TCA cycle inhibition to
produce the acid burst. Future work could address the role of Ald4p in the acid burst
more closely by examining its expression, protein levels, and activity under glucose
starvation conditions.
Our work is the first to examine the mechanism by which yeast switch from
periods of media alkalinization to acidification during glucose starvation. It would be
interesting to examine cells of other organisms to determine if a similar mechanism
exists. At a minimum this is likely to be a general feature of yeasts, just as the ammonia
signaling during the period of alkalinization was shown by Palkova and colleagues to
occur in a variety of yeast strains [7]. However, it would also be interesting to examine
the intersection of nutrient deprivation, mitochondrial ROS, and growth in mammalian
cells. A number of similarities between cancer development, which is essentially an
overgrowth phenotype, and the acid burst of yeast suggest a similar pathway may exist in
mammalian cells. For one, defects in the TCA cycle initiate the acid burst and are also
known drivers of cancer formation [11]. Furthermore, when SDH mutations that are
known to cause cancer in humans are introduced into yeast grown on low glucose, the
resulting acid produced is very similar to the acid burst in both degree and composition
(i.e. high organic acid secretion, with particularly high levels of acetate) [12,13].
Secondly, increased mitochondrial ROS are associated with cancer formation [14,15].
Third, at least some studies show that glucose deprivation contributes to increased
mitochondrial ROS production in mammalian cells [14,16-19]. Few if any studies have
focused on the intersection between these factors and the potential role of extracellular
acidification in promoting growth.
154
2.2. Glucose-dependent Sod1p regulation of Pma1p (Chapter 3)
Rescue of T912D sod1Δ aerobic lethality by Mn anti-oxidants and anaerobic
growth, along with the viability of T912D akr1Δ strongly suggests that the T912D sod1Δ
synthetic lethality is a result of oxidative damage and not misregulation of Yck1p (Fig. 3-
5)[2]. However, whether Sod1p protects Pma1p directly or indirectly remains unclear.
To determine if Pma1p is directly damaged by oxidative stress, Pma1p WT and T912D
could be immunoprecipitated from sod1Δ and SOD1+ cells and examined in western blots
to identify carbonyl formation (Oxyblot). Significantly more carbonyls in sod1Δ cells
would be indicative of oxidative damage to Pma1p. Comparison of Pma1-T912D and
WT carbonylation in sod1Δ cells might also indicate whether the Pma1-T912D mutant is
more sensitive to oxidative damage than WT. In a similar manner, the oxidation state of
the five cysteines located in the catalytic domain of Pma1p could be compared in sod1Δ
versus SOD1+ cells. To do this, WT and T912D Pma1p would be pretreated with
iodoacetamide (protects reduced cysteines) and run on SDS-PAGE gels under reducing
and non-reducing conditions. sod1Δ-induced changes in cysteine oxidation would appear
as a gel shift in the reduced versus non-reduced samples. As before, this may also allow
the comparison of cysteine oxidation in Pma1-T912D relative to WT to determine if the
mutant is more sensitive. Finally, oxidative modifications to Pma1p in sod1Δ versus
SOD1+ cells could also be examined more generally using mass spectrometry.
Our work suggests the Pma1-T912D mutant is sufficiently sensitive to oxidative
damage that it can be totally inactivated in sod1∆ null cells. However, the effect of
oxidative damage on WT Pma1p activity was not explored. Previously, the activity of
WT Pma1p was shown to be reduced ≈ 80% in sod1∆ null cells [2]. This reduction was
155
attributed to destabilization of the Pma1p regulatory kinase, Yck1p, but no direct
evidence for this was provided and at least a portion is likely due to oxidative damage.
Future work could determine the extent that Sod1p regulation of Yck1p and Sod1p
oxidative stress protection play in promoting glucose-induced Pma1p activation. The
effect of oxidative damage could be explored by measuring Pma1p activity in sod1∆ null
cells that are grown in the presence of Mn to prevent oxidative damage, as this will not
rescue Yck1p destabilization. Conversely, Pma1p activity in sod1∆ null cells expressing
a mutant of Yck1p that is not degraded (YCK1-K383R/K386R/K390R) [2] would
indicate how important Sod1p regulation of Yck1p is to Pma1p activation.
The importance of Sod1p in regulating Pma1p, particularly by preventing
oxidative damage, was previously unexplored. This role of Sod1p may not be limited to
Pma1p and Sod1p is likely to play a role in regulating other transporters in both yeast and
mammalian cells. To identify transporters protected by Sod1p from oxidative damage, a
western blot for protein carbonyls could be used to identify membrane proteins modified
specifically in sod1∆, but not WT, yeast cells after crude membrane extraction and
protein separation by 2-dimensional gel. A similar approach could also be used in
mammalian cells. Interestingly, at least one study suggests that the plasma membrane
Na+/H+ exchanger, NHE1, that maintains pHi in mammalian cells (analogous to the role
of Pma1p in yeast) is sensitive to oxidative insult [20]. This suggests the possibility that
mammalian SOD1 also regulates NHE1 which to our knowledge has not been explored in
detail.
156
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J. ALLEN BARON 634 Melvin Dr. • Baltimore, MD 21230
Phone: 410-807-2955 • Email: [email protected]
EDUCATION Sept 2013, expected Ph.D. in Biochemistry and Molecular Biology Johns Hopkins University–Bloomberg School of Public Health Baltimore, MD
Dec 2005 B.S. in Biology (summa cum laude) Brigham Young University – Idaho Rexburg, ID RESEARCH EXPERIENCE 2007 – present Graduate research in the lab of Dr. Valeria C. Culotta. Cellular Redox and pH Signaling
• Biochemical characterization of cellular metabolites and enzyme activities central to energy metabolism and pH homeostasis affected by reactive oxygen species signaling
• Fractionation and isolation of subcellular components • Extensive experience in cloning and yeast molecular genetics • Fluorescence microscopy of protein localization using
immunofluorescence
2007 – 2010 Graduate research in the lab of Dr. Eric Grote. Calcium Regulation of Cell-Cell Fusion
• Used steady-state and time-lapse fluorescence microscopy to monitor cell-cell fusion events and dynamics of calcium signaling
• Developed a facile and high-throughput genetic screen in yeast to identify factors required for membrane fusion
2006 Five 7-week rotations during first year of Ph.D. introduced me to: • Mammalian cell culture • PCR-based genomic screening and genetic screens in plants • Rotations were in the labs of Drs. Terry Brown, Roger McMacken,
Judith Bender, Eric Grote, and William Wright
2005, summer IDeA Network of Biomedical Research Excellence (INBRE) Summer Fellow in the lab of Douglas G. Cole, Department of Biological Sciences, University of Idaho. Expression and Protein Interaction Analysis of Two Chlamydomonas Intraflagellar Transport Proteins
TEACHING EXPERIENCE 2012 Teaching Assistant, Principles of Cell Biology (Master’s level), Johns
Hopkins University
160
2011 – 2013 Research mentor for Janice Chen (undergraduate), Johns Hopkins University
• Working with Pamela Silver, Ph.D. (Harvard) and applying to MD/PhD programs
2008 Research mentor for Preston Kramer (undergraduate), Johns Hopkins University
2007 Tutor, Molecular Biology and Genomics (Ph.D. level), Johns Hopkins University
2005 Tutor, Genetics, Brigham Young University – Idaho • College Reading & Learning Association Certified - Advanced Tutor
2004 Lab Assistant, General Microbiology, Brigham Young University – Idaho
PUBLICATIONS 2. Baron, J.A., Laws, K.M., Chen, J.S., and Culotta, V.C. (2013) Superoxide triggers
an acid burst in Saccharomyces cerevisiae to condition the environment of glucose-starved cells. J. Biol. Chem. 288, 4557-4566.
1. Leitch, J.M., Li, C.X., Baron, J.A., Matthews, L.M., Cao, X., Hart, P.J., and Culotta, V.C. (2012) Post-translational modification of Cu/Zn superoxide dismutase under anaerobic conditions. Biochemistry. 51, 677-685.
CONFERENCE PRESENTATIONS 3. Mitochondrial superoxide mimics the cellular response to nutrients in the yeast
Saccharomyces cerevisiae. Baron, J. Allen and Culotta, Valeria C. Society for Free Radical Biology and Medicine, 18th Annual Meeting (Nov 2011). Atlanta, GA, United States.
2. Ca2+ regulation of fusion/lysis in S. cerevisiae during plasma membrane fusion. Baron, J. Allen and Grote, Eric. Gordon Research Conference: Cell-Cell Fusion (Aug 2009). University of New England, Biddeford, ME, United States.
1. Expression and Protein Interaction Analysis of Two Chlamydomonas Intraflagellar Transport Proteins. Baron, J. Allen; Behal, Robert; Qin, Hongmin; Rosenbaum, Joel L.; Cole. Douglas G. IDeA Network of Biomedical Research Excellence (INBRE) Summer Research Conference (Aug 2005). Northwest Nazarene University, Nampa, ID, United States.
